Markus Montag - A Practical Guide To Selecting Gametes and Embryos (2014, Taylor and Francis, CRC Press)

a Practical guide to
Selecting
gameteS and
embryoS
EditEd by Markus Montag

Selecting
gameteS and
embryoS
Selecting
gameteS and
embryoS
Edited by
Markus Montag, PhD
iLabCoMM GmbH, International Reprolab Consulting
St. Augustin, Germany
Boca Raton London New York
CRC Press is an imprint of the

Taylor & Francis Group, an informa business
CRC Press
Taylor & Francis Group
6000 Broken Sound Parkway NW, Suite 300
Boca Raton, FL 33487-2742
© 2014 by Taylor & Francis Group, LLC

CRC Press is an imprint of Taylor & Francis Group, an Informa business
No claim to original U.S. Government works

Version Date: 20140224
International Standard Book Number-13: 978-1-84214-548-7 (eBook - PDF)
This book contains information obtained from authentic and highly regarded sources. While all reasonable efforts have been made to
publish reliable data and information, neither the author[s] nor the publisher can accept any legal responsibility or liability for any errors
or omissions that may be made. The publishers wish to make clear that any views or opinions expressed in this book by individual editors,
authors or contributors are personal to them and do not necessarily reflect the views/opinions of the publishers. The information or guid-
ance contained in this book is intended for use by medical, scientific or health-care professionals and is provided strictly as a supplement to
the medical or other professional’s own judgement, their knowledge of the patient’s medical history, relevant manufacturer’s instructions
and the appropriate best practice guidelines. Because of the rapid advances in medical science, any information or advice on dosages, pro-
cedures or diagnoses should be independently verified. The reader is strongly urged to consult the drug companies’ printed instructions,
and their websites, before administering any of the drugs recommended in this book. This book does not indicate whether a particular
treatment is appropriate or suitable for a particular individual. Ultimately it is the sole responsibility of the medical professional to make
his or her own professional judgements, so as to advise and treat patients appropriately. The authors and publishers have also attempted
to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in
this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in
any future reprint.
Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by
any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in
any information storage or retrieval system, without written permission from the publishers.
For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.
com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-
profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy
license by the CCC, a separate system of payment has been arranged.
Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and
explanation without intent to infringe.
Visit the Taylor & Francis Web site at
http://www.taylorandfrancis.com
and the CRC Press Web site at

http://www.crcpress.com
Contents
Preface��vii
Contributors��ix
1. Handling Gametes and Embryos: Sperm Collection and Preparation Techniques��1

Verena Nordhoff, Con Mallidis, and Sabine Kliesch
2. Handling Gametes and Embryos: Oocyte Collection and Embryo Culture�� 17

Lars Johansson
3. Handling Gametes and Embryos: Quality Control for Culture Conditions�� 39

Jason E. Swain
4. Morphological Selection of Gametes and Embryos: Sperm�� 59

Pierre Vanderzwalmen, Magnus Bach, Olivier Gaspard, Bernard Lejeune, Anton Neyer,
Françoise Puissant, Maximilian Schuff, Astrid Stecher, Sabine Vanderzwalmen,
Barbara Wirleitner, and Nicolas H. Zech
5. Morphological Selection of Gametes and Embryos: Oocyte�� 81

Başak Balaban and Thomas Ebner
6. Morphological Selection of Gametes and Embryos: 2PN/Zygote��97

Martin Greuner and Markus Montag
7. Morphological Selection of Gametes and Embryos: Embryo�� 115

Gayle Jones and M. Cristina Magli
8. Morphological Selection of Gametes and Embryos: Blastocyst�� 131

Thomas Ebner
9. Noninvasive Techniques: Gamete Selection—Sperm�� 143

Victoria Sánchez, Joachim Wistuba, and Con Mallidis
10. Noninvasive Techniques: Gamete Selection—Oocyte�� 155

Laura Rienzi, Benedetta Iussig, and Filippo Maria Ubaldi
11. Noninvasive Techniques: Embryo Selection by Oxygen Respiration�� 165

Alberto Tejera, Belén Aparicio, Carmela Albert, Arancha Delgado, and Marcos Meseguer
12. Noninvasive Techniques: Embryo Selection by Time-Lapse Imaging�� 177

Alison Campbell
13. Noninvasive Techniques: Embryo Selection by Transcriptomics, Proteomics,

and Metabolomics��191
Asli Uyar and Emre Seli
v
vi Contents
14. Invasive Techniques: Polar Body Biopsy��209

Markus Montag, Jana Liebenthron, and Maria Köster
15. Invasive Techniques: Embryo Biopsy at the Cleavage Stage�� 219

Anick De Vos
16. Invasive Techniques: Blastocyst Biopsy�� 233

Steve McArthur
17. Invasive Techniques: Aneuploidy Testing by FISH�� 241

Semra Kahraman and Çağrı Beyazyürek
18. Invasive Techniques: Aneuploidy Testing by Array-CGH�� 255

Alan R. Thornhill, Christian Ottolini, Gary Harton, and Darren Griffin
19. Summary: Comprehensive Summary of Main Points by Topic��269

Markus Montag
Preface
The development of current technologies to identify the most viable oocyte, sperm, or embryo is one of the most
discussed topics in assisted reproduction conferences and symposia. However, most of these, sometimes fancy,
technologies are yet to be introduced in the daily routine of an in vitro fertilization laboratory, and the reasons
for this are many.
So, from the perspective of the laboratory, it would be helpful to have an overview of ready-to-use assisted
reproduction technologies, with practical guidance from those who have knowledge and expertise in the relevant
field. The resultant textbook, A Practical Guide to Selecting Gametes and Embryos, has compiled such practical
tips, accompanied by numerous figures and descriptions of methods. This book differentiates between noninva-
sive and invasive techniques; however, to my understanding, even the most sophisticated selection strategies may
fail if they are not based on a sound start. It happens frequently—that is, by looking at the most ideal endpoint,
we simply overlook the way to reach that endpoint. Practically speaking, the starting point involves good practice
in the laboratory setting in handling gametes and embryos with special focus on quality measures. This starting
point is an absolute prerequisite to be able to apply selection strategies that make sense.
I thank all the authors who have devoted their time to contribute to this illustrated textbook. They are all active
in their respective fields with numerous obligations; hence, taking on the additional task of contributing to this
book is a commitment that cannot be praised enough.
I also thank Robert Peden (Senior Editor for Reproductive Medicine, CRC Press) for his enthusiasm, support,
and continuous efforts in bringing this project to life.
Markus Montag
vii
Contributors
Carmela Albert Martin Greuner

Instituto Valenciano de Infertilidad Fertility Center IVF-Saar
Universidad de Valencia Saarbrücken-Kaiserslautern, Germany
Valencia, Spain
Darren Griffin
Belén Aparicio School of Biosciences
Instituto Valenciano de Infertilidad University of Kent
Universidad de Valencia Canterbury, United Kingdom
Valencia, Spain
Gary Harton
Magnus Bach School of Biosciences
IVF Centers Prof. Zech University of Kent
Bregenz, Austria Canterbury, United Kingdom
Başak Balaban Benedetta Iussig
Assisted Reproduction Unit GENERA Centres for Reproductive Medicine
VKF American Hospital Rome, Italy
Istanbul, Turkey
Lars Johansson
Çağrı Beyazyürek Origio a/s
Reproductive Endocrinology and Genetics Center Måløv, Denmark
Istanbul Memorial Hospital
Istanbul, Turkey Gayle Jones
Centre for Human Reproduction
Alison Campbell Genesis Hospital
CARE Fertility Athens, Greece
Nottingham, United Kingdom
and
Arancha Delgado Department of Anatomy and Developmental Biology
Instituto Valenciano de Infertilidad Monash University, Clayton
Universidad de Valencia Victoria, Australia
Valencia, Spain
Semra Kahraman
Anick De Vos Reproductive Endocrinology and Genetics Center
Centrum voor Reproductieve Geneeskunde Istanbul Memorial Hospital
Universitair Ziekenhuis Brussel Istanbul, Turkey
Brussels, Belgium
Sabine Kliesch
Thomas Ebner Department of Clinical Andrology
Landes-, Frauen- und Kinderklinik Centre of Reproductive Medicine and Andrology
Kinderwunsch Zentrum University Hospital of Münster
Linz, Austria Münster, Germany
Olivier Gaspard Maria Köster

Department of Gynaecology and Obstetrics Department of Gynecological Endocrinology and
Centre of AMP (CPMA) Reproductive Medicine
University of Liège University Women’s Hospital Bonn
Liège, Belgium Bonn, Germany
ix
x Contributors
Bernard Lejeune Françoise Puissant

Assisted Reproductive Department Assisted Reproductive Department
Centre Hospitalier Inter Régional Cavell (CHIREC) Centre Hospitalier Inter Régional Cavell (CHIREC)
Braine l’Alleud, Belgium Braine l’Alleud, Belgium
Jana Liebenthron Laura Rienzi

Department of Gynecological Endocrinology and GENERA Centres for Reproductive Medicine
Reproductive Medicine Rome, Italy
University Women’s Hospital Bonn
Bonn, Germany Victoria Sánchez
Centre of Reproductive Medicine and Andrology
M. Cristina Magli University Hospital of Münster
Società Italiana Studi Medicina della Riproduzione Münster, Germany
Bologna, Italy
Maximilian Schuff
Con Mallidis IVF Centers Prof. Zech
Centre of Reproductive Medicine and Andrology Bregenz, Austria
University Hospital of Münster
Münster, Germany Emre Seli
Department of Obstetrics, Gynecology, and
Steve McArthur Reproductive Sciences
Genea Fertility Yale School of Medicine
Sydney, Australia New Haven, Connecticut
Marcos Meseguer Astrid Stecher

Instituto Valenciano de Infertilidad IVF Centers Prof. Zech
Universidad de Valencia Bregenz, Austria
Valencia, Spain
Jason E. Swain
Markus Montag Fertility Lab Sciences
iLabCoMM GmbH (International Reprolab Consulting) National Foundation for Fertility Research
St. Augustin, Germany Colorado Center for Reproductive Medicine
Lone Tree, Colorado
Anton Neyer
IVF Centers Prof. Zech Alberto Tejera
Bregenz, Austria Instituto Valenciano de Infertilidad
Universidad de Valencia
Verena Nordhoff Valencia, Spain
Department of Clinical Andrology
Centre of Reproductive Medicine and Andrology Alan R. Thornhill
University Hospital of Münster Assisted Conception Unit
Münster, Germany Guy’s Hospital
London, United Kingdom
Christian Ottolini and
The London Bridge Fertility, Gynaecology and
Genetics Centre School of Biosciences
London, United Kingdom University of Kent
Canterbury, United Kingdom
and
School of Biosciences Filippo Maria Ubaldi
University of Kent GENERA Centres for Reproductive Medicine
Canterbury, United Kingdom Rome, Italy
Contributors xi
Asli Uyar Barbara Wirleitner

Department of Obstetrics, Gynecology, and IVF Centers Prof. Zech
Reproductive Sciences Bregenz, Austria
Yale School of Medicine
New Haven, Connecticut Joachim Wistuba
Centre of Reproductive Medicine and Andrology
Pierre Vanderzwalmen University Hospital of Münster
IVF Centers Prof. Zech Münster, Germany
Bregenz, Austria
and Nicolas H. Zech
IVF Centers Prof. Zech
Assisted Reproductive Department
Bregenz, Austria
Centre Hospitalier Inter Régional Cavell (CHIREC)
Braine l’Alleud, Belgium
Sabine Vanderzwalmen
Assisted Reproductive Department
Centre Hospitalier Inter Régional Cavell (CHIREC)
Braine l’Alleud, Belgium
1
Handling Gametes and Embryos: Sperm
Collection and Preparation Techniques
Introduction
Spermatozoa are an integral part of assisted reproduction techniques (ARTs). As such, the collection and
preparation of spermatozoa should be of high quality standards to ensure the best possible treatment outcome.
It is therefore important that the protocols for the preparation and the use of sperm are standardized and
dependable. A series of such standards exists in the form of guidelines and procedures comprising the World
Health Organization (WHO) laboratory manual [1].
It is important therefore that protocols for the preparation and the use of sperm are standardized
and dependable.
The following sections provide a comprehensive description of the steps undertaken for the analysis of
a semen sample. They are not meant to be as detailed as the WHO manual nor replace the same [1]; rather, they act as
an adjunct. For specific details and explanations for the choice of techniques, the WHO manual should be referred to.
Semen Analysis
Semen consists of two major fractions: spermatozoa from the testis, making up a small fraction of the ejaculate,
and seminal fluid from the accessory glands, making up the major portion of the ejaculate [2].
The proper analysis of the semen sample necessitates the evaluation of the following parameters:
• Volume (or weight)

• Liquefaction
• Viscosity
• Appearance
• pH
• Aggregation or agglutination
• Motility
• Concentration
• Morphology
For ARTs, the most important of the aforementioned parameters are motility, concentration, and morphology.
Normozoospermia is defined as a sample containing more than 39 million spermatozoa/ejaculate or
15 million/mL, with >32% of the spermatozoa being progressively motile (PR) and >4% having normal forms.
If the total sperm count is below the reference value, the sample is classified as oligozoospermic (O); if motility is
1
2 A Practical Guide to Selecting Gametes and Embryos
lower, it is considered as asthenozoospermic (A), and if morphology is less than the defined cutoff, it is d esignated
teratozoospermic (T). Combinations of the conditions exist and are classified accordingly (e.g., OA, OT, OAT).
The most important sperm parameters for ARTs are motility, concentration, and morphology.
Biochemical markers of seminal fluid can be determined to evaluate the function of the epididymis, seminal
vesicles, and prostate [1]. Biochemical markers may help to differentiate between obstructive and nonobstructive
restrictions of semen quality.
Analysis of the Native Ejaculate

For assisted reproduction, the native semen must be collected in a sterile container to allow either the exact
measurement of the volume (e.g., a cylinder with scale) or the determination of the weight of a sample. Directly
after collection, semen is normally coagulated, but it liquefies after 30–60 min at room temperature (37°C).
If liquefaction takes >60 min, this should be noted. If no liquefaction is obvious, then the use of a proteolytic
enzyme such as bromelain may be necessary. A process that involves mixing equal volumes of ejaculate and a
10 IU/mL bromelain/medium (e.g., an HEPES-buffered medium) solution followed by incubation at 37°C for
10 min. If this process is undertaken, it must be duly noted because it might affect other semen parameters, most
notably motility and final sperm concentration (due to the dilution, the original concentration is halved).
Because semen is very heterogeneous, it is extremely important that the sample is mixed thoroughly before
starting the different facets of the analysis. Only a very well mixed sample will provide data that are rep-
resentative and reproducible (i.e., when duplicate samplings are taken). After liquefaction, the semen sample
should be evaluated for its viscosity and appearance. Viscosity is determined by using a glass rod to mix the
sample. If threads of >2 cm can be seen after pulling the rod out of the semen, then the viscosity is higher than
what is considered normal, and represents a finding that should be recorded. Normal-appearing semen is gray-
opalescent; any variation from this color must be noted because it can be indicative of some underlying pathol-
ogy. For example, a brown color is symptomatic of the presence of blood cells. The final visual assessment is that
of pH, an assessment that must be performed quickly after liquefaction because the pH of semen changes with
time. The process involves placement of one drop of mixed semen unto pH paper with a range of 6.0–10.0. A pH
value between 7.2 and 8.0 is accepted as normal.
Only a very well mixed semen sample will provide representative and reproducible data.
For the initial microscopic assessments, a wet preparation is performed whereby 10 μL of native ejaculate is
placed onto a microscope slide, which is then covered by a 22 × 22 mm coverslip forming a 20 μm-deep chamber.
It should be noted that all aliquoting of semen must be performed using a positive displacement pipette, because
the viscosity of the ejaculate is such that accurate and reliable sampling is precluded by the use of other forms of
pipettes. The microscope for evaluation has to be equipped with a phase contrast system and should have objec-
tives of 4×, 10×, 40×, and 100× (oil) magnifications. The first microscopic evaluation entails noting the occurrence
of cells other than spermatozoa (e.g., spermatogenic or epithelial cells) and the presence of sperm aggregation
and agglutination. Although aggregation is rather unspecific and may be the incidental binding of spermatozoa to
other cells or debris, agglutination refers to motile spermatozoa that adhere to one another. The extent and type of
agglutination are of importance and must be noted, preferably using the four-grade c lassification system recom-
mended by the WHO (for examples and illustrations, please refer to the relevant section of ref. [1]). If agglutina-
tions are present, additional testing for antibodies against spermatozoa are indicated.
Next, sperm motility should be assessed. It must be kept in mind that if the sample is left on the bench
(i.e., at room temperature) for a long time, changes in the sample may occur that could lead to mistaken
Sperm Collection and Preparation Techniques 3
determinations of motility. Therefore, it is important that the assessment is performed quickly after liquefaction.
Duplicate wet preparations (as described above) should be prepared and evaluated by counting and grading the
motility or otherwise of 200 spermatozoa at 200× or 400× magnification. Three categories of movement may
be detected: (1) spermatozoa that progress actively regardless of their type of movement are designated the PR
fraction, (2) spermatozoa that have slow movements without any progression are designated the nonprogressive
(NP) motile fraction, and (3) spermatozoa with no movement whatsoever are d esignated the immotile (IM) frac-
tion. If the agreement of the duplicate measurements is not within an acceptable range (for tables, see ref. [1]), a
new set of samples has to be prepared. For an accurate assessment, a minimum of five fields per slide should be
evaluated. Often, the motile fraction is overestimated because the eye pays more attention to movements. A solu-
tion to this problem is to first count the immotile spermatozoa and then the motile fraction in the same field. The
necessity of counting a minimum of 200 spermatozoa helps to minimize the sampling error to a 95% confidence
interval (see ref. [1] for acceptable differences) and to fulfill internal quality-control standards. The lower normal
reference limit for normozoospermia is determined by at least 32% PR spermatozoa (5% percentile).
For ARTs, only living spermatozoa are of importance and are assessed using dyes that are able to pass frag-
mented membranes (i.e., those of dead cells) but not intact membranes (i.e., living cells). Because the number of
viable cells may vary over time, this assessment should be done quickly after liquefaction. The recommended stains
are either eosin alone or eosin in combination with nigrosin (see ref. [1] for preparation of the staining dyes). As with
motility, 2 × 200 spermatozoa are assessed to achieve a precise count and a low sampling error with the percent-
age of spermatozoa not stained constituting the living fraction. At least 58% vital spermatozoa should be available.
Sperm concentration and total sperm count are important parameters because they may give an indication as
to which of the repertoire of ART techniques is to be used. The most important consideration for the determina-
tion of the concentration is the choice of counting chamber. The 100 μm-deep hemocytometer (i.e., improved
Neubauer) chamber is highly recommended, although other deep chambers may also be used once certain con-
siderations have been taken into account, namely the differences in chamber volumes, grid patterns, and calcula-
tions needed to derive the final concentration. Importantly, whatever the type of chamber to be used, it must first
be validated against the approved improved Neubauer.
The improved Neubauer has two counting chambers with a grid in each chamber; when these are covered by a
thick coverslip (i.e., 0.44 mm), an underlying chamber exactly 100 μm in depth is created. When Newton’s rings
can be seen between the chamber and the coverslip, it is ready for loading.
Sperm concentration and total sperm count may give an indication as to which of the repertoire of
ART techniques is to be used.
The counting grids are 3 × 3 mm and contain nine large squares (Figure 1.1). The central of these nine squares
has five rows containing five squares each, all of which are surrounded by three lines. The middle of the three
lines represents the boundary of the square. Each of these 25 squares is further subdivided into 16 smaller
squares. Normally, counting the number of sperm present in the central 25 squares is sufficient for a reliable
measurement; however, if sperm numbers are low, then counts of the contents of the neighboring horizontal grids
or even all nine big grids should be performed.
Only intact spermatozoa showing clearly distinguishable heads and tails are counted, and then only if more
than half of the head lays over the boundary of a square. As mentioned, a minimum of 200 spermatozoa should be
counted in each of the replicates (i.e., a total of 400 spermatozoa). To circumvent the distraction of moving sperm,
an appropriate fixative (refer to ref. [1]) should be used, ideally at a 1:1 (1+1) dilution. Because semen samples
with high concentrations may be difficult to count (e.g., spermatozoa may be lying on top of each other) and there-
fore a site of possible errors, the dilution can be adjusted (e.g., 1:5 [1+4], 1:20 [1+19], even 1:50 [1+49]) to provide
numbers that are easily countable. If the chosen dilution results in numbers that are too high or too low, it can
be amended (i.e., a higher or lower dilution can be used); in such cases, it is important that a new dilution of the
sample must be prepared (i.e., not a dilution of the already diluted sample), otherwise counting errors can arise.
Whatever dilution is used, it must be taken into account when calculating the final concentration of the sample.
FIGURE 1.1 Grids of the improved Neubauer chamber. (a) All nine big grids. (b) Only the middle grid. (c) Twenty-five squares for
counting concentration, keeping the middle of the three lines as the boundary of each small square.
Once the sample has been loaded into the chamber, the spermatozoa must be given time to settle onto the grid.
The contents of each side of the chamber are then counted, with the process always finishing a row even if >200
spermatozoa are counted; stopping elsewhere will introduce counting errors. Each evaluation is repeated on the
second side of the chamber, and if the difference between the counts is within acceptable levels (refer to ref. [1]),
the concentration can be calculated. To calculate total sperm count per ejaculate, concentration is multiplied with
seminal fluid; thus, correct measurements are prerequisites for further use of semen samples.
Low Sperm Numbers

If the initial inspection of the wet preparation shows the presence of any spermatozoa, then the sample might
be azoospermic. For this suspicion to be substantiated, the sample must first be centrifuged at high velocity and
force (e.g., 3000g for 15 min) so as to pellet the contents of the semen. After removal of the supernatant, the pel-
let should be resuspended in 50 μL of seminal plasma from which two aliquots of 10 μL each are placed under
coverslips (22 × 22 mm) and the contents scanned for the presence of spermatozoa. Absence of spermatozoa most
likely indicates azoospermia; however, for this finding to be confirmed, all the contents of the pellet need to be
examined. If spermatozoa are found during this scanning procedure, the sample is most likely cryptozoosper-
mic. The fewer the sperm found, the greater the sampling error and the more likely that it will exceed the accept-
able 5%. If fewer than 25 spermatozoa are found, the concentration will most likely be <56,000 spermatozoa/
mL [1], and as such a definite measure cannot be given in such cases but must be recorded in both the assessor’s
lab book and in the patient’s file. If cryptozoospermic samples are to be used for ARTs, the absolute number of
motile spermatozoa is decisive. To evaluate motile spermatozoa, high-speed centrifugation is contraindicated [1].
Azoospermia can be confirmed only after centrifugation and screening of the pellet for the
presence of sperm.
The final analysis is the assessment of sperm morphology. For this purpose, replicate air-dried smears are
prepared: one smear for evaluation and the other smear serving as a safeguard should an unforeseen incident
occur (i.e., breakage of the slide or staining not working). The smears are made by quickly placing an aliquot of
well-mixed semen onto the first slide. The replicate is similarly prepared but by using an aliquot of semen that
has again been mixed. Aliquots of 5 μL should be used for samples with high concentrations, whereas larger
volumes (up to 10 μL) should be used for low concentrations. If a sample has a very low concentration, it is
advisable that it is first centrifuged (e.g., 600g for 10 min), the supernatant removed, and the pellet resuspended
with a suitable volume of the supernatant. If the semen is too viscous, a washing step might be considered before
preparing the smear. The smear itself is prepared using a second glass slide that is pulled at an angle of about
45° over the slide with the aliquoted sample. Slides are then left to air dry for 1–4 hr before staining.
The staining methods recommended by the WHO [1] are Papanicolaou, Shorr, or Diff-Quik, all of which
produce a pale blue staining in the acrosomal region and a dark blue staining in the post-acrosomal region.
The midpiece might appear red and the tail red to blue. Excess cytoplasm in the area around the midpiece (i.e.,
cytoplasmic droplet) might occur on some sperm and will stain red with Papanicolaou and orange to red with
the Shorr procedure. For more details regarding the preparation, staining, and particularly the exact grading of
sperm morphology, please refer to the WHO manual [1]. As with all evaluations, duplicate counts of a minimum
of 200 spermatozoa are needed to decrease the extent of possible counting error. The lower normal limit for
morphology is as low as at least 4% of normal spermatozoa. Morphological criteria are strict and intend to define
those spermatozoa as normal that are presumably capable of fertilizing [1].
Other tests that may be conducted are the differentiation of round cells using peroxidase staining. Because only
leucocytes contain this enzyme, they can be recognized from other round cells (e.g., immature germ cells) that
do not stain. The presence of antisperm antibodies can be detected using the mixed antiglobulin reaction (MAR)
whereby immunoglobulin A (IgA)- and G (IgG)-coated beads attach to sperm possessing the antigen and thereby
restrict their movement. 5 μl of native semen are mixed with 5 μl of anti-IgG/IgA and 5 μl of erythrocytes (0 Rh pos)
and incubated for 3 minutes. After 3 and 10 minutes only motile spermatozoa are evaluated. The test is positive with
at least 10% IgG- and/or IgA-bound spermatozoa. However, only >50% IgG- and/or IgA-bound spermatozoa are
considered clinically relevant. Instead of erythrocytes, commercially available immunobeads can be used.
Preparation Techniques
In nature, the mucus of the female reproductive tract actively selects fertile spermatozoa, whereas debris, non-
sperm cells, and the constituents of seminal plasma are excluded. In addition, the essential physiological change
of a spermatozoon, capacitation, is initiated during passage through the cervical mucus. These important steps
are omitted during ARTs, and as a consequence, sperm preparation techniques are necessary.
First, it is necessary to separate spermatozoa from the seminal fluid within 1 hr of ejaculation to minimize
possible damage (e.g., oxidative attack) coming from nonspermatogenic cells contained in semen. The second
aim of preparing the semen is to separate cells that would normally never enter the female tract because they
cannot cross the cervical mucus. However, because ARTs circumvent this natural border, their presence must
be dealt with accordingly. The third benefit is the separation and concentration of motile and normally formed
spermatozoa that are most likely to fertilize an oocyte.
Several techniques for preparation of spermatozoa are available. The most commonly used techniques are simple
washing, “swim-up,” and density gradient (DG) centrifugation. The technique chosen greatly depends on the nature
of the semen sample. If the sample is within the normal range, according to ref. [1] reference values, a swim-up is
preferred. If the sample has low concentration, motility, or morphology, then often DG centrifugation is the method
of choice (for comparison, see Table 1.1). It is important to note that swim-up does not give as high recovery rates
of motile spermatozoa as centrifugation, but it is quick, easy to perform, and yields a fraction enriched with motile
spermatozoa because it is dependent on the movement of the sperm for selection [3,4]. DG centrifugation yields high
numbers of motile spermatozoa, but the choice of appropriate centrifugal force is often difficult because samples vary
and the pellet containing the spermatozoa may be too loose and may detach too quickly from the bottom of the tube.
For all preparation techniques, a medium based on a balanced salt solution supplemented with proteins should
be used. If the procedure is not conducted in an incubator of 37°C without CO2, then the medium used should be
supplemented with HEPES or a similar buffer for pH stability, and a preincubation of the medium without CO2
should be performed. If a 37°C incubator with CO2 is used, then the medium is usually buffered with sodium
bicarbonate, and the cap of the tube should be loosely closed to allow gas exchange for optimal pH.
Simple Washing
This technique is often used for normozoospermic semen samples. It is quick, easy, and yields high quantities of
spermatozoa (Figure 1.2).
TABLE 1.1
Comparison of Different Sperm Preparation Techniques
Type of Ejaculate Quality Pros Cons
Washing (W) Normozoospermia Quick-and-easy technique Only for sperm counts with high
Yields high numbers of sperm numbers
spermatozoa Not useful for samples with
contamination with other cells,
debris, or blood cells
Swim-up (SU) Normozoospermia Quick and easy to perform Lower recovery rates compared
Moderate OAT High rates of highly progressive with DG
motile spermatozoa
Density gradient (DG) Moderate-to-severe OAT High numbers of motile Difficult to standardize
spermatozoa High-molecular-weight
Best removal of debris or other cell compounds of unknown influence
types occurring in the semen Costs more because DG solutions
sample have to be purchased
Sperm medium
Ejaculate
Washing Centrifuge Add medium,

resuspend
FIGURE 1.2 Sketch of protocol for simple washing.
BOX 1.1 SIMPLE SPERM WASHING

Materials
• A balanced salt solution or a commercially available sperm preparation medium supplemented
with protein (usually human serum albumin [HSA]) according to the manufacturer’s protocol;
ready-for-use media are also available.
• Test tubes of 6 or 13 mL volume.
• A centrifuge that is able to reach at least 500g.
• Pipettes with variable volumes (e.g., 2–200 μL and 100–1000 μL).
Protocol
• The semen sample is mixed well (e.g., stirring with a glass rod).
• Transfer the sample to a test tube (if the sample is of high volume, more test tubes may be used).
• Add an equal amount of medium to the semen sample.
• Mix gently by tilting the closed test tube.
• Centrifuge the mixture at 300–500g for 5–10 min.
• Gently aspirate or decant the supernatant.
• Add 1 mL of preincubated equilibrated medium to the pellet.
• Gently resuspend the pellet by flicking the tube or by gentle pipetting of the pellet.
• Centrifuge again at 300–500g for 3–5 min.
• Again gently aspirate or decant the resulting supernatant.
• Resuspend the pellet with medium, normally the final volume is 0.5 mL. (The volume is impor-
tant! If the sample has to be used for intrauterine insemination (IUI), the amount should not be
>0.5 mL; for other purposes, higher volume can be used.)
• Determine sperm concentration and if necessary the motility and morphology (according to
ref. [1]).
Direct Swim-Up
The direct swim-up is a quick-and-easy technique using the unique ability of spermatozoa to move forward. The
basis of the procedure is the layering of a medium over the sample; the motile spermatozoa then swim into the
medium. Although the final number of spermatozoa might be not very high, it is compensated by the enrichment
of the number of spermatozoa with the highest forward progression (Figure 1.3).
Sperm medium
Collect
Ejaculate upper layer
Overlay/ Swim-up Enriched

underlay (45°) fraction
FIGURE 1.3 Sketch of protocol for direct swim-up.
BOX 1.2 DIRECT SWIM-UP

Materials
with protein (usually HSA) according to the manufacturer’s protocol; ready-for-use media are
also available.
Protocol
• The semen sample is mixed well by stirring with a glass rod.
• Ejaculate (1 mL) is transferred to a test tube (if the sample is of high volume, more tubes may
be used).
• Medium (1.0–1.2 mL) is pipetted onto the semen sample (it is also possible to pipette the medium
under the ejaculate; however, this is more difficult than overlaying it).
• Place the test tube bevelled at an angle of 45° in the incubator (this is necessary to reach a higher
surface area where spermatozoa can swim into the medium).
• Incubate for 1 hr at 37°C. (The duration of incubation can be reduced if the semen sample is of
very high concentration. Again, the final application will determine the time span: high concen-
tration, low incubation time; low concentration, high incubation time. Regardless, it is prudent
not to exceed 1 hr because zinc and the other components of the seminal plasma may diffuse into
the medium and, in turn, might create problems for the spermatozoa [5].)
• After incubation, gently aspirate the top 0.5–1.0 mL of the medium.
• If the sample is high in concentration, a dilution might be necessary. (Again, if the sample is
used for IUI, it might be better to decrease the incubation time than to use an additional dilution
because the amount of fluid that can be inseminated is limited.)
• A variation of the procedure is to centrifuge the sample at 300–500g for 5 min and to resuspend
the pellet in 0.5 mL of medium.
• Determine sperm concentration and if necessary the motility and morphology (according to
ref. [1]).
Swim-Up with an Additional Washing Step

This technique is equivalent to the direct swim-up and differs only in the introduction of a washing or dilution
step (Figure 1.4). The washing step is an elegant way to minimize or remove substances from the seminal plasma
(e.g., zinc) that may have a negative influence on spermatozoal quality.
Sperm medium Sperm medium
Collect
upper
Ejaculate layer
Dilution/ Centrifuge Swim-up Enriched

washing (45°) fraction
FIGURE 1.4 Sketch of protocol for swim-up with dilution/washing.
BOX 1.3 SWIM-UP WITH AN ADDITIONAL WASHING STEP

Materials
also available.
Protocol
• The semen sample is mixed very well by stirring with a glass rod.
• Transfer the sample to a test tube.
• Add an equal amount of medium.
• Mix gently by tilting the closed test tube.
• Centrifuge the mixture at 300–500g for 5–10 min.
• Carefully aspirate or decant the supernatant.
• Add 1 mL of preincubated equilibrated medium to the pellet.
• Place the test tube bevelled at an angle of 45° in the incubator (this is necessary to reach a higher
surface area where spermatozoa can swim into the medium).
• Incubate for 1 hr at 37°C. (The duration of incubation can be reduced if the semen sample is of
very high concentration; again, the final application will determine the time span: high concen-
tration, low incubation time; low concentration, high incubation time. Regardless, it is prudent
not to exceed 1 hr because zinc and the other components of the seminal plasma may diffuse into
the medium and, in turn, might create problems for the spermatozoa [5].)
• After incubation, gently aspirate the top 0.5–1.0 mL of the medium.
• If the sample is high in concentration, a dilution might be necessary. (Again, if the sample is used
for IUI, it might be better to decrease the incubation time rather than use an additional dilution
because the amount of fluid that can be inseminated is limited.)
• As with the previous method, the procedure can be varied, namely, centrifuge the sample at
300–500g for 5 min and then resuspend the pellet in 0.5 mL of medium.
• Determine concentration and if necessary also motility and morphology (according to ref. [1]).
Discontinuous Density Gradient (DG) Centrifugation

A discontinuous DG is often the method of choice if a sample is of low concentration and is to be used for in
vitro fertilization (IVF) and intracytoplasmic sperm injection (ICSI). The technique yields the optimal s election
of good spermatozoa while simultaneously reducing debris, leucocytes, and other cells present in the ejaculate.
For this preparation, two solutions of different densities are used, both consisting of colloidal silica coated with
silane of high molecular mass and low osmolality. Usually, an upper gradient with 40% (v/v) and a lower gradient
with 80% (v/v) are used, and the semen is placed on top. The layers are then centrifuged, and the enriched frac-
tion of spermatozoa is retrieved from the loose pellet at the bottom of the tube (Figure 1.5).
Ejaculate Sperm medium

Gradient
solutions
Collect
upper layer
40 v/v
80 v/v
Overlay Centrifuge Washing/ Enriched
centrifuge fraction
FIGURE 1.5 Sketch of protocol for density gradient centrifugation.
BOX 1.4 DISCONTINUOUS DG CENTRIFUGATION

Materials
also available.
• Gradient media (these are commercially available and should be used according to the manufac-
turer’s protocol); often, they are ready to use or need to be diluted with an iso-osmotic medium
that resembles the fluids of the female reproductive tract.
Protocol
• First, prepare the DG layers: place 1 mL of the 80% gradient medium into a test tube and then
1 mL of the 40% medium on top of the first layer.
• The semen sample is mixed well by stirring with a glass rod.
• Semen (1 mL) is gently transferred to the prelayered gradient media (if the sample is of high
volume, more tubes may be used).
• Place the test tube carefully into the apparatus and centrifuge at 300–400g for 15–30 min.
• Aspirate the supernatant leaving the sperm pellet.
• The sperm pellet is washed by adding 5 mL of preincubated equilibrated medium and r esuspended
gently by pipetting.
• Centrifuge again at 200g for 4–10 min.
• Resuspend the pellet in 0.5–1 mL of medium (depending on the ART to be performed).
• Determine concentration and if necessary also motility and morphology (according to ref. [1]).
Epididymal Spermatozoa
In cases of nonreconstructible obstructive azoospermia, it is possible to surgically retrieve spermatozoa by the
aspiration of fluid from expanded epididymal tubules. This fluid should be harvested with as few as possible
contaminating erythrocytes or other cell types originating from the operation site. The type of sample prepara-
tion technique depends on the number of spermatozoa retrieved, that is, if a high number has been found, a DG
centrifugation or a swim-up can be undertaken. However, if only few spermatozoa are seen, then a simple wash-
ing step is adequate.
Testicular Spermatozoa
Spermatozoa can be harvested from material obtained by open biopsy using either the conventional testicu-
lar sperm extraction (TESE) procedure in case of obstruction or its microsurgical variant mTESE in case
of non-obstructive azoospermia with severe testicular damage (Figure 1.6) [6,7]. As TESE material is often
contaminated with high numbers of erythrocytes, a washing step is always necessary.
FIGURE 1.6 Microscopic picture of a testis during operation. Thicker tubuli (in the middle) are identified for mTESE.
FIGURE 1.7 Mechanical maceration by fine needles.
During the operation, without applying force and squeezing, the testicular tissue is gently retrieved from
different parts of the segmentally structured testis and placed by nontouch technique into a HEPES-buffered
medium (e.g., a balanced salt solution with HEPES or a commercially available sperm preparation medium).
To have several samples for cryopreservation and thawing, the tissue samples should be distributed to four to
eight different tubes per testis. To release and collect the mostly tissue-bound spermatozoa, either a mechanical
(Figure 1.7) or an enzymatic preparation technique is used.
Mechanical Preparation of Testicular Sperm
BOX 1.5 MECHANICAL PREPARATION OF TESTICULAR SPERM

Materials
• A balanced salt solution or a commercially available sperm preparation medium supplemented with
protein (usually HSA) according to manufacturer’s protocol; ready-for-use media are also available.
• Two tuberculin syringes with fine needles or glass slides.
• Petri dish of 6–10 cm.
• A centrifuge able to reach at least 500g.
Protocol
• The freshly retrieved or frozen tissue is incubated in preequilibrated medium for up to 30 min to
remove contaminating red blood and other cells.
• The tissue is removed from the test tube and placed into a Petri dish and if necessary the culture
medium is augmented.
• The tissue is macerated either with the fine needles or by glass slides until a suspension of small
pieces of tissue has been achieved.
• The suspension is transferred to tube and centrifuged at speed between 100 and 300g for 10 min.
• The supernatant is gently discarded.
• Gently resuspend the pellet by flicking the tube or by pipetting.
• Centrifuge again at speed between 100 and 300g for 10 min.
• Resuspend the pellet either in the remaining culture medium or by adding 0.2–0.5 mL of fresh
medium.
Enzymatic Preparation of Testicular Sperm

An alternative procedure for the retrieval of testicular spermatozoa is digestion of either a piece of whole tissue
or of single tubules with the enzyme collagenase (from Clostridium histolyticum, type 1A) (Figure 1.8).
FIGURE 1.8 Digested testicular tissue after enzymatic treatment.
BOX 1.6 ENZYMATIC PREPARATION OF TESTICULAR SPERM

Materials
• Collagenase type 1A from Clostridium histolyticum (commercially available).
with protein (usually HSA) according to manufacturer’s protocol; ready-for-use media are also
available.
Protocol
• Incubate the freshly retrieved or frozen tissue in preequilibrated medium at 37°C for 30 min.
• Prepare the collagenase solution by dissolving 0.8 mg of collagenase type 1A in 1 mL of culture
medium.
• Add the tissue to the collagenase and incubate at 37°C for 1.5–2 hr, vortex every 30 min (it might
still be that remnants of the tissue are not resolved; often these are remnants of the tubular struc-
ture in cases of testicular dysfunction).
• Centrifuge at 100–300g for 10 min.
• The supernatant is gently discarded.
• Wash the pellet to remove any collagenase by adding 1 mL of culture medium.
• Gently resuspend the pellet by flicking at the tube or by pipetting.
• Centrifuge again at 100–300g for 10 min.
• Resuspend the pellet either in the remaining culture medium or by adding 0.2–0.5 mL of fresh
medium.
Collecting Testicular Spermatozoa for ICSI
BOX 1.7 COLLECTING TESTICULAR SPERMATOZOA FOR ICSI (FIGURE 1.9)

Protocol
• Place 5–10 μL of the prepared suspension into an ICSI dish.
• Allow the suspension time to settle to the bottom of the dish.
• Retrieve motile, or if none are available, immotile spermatozoa (for rationale, see the following
section) using an ICSI needle.
• Place into a drop of polyvinylpyrrolidone (PVP) until injected.
Vitality Tests for Immotile Spermatozoa

It has been shown that immotile spermatozoa either from the ejaculate [8] or extracted from a testicular biopsy [9]
are capable of fertilizing an oocyte and producing viable healthy pregnancies.
Using light microscopy, dead spermatozoa are undistinguishable from spermatozoa that are alive but immotile;
consequently, several tests have been developed to differentiate between them ([10]; for comparison, see Table 1.2).
A long-established technique is the hypoosmotic swelling (HOS) test wherein viable spermatozoa are selected based
on their response to exposure to a hypoosmotic medium [11,12]. In such medium, the tails of viable sperm become
curved or swollen, thus making them recognizable for selection. This technique in combination with an ART has
been successfully implemented with ejaculated and testicular samples [13,14]. The technique as such is useful,
although the time frame in which spermatozoa are exposed to the hypoosmotic medium should be as low as possible.
If spermatozoa are exposed too long, the curved tail will result in steric problems while getting the spermatozoon
into an ICSI needle. In addition, this test is time-consuming and in cases of, for example, severe TESE-ICSI, the
retrieval of every individual spermatozoon already is cumbersome and time-consuming and thus not applicable.
Oocytes can be fertilized by immotile sperms if these sperms are viable.
Other tests available use xanthine derivatives (e.g., pentoxifylline or theophylline) that induce tail movements
of immotile ejaculated or testicular spermatozoa or that enhance the movement of spermatozoa with very low
motility [15–18]. The application of such compounds is quite easy, but in humans it is not known whether their use
might interfere with later embryonic development. Therefore, this procedure cannot be recommended for routine
clinical use.
For experienced embryologists, the mechanical touch technique can be used, wherein spermatozoa are gently
agitated with an ICSI needle and the flexibility of the tail is taken as a sign of vitality [19,20]. But for routine use,
FIGURE 1.9 Mechanically derived suspension (a), enzymatic digestion (b), and collection of individual spermatozoa (c) in an ICSI
dish.
TABLE 1.2
Pros and Cons of Different Spermatozoal Vitality Tests
Type of Spermatozoa Pros Cons
HOS test All types (ejaculate, Easy recognition of living Time of exposing spermatozoa to the
MESA, TESE) spermatozoa because their tails hypo-osmotic medium should be low
swell Steric problems from the tail swelling
Solution has to be purchased
Chemical All types Vital spermatozoa that are able to In humans, it is not known whether these
compounds bend their tails can be visualized compounds may have an influence on
very quickly further development
Useful for ejaculate and TESE Solution has to be purchased
spermatozoa Vital spermatozoa that are not able to bend
their tails will not be identified
Mechanical All types Quick technique without need for Only for experienced lab personnel
touch technique further solutions
Laser-assisted All types Quick-and-easy technique without ICSI microscope has to be equipped with a
immotile sperm need for further solutions laser and a 40× laser objective
selection Easy to learn
this technique is not reproducible enough. A refinement of this technique is the use of a laser that when directed
onto the tip of the viable spermatozoon’s flagellum causes it to coil at the site of impact [21,22]. This technique
seems to be quick and easy, although the microscope has to have a laser and also a 40× laser objective has to be
available. The laser-assisted TESE-ICSI in severe nonobstructive azoospermia has been recently introduced into
clinical routine and results in increased fertilization rates [22].
Summary and Essentials

The analysis of a semen sample is a crucial point for the treatment decision in ARTs. The sequence of analyzing
each individual parameter should always be the same because some parameters are susceptible to change. After
liquefaction, the volume and appearance should be recorded. The next step should always be the measurement
of the pH, followed by motility assessment, identification of aggregation or agglutination, and determination
of vitality. Because these parameters may change over time, it is important to start with these measurements.
Especially pH, motility, and vitality are prone to variation due to external influences, for example, temperature
or air. The final steps are normally the evaluation of concentration and morphology. These factors are the least
susceptible to change because sperm numbers are not affected by external influences and morphology is assessed
on fixed, stained slides.
Only after appropriate analysis of the native semen sample can the right technique for its preparation be
chosen. Sperm preparation is as important as that of oocytes. A preparation method that is able to select the most
motile and morphologically normal spermatozoa from a semen sample is crucial, because the quality of the
gametes used constitutes the determinants for the outcome of the ART treatment.
The most used techniques are simple washing, swim-up, and DG centrifugation. Semen samples that possess
sperm with parameters within the normal range are processed primarily by swim-up; those with lower concen-
trations benefit from DG preparation. Although the swim-up procedure does not provide as high recovery rates
as gradient centrifugation, it is quick, easy to perform, and yields a population enriched with the best motile
spermatozoa. Gradient centrifugation results in the retrieval of high numbers of motile spermatozoa; however,
standardization of the method is difficult due to the inherent variation of semen samples.
Testicular or epididymal spermatozoa can be retrieved relatively easily using standard surgical techniques.
Sperm retrieval rates from testicular tissue depend on the surgical technique applied, the status of the testis,
and its underlying spermatogenic state. In cases of obstructive azoospermia, high retrieval rates are achieved.
The method of choice for surgical sperm retrieval in nonobstructive azoospermia is the microsurgical method
wherein focal areas with spermatogenic activity are selected. In nonobstructive cases where spermatogenic
abnormalities are the expected cause, spermatozoa retrieval might be laborious and time-consuming and should
always be handled with care.
All sperm preparation methods are necessary and should be done in a quality-controlled manner. By preparing
spermatozoa, the best selection is possible and guarantees high success rates in an ART program.
REFERENCES
1. WHO. WHO Laboratory Manual for the Examination of Human Semen, 5th edn. Cambridge: Cambridge
University Press, 2010.
2. Björndahl L, Kvist U. Sequence of ejaculation affects the spermatozoon as a carrier and its message. Reprod
Biomed Online. 2003;7:440–8.
3. Mortimer D. Laboratory standards in routine clinical andrology. Reprod Med Rev. 1994;3:97–111.
4. Mortimer D. Practical Laboratory Andrology. Oxford: Oxford University Press, 1994.
5. Björndahl L, Mohammadieh M, Pourian M, Söderlund I, Kvist U. Contamination by seminal plasma factors dur-
ing sperm selection. J Androl. 2005;26:170–3.
6. Marconi M, Keudel A, Diemer T, Bergmann M, Steger K, Schuppe HC, et al. Combined trifocal and microsurgi-
cal testicular sperm extraction is the best technique for testicular sperm retrieval in “low-chance” nonobstructive
azoospermia. Eur Urol. 2012;62:713–19.
7. Ramasamy R, Yagan N, Schlegel PN. Structural and functional changes to the testis after conventional versus
microdissection testicular sperm extraction. Urology. 2005;65:1190–4.
8. Barros A, Sousa M, Andrade MJ, Oliveira C, Silva J, Beires J. Birth after electroejaculation coupled to intracyto-
plasmic sperm injection in a gun-shot spinal cord-injured man. Arch Androl. 1998;41:5–9.
9. Shulman A, Feldman B, Madgar I, Levron J, Mashiach S, Dor J. In-vitro fertilization treatment for severe male
factor: The fertilization potential of immotile spermatozoa obtained by testicular extraction. Hum Reprod.
1999;14:749–52.
10. Ortega C, Verheyen G, Raick D, Camus M, Devroey P, Tournaye H. Absolute asthenozoospermia and ICSI: What
are the options? Hum Reprod Update. 2011;17:684–92.
11. Bourne H, Richings N, Liu DY, Clarke GN, Harari O, Baker HW. Sperm preparation for intracytoplasmic sperm
injection: Methods and relationship to fertilization results. Reprod Fertil Dev. 1995;7:177–83.
12. Casper RF, Meriano JS, Jarvi KA, Cowan L, Lucato ML. The hypo-osmotic swelling test for selection of
viable sperm for intracytoplasmic sperm injection in men with complete asthenozoospermia. Fertil Steril.
1996;65:972–6.
13. Ved S, Montag M, Schmutzler A, Prietl G, Haid G, van der Ven H. Pregnancy following intracytoplasmic sperm
injection of immotile spermatozoa selected by the hypo-osmotic swelling-test: A case report. Andrologia.
1997;29:241–2.
14. Sallam HN, Farrag A, Agameya AF, El-Garem Y, Ezzeldin F. The use of the modified hypo-osmotic swelling test
for the selection of immotile testicular spermatozoa in patients treated with ICSI: A randomized controlled study.
Hum Reprod. 2005;20:3435–40.
15. Nassar A, Mahony M, Morshedi M, Lin MH, Srisombut C, Oehninger S. Modulation of sperm tail protein tyrosine
phosphorylation by pentoxifylline and its correlation with hyperactivated motility. Fertil Steril. 1999;71:919–23.
16. Kovacic B, Vlaisavljevic V, Reljic M. Clinical use of pentoxifylline for activation of immotile testicular sperm
before ICSI in patients with azoospermia. J Androl. 2006;27:45–52.
17. Yildirim G, Ficicioglu C, Akcin O, Attar R, Tecellioglu N, Yencilek F. Can pentoxifylline improve the sperm
motion and ICSI success in the primary ciliary dyskinesia? Arch Gynecol Obstet. 2009;279:213–15.
18. Ebner T, Tews G, Mayer RB, Ziehr S, Arzt W, Costamoling W, et al. Pharmacological stimulation of sperm motil-
ity in frozen and thawed testicular sperm using the dimethylxanthine theophylline. Fertil Steril. 2011;96:1331–6.
19. Soares JB, Glina S, Antunes N, Jr, Wonchockier R, Galuppo AG, Mizrahi FE. Sperm tail flexibility test: A simple
test for selecting viable spermatozoa for intracytoplasmic sperm injection from semen samples without motile
spermatozoa. Rev Hosp Clin Fac Med Sao Paulo. 2003;58:250–3.
20. de Oliveira NM, Vaca Sanchez R, Rodriguez Fiesta S, Lopez Salgado T, Rodriguez R, Bethencourt JC, et al.
Pregnancy with frozen-thawed and fresh testicular biopsy after motile and immotile sperm microinjection, using
the mechanical touch technique to assess viability. Hum Reprod. 2004;19:262–5.
21. Aktan TM, Montag M, Duman S, Gorkemli H, Rink K, Yurdakul T. Use of a laser to detect viable but immotile
spermatozoa. Andrologia. 2004;36:366–9.
22. Nordhoff V, Schüring AN, Krallmann C, Zitzmann M, Schlatt S, Kiesel L, et al. Optimizing TESE-ICSI by laser-
assisted selection of immotile spermatozoa and polarization microscopy for selection of oocytes. Andrology.
2013;1:67–74.
2
Handling Gametes and Embryos: Oocyte
Collection and Embryo Culture
Lars Johansson
Introduction
A couple that undergoes a treatment in an assisted reproduction technique (ART) clinic expects that the clinic under-
takes all necessary pre-cautions to ensure that their embryos are cultured in a clean and safe environment, generating
embryos of a high implantation potential and the delivery of a healthy baby after a normal gestational period.
All efforts have been taken by the clinic to prevent the exposure of oocytes or embryos to embryo-toxic
pollutants from the surrounding environment (e.g., traffic, industrial hazards) or from within the clinic
(e.g., construction materials, ventilation system, furniture, lightning, gas cylinders, disposables, detergents, and
cloths). Accumulation of toxins, especially volatile organic compounds (VOCs) [1–6], in the culture media could
affect the embryos’ developmental and implantation capacities, increasing miscarriage rates and reducing a
couple’s chances of a successful treatment.
Oocyte Collection
Clinical Part
For the best outcome, the clinic should implement a mild stimulation protocol promoting recovery of high-
quality and mature oocytes from large follicles and an endometrium that supports implantation. In contrast,
excessive stimulation and high serum estradiol levels influence oocyte quality, increase oocyte dysmorphism
and epigenetic problems, decrease endometrial receptivity, and increase miscarriage rates [7–16]. The couple
should also be informed about the age-dependent decline in oocyte quality and quantity and how it affects
the outcome [17] before they sign the consent form.
Before an egg collection is undertaken, the nurse and the clinician verify together the identity of the patient and
inform the laboratory. Furthermore, they control the settings and functions of the equipment in the oocyte collection
room; this duty should at least include control of the pressure (90–120 mmHg) [16] and the flushing mechanism of
the aspiration pump. The ultrasound machine and probes are controlled, and the test tube warmer is loaded with
prewarmed test tubes which are kept at the correct temperature (36.5°C–36.9°C) [18]. Some clinics also preload the
warm test tubes with a low volume of flushing media containing heparin, to avoid formation of blood clots.
The oocyte collection is usually performed 36–38 hours post–human chorionic gonadotropin (hpHCG),
using a short-acting conscious sedation regime to avoid detrimental effects of anesthesia on oocytes, fertiliza-
tion, embryo development, and pregnancy rates [19–30]. Just before the oocyte collection starts, the needle is
rinsed with flushing media for removal of potential debris or contaminations from the manufacturing procedure.
Be aware of not touching the needle tip against the wall of the test tube, because a blunt tip will negatively affect
the ability to penetrate the follicle and may induce pain and bleeding in the patient.
Nowadays, most clinics retrieve the follicular fluid (FF) via transvaginal ultrasound guidance [31] and a thin
single-lumen needle (17–21 gauge) [32] that is connected to a pressure-controlled aspiration pump, thus avoiding
flushing of follicles.
17
FIGURE 2.1 Collection of follicular fluid (FF). Deep heat FIGURE 2.2 Flushing of follicles. Do not flush follicles
block is needed for keeping prewarmed test tubes at an with media in hand-held syringes because the media cool
acceptable temperature during the collection of the FF. Be down quickly, and the lower temperature might damage the
sure also that the transparent plug is properly inserted into the temperature-sensitive meiotic spindle in the oocyte.
test tube, so that the aspiration pressure can be maintained.
Also, see to that the tubings between the needle and the FF
collecting test tube are kept as short as possible to prevent a
drop in temperature.
Oocyte retrieval should be performed in a way that avoids damaging the oocyte via aspiration that
is too strong or exposure to suboptimal temperatures.
The oocyte collection is therefore quick and efficient, and it reduces the costs for anesthesia, disposables, and
flushing media. A thin aspiration needle also reduces the patient’s experience of pain and causes less bleeding,
thereby facilitating the laboratory being able to find and retrieve the cumulus–oophorous complexes (COCs)
from the almost clear FF [33].
The length of the tubing between the aspiration needle and the prewarmed test tube should be as short as pos-
sible to reduce temperature fluctuations that damage the meiotic spindle [34], increase aneuploidy rates, or entrap
the COC within a blood clot (Figure 2.1). If the latter occurs, the COC can be released from the blood clot by
cutting it with two large cannulas.
Double-lumen aspiration needles can also be used as long as the temperature of the flushing solution can
be kept within an acceptable range (36.5°C–36.9°C; [18]). Unfortunately, many clinics flush the follicles with
media kept in hand-held syringes in which acceptable temperatures cannot be maintained, causing unnecessary
damage to meiotic spindles and induction of aneuploidies (Figure 2.2) [35]. It also increases the risk of exposing
the oocytes to toxic lubricants from the syringes; if not embryo tested, one-component syringes are used [36].
In addition, COCs are mechanically damaged due to excessive pressure [37]. There is also evidence that a
brief exposure of the COCs to an inappropriate flushing buffer during the egg collection, such as phosphate-
buffered saline (PBS), might compromise the function of the oocyte and reduce embryo development and
implantation rate [38]. Regardless, a high proportion of clinics still use a double-lumen needle by which they
seem to retrieve more oocytes [32].
Laboratory Part
Every morning, the laboratory staff changes their clothes and places all their personal belongings (e.g., mobile
phones, jewelry, pens) in their lockers. After entry into the first coded, controlled prelaboratory room, the staff
Oocyte Collection and Embryo Culture 19
change their shoes, dress in lint-free and antistatic nonwoven gowns that cover the skin and hair, and perform
hand decontamination before entering the second prelaboratory room through an airlock chamber (air shower).
In this room, the laboratory stores unwrapped and fumed-off embryo-tested disposables in ventilated cupboards
and culture media in medical refrigerators.
From the latter room, the staff enter the cleanest, most sensitive and restricted area, the culture room, ideally
via a foot-controlled door opener. First, the embryologist performs a quality control (QC) of the facility and the
equipment (e.g., temperature, CO2, O2, pH, and gas inflow). The QC can either be performed via 24 hr surveil-
lance systems or by physically writing down the numbers.
In the afternoon, on the day before the egg collection, the laboratory turns off the heating of the laminar
airflow (LAF) bench and reduces its fan speed to prevent evaporation of media and potential increase in
osmolality. Equally important is that a turned off LAF bench is not restarted at the time of the prepara-
tion of the dishes and disposables since the filter initially releases dirt that contaminates the bench surface
area. If the LAF bench has been shut down overnight, it must run for at least 15–20 min before it is taken
into use.
Clean the LAF bench surface area with water for injection and lint-free clean-room wipes. The embryologist
puts on embryo-tested powder-free gloves, and another staff member retrieves fumed-off collection and culture
dishes, round-bottomed FF collection test tubes, serological pipettes, and combi- and filter-tips and places them
aseptically on the LAF bench. The number of test tubes and dishes (e.g., collection, insemination, denudation,
holding, and culture dishes) (Figure 2.3) are prepared in accordance with the expected number of retrieved
COCs, treatment, and culture techniques. Each disposable is labeled with the couple’s specific nontoxic identity
code by a diamond marker pen, cannula, or transparent name and barcoded sticker (www.mtg-de.com, www.
fertgms.com, www.research-instruments.com). More recently, a new laser-engraving system of the patient’s
name and barcodes is available (www.ankdatasystems.com). Both the lid and the bottom of the dishes are labeled
with the couple’s specific identifier.
After labeling of all test tubes and dishes, one of the culture media products is retrieved from the refrigerator.
The expire date of the product is entered into the QC overview of disposables and on the patient’s protocol, and
then the retrieved product is placed in the lid of a Petri dish to prevent the adherence of contaminants to the prod-
uct. This is especially important if preincubated products, such as liquid paraffin oil (LPO), are to be returned
back into the incubator. It is recommended that LPOs are stored at the dark and at 4°C–6°C to prevent excess for-
mation of peroxides or VOCs that could affect embryo quality and reduce implantation rate [39,40]. Therefore,
FIGURE 2.3 Selection of embryo-tested disposables. (Courtesy of Thermo Fisher.)

avoid prolonged storage of large volumes of LPO in incubators. Equally important is to not aliquot and transfer
media products into nonembryo-tested vials or flasks that potentially could contaminate or alter the product.
Prepare only one dish at a time if micro-drop cultures are to be used and avoid repeated retrieval
of culture media from the vial or flask.
Prerinse all disposables with excess media, just before they are to be used, so that potential toxins from the
manufacturing of the disposables are removed and the drying out of media, within the pipette, and a potential
increase in osmolality are avoided. Preferably, a multipipette, fitted with a prerinsed DNA, RNA, and pyrogenic-
free syringe, should be used for quick and safe dispensing of the culture media. Thus, because the multipipette is
filled only twice, it reduces the risk for contamination of the vial or the flask, allows quick, reproducible p ipetting
of droplets from a sealed unit, thereby prevents increase in osmolality and reduces; and reduces the laboratory
workload. If only one dish is prepared at a time, culture media droplets <40 μL–50 μL could be prepared, if
the droplets are quickly covered with preincubated LPO [41,42]. Avoid a too thick overlay of LPO over the
media droplets because this would require a longer preincubation of the media before it reaches its optimal pH.
Be aware that variability in pH during egg collection disrupts mitochondrial filaments and actins [41], and during
culture reduces cell numbers and formation and hatching of blastocysts, increases cell apoptosis, and decreases
fetal weights [42–44].
Clinics collect and rinse COCs in many different types of embryo-tested dishes, in combination with either
CO2-dependent or bench work media. The CO2-dependent media are preferably preincubated overnight in mini-
incubators, at low oxygen tension, wherein the media in the center or four-well dishes have been covered with a
thin film of LPO. In contrast, bench work media only need to be pipetted into test tubes; their corks should be
properly pushed down and prewarmed overnight in a bacteriological cabinet.
The patient’s dishes and test tubes are allocated to the patient’s incubator and bacteriological cabinet, respec-
tively, as noted in the patient’s embryo protocol. A nonsticky magnetic labeler on the incubator, verifying the
location of the dishes, reduces incubator openings and thereby negative effects on culture conditions.
Finally, the embryologist cleans the surface of the LAF workbench with water for injection and fumed off
empty collection and rinsing dishes and collector devices (e.g., embryo-tested Pasteur pipettes, yellow Eppendorf
Biopur filter tips, Stripper and glass denudation tips, and plastic tips or syringes) (Figure 2.4) are placed on its
surface.
Use embryo-tested material for all steps.
The decision of what type and size of embryo-tested collection and rinsing dishes (e.g., Nunc, Corning,
VitroWare) are to be used is that of the laboratory. For example, if a large dish is used, the full content of a test
tube can be poured in giving a thin layer of FF for easy localization of the COCs. However, the large search area
prolongs the search time and COCs might cool down. If a smaller Petri dish is used, the content of the test tube
must be divided into two dishes, thereby generating a thicker layer of FF with less fluctuation in temperature; but
again, this prolongs the search time. Remember that the surface temperature of the LAF bench is set such that it
generates a temperature of approximately 37°C within the selected dishes. Fluctuation in temperature should be
kept to a minimum because it causes irreversible damage to the meiotic spindle [35].
The optimal temperature must be checked at the location of the oocyte: in a dish/tube/….
On the oocyte collection day, the staff perform QC of the area and equipment, including a Køhler adjustment
of stereomicroscopes. The stereomicroscopes should be set at their lowest magnification for a quicker overview,
localization, and evaluation of the COCs within the FF. A light-emitting diode (LED) source, complemented
FIGURE 2.4 Devices for collection of cumulus–oophorous complexes (COCs). There are many different “collectors” of COCs
available in the market. In the LAF, you see, from the left, a sterile Pasteur pipette, Eppendorf yellow filter tip connected to an
Eppendorf pipette, and a Stripper with a large tip.
with different filters, reduces light-induced formation of reactive oxygen species (ROS) in the media, gives more
individual options, and facilitates the evaluation of the COCs [45,46].
After verification of the identity of the patient (e.g., double identification via wristband), the gas humidifier
(dish gasser), which delivers a media- and altitude-adjusted mix of humidified tri-gas over the preincubated
dishes, is turned on in the LAF bench. The clinician prepares the patient for the oocyte collection and rinses
the oocyte retrieval needle with flushing media. When the patient has been properly sedated, the laboratory
staff put on embryo-tested gloves, control the identity (www.fertqms.com, www.research-instruments.com) of
the two collection dishes (Figure 2.5), and place them under the dish gasser. Some clinics replace the media
droplets with preincubated media to remove potential toxins that have accumulated in the culture media via
the oil overlay.
When the first test tube with FF is placed in the block heater, the embryologist prerinses the collector and
pours out the FF into a prewarmed collection dish. The temperature-controlled high-efficiency particulate air
(HEPA) and VOC filter ventilation system might generate a slight draft through the hatch. The laboratory should
therefore avoid placing any heated blocks for test tubes within the passage of the hatch because the overpressure
and draft will cool them down. It is better to place the calibrated block heaters, for storage of FF-filled test tubes,
below the hatch, close to the LAF bench.
The FF is scanned and when a COCs is found one of the dishes from underneath, the dish gasser is moved next
to the collection dish. Preincubated media from the outer ring of the center-well dish is aspirated up into the col-
lector and flushed over the COC to clear the area around it from debris. The COC is taken up in the collector and
repeatedly washed in the outer ring of the center-well dish and when clean it is transferred into the center well.
The coloration of the FF is dependent on the nutritional status of the female, the vascularization of the follicle,
and contamination within the FF (Figure 2.6). Good blood supply to the follicles generates more oxygenation and
reduces aneuploidy rates and cytoplasmic abnormalities in the oocytes [47]. In fact, retrieval of FF from follicles
with a good blood supply, via Doppler imaging, is correlated to high oocyte quality developing into top-quality
embryos with high implantation rates [48].
Large blood clots attached to or entrapping the COC should be removed because they can reduce fertilization
rates [49] (Figure 2.7), whereas blood within the cumulus, generated during folliculogenesis, is an indication of
low oocyte quality and developmental capacity [50].
FIGURE 2.5 Follicular fluid. The color of the follicular fluid is FIGURE 2.6 Blood clot in a cumulus–oophorous
dependent on the nutritional status, food intake, and blood content. complex (COC). If a too slow aspiration of follicular
fluid is performed, a long blood clot can be formed that
adheres and entraps the COC.
The practice of reducing the cumulus cloud, by cutting, should be omitted because it increases the time that
the COCs are exposed to the environment, but perhaps more seriously damages the transzonal projections
(TZPs) that symbiotically and bidirectionally transfer information and nourishment between the cumulus and
the oocyte [51] (Figure 2.8).
After prewash, the COC is transferred into the center well, and the dish is placed under the dish gasser. When
the next COC is found, the second dish from underneath the dish gasser is used, thus alternating between the
dishes until you have a maximum of five COCs per dish or the oocyte collection is finished. The number and
quality of the COCs are evaluated (Table 2.1, Figures 2.8–2.15), and the dishes are placed in the patient’s allo-
cated mini-incubator that quickly recovers the temperature and pH of the culture media. The COCs are left in
the incubator, awaiting in vitro fertilization (IVF) insemination or partial denudation for intracytoplasmic sperm
injection (ICSI).
Work fast and concentrated and avoid too long exposure of COC/oocytes to the environment.
Retrieval of oocytes from small follicles, too early after HCG or to prevent OHSS, might generate morphologi-
cally good embryos, but it gives a much lower clinical pregnancy rate and a lower birth weight [52]. Cysts should
be emptied last to avoid contamination of disposables and culture media [53]. Finally, the surface area of the LAF
bench is cleaned with lint-free wipes and water for injection, gloves are discarded and all information is written
down. The area and staff are ready for the next oocyte collection.
It is very important that the COCs, and the developing embryos and blastocysts, are cultured at low oxy-
gen tensions, and preferably in mini-incubators with quick recovery of the culture conditions. Culture at low
oxygen tension is very beneficial because it lowers ROS formation, aneuploidy rates, and apoptosis; improves
embryonic gene expression, cleavage rate, and speed; generates a higher percentage of top quality embryos
and blastocysts; and increases pregnancy, implantation, cryopreservation, and baby take-home rates [54–60].
For this to become successful, the clinic must adjust the number of incubators to the number of patients so
that the frequency of incubator lid openings and embryo evaluation is kept to a minimum, or perhaps also
by using a reliable 24 hr embryo culture surveillance system. However, it also requires that the CO2 and O2
FIGURE 2.7 Culture techniques. Cultures can be FIGURE 2.8 Immature (I) cumulus–oophorous complex (COC).
performed in either small microdrops or in open large Immature COC with a compact nonexpanded cumulus and a nonvisible
dishes. oocyte. The COC is adhered to the dark follicle wall cells (DFWCs).
(Courtesy of Dr. Maria Köster, University of Bonn.)
TABLE 2.1
Classification of Cumulus Oophorous Complexes
Quality Cumulus Corona Oocyte
Immature (I) Small, compact, grey, Dark ring around the zona pellucida Not visible, if visible GV
(Figure 2.8) non-expanded stage
Mature (M) Small, bright, expanded Not fully radiated Partially visible
(Figure 2.9–2.11)
Excellent (E) Large, bright, expanded Radiated Clear and visible
(Figure 2.12–2.13)
Overmature (O) Small, thin, patches of dark cells Light or dark Visible, granular and dark
(Figure 2.14–2.15)
Atretic (A) Patches of dark cells or absent None or clumps Dark
concentrations have been adjusted to the requirement of the culture media, stage of development, and altitude
of the clinic.
Exposure of oocytes to variable pHs during handling, denudation, or ICSI affects embryo development and
blastulation rates [44]. For example, completely denuded oocytes, before ICSI or cryopreservation, cannot regu-
late their intracellular pH (pHi) and are highly dependent on the extracellular pH (pHe), whereas the later stages
of development are less sensitive, probably due to the formation of tight junctions [61–69]. In addition, prolonged
exposure of oocytes to high concentrations of sperm in IVF lowers the fertilization rate because the metabo-
lism and decomposition of sperm lowers the pHe [70] and excess formation of ROS affects the development of
the embryo [60]. However, inclusion of amino acids in handling and culture media could preserve the pHi and
protect gametes and embryos [71–73].
FIGURE 2.9 Mature (M) COC surrounded with expanded or FIGURE 2.10 Mature (M) COC surrounded with expanded or
nonexpanded cumulus. A mixture of mature COC with nonex- nonexpanded cumulus. A mixture of mature COC with nonex-
panded (MNE) or expanded (ME) cumulus. The dark patches panded (MNE) or expanded (ME) cumulus. The dark patches
within the cumulus cells represent blood (B). (Courtesy of within the cumulus cells represent blood (B). (Courtesy of
Dr. Maria Köster, University of Bonn.) Dr. Maria Köster, University of Bonn.)
FIGURE 2.11 Mature (M) COC surrounded with FIGURE 2.12 Excellent (E) mature COC. Large, bright expanded
expanded or nonexpanded cumulus. A mixture of cumulus with radiated corona (CR) and visible oocyte with polar body
mature COC with nonexpanded (MNE) or expanded (PB). (Courtesy of Lev Levkov.)
(ME) cumulus. The dark patches within the cumulus
cells represent blood (B). (Courtesy of Dr. Maria Köster,
University of Bonn.)
FIGURE 2.13 Excellent (E) mature COC. Large, bright expanded FIGURE 2.14 Overmature (O) COC. An overmature
cumulus with radiated corona (CR) and visible oocyte with polar body COC with a small cumulus, patches of dark cumulus cells
(PB). (Courtesy of Lev Levkov.) (DCCs), and a v isible oocyte. (Courtesy of Dr. Maria
Köster, University of Bonn.)
FIGURE 2.15 Overmature (O) COC. An overmature COC with a small cumulus, patches of dark cumulus cells (DCCs), and a visible
oocyte. (Courtesy of Dr. Maria Köster, University of Bonn.)
Control and maintenance of proper pH and temperature are key to success.
If oocytes and embryos also are cultured in dishes with a flat surface that is in direct contact with the heated
surface of the incubator, there will be less variation in temperature and less damage to meiotic spindles, aneu-
ploidy rate, and delay in embryo development.
After fertilization and scoring of the quality of the zygotes, they are either cultured singly or in
groups, and preferably in droplets. In single culture, the embryo development and quality can be succes-
sively evaluated, facilitating the selection of the best embryos for transfer. Group culture, in contrast, is
sought to improve implantation rate and to rescue borderline-quality embryos for cryopreservation [74].
Group culture in concavely shaped grooves separated by a low barrier allows both the exchange of nutrients
between the embryos and the successive evaluation of embryos [75–79]. In this embryo-friendly culture
environment, high concentrations of autocrine and paracrine embryotrophic and detoxifying factors sur-
round the embryos [77], but the exchange of nutrients and metabolites is limited by diffusion, which is
estimated to be 80 μm–120 μm [76]. The exchange of nutrients between the embryos and the proliferation
of cells are thought to be enhanced by tilting the incubators or dishes and by mechanical or indirect vibra-
tion of dishes [80–82], thereby mimicking the movements of the fallopian tubes and the uterus [83,84]
and generating a higher proportion of high-quality blastocysts and higher implantation and live birth rates.
A high density of embryos generates higher concentrations of embryotrophic factors, but it also gener-
ates a potential enrichment of detrimental waste products that could be dependent on media product and
on culture technique. The combination of group culture and coculture of embryos with autologous cumu-
lus or endometrial cells m imics the m icroenvironment that surrounds the embryos; generates growth
factors that induce better embryo growth and earlier compaction of embryos; reduces apoptosis; increases
cell numbers, blastocyst formation, hatching ability and pregnancy rate; and makes the blastocysts more
resistant to cryo-damage [85–94].
The choice of culture media products [95] or the day of embryo transfer [96,97] does not seem to affect the
pregnancy rate, even though higher implantation and lower aneuploidy rates are seen in embryos that reach the
blastocyst stage [98].
There is also insufficient clinical evidence favoring the single or the sequential media systems. The single-
step culture media system decreases environmental stress and removal of valuable nutrition (e.g., autocrine and
paracrine factors), it is less laborious and costly, and it generates more blastocysts. In contrast, a sequential media
system takes into consideration the changes in metabolism and the environment that the embryo encounters
during passage through the female reproductive tract [99,100].
Practical Guide in Preparation for Oocyte Collection and Culture Techniques

Every morning and before entering the restricted culture area, the staff must be properly dressed. In the
pre entry room, staff redress according to the culture room dress code and perform hand decontamina-
tion. Upon entry into the culture room, the staff perform QC of the area and equipment before starting the
day’s work.
BOX 2.1 EQUIPMENT AND MATERIALS

The equipment and materials suggested give consistent, good results in clinics audited worldwide by the
author.
The disposables are sterile, nonembryotoxic, and nonpyrogenic, but there are many additional regional
alternatives that can be used.
The following protocols should be considered only as a guide to avoid the most frequent problems
during egg collection and culture routines.
Equipment and materials

1. LAF bench, class II cabinet, or IVF chamber (hereafter called LAF bench)
a. A dish gasser that purges a temperature-controlled, humidified, pH- and altitude-adjusted
tri-gas mix over the dishes is recommended instead of using a small CO2 incubator within
the LAF bench. The recovery of the culture environment within the latter incubator type
is slow and affects the culture conditions, thereby potentially harming the COCs and the
development of the embryos.
2. Labeling machine, diamond marker pen, or cannula
3. Incubators
a. Large (60 L) with small inner doors and low oxygen
i. Dishes kept in metal blocks for uniform heating
b. Benchtop mini-incubator, premixed gas or gas mixer, and low oxygen
i. Most types of dishes fit within the incubator, and they are in direct contact with the
heated metal surface
4. Pipette aid, pipetter, and pipettes (www.eppendorf.de)
a. EasyPet 4421: for transfer of preincubated oil or large volumes of culture media with sero-
logical pipettes (Nunc)
b. Repeater Plus or Repeater Xstream: the pipette is used in combination with Combi Plus Tips
of Biopur quality (RNA, DNA, and pyrogen free)
i. Use single-wrapped repeater tips of 0.1–50 mL sizes
c. Pipettes: Research Plus kit, possible to autoclave
i. These pipettes are used with filter tips of Biopur quality
Materials
1. Gloves
a. Embryo-tested, powder-free sterile gloves without chemical additives, accelerants, or
emulsifiers
i. BioClean N-Plus, sterile nitrile gloves for clean-room
ii. Ansell, TNT 92-760
iii. Ansell, Derma Prene Type 2-RT
iv. Kimberly-Clark, Safeskin NXT 62992
2. Collection pipettes (hereafter called collectors)
a. Pasteur pipettes with cotton plug (Humagen, Hunter), stripper, or Flexipet tips (600 μL);
glass pipettes (Humagen), or Eppendorf Yellow Biopur filter tips
3. Cleaning solution for surfaces in-between patients
a. Wipe only with water for injection
b. Soiled surfaces are cleaned with embryo-tested detergents that should be used only after all
work has been done to avoid exposure of gametes, embryos, and environment to potentially
toxic components
i. Oosafe, hydrogen peroxide, and 70% ethanol are the most frequently used detergents,
but they need extensive cleaning with water for injection after use (Catt, ESHRE 2013,
O–240).
4. Lint-free wipes for clean-room work (www.techniwipe.com)
5. Round-bottom test tubes for collection of FF
a. Nunc, 150268, VitroWare, Corning
6. Serological pipettes
a. 5 and 10 mL (Nunc, 159625 and 159633)
7. Collection and culture dishes
a. Select the choice of preference as long as they are embryo tested with several different meth-
ods, preferably gamma-irradiated and of clear plastic, facilitating inspection and evaluation
of gametes and embryos (e.g., Nunc, LifeGlobal, Corning, VitroWare).
8. Culture media of your choice
BOX 2.2 PROCEDURE

Day 1: Preparations on the day before oocyte collection
1. Turn off the heating of the LAF bench where the dishes are to be prepared.
a. The ventilation of the LAF bench should have been turned on before the area is cleaned to
prevent release of dirt from the filter during the initial start of the LAF bench.
Note: Use a low fan speed to reduce evaporation rate of the media during preparation of the
dishes.
b. Clean the area with water for injection and lint-free clean-room wipes.
c. Put on embryo-tested gloves and work aseptically.
2. Place fumed-off collection and culture dishes, round-bottom FF collection test tubes, serological
pipettes, and Combitips and filter tips on the surface of the LAF bench.
a. Clean the outer part of the Pipette Aid, Repeaters, and metal blocks (for large incubators)
with wipes moist with water for injection.
b. Adjust the number of dishes according to the number of expected COCs per patient and
culture techniques.
c. Label dishes, both lid and bottom, with the couple’s specific barcode identifier.
Note: Prerinse all disposables, with their designated media, just before use to avoid drying
of media and potential increases in osmolality.
3. Take out the media of choice, one at a time, for rinsing of COCs, insemination (IVF), denudation
(ICSI), and culture, from the refrigerator and place the vials, flasks, or bottles are placed in a
Petri dish lid on the cold and clean surface of the LAF bench.
a. Also, take out the preincubated liquid paraffin oil (PLPO) from the incubator.
4. Preparation of rinsing and collection media
Bench work media (HEPES or MOPS)
1. Place four round-bottomed test tubes in a metal heat block with holes of a depth that covers
most of the length of the test tubes.
2. Fill the test tubes completely with media for cleaning of the aspiration needle, before and
after the oocyte collection, and for rinsing of the COCs during the collection.
3. Push down the cork of the test tubes and place the metal block, containing the test tubes, for
warming overnight in a bacteriological cabinet (37°C).
Note: The day before the egg collection, the nurse places round-bottom test tubes for pre-
warming in the bacteriological cabinet of the oocyte retrieval room. Additional test tubes
with bench work media can be prepared for unforeseen additional work.
CO2-dependent media
1. Prepare and place two round-bottom test tubes in the metal block for cleaning of the aspira-
tion needle before and after oocyte collection.
2. Prepare, in accordance to the number of large follicles, several center-well dishes containing
CO2-dependent media for holding collected COCs.
a. Aspirate and rinse a suitable-sized Combitip, refill it, and adjust the dispense volume.
b. Fill the outer ring with 1.5 mL and the inner well with 600 μL of holding media, usually
the fertilization media.
c. Cover the inner center well with a thin layer of PLPO.
i. Preincubate the dishes overnight or at least for 8 hr.
ii. When five COCs have been collected in each dish, they are returned to the
mini-incubator.
Note: These dishes are ready for insemination (IVF).
iii. Select a flat-bottom dish for insemination of COCs in microdrops. Number each
droplet.
iv. Aspirate and rinse the selected Combitip, refill it, and adjust the dispense volume to
the required droplet size (40 μL–50 μL).
v. Dispense the droplets according to your preference, preferably four prerinsing
droplets in the middle and insemination droplets at the prelabeled numbers (1–5).
vi. Cover the droplets immediately with PLPO to avoid evaporation and detrimental
changes in osmolality.
Note: Only prepare one dish at a time!
Note: Do not place the droplets at the edge of the dish because it causes optical
interference and a drop in temperature toward the edge of the dish.
5. For ICSI patients, you also prepare dishes for the following.
1. Denudation
a. Four-well dish
i. Aspirate and rinse a Combitip, refill it, and dispense 500 μL of fertilization medium
in all four wells.
Note: One four-well dish per every five COCs.
ii. Place the dishes in the incubator for preincubation overnight.
b. Flat-bottom 35 mm Petri dish
i. Rinse a Combitip with fertilization media, refill it, and adjust the volume to
100 μL–200 μL and dispense the media in the upper part of the dish.
ii. Underneath the large droplet, dispense three additional 50 μL droplets of fertil-
ization media to be used for rinsing and scoring of the partially denuded oocytes
before they are transferred to the holding dish of the oocytes.
iii. Cover the droplets immediately with PLPO.
iv. The next day, the large upper droplet is aspirated with a yellow Eppendorf filter tip
and is replaced with 100 μL–200 μL of prewarmed nontoxic hyaluronidase (ICSI
Cumulase).
2. Holding dishes for denuded oocytes.
3. Oocyte injection dishes (Nunc, Corning, or VitroWare)
Notes:
• The ICSI Injection dishes can be prepared in many different ways, but the method should
be standardized within the laboratory.
• Select a flat-bottom ICSI dish that is in direct contact with the heated surface of the
inverted microscope.
• A dish with a low wall does not restrict the movement of the ICSI pipettes!
• If the media droplets are placed in the middle of the dish, it is easier and quicker to
perform the ICSI procedure.
• Minimize the exposure of the oocytes to the outer environment by keeping the number
of oocytes per dish at a minimum, which speeds up the ICSI procedure.
a. Quickly cover the fertilization media droplets with PLPO.
b. The next day, just before the ICSI procedure starts one of the droplets is replaced
by prewarmed polyvinylpyrrolidone (PVP) or Sperm-Slow (SS).
c. Return the dish to the incubator or preferably to the dish gasser for short rewarming
and preincubation before the sperm and oocytes are introduced.
4. Culture dishes for injected (ICSI) or short-time (1–4 hr) inseminated oocytes (IVF).
6. Place the dishes in the patient’s allocated mini-incubator for preincubation overnight. Fill in the
location of the dishes on the patient’s embryo protocol (paper or software).
7. Confirm that you have enough FF collection dishes (Nunc, 35 mm) for warming and a eration on
the heated and calibrated surface of the LAF bench.
a. Warm additional back-up dishes in the bacteriological cabinet overnight.
8. Prepare the oocyte collectors and place them in the LAF bench for aeration overnight.
Note: Avoid mouth-pipetting, for safety reasons, and pulling of denudation pipettes, which will
vary in dimensions, because they can damage oocytes and prevent standardization.
BOX 2.3 OOCYTE COLLECTION AND INSEMINATION

Day 0: Oocyte collection and insemination
1. Adjust the light source and perform Køhler adjustment of the stereomicroscope.
Note: The more efficiently you work, the less exposure of the COCs to the outer environment and
the better the embryo development.
2. Verify the identity of the patient (double identification, wristband).
3. When anesthesia has been given, two prewarmed test tubes containing bench work media are
placed in the test tube warmer in the oocyte retrieval room.
4. Turn on the gas humidifier in the LAF bench that delivers humidified, premixed gas to the dish
gasser.
5. The clinician prerinses the aspiration needle with the bench work media and controls simultane-
ously the pump pressure.
Note: Perform oocyte collection 36–38 hpHCG
6. Meanwhile, the embryologist puts on the correct-size embryo-tested nontoxic gloves and places
two of the patient’s center-well dishes under the dish gasser.
7. The embryologist prerinses one of the collectors with culture media, from the outer ring of the
center-well dish, when the first test tube with FF is placed in the heat block next to or inside
the LAF bench.
8. The contents of the test tube are poured out into prewarmed Petri collection dishes, and the test
tube is discarded.
9. The FF is scanned through and when a COC is found
A. CO2-dependent media
i. One of the dishes, underneath the dish gasser, is moved to the side of the collection dish,
and preincubated media from the outer ring of the center-well dish is aspirated up into
the prerinsed collector.
ii. Preincubated media are flushed over the COC, to clear the area around the COC from
debris, and the COC is taken up into the collector with as little contamination as
possible.
iii. The COC is repeatedly washed in the outer ring of the center-well dish and when clean,
it is transferred into the center well.
iv. The dish is then returned to its position under the dish gasser, and the collection dish is
discarded in the waste bin.
v. When the next COC is found, the second dish under the dish gasser is used, thus alter-
nating between the dishes until you have a maximum of five COCs per dish or the oocyte
collection is finished.
vi. When the oocyte collection is finished, the number and quality of the COCs
are evaluated (Table 2.1), and the dishes are placed in the patient’s allocated
mini-incubator.
vii. The bench surface area is cleaned with lint-free wipes and water for injection, and the
gloves are discarded.
viii. Fill in all paper or software protocols.
ix. The area and staff are ready for the next oocyte collection.
Alternatively you can use the following protocol.

B. Bench work media (HEPES or MOPS)
i. Follow the instructions in points 7 and 8.
ii. Pour out the FF of the first test tube in the prewarmed collection dishes.
iii. Quickly pour in prewarmed media, from the bacteriological cabinet, into the outer and
inner rings of a prewarmed center-well dish, for rinsing and collection of COCs.
iv. Locate the COC and aspirate it with the prerinsed collector.
v. Wash the COC repeatedly in the outer ring of the center-well dish, and when clean it is
transferred into the center well.
vi. When four or five COCs have been collected or when the dish has been on the LAF
bench for 3–5 min, one of the patient’s preincubated culture dishes is placed on the sur-
face of the LAF bench.
a. Wash the COCs repeatedly in the outer ring of the culture dish, and when clean
they are transferred into the center well and placed in the patient’s allocated
mini-incubator.
10. Let the COCs “rest” in the mini-incubator until it is time for insemination (IVF) or denudation
(ICSI).
11. Insemination (IVF)
A. Short-time insemination (1–4 hr) in droplets (40–42 hpHCG)
i. Verify the identity of the couple’s prepared, diluted, and well-mixed sperm suspension.
ii. Place one of the culture dishes with microdroplets on the heated surface of the LAF
bench.
iii. Prerinse a gray filter tip (1 μL–10 μL) and take up a standard aliquot of the sperm sus-
pension and inseminate the microdroplets (20,000 motile sperm per droplet).
a. Confirm that the droplets have been properly inseminated and incubate the dish for
30 min before the COCs are transferred into the droplets.
iv. Place one culture and insemination dish close to each other on the heated surface of the
LAF bench.
v. Quickly prerinse a yellow filter tip (Biopur quality) with culture media from
one of the centrally located droplets and aspirate one of the COCs from the
center-well dish.
vi. Wash the retrieved COC in two of the centrally located droplets, and then put it into one
of the inseminated microdroplets.
vii. Place the dish in the patient’s allocated mini-incubator.
viii. Incubate the dish in accordance to the preference of the laboratory.
ix. After the selected insemination time, the dish is retrieved from the mini-incubator and
placed on the heated surface of the LAF bench.
x. Prerinse a stripper tip (175 μm), quickly aspirate the oocytes, and then expel them into
the unused third centrally located droplet. Finally, place them in the fourth droplet with
as little debris as possible.
Note: Do not denude the oocytes; just aspirate and transfer them to a clean droplet.
xi. Return the dish to the patient’s allocated mini-incubator.
B. Overnight insemination in a center-well dish
i. Verify the identity of the couple’s prepared, diluted. and well-mixed sperm suspension.
ii. Prerinse a gray filter tip (1 μL–10 μL), aspirate a standard aliquot of the sperm suspen-
sion, and expel it into the collection dish containing the COCs (100,000 motile sperm
per dish and five oocytes).
iii. Place the dish in the patient’s allocated mini-incubator overnight.
12. Denudation of oocytes (40–41 hpHCG) followed by ICSI within 30–60 min
A. Four-well dish
i. Prewarm a vial of ICSI Cumulase for 30 min in a block heater.
ii. Retrieve the four-well dish containing the fertilization media from the incubator.
iii. Quickly replace the fertilization media in the first well with prewarmed ICSI Cumulase
and place the dish under the dish gasser.
iv. Retrieve one of the patient’s collection dishes with the COCs from the incubator and
place it close to the light source of the LAF bench.
v. Move the denudation dish from underneath the dish gasser, and put it close to the collec-
tion dish.
vi. Quickly prerinse an Eppendorf yellow filter tip and a 150 μm Stripper tip with ICSI
Cumulase.
vii. Transfer the five COCs to the first well of the denudation dish, with the yellow filter tip,
and gently denude the COCs by repeatedly aspirating and expelling the COCs into the
ICSI Cumulase solution.
−− When a few layers of the cumulus are left, shift to the Stripper tip and transfer the
partially denuded oocytes to the second well and continue the denudation.
−− Wash the partially denuded oocytes in the remaining wells.
−− Score the quality and maturation of the oocytes according to Table 2.2.
−− Take out a holding dish (see Holding dishes (2.4.5- CO2 dependent media, page 32,
#2) for denuded oocytes in Box 2.2) from the mini-incubator and transfer the
denuded oocytes to the dish.
−− Place the dish in the patient’s allocated space in the mini-incubator.
B. Denudation of oocytes in droplets
i. Prewarm a vial of ICSI Cumulase for 30 min in a block heater.
ii. Retrieve the droplet denudation dish from the incubator.
iii. Quickly replace the upper big fertilization media droplet with prewarmed ICSI Cumulase
and place the dish under the dish gasser.
iv. Retrieve one of the patient’s collection dishes with the COCs from the incubator and
place it close to the light source of the LAF bench.
v. Move the denudation dish from underneath the dish gasser and put it close to the collec-
tion dish.
vi. Quickly prerinse an Eppendorf yellow filter tip and a 150 μm Stripper tip with ICSI
Cumulase.
vii. Transfer five COCs to the large ICSI Cumulase droplet, with the yellow filter tip, and
gently denude the COCs by repeatedly aspirating and expelling the COCs into the ICSI
Cumulase solution.
−− When a few layers of the cumulus are left, shift to the Stripper tip and transfer the
partially denuded oocytes to the first rinsing droplets and continue the denudation
as described in “Four-well dish.”
TABLE 2.2
Classification of the Oocyte Maturational Stages
Stage Polar Body Nucleus
Germinal vesicle (GV) None Large halo with nucleoli
Meiosis I (MI) None None
Meiosis II (MII) Present None
TABLE 2.3
Timing of Procedures in Relationship to Injection of HCG (hpHCG)
Procedure Egg Collection Oocyte Vitrification Insemination (IVF) Denudation ICSI
hpHCG 36–38 38–40 40–41 40–41 41–42
13. Final preparation of the ICSI dishes

Note: The dishes can also be prepared the same day; the oocyte-holding droplets are replaced
by prewarmed HEPES- or MOPS-buffered media.
A. Take out the preprepared ICSI dishes from the mini-incubator and exchange one of the drop-
lets against PVP or SS.
B. Add a low volume of highly motile sperm suspension to the edge of PVP or SS for swim-out
and selection of sperm.
C. Add a few partially denuded oocytes and perform ICSI as quickly as possible.
D. After ICSI, the oocytes are transferred back to the patient’s culture dish.
i. Wash the oocytes in the centrally located droplets and keep the oocytes in the outer
numbered droplets.
ii. Place the dish in the patient’s allocated incubator.
iii. Repeat the procedure until all oocytes have been injected. (See Table 2.3, Timing of
Procedures.)
14. Preparation of dishes for culture of fertilized oocytes (Zygotes, 2PN).
A. The culture dishes are prepared in accordance to the steps discussed in Box 2.2, but for
culture of zygotes to Day 2–3 or for Day 4–6 embryo (morula and blastocysts) cleavage,
respectively, blastocyst culture media are used.
i. There are also single-step media available that can be used from the zygote to the blas-
tocyst stage without the need for changing of the media.
BOX 2.4 FERTILIZATION AND CLEAVAGE

Day +1: Control of fertilization (16–18 hpInsemination [hpInsem])
1. Clean the area with water for injection and lint-free clean-room wipes.
2. Verify the location of the couple’s dishes.
3. Put on embryo-tested gloves and work aseptically.
4. Adhere a plastic or a glass denudation tip to a stripper (150 μm) or a Humagen handle.
5. Take out the patient dish from the incubator and verify the identity of the dish.
6. Place the dish under the dish gasser and retrieve the patient’s preincubated culture dish, containing
cleavage stage media, from the mini-incubator.
7. Prerinse the denudation pipette with media, from one of the droplets in the dish containing the
oocytes and strip the oocytes of their remaining cumulus cells.
8. Gently roll the oocytes while evaluating the fertilization.
9. Retrieve all normally fertilized oocytes (2PN) and wash them in the centrally located droplets of
the cleavage stage media and then group culture them in one of the outer droplets.
i. Separate unfertilized, single and multinucleated oocytes and transfer them to their allocated
droplets.
ii. Note in which droplet they are cultured.
iii. Score early cleavage 25–27 and 27–29 hpInsem for ICSI and IVF, respectively.
Day +2–3: Control of embryo cleavage and quality (43–45 resp. 67–69 hpInsem)
Day +4–5: Control of compaction and blastocyst stage (90–94 resp. 114–118 hpInsem)
For information on how to perform and morphologically evaluate oocytes and perform scoring of
zygotes, embryos, and blastocysts, see Chapters 5 through 8 in this book.
REFERENCES
1. Doshi A, Karunakaran S, Worrilow K, et al. What makes an IVF laboratory successful? In: A Practical Guide
to Setting Up an IVF Lab, Embryo Culture Systems and Running the Unit (eds. Varghese A, Sjoblom P,
Jayaprakasan K). New Delhi: Jaypee Brothers Medical Publishers, 2013, pp. 13–23.
2. Johansson L. Establishment of an ART clinic: Location, construction and design. In: A Practical Guide to Setting
Up an IVF Lab, Embryo Culture Systems and Running the Unit (eds. Varghese A, Sjoblom P, Jayaprakasan K).
New Delhi: Jaypee Brothers Medical Publishers, 2013, pp. 24–30.
3. Schimmel T, Gilligan A, Garrisi GJ, et al. Removal of volatile organic compounds from incubators used for gam-
ete and embryo culture. Reprod Fertil 1997;68(Suppl 1):S52–S53.
4. Khoudja RY, Xu Y, Li T, et al. Better IVF outcomes following improvements in laboratory air quality. J Assist
Reprod Genet 2012;30(1):69–76.
5. Worrilow KC. The Impact of UVC Irradiation on Clinical Pregnancy Rates in an ISO 5 Cleanroom In Vitro
Fertilization Laboratory. ASHRAE IAQ Guide, ASHRAE, Atlanta, 2009, pp. 4–6.
6. Worrilow KC, Huynh HT, Bower JB, et al. A retrospective analysis: Seasonal decline in implantation
rates (IR) and its correlation with increased levels of volatile organic compounds (VOC). Fertil Steril
2002;78(Suppl 1):S39.
7. Baart EB, Martini E, Eijkemans MJ, et al. Milder ovarian simulation for in vitro fertilization reduces aneuploidy
in the human preimplantation embryo: A randomized controlled trial. Hum Reprod 2007;22:980–988.
8. Pena JE, Chang PL, Chan LK, et al. Supraphysiological estradiol levels do not effect oocyte and embryo quality
in oocyte donation cycles. Hum Reprod 2002;17:83–87.
9. Hohmann FP, Macklon NS, Fauser BC. A randomized comparison of two ovarian stimulation protocols with
gonadotropin-releasing hormone antagonist cotreatment for in vitro fertilization commencing recombinant fol-
licle stimulating hormone on cycle 2 or 5 with standard long GnRH agonist protocol. J Clin Endocrinol Metab
2003;88:166–173.
10. Fauser BC, Devroey P, Macklon NS. Multiple birth resulting from ovarian stimulation for subfertility treatment.
Lancet 2005;365:1807–1816.
11. Macklon NS, Fauser BC. Impact of ovarian hyperstimulation on the luteal phase. J Reprod Fertil 2000;55:101–108.
12. Howles CM, Macnamee MC, Edward RG, et al. Effect of high tonic levels of luteinizing hormone on the outcome
of in-vitro fertilization. Lancet 1986;2(8505):521–522.
13. Homburg R, Armar NA, Eshel A, et al. Influence of serum luteinizing hormone concentrations on ovulation,
conception and early pregnancy loss in polycystic ovary syndrome. Br Med J 1988;297:1024–1026.
14. Regan L, Owen EJ, Jacobs HS. Hypersecretion of luteinizing hormone, infertility and miscarriage. Lancet
1990;336:1141–1144.
15. Outsuki J, Okada A, Morimoto K, et al. The relationship between pregnancy outcome and smooth endoplasmic
reticulum clusters in MII human oocytes. Hum Reprod 2004;19:1591–1597.
16. Hassan-Ali H, Hisham-Saleh A, El-Gezeiry D, et al. Perivitelline space granularity: A sign of human menopausal
gonadotropin overdose in intracytoplasmic sperm injection. Hum Reprod 1998;13:3425–3430.
17. Nagaoka I, Hassold TJ, Hunt PA. Human aneuploidy: Mechanisms and new insights into an age-old problem.
Nat Rev Genet 2012;13:493–504.
18. Wikland M. Oocyte retrieval. In: In Vitro Fertilization – A Practical Approach (ed. Gardner D). New York: Informa
Health Care Inc., 2007, pp. 117–126.
19. Higdon H, Blackhurst D, Boone W. Incubator management in an assisted reproductive technology laboratory.
Fertil Steril 2008;89:703–710.
20. Endler GC, Stout M, Magyar DM, et al. Follicular fluid concentrations of thiopental and thiamylal during lapa-
roscopy for oocyte retrieval. Fertile Steril 1987;48:828–833.
21. Hayez MF, Sacco AG, Savoy-Moore RT, et al. Effect of general anaesthesia on fertilization and cleavage of
human oocytes in vitro. Fertil Steril 1987;48:975–981.
22. Gonen O, Schulman A, Gethler Y, et al. The impact of different types of anaesthesia on in vitro fertilization-
embryo transfer treatment outcome. J Assist Reprod Genet 1995;12:678–682.
23. Matt DW, Steingold KA, Dastvan CM, et al. Effects of sera from patients given various anaesthetics on pre-
implantation mouse embryo development in vitro. J In Vitro Fert Embryo Transf 1991;8:191–197.
24. Soussis I, Boyd O, Paraschos T, et al. Follicular levels of midazolam, fentanyl and alfentanil during transvaginal
oocyte retrieval. Fertil Steril 1995;64:1003–1007.
25. Wilhelm W, Hammamed ME, White PF, et al. General anaesthesia versus monitored anaesthesia care
with remifentanil for assisted reproductive technologies: Effect on pregnancy rate. J Clin Anesthesia
2002;14:1–5.
26. Bokhari A, Pollard BJ. Anaesthesia for assisted conception: A survey of UK practice. Eur J Anesthesiol
1999;16:225–230.
27. Wikland M, Evers H, Jacobsson AH, et al. The concentration of lidocaine in follicular fluid when used for para-
cervical block in a human IVF-ET programme. Hum Reprod 1990;5:920–923.
28. Ben-Shlomo I, Moskovich R, Golan J, et al. The effect of propofol anaesthesia on oocyte fertilization and early
embryo quality. Hum Reprod 2000;15:2197–2199.
29. Hadimiglou N, Titz T, Dosemeci L, et al. Comparison of various sedation regimes for transvaginal oocyte
retrieval. Fertil Steril 2002;78:648–649.
30. Jain D, Kohli A, Gupta L, et al. Anaesthesia for in vitro fertilization. Indian J Anaesth 2009;53(4):408–413.
31. Garg R, Dali JS. Assisted reproductive technology and anesthetic considerations: Review of literature. Int
J Anesthesia 2008;18(2).
32. Steiner HP. Optimizing technique of follicular aspiration and flushing. In: Minimal Stimulation IVF – Milder
Mildest or Back to Nature (eds. Chaves-Badiola A, Allahbadia GN). New Delhi: Jaypee Brothers Medical
Publishers, 2011, pp. 98–102.
33. Awonuga A, Waterstone J, Ovesanya O, et al. A prospective randomized study comparing needles of different
diameters for transvaginal ultrasound-directed follicle aspiration. Fertil Steril 1996;65:109–113.
34. Wikland M, Blad S, Bungum L, et al. A randomized controlled study comparing pain experience between a
newly designed needle with a thin tip and a standard needle for oocyte aspiration. Hum Reprod 2011;26(6):
1377–1383.
35. Pickering SJ, Braude PR, Johnson MH, et al. Transient cooling to room temperature can cause irreversible
disruption of the meiotic spindle in the human oocyte. Fertil Steril 2009;54:102–108.
36. Nijs M, Franssen K, Cox A, et al. Reprotoxicity of intrauterine insemination and in vitro fertilization-embryo
transfer disposables and products: A 4-year survey. Fertil Steril 2009;92(2):527–535.
37. Cohen J, Avery S, Campbell S, et al. Follicular aspiration using a syringe suction system may damage the zona
pellucida. J In Vitro Fert Embryo Transf 1986;3(4):224–226.
38. Will M, Clark N, Swain J. Biological pH buffers in IVF: Help or hindrance to success. J Assist Reprod Genet
2011;28(8):711–724.
39. Morbeck DE, Khan Z, Barnidge DR, et al. Washing mineral oil reduces contaminants and embryotoxicity. Fertile
Steril 2010;94(7):2747–2752.
40. Otsuki J, Nagai Y, Chiba K. Damage of embryo development caused by peroxidized mineral oil and its associa-
tion with albumin in culture. Fertil Steril 2009;91(5):1745–1749.
41. Wagner-Coughlin CM, Maravilla AE, Nikurs AR, et al. Microdroplets under oil improves embryo quality even
when an isolette is used. Fertil Steril 1994;62(Suppl 1):S177.
42. Swain JE, Cabrera L, Xu X, et al. Microdrop preparation factors influence culture-media osmolality which can
impair mouse embryo preimplantation development. Reprod Biomed Online 2012;24(2):142–147.
43. Swain JE. Media composition: pH and buffers. In: Embryo Culture Methods and Protocols (eds. Smith G, Swain
JE, Pool TB). New York: Humana Press, Springer Science + Business Media, 2012, pp. 161–175.
44. Koustas G, Sjøblom Ø. Epigenetic consequences of pH stress in mouse embryos. Hum Reprod 2011;26:i78.
45. Ottesen LD, Hindkjaer J, Ingerslev J. Light exposure of the ovum and pre-implantation embryo during ART
procedures. J Assist Reprod Genet 2007;24(2–3):99–103.
46. Takenaka M, Horiuchi T, Yanagimachi R. Effects of light on development of mammalian zygotes. Proc Natl Acad
Sci USA 2007;105(36):14289–14293.
47. Van Blerkom J, Antczak M, Schrader R. The developmental potential of the human oocyte is related to the
dissolved oxygen content of follicular fluid: Association with vascular endothelial growth factor levels and
perifollicular blood flow characteristics. Hum Reprod 1997;12:1047–1055.
48. Huey S, Abuhamad A, Barros G, et al. Perifollicular blood flow Doppler indices, but not follicular pO2, pCO2 or
pH predict oocyte developmental competence in in vitro fertilization. Fertil Steril 1999;72(4):707–712.
49. Daya S, Kohut J, Gunby J, et al. Influence of blood clots in the cumulus complex on oocyte fertilization and cleav-
age. Hum Reprod 1990;5:744–746.
50. Ebner T, Moser M, Shebl O, et al. Blood clots in the cumulus-oocyte complex predict poor oocyte quality and
post-fertilization development. Reprod Biomed Online 2008;16:801–807.
51. Gilchrist RB, lane M, Thompson JG. Oocyte-secreted factors: Regulators of cumulus cell function and oocyte
quality. Hum Reprod Update 2008;14(2):159–177.
52. Montag M, Køster M, Van der Ven H. Polarization microscopy. In: Practical Manual of In Vitro Fertilization, Advanced
Methods and Novel Devices (eds. Nagy ZP, Varghese AC, Agarwal A). New York: Springer, 2012, pp. 121–125.
53. Pelzer ES, Allan JA, Cunningham K, et al. Microbial colonization of human follicular fluid: Alterations in cyto-
kine expression and adverse assisted reproductive technology outcome. Hum Reprod 2011;26(7):1799–1812.
54. Kovacic B, Vlaisavljevic V. Influence of atmospheric versus reduced oxygen concentrations on development of
human blastocysts in vitro: A prospective study on sibling oocytes. Reprod Biomed Online 2008;17:229–236.
55. Waldenstrøm U, Engstrøm A, Hellberg D, et al. Low-oxygen compared with high-oxygen atmosphere in blasto-
cyst culture, a prospective randomized study. Fertil Steril 2009;91:2461–2465.
56. Meintjes M, Chantilis SJ, Douglas JD, et al. A controlled randomized trial evaluating the effect of lowered
incubator oxygen tension on live births in a predominantly blastocyst transfer program. Hum Reprod
2009;24:300–307.
57. Catt J, Henman M. Toxic effects of oxygen on human embryo development. Hum Reprod 2000;15(Suppl 2):
199–206.
58. Gardner D, Lane M. Ex vivo early embryo development and effects on gene expression and imprinting. Reprod
Fertil Dev 2005;17(3):361–370.
59. Katz-Jaffe MG, Linck DW, Schoolcraft WB, et al. A proteomic analysis of mammalian preimplantation embry-
onic development. Reproduction 2005;130(6):899–905.
60. Enkhmaa D, Kasai T, Hoshi K. Longtime exposure of mouse embryos to the sperm produces high levels of
reactive oxygen species in culture medium and relates to poor embryo development. Reprod Domest Anima
2009;44:634–637.
61. Fitzharris G, Baltz KM. Granulosa cells regulate pH of the murine growing oocyte via gap junctions: Development
of independent homeostasis during oocyte growth. Development 2006;133(4):591–599.
62. Fitzharris G, Sitanov V, Baltz KM. Granulosa cells regulate oocyte intracellular pH against acidosis in preantral
follicles by multiple mechanisms. Development 2007;134(23):4283–4295.
63. Phillips KP, Petrunewich MA, Collins JL, et al. The intracellular pH-regulatory HCO3/Cl exchanger in the mouse
oocyte is inactivated during the first meiotic metaphase and reactivated after egg activation via the MAP kinase
pathway. Mol Biol Cell 2002;13(11):3800–3810.
64. Lane M, Lyons EA, Bavister BD. Cryopreservation reduces the ability of hamster 2-cell embryos to regulate
intracellular pH. Hum Reprod 2000;15(2):389–394.
65. Phillips KP, Leveille MC, Claman P, et al. Intracellular pH regulation in human preimplantation embryos. Hum
Reprod 2000;15(4):896–904.
66. Swain JE. Optimizing the culture environment in the IVF laboratory: Impact of pH and buffer capacity on gamete
and embryo quality. Reprod Biomed Online 2010;21(1):6–16.
67. Swain JE, Pool TB. New pH-buffering system for media utilized during gamete and embryo manipulations for
assisted reproduction. Reprod Biomed Online 2009;18(6):799–810.
68. Pool TB. Optimizing pH in clinical embryology. Clin Embryol 2004;7:1–7.
69. Ebner T, Moser M, Sommergruber M, et al. Incomplete denudation of oocytes prior to ICSI enhances embryo
quality and blastocyst development. Hum Reprod 2006;21:2972–2977.
70. Cunningham KJ, McCarthy CB, Russell JB. The pH change in a microdroplet system under oil with different
numbers of sperm added for insemination. Fertil Steril 1994;62(Suppl 1):S47.
71. Edwards LJ, Williams DA, Gardner DK. Intracellular pH of the mouse preimplantation embryo: Amino acids act
as buffers of intracellular pH. Hum Reprod 1998;13(12):3441–3448.
72. Baltz JM. Media composition: Salts and osmolality. In: Embryo Culture: Methods and Protocols (eds. Smith GD,
Swain JE, Pool TB). New York: Humana Press, Springer Science + Business Media, 2012, pp. 61–80.
73. Kolajora M, Baltz JM. Volume-regulated anion and organic osmolyte channels in mouse zygotes. Biol Reprod
1999;60:964–972.
74. Quinn P, Margalit R. Beneficial effects of co-culture with cumulus cells on blastocyst formation in a prospective
trial with supernumerary human embryos. J Assist Reprod Genet 1996;13:9–14.
75. Ebner T, Shebl O, Moser M, et al. Group culture of human zygotes is superior to individual culture in terms of
blastulation, implantation and life birth. Reprod Biomed Online 2010;21:762–768.
76. Brison DR, Schultz RM. Apoptosis during mouse blastocyst formation: Evidence for a role for survival factors
including transforming growth factor alpha. Biol Reprod 1997;56(5):1088–1096.
77. Gardner DK, Lane M. Embryo culture. In: Textbook of Assisted Reproductive Techniques. Laboratory and
Clinical Perspectives (eds. Gardner D, Weismann A, Howles CM, Shoham Z). London: Martin Dunitz, 2008;
pp. 219–240.
78. Behr B, Pool TB, Milki AA, et al. Preliminary clinical experience with human blastocyst development in vitro
without co-culture. Hum Reprod 1999;14:454–457.
79. Almagor M, Bejar C, Kaftka I, et al. Pregnancy rates after communal growth of preimplantation human embryos
in vitro. Fertil Steril 1996;66:394–397.
80. Smith G, Bormann C, Takayama S. Microfluidics for gamete manipulation and embryo culture. In: Practical
Manual of In Vitro Fertilization: Advanced Methods and Novel Devices (eds. Nagy ZP, Varghese AC, Agarwal A).
New York: Springer Science + Business Media, 2012, pp. 171–181.
81. Swain JE, Smith GD. Advances in embryo culture platforms: Novel approaches to improve preimplantation
embryo development through modifications of the microenvironment. Hum Reprod Update 2011;17:541–557.
82. Matsuura K, Hayashi N, Kuroda Y, et al. Improved development of mouse and human embryos using a tilting
embryo culture system. Reprod Biomed Online 2010;20:358–364.
83. Isachenko E, Maettner R, Isachenko V, et al. Mechanical agitation during the in vitro culture of human pre-
implantation embryos drastically increases the pregnancy rate. Clin Lab 2010;56(11–12):569–576.
84. Isachenko V, Maettner R, Sterzik K, et al. In-vitro culture of human embryos with mechanical micro-vibration
increases implantation rates. Reprod Biomed Online 2011;22(6):536–544.
85. Guerin P, El Mouatassim S, Menezo Y. Oxidative stress and protection against reactive oxygen species in the pre-
implantation embryo and its surroundings. Hum Reprod Update 2001;7:175–189.
86. Menezo Y, Sakkas D. Monozygotic winning, is it related to apoptosis in the embryo? Fertil Steril
2002;17:247–248.
87. Desai N, Kattal N, AbdelHafez F, et al. Granulocyte-macrophage colony-stimulating factor (GM-CSF) and
co-culture can effect post thaw development and apoptosis in cryopreserved embryos. J Assist Reprod Genet
2007;24:215–222.
88. Sjøblom C, Wikland M, Robertson S. Granulocyte-macrophage colony-stimulating factor promotes human blas-
tocyst development in vitro. Hum Reprod 1999;14:3069–3076.
89. Kaufman RA, Menezo Y, Hazout A, et al. Cocultured blastocyst cryopreservation: Experience of more than
500 transfer cycles. Fertil Steril 1995;64:1125–1129.
90. Kim D, Kim M, Kang H, et al. The supplementation of granulocyte-macrophage colony-stimulating fac-
tor (GM-CSF) in culture medium improves the pregnancy rate in human ART programs. Fertil Steril 2001;76:s6.
91. Shapiro B, Richter K, Daneshmand S, et al. Granulocyte-macrophage colony-stimulating factor enhances human
embryo development to the blastocyst stage, a randomized study. Fertil Steril 2003;79:15–16.
92. Milki A, Jun S, Hickley M, et al. Incidence of monozygotic twinning with blastocyst transfer compared to
cleavage-stage transfer. Fertil Steril 2003;79(3):503–506.
93. Behr B, Fisch J, Racowsky C, et al. Blastocyst-ET and monozygotic twinning. J Assist Reprod Genet
2000;17(6):349–351.
94. Wright V, Schieve L, Vahratian A, Reynolds M. Monozygotic twinning associated with day 5 embryo transfer in
pregnancies conceived after IVF. Hum Reprod 2004;19(8):1831–1836.
95. Mantikou E, Youssef MA, van Wely M, et al. Embryo culture media and IVF/ICSI success rates: A systematic
review. Hum Reprod Update 2013;19(3):210–220.
96. Aoki VW, Wilcox AL, Peterson CM. Comparison of four media types during 3-day human IVF embryo culture.
Reprod Biomed Online 2005;10(5):600–606.
97. Kovacic B, Vlaisavljevic V, Reljic M, et al. Clinical outcome of day 2 versus day 5 transfer in cycles with one or
two developed embryos. Fertil Steril 2002;77(3):529–536.
98. McArthur SJ, Leigh D, Marshall JT, et al. Blastocyst trophectoderm biopsy and preimplantation genetic diagnosis
for familial monogenetic disorder and chromosomal translocations. Prenat Diagn 2008;28:434–442.
99. Reed ML, Hamic A, Thompson DJ, et al. Continuous uninterrupted single medium culture without medium
renewal versus sequential media culture: A sibling embryo study. Fertil Steril 2009;92(5):1783–1786.
100. Sepulveda S, Garcia J, Arriaga E, et al. In vitro development and pregnancy outcomes for human embryos
cultured in either a single medium or in a sequential media system. Fertil Steril 2009;91:1765–1770.
3
Handling Gametes and Embryos: Quality
Control for Culture Conditions
Jason E. Swain
Introduction
Quality control (QC) in the culture system is a crucial factor in ensuring success within the in vitro fertilization
(IVF) lab. Without proper QC or quality assurance (QA), no combination of culture media, protein, and novel
culture platforms of implementation of cutting-edge lab technology will yield acceptably high outcomes.
When it comes to optimizing results within the IVF laboratory, the devil really is in the details.
When optimizing IVF results in the laboratory, the devil really is in the details.
Listing all areas for QC and QA within the lab is a monumental task, and it is not feasible to address all of them
within the confines of this chapter. However, a general area that deserves extra attention and focus is laboratory
environmental control; specifically those environmental variables that directly affect the culture system. Various
environmental parameters can act as stressors within the culture system and compromise gamete and embryo
development and function. Only through accurate monitoring of these variables can proper implementation be
achieved. Of particular importance are those variables that can be adjusted or altered directly within the labora-
tory, sometimes inadvertently, resulting in compromised embryo development.
Contact Material Testing

One of the most important aspects of laboratory QC and QA with respect to the culture system is the testing of all
contact materials to avoid introduction of contamination or toxicity into the culture environment. It is well docu-
mented that not all products, despite being packaged or sold as sterile, are inert in terms of impact on embryo
development [1,2]. Thus, verification of material safety is required before clinical use. This verification is usually
performed using a relevant bioassay, such as the mouse embryo assay (MEA) (Appendix 3A) or the human sperm
survival assay, also known as the human sperm motility assay (Appendix 3B).
Comparisons and the merits of these two assays have been discussed previously [3–9], and each can be useful
in its own way, with factors such as cost and availability warranting attention. Perhaps more importantly, the
sensitivity of the bioassay is imperative, and approaches can be modified to increase the sensitivity to ensure
that subtle material toxicities can be detected. Examples of approaches used to increase sensitivity are the one-
cell versus the two-cell MEA, outbred versus inbred versus hybrid mouse strains, blastocyst cell counts versus
simple blastocyst formation, exclusion of protein from media versus protein inclusion, and simple media versus a
more robust complex media [10–12]. Notably, thresholds must be set for an assay to “pass,” and these thresholds
should be set to ensure rigorous criteria. Each laboratory can determine which assay and threshold suit their
needs regarding sensitivity and cost. A commonly used and often recommended sensitive assay includes using
the one-cell MEA, noting time-appropriate embryo development with rate of expanded or hatching blastocyst
>70% at 96 hr (4 days) of culture. Other useful endpoints to assess material biocompatibility may include early
39
APPENDIX 3A: CONTACT MATERIAL MEA ASSAY

Purpose: All contact materials that are used during the collection, incubation, and transfer of gametes
and embryos must be tested before use for their ability to support mouse blastocyst development from
the one-cell stage. Untested material must remain in packing boxes and not be placed into lab circulation
until testing is complete and passed. If materials are pretested by the manufacturer using an approved
assay with appropriate sensitivity, and the laboratory has confidence in the pretesting procedures from
the manufacturer, these materials may be exempt from further laboratory testing (this decision is at the
discretion of each laboratory).
Controls
A. Positive control
1. A positive control is set up for each item being tested. This usually entails use of m
aterials/
lot numbers already in use, having previously passed screening (>70% blastocyst
formation).
Materials and Equipment (not all inclusive)

Culture dishes
Collection dishes
Test tubes (all sizes)
Pipettes (all sizes)
Catheters
Specimen collection containers
Pipette tips
Filters and receivers
Syringes
Media (all types)
Mineral oil
Procedure
A. Grouping of items helps conserve resources and cost. No more than three items should be
grouped together at one time to facilitate ease of retesting and identifying the offending item if
a test has failed. An example procedure is as follows:
1. Draw up culture media into new lot of serological pipettes. Hold for 5 min to allow adequate
exposure.
2. Expel media from pipette into new lot of test tubes. Hold media for 5 min.
3. Draw up media from new test tubes and expel appropriate amount on new lot of culture
dishes per laboratory protocol to permit embryo culture (i.e., 50 μL microdrop covered with
5 mL of oil).
4. Place at least 10 one-cell mouse embryos into the test drop of media and culture for
96 hr, checking development at appropriate intervals to verify time-appropriate devel-
opment. Embryos can be obtained fresh if a mouse colony is available, or frozen one-
cell embryos may be used. The strain of mouse is flexible, although F1 hybrids are less
sensitive.
5. Prepare culture media using currently used, prior tested, or approved lots as per lab protocol
to serve as the positive control. Place at least 10 embryos into the medium and incubate in
the same incubator as the test sample made in step 4.
Quality Control for Culture Conditions 41
Results
A. If the tested material yields expanded and hatching blastocyst formation >70%, it passes the
MEA. This should be within 10%–20% of the positive control results. Materials can be released
into lab circulation. Also important to note is that time-appropriate development should be appar-
ent between test and control materials, paying particular attention to the percentage of advanced
cell development at each time point to ensure no lag is present with the new test lots (e.g., early
compaction at 48 hr and early blast formation at 72 hr should be within 20% of controls).
B. If out of limits, various options can be explored:
1. Retest in same manner/group
a. If the test passes, materials can be released or used.
b. If the test fails, items can be retested individually to identify the offending product.
If the toxic item is identified from the failed test, return the lot to the manufacturer and request new lot
number.
If the specific item cannot be identified when testing individually, a more sensitive test can be performed
(no protein, less robust medium) or all items from the failed test batch can be returned and new items
ordered or tested.
C. Record all tests and results in a contact material log. Also note when new lots are placed into
circulation or used to permit tracking.
D. Label tested material with test date and technologist’s initials and place into circulation if passed.
Date/Time Thawed:
Items/Lots Tested:
Date: Date: Date: Date: Date:
Pos Con Test Pos Con Test Pos Con Test Pos Con Test Pos Con Test
8hpt n= n= 30hpt n= n= 48hpt n= n= 72hpt n= n= 96hpt n= n=
1-cell deg deg deg deg
2-cell 1-cell 1-cell 1-cell arrested cleavage
>2-cell 2-cell 2–4 cell 2–4 cell early morula
3-cell 5–7 cell 5–7 cell morula
4-cell 8-cell 8-cell early blast
>4-cell early morula early morula blast
morula morula expanded blast
early blast hatching blast
hatched
%2-cell % >3-cell % compact % > compact % > expanded
(limits: >70%)
pass
fail
Supervisor/Director Review: Date:
FIGURE 3A.1 Material testing and sample MEA development tracking form.
compaction or early blastocyst formation at 48 and 72 hr, respectively, to ensure new materials yield similar
results to controls (Figure 3A.1). Indeed, real-time assessment of morphokinetics has been used to improve
sensitivity of the MEA [13].
One point worthy of note is that many materials from manufacturers can now be purchased as “pretested,”
having passed some manner of bioassay. However, these manufacturers’ bioassays may not meet the sensitivity
criteria of individual labs. Furthermore, factors affecting material quality can convey detrimental effects at a
point after the initial successful passing of the bioassay. For example, improper storage conditions in warehouses
or delayed effects from prolonged storage can cause a “prescreened” item to later become toxic once received
in the laboratory. A common example of this situation is the toxicity associated with mineral oil [1,13,14]. Also,
potential toxicity from a single item could be very minor, but when used in the context of several sequential
items in a particular laboratory culture system, this toxicity could be c ompounded due to interactions with other
system components. This occurrence is noted anecdotally across several labs, where “tested” materials are later
found to have a detrimental impact. Therefore, more stringent labs retest c ertain pretested materials, or they test
them in a manner similar to their use within the lab to help ensure safety.
APPENDIX 3B: CONTACT MATERIAL HUMAN SPERM MOTILITY ASSAY

Principle: All contact materials that are used during the collection, incubation, and transfer of gametes
and embryos must be tested before use for their effect on sperm motility. Untested material must remain
in packing boxes and not be placed into lab circulation until testing is complete, with a positive outcome.
If materials are pretested by the manufacturer using an approved assay with appropriate sensitivity, and
the laboratory has confidence in the pretesting procedures from the manufacturer, these materials may be
exempt from further laboratory testing (this is at the discretion of each laboratory).
Controls
A. Positive control
1. A positive control is set up for each item being tested.
2. The positive control is the current lot number being used.
B. Negative control
1. A negative control is set up for each batch of material being tested.
2. A small piece of powdered glove serves as the negative control.
3. The glove piece is placed in a small snap top tube that contains a density gradient processed
sperm sample (10 × 106/mL and ≥70% motility).
4. Motility is observed after 24 hr incubation.
a. Expected value: 0% motility
Materials and Equipment

Culture dishes
Collection dishes
Test tubes (all sizes)
Pipettes (all sizes)
Catheters
Specimen collection containers
Pipette tips
Filters and receivers
Syringes
Media (all types)
Mineral oil
Procedure: Sperm should be prepared using density gradient separation to obtain a final motility >70%.
Several sperm samples can be combined to form a high concentration of a homogenous mixture and 1 mL
aliquots made for testing of multiple items.
A. Culture dishes
1. Fill dish with processed sperm sample (10 × 106/mL and ≥70% motility).
2. Repeat for positive controls of each item being tested.
3. For the small tissue culture dishes, place sperm sample on dish as microdrops and cover with oil.
4. Incubate for 24 hr in CO2 incubator.
5. Check motility and forward progression.
B. Tubes
1. Fill tubes with 1 mL of processed sperm sample (10 × 106/mL and ≥70% motility).
C. Pipettes, catheters, syringes, and pipette tips

1. Aspirate processed sperm (10 × 106/mL and ≥70% motility) into material being tested.
3. Allow to sit for 30 min.
4. Expel sample into tested test tube.
D. Specimen containers
1. Fill containers with 1 mL of processed sperm sample (10 × 106/mL and motility ≥70%).
2. Repeat for positive controls for each item being tested.
4. Transfer sample into tested small snap top tube (Falcon 2054).
E. Filter units
1. Filter media through filter unit.
2. Add appropriate amount of sperm with ≥70% motility to filtrate to obtain 10 × 106/mL.
5. Transfer a portion of sample into tested small snap top tube (Falcon 2054).
Results
A. If the tested material has sperm motility within 10% of the positive control and the same forward
progression, it is within acceptable limits for use.
B. If out of limits, perform the following steps.
1. Retest.
2. If material is still out of limits, return the lot to the manufacturer and request a new lot number.
C. Record all tests and results in a contact material log. Also note when new lots are placed into
circulation or used to permit tracking.
D. Label tested material with test date and the technologist’s initials and place into circulation if passed.
Air Quality
Air quality is another environmental variable that deserves attention when performing QC in the IVF laboratory
culture system [15], and it can be inadvertently influenced. All modern IVF labs should have laminar flow hoods
in which to prepare culture dishes or observe embryos to avoid possible contamination, and the these hoods should
undergo routine annual maintenance to ensure proper functioning. Many labs also have specialized, dedicated
air filtration units that not only provide positive pressure and limit particle counts through use of high-efficiency
particulate absorption (HEPA) filters, but also are outfitted with charcoal or potassium permanganate filters to
remove volatile organic compounds (VOCs) from circulation. Use of ultraviolet (UV) photocatalytic oxidation
may also be used to reduce VOC levels [16,17]. The presence of VOCs, such as aldehydes or toluene, within the
air of IVF labs has been well documented [18–20], and it is well accepted that poor air quality can compromise
the culture system, whereas measures to improve air quality result in improved outcomes [21], presumably due to
the detrimental effects of the VOCs on the preimplantation embryo [18,20,22,23]. Importantly, these VOCs may
stem from materials used in lab construction or cleaning, such as cabinet or flooring materials, various adhesives,
TABLE 3.1
Recommended Monitoring and Filter Exchange Schedules to Maintain or Improve IVF Lab Air Quality
Equipment Recommended Maintenance
Heating, ventilating, and air-conditioning (HVAC)/air handler (positive pressure, if present) Confirm daily
HVAC/air handler (HEPA filters, charcoal/permanganate filters) Replace 3–6 months or as needed
Incubator HEPA filters Replace 6–12 months
Incubator/gas line VOC filters Replace 3–6 months
Laminar flow hood prefilters Replace or clean annually or as needed
Laminar flow hood (flow rate inspection) Inspect annually
Lab air quality assessment (particle counts, VOC levels) Inspect annually
Note: Lab air exchange rates may be modified to improve certain variables, such as particle counts. Additional maintenance may be
required to maintain air quality if other systems are in place (i.e., UV photocatalytic oxidation).
solvents, cleansers, or paint or from plastics and other materials stored in the lab. Thus, careful selection of items
used in and around the laboratory is essential to prevent accidental introduction of harmful VOCs.
Poor air quality in the lab can compromise embryo development and outcome, but air quality in
the incubator should also be addressed.
An initial assessment of air quality within the IVF lab is recommended to determine whether a problem
exists, and whether preventative measures are needed to improve air quality (i.e., reduce VOC levels). Methods
for air quality assessment can be found elsewhere and include particle count testing, VOC level determination,
and other methods [15,24]. Unfortunately, threshold limits for detrimental impact of air quality are difficult
to verify. Particle counts are often recommended to fall below standards for International Organization for
Standardization (ISO) class 7, and harmful levels of VOCs are undefined. Regardless of lab levels, regular
assessment of laboratory air quality, including particle count and VOC levels, and routine replacement of rel-
evant air filters and adjustment of air exchange rates is prudent to ensure air quality conditions have not changed
or deteriorated dramatically over time; such assessments also may indicate inappropriate procedures (e.g., new
cleaning agents, improper storage of lab supplies) or improper equipment or filter functioning (Table 3.1).
Perhaps, more importantly, air quality within the laboratory incubator should also be addressed. Air quality
within the laboratory incubator, in part, is influenced by the quality of the room air. However, VOCs can accu-
mulate inside of incubators due to presence of dishware at elevated temperatures and lack of routine or thorough
air exchange within the closed environment. Interestingly, use of low-oxygen incubators entails the use of ~90%
nitrogen in place of room air or premixed tri-gas cylinders. Thus, this may offer an opportunity to provide an initial
purer gas quality within the laboratory incubator, if the gas supply is free from contaminants. However, this may not
always be the case, because gas cylinders can be the source of harmful VOCs, such as benzenes or alcohols [18,19].
Thus, inclusion of specialized inline filters or filters placed within the incubator [23,25–27] are recommended to
remove VOCs from the gas supply or incubator and to further improve air quality inside. These incubator filters
should be replaced at regular intervals, with recommendations often being every 3–6 months (Table 3.1).
Dish Setup
It is known that media osmolality affects embryo development [28–32], although tolerances and optimal values
vary between cell stages and are dependent upon media formulation. When osmolality increases >~300 mOsm,
development is compromised. Thus, embryo culture media is usually formulated in the range of 260–290 mOsm.
In addition, inclusion of organic osmolytes, including key amino acids, helps regulate cell volume and combat
media osmolality changes [28–31,33–37]. Although it is not required to retest commercial media osmolality,
because these media are generally pretested using an MEA before release by the manufacturer, QC measures
should be taken to ensure that detrimental shifts in osmolality do not inadvertently occur within the laboratory.
This measure is achieved via avoidance of evaporation that can occur primarily during improper setup of culture
dishes or improper culture conditions.
It has been shown that temperature, volume of media, and drop preparation technique during dish prepara-
tion with microdrops can influence media osmolality [32] (Figure 3.1). When combined with airflow inside the
working sterile hood, these four variables can raise media osmolality to a point that capable of inhibiting embryo
development. Thus, care should be taken when preparing dishes.
Similar rises in media osmolality were recorded when 3 mL of media was left in 30 mm dishes for 30–60 min
in a nonhumidified environment without oil overlay (5%–14% increase), a condition that might be encountered
during oocyte retrieval or other procedures [38]. In these cases, media should be warmed within a closed test tube
to prevent evaporation and then aliquoted into the dish immediately before use, rather than warming the media
for extended periods within the dish. When warming media within a dish, condensation can be seen on the lids,
a clear indication that evaporation has occurred and osmolality has thus increased. If an osmometer is available,
initial testing can be performed to determine whether media osmolality is altered after routine dish preparation.
If elevations are detected, procedures should be modified to avoid osmolality increase. Although airflow in the
sterile hood is preferable for sterility during dish preparation, preparation should be performed on an unheated
surface if possible. In addition, if using small volumes of media, alteration of microdrop preparation technique
may be explored, including use of a “wash drop,” where media are removed and fresh media are added after oil
overlay. Another easy modification to help prevent osmolality shifts due to media evaporation includes imple-
menting a protocol limiting the number of dishes prepared at one time before oil overlay is added (Appendix 3C).
10, 20, or
40 μL drop
Air flow No flow
37°C 23°C 37°C 23°C
Std Wash Std Wash Std Wash Std Wash
(a)
p < 0.001
25 p < 0.0001 360
p < 0.0001
Osmolality (mOsm/kg)
20 340
Osmolality increase
15 320
10 300
5 280
0 260
23°C 37°C No air Air Wash Std 40 μL20 μL10 μL Control 40 μl/23°C/no air 10 μl/37°C/air
flow flow bottle media flow/wash flow/std
Temp Air flow Drop type Drop volume
(b) (c)
FIGURE 3.1 Conditions present during culture dish preparation can result in evaporation and affect media osmolality, impairing
subsequent embryo development. (a) Various conditions tested for affect on media osmolality. (b) Impact of individual conditions
on media osmolality. Temperature, drop volume, and drop preparation techniques all significantly increased media osmolality.
(c) Demonstration of how combining several suboptimal preparation conditions can cause an osmoality increase high enough to
impair embryo development (>300 mOsm). Different superscript symbols represent significant differences (p < 0.05).
APPENDIX 3C: CULTURE DISH PREPARATION

Purpose: To prepare embryo culture dishes in a manner so as to avoid evaporation and the resultant
detrimental elevations in media osmolality that can impair subsequent embryo development.
Equipment and Materials

Airflow hood
Embryo culture dishes
Serological pipettes
Pipetman/pipettors
Pipette tips
Culture medium/protein
Mineral oil
Procedure
A. To help ensure sterility, all procedures should be performed within the confines of the airflow
hood. Procedures should be performed away from the front of the hood, toward the center, and all
items and packaging should be opened and closed within the hood when possible to ensure steril-
ity. If the surface within the airflow hood is heated, dishes should be prepared on the unheated
portion of the work surface.
1. Set out all necessary pipettes, culture dishes, media, and oil into the airflow hood.
2. Prepare media/protein as necessary based on the procedure.
a. If protein is presupplemented, media are ready to use.
3. When media are ready, lay out two to four culture dishes.
a. If more dishes are to be made, these can also be set aside from their storage bags, but
no more than two to four dishes should ever be prepared with media at any one time to
avoid excess evaporation.
4. Prepare two to four dishes by aliquoting the appropriate amount of medium and then imme-
diately cover the dishes with the appropriate amount of mineral oil.
a. Amounts of media and oil will vary based on individual lab protocols.
b. If using microdrops (volumes <100 μL), it is recommended that no more than two dishes
be made at any given time before adding oil overlay. It is also recommended that “wash”
drops be used, where approximately half of the total final drop volume is first placed on
the dish, covered with oil, and the remaining half of the drop volume is then added.
Care should be taken not to form too many microdrops per plate. Limiting drop num-
ber to fewer than eight helps ensure rapid processing and avoids evaporation.
c. If using four-well dishes or other larger volume methods (volumes >500 μL), up to four
dishes may be made, adding the entire media volume to each well, before oil overlay.
B. If an osmometer is available, prepared media can be tested to confirm that osmolality is not
elevated under laboratory preparation conditions. This need not be repeated each time, but may
be useful during initial procedure implementation, during new technician training, or for routine
monitoring. Osmolality of media from prepared dishes should be compared with control media
out of the bottle. Differences in osmolality should be <10 mOsm.
N O T E: For other laboratory procedures that do not use oil overlay, such as oocyte retrieval and washing,
embryo transfer, or cryopreservation, care must also be taken to avoid evaporation and increases in osmolality.
For these procedures, media should be warmed within a closed test tube, rather than warming within a plate on
a warmed surface. This avoids evaporation. Any time condensation is observed on the lid of a dish, evapora-
tion has occurred. Larger volumes of media can be used to combat this and care should be taken to use the lid
when possible due to the confounding effect of the airflow hood on facilitating evaporation, as well as heat loss.
Routine monitoring of osmolality after dish preparation can help ensure procedures are being followed and may
be especially important in larger labs that make up numerous dishes on a daily basis, or when training new per-
sonnel to ensure proper technique.
Gas Monitoring and pH

Another aspect of culture media that deserves special attention with regard to culture system QC is pH. Although
embryos have membrane-bound transporters that regulate their internal pH and permit them to develop a
range of media pHs, improper media pH can stress embryos as they try to compensate [39–41]. Furthermore,
some cell types, such as sperm, oocytes, and frozen-thawed embryos, lack robust intracellular pH regulators and
are thus especially sensitive to media pH [42–45]. Therefore, adherence to an appropriate and narrow pH range
is prudent.
The pH of culture media is established primarily via the balance of sodium bicarbonate in the medium and
CO2 levels from the incubator. Because bicarbonate levels are often set by the commercial manufacturer, pH
is most easily adjusted via altering incubator CO2 concentrations. Although daily checks of CO2 concentration
in incubators are often performed using Fyrite or any another measuring system, this is a poor substitute for
actually measuring pH.
CO2 measurement is a poor substitute for actually measuring pH.
Not all CO2 measurement devices are accurate, and they can vary in their readings [46,47] (Figure 3.2).
In addition, not all media are formulated in the same manner, and they may have differing bicarbonate concen-
trations. As a result, different media may yield different pH even if used in the same incubator. Furthermore,
different protein supplements and varying amounts used can affect pH, either by impacting bicarbonate concen-
trations through dilution, or because protein supplements themselves are slightly acidic (Figure 3.3).
Finally, location of a particular laboratory at high altitude can impact the amount of CO2 needed to achieve
a desired pH, as labs at higher elevation require higher CO2 levels due to the decreased atmospheric pressure.
As a result, a single CO2 concentration cannot be used reliably between laboratories to give the same medium
pH. Thus, although daily measurement of incubator CO2 concentration may be suitable to track deviations in
7.5
7.0 Fyrite CO2 Avg. = 5.9 ± 0.042a

IR CO2 Avg. = 5.6 ± 0.036b
Incubator CO2 Avg. = 6.0 ± 0.0a Fyrite CO2
% CO2 or pH
6.5 IR CO2
Incubator CO2
6.0 pH
5.5
5.0
1 2 3 4 5 6 7 8 9 10 11 12 13
Day
FIGURE 3.2 Different CO2 measuring devices can yield different CO2 readings, so pH should be mesured on all incubators to help
determine the optimal CO2 value. Simply relying on a CO2 measuring device to set a specific CO2 value could result in improper pH
and improper growth conditions. Different superscripts represent a statistically significant difference in average CO2 readings, p < 0.05.
Post-supplemented Pre-supplemented
7.45 7.34
7.4 7.32
7.35 7.3
Media pH
Media pH
7.3 7.28
7.25 7.26
7.2 7.24
7.15 7.22
6.0% CO2 6.5% CO2 5% Protein 10% Protein
(a) (b)
FIGURE 3.3 Different media can yield different pH values in the same incubator. (a) A medium presupplemented with protein by
the manufacturer yields a different pH in the same incubator from exactly the same medium supplemented with the same protein and
concentration by the lab. Supplementation of protein by the lab resulted in a dilution in media component concentrations, including
bicarbonate, thereby lowering the pH at the same CO2 concentration. (b) Similarly, differing amounts of protein can change the pH
of the same medium in the same incubator by the dilution effect and also by the the fact that protein supplements are slightly acidic.
Higher protein supplementation by the lab results in a lower pH value in the same incubator. As a result, all media should be tested
for pH in each incubator to determine the optimal CO2 concentrations required to obtain the desired pH.
incubator function from day to day, at some point pH should be measured to ensure that the incubator gas
concentration is set correctly for a particular lab. At a minimum, pH measurement should be done on all new lots
of media, but more routine monitoring may also be useful. Although there is no agreed upon “optimum” pH [40],
and no proven need to change pH during culture has been demonstrated [40], following manufacturers’ recom-
mendations is prudent, and maintaining a narrow acceptable range is essential to reduce variability within the
laboratory. A common acceptable pH range is 7.2–7.4, although a narrower target range of around 7.25–7.35
ensures less variation in culture conditions and tighter QC (Figure 3D.2).
A crucial aspect of pH and QC entails accurate measurement of the variable (Appendix 3D and Figure 3D.1).
pH measurement itself can vary; thus, careful attention should be paid to measure pH using the correct
methods. For example, temperature not only affects accuracy of a pH meter but also changes the actual pH
of certain buffered media. Thus, the impact of temperature should be considered when taking pH measure-
ments. In addition, some pH meters or electrodes may differ in their readings compared with other instruments
(Figure 3.4); thus, establishment of baseline function of a particular device and an initial validation before
clinical application is helpful when beginning pH measurements. Failure to validate pH measuring devices
and probes could result in improper CO2 values being set, thereby creating a potentially compromising growth
environment for embryos. Validation should be performed again when probes are replaced on a regular basis
to ensure proper functioning. Glass, double-junction, KCl-filled electrodes are often recommended for use with
culture media due to protein content and organic buffers that can eventually clog the junction of the e lectrode.
Other “in-incubator” pH systems are also available that measure pH inside the incubator or in real time, although,
as mentioned, the accuracy of these newer systems should be validated before clinical implementation.
Temperature Control
Temperature is another environmental variable controlled within the lab that is crucial to the culture system
and should be routinely monitored. It is also well known that temperature can affect various aspects of gam-
ete and embryo function, most notably, meiotic spindle stability [48–50] and possibly embryo metabolism [51].
APPENDIX 3D: pH MEASUREMENT

Purpose: To measure pH accurately to confirm adequate culture conditions and aid in proper incubator
settings (CO2 levels).
Materials and Equipment

Snap top test tube
pH meter
pH electrode (semimicro, glass, double-junction KCl-filled recommended)
ATC probe (optional)
pH standards (7 and 10)
Electrode storage solution (KCl)
Warming block (37°C)
Procedure
A. For culture media used in the incubator, aliquot the medium being tested into a loosely capped
test tube and place the test tube into the incubator. The same medium and protein content that
are used clinically should be used for pH testing. Volume of medium will vary depending on
tube size used and size of the pH probe or device, but 3–5 mL is usually adequate. Place tube
loosely capped into each incubator and allow >12 hr to ensure adequate temperature and gas
equilibration.
B. Immediately before measuring pH, the meter and probe should be calibrated according to manu-
facturer’s instructions.
1. It is important that temperature is accounted for to obtain the most accurate readings
possible. Thus, the pH meter should be set to ~37°C, or an external ATC probe should be
used. If an ATC probe is used, it should be placed into an ~37°C warming block. A standard
benchtop pH meter and a semimicro-sized, double-junction glass KCl–filled electrode is
recommended for use due to protein content of culture media and for use with organic buf-
fers. This helps prevent clogging of the junction and ensures rapid and efficient readings.
However, other electrodes will suffice in many cases, although care may be needed in clean-
ing and restoring electrodes to ensure rapid response and accurate readings. Regardless of
the electrode, proper storage conditions and cleaning of electrodes are required for optimal
performance. In this case, electrodes should be stored in a 3M KCl solution, not water, and
the electrode should remain filled with 3M KCl. Before calibration and use, ensure electrode
is filled with proper solution.
2. It is helpful if there is a tight fit or seal between the electrode and the walls of the test tube.
This can be accomplished by fitting a rubber gasket around the electrode to fill the gap when
placed into the test tube to help stabilize the reading. Alternatively, a small hole can be made
in a test tube cap and the electrode slipped through this hole. The cap/electrode can then be
fitted onto the test tube to create a seal. This setup can be used during electrode storage, and
if the same-size test tubes are used, the cap/electrode can be easily fitted over the test sample
test tubes to create a seal and help stabilize pH readings.
3. It is important to use freshly aliquoted calibration standards before each calibration and
measurement to ensure accurate calibration, because pH of standards can drift after
extended exposure to air. Use of a two-point calibration with standards of pH 7 and 10
is sufficient, although a three-point calibration using pH 4 can also be used if desired.
Standards should be aliquoted into test tubes, capped, and then warmed in a water bath or
heating block to ~37°C.
C. After equilibration of the medium and calibration of the pH meter, quickly remove a test sample
tube from the incubator and snap the cap closed. Quickly move the tube to the pH meter that is
adjacent to the incubator and place the pH electrode into the medium. This should be done first
thing in the morning before any incubator openings to ensure accurate pH readings. This is made
easier if the pH meter is set up near the incubators. Allow the reading to stabilize and record the
reading. Repeat for each incubator and each medium used in that particular incubator. The read-
ings should stabilize in <5–10 s.
Results
A. Record reading on pH log sheet.
1. If pH readings in each incubator are in range (acceptable range set by lab based on manufac-
turer’s recommendations, i.e., 7.25–7.35), no further action is required.
2. If pH of a particular incubator is out of range, the incubator CO2 levels should be adjusted
according to manufacturer’s instructions to raise or lower pH. Raising CO2 levels lowers the
pH and lowering CO2 levels raises the pH. Adjust CO2 accordingly and repeat pH measure-
ment the next day or until values fall within the specified range.
NOTE:
A. pH measurements can be performed on a weekly or daily basis, based on lab protocols. Weekly
measurement may be more practical when one considers cost of culture media and other variables.
B. Importantly, if testing a new lot of media, perform the pH reading in a side-by-side manner with
the old verified lot. If the pH of the new lot is out of range and cannot be easily brought within
range by a small adjustment of CO2 levels, repeat to verify before contacting the media company
to request a new lot of media. In addition, all new lots of culture media should have been pH
verified before clinical implementation.
C. Handling media with HEPES or MOPS should never be placed inside the incubator. When
measuring pH of these media, warm to 37°C in a water bath or warming block for >30 min
in a capped test tube before measuring pH. Changes in temperature change the pH of these
buffered media.
Electrode
Cap
Sample/
tube
FIGURE 3D.1 Example setup of a pH meter and electrode for measurement of culture media pH. A warming block is used
to account for the impact of temperature and is set to ~37ºC. The electrode is placed through the cap to prevent excessive gas
exchange that could affect readings. The electrode/cap is removed from the storage solution tube, calibrated, and then quickly
placed into the tube of medium for measurement.
pH Recording Form
Month:
Calibration
6.98–7.01 Corrective
Media/ 9.88–10.01 Temp Inc #1 Inc #2 Inc #3 Inc #4 action/
Date Tech lot # Slope >98 3.65°C–37.5°C 7.25–7.35 7.25–7.35 7.25–7.35 7.25–7.35 comment
Fert: Fert: Fert: Fert:

Culture: Culture: Culture: Culture:




FIGURE 3D.2 Sample recording sheet for routine pH measurements.
23°C 37°C
7.60
7.55 c
7.50
7.45
7.40 d
b b
7.35 b
7.30
7.25 a
7.20
7.15
7.10
Blood gas Benchtop Solid state unit
analyzer meter/probe
FIGURE 3.4 Validation of a pH meter is required before clinical implementation to verify accuracy. Failure to validate a pH meter
could result in the improper CO2 setting on an incubator, thereby providing an improper pH and possibly compromising growth
conditions. Three pH measuring devices were compared using the same medium at two different temperatures. Different superscript
letters represent significant differences (p < 0.05).
Thus, one aspect of temperature monitoring entails the laboratory incubator. The majority of IVF labs set incuba-
tor t emperature at 37.0°C, with this value being presumably based on human core body temperature. However, it
should be noted that this may not be entirely accurate. Regarding body temperature, Cox in 1998 reported that for
decades it was thought that the normal body temperature was 98.6°F [52,53]. This number was calculated from
a study in Germany that reported normal body temperature at 37°C. What was not known was that this number
was an average rounded to the nearest degree. In other words, it was accurate only to two significant digits, not
the three we have with 98.6. Scientists today know that normal is actually 98.2°F (±0.6°F), that is to say anything
in the range of 97.6°F–98.8°F should be considered normal. Furthermore, it is thought that the temperature of
37
+ oil + lid
36 + oil – lid
– oil + lid
Temperature (°C)
– oil – lid
35
34
33
32
Nunc 4-well Center well Microdrop
500 μL media 1 mL media 50 μL media
+
– 300 μL oil +
– 1 mL oil +
– 7 mL oil
Surface temp 37.2°C
FIGURE 3.5 Temperature within a culture dish on a warmed surface can be impacted by various factors in the lab. The type of
dish and its surface contact with the warmed surface, the presence of a lid, and the presence of an oil overlay can all impact the
temperature of culture media within the confines of a sterile airflow hood. Surface temperature may need to be slighlty higher than
the target temperature to achieve the correct temperature within the dish itself.
the human follicle is ~2.3°C cooler than core body temperature and that the fallopian tube from animal models
carries a temperature gradient ~1.5°C cooler [54–59]. As a result, whether strict adherence to a temperature set
point of 37°C within the incubator is required is questionable. The question has been poised “of whether human
IVF and related procedures should be carried out at, say, 35.5°C–36°C rather than at 37°C” [51]. Use of a 1.5ºC
lower temperature would presumably lower the metabolic rate of the embryo by ~15% and may result in a more
“quiet” metabolism, thereby possibly benefiting embryo development [51]. Indeed, a recent preliminary study
demonstrated that culture of human embryos at 36°C had no appreciable negative effect on blastocyst develop-
ment or chromosomal compliment compared with 37°C, though no benefit was noted and no impact on outcomes
was examined [60]. At least one study suggests that a temperature <37°C may improve outcomes, when increased
pregnancy rates were obtained from incubators with a temperature of 36.96°C ± 0.13°C compared with those
with a temperature of 37.03°C ± 0.13°C [25]. However, with such a narrow margin and limited accuracy of ther-
mometers, it is not clear how meaningful this observation is. More detailed analysis of the optimal temperature
for IVF is required.
In spite of the optimal temperature at which to culture embryos being debatable, a strict adherence to a nar-
row acceptable temperature range should be followed in the lab to reduce variability. Toward this end, surface
temperatures in particular require careful monitoring. Different working surfaces in the laboratory, including
microscope stages, may be warmed to reduce media temperature excursions outside the incubator for observa-
tion. Importantly, different warming surfaces may yield differing temperatures and should be monitored [49].
Similarly, different dishes may vary in their surface contact due to the conformity of the dish bottom. This can
affect heat exchange and thus requires a differential surface temperature to achieve the desired temperature
within the dish or media (Figure 3.5).
In addition, when used in sterile airflow hoods, volume of media and presence or absence of oil overlay or lids
can also affect heat transfer and temperature (Figure 3.5). Similar findings have been reported previously [61].
Thus, a calibrated thermocouple should be used to measure temperature inside the various dishes used on the
warmed surface and the temperature can then be adjusted to give the desired media temperature. For example,
if aiming for 36°C–37°C in culture media, the surface temperature may have to be raised to 37°C–38°C. In addi-
tion, temperature within the incubator can also vary slightly from shelf to shelf. Thus, some have suggested that
determining whether variation exists may be prudent, and preferential placement of dishes on a particular shelf
may be helpful in optimizing incubator management [25]. As an added note, use of milled aluminum plates or
tube holders can be used inside the incubator or on heated work surfaces to maintain thermal stability during
repeated openings and closings [61].
Lab temperatures should be monitored daily using calibrated thermometers. This can be achieved through use
of a National Institute of Standards and Technology (NIST) traceable device that has been verified for accuracy,
or by comparing equipment to a calibrated NIST device, making sure to note any offsite or discrepancy to permit
compensation when setting equipment temperatures.
Incubator Management
On a final note, one of the most important aspects of QC in the culture system is incubator management. Because
gametes and embryos spend the majority of their time within the laboratory incubator, and the incubator con-
trols multiple environmental variables, proper management of the incubator is an extremely important factor
to consider in maximizing efficacy of the IVF lab. Even the most cutting-edge incubator with the more current
technology can be made inefficient if managed improperly.
One of the most important aspects of QC in the culture system is incubator management.
Incubator management involves more than just selecting an appropriate incubator, setting it up, ensuring
that it is off-gassed to reduce VOCs, measuring and setting proper gas levels (proper pH), and monitoring air
quality and temperature, as mentioned previously. Proper incubator management also entails careful attention
to caseload and workflow to help maintain a stable growth environment. IVF patients should be spread out
among various incubators for a variety of reasons. Practically, having several incubators in which to spread
out patients avoids catastrophic affects if patients are all placed within a single unit and it fails. Furthermore,
spreading out of patients among several incubators also prevents excessive entry into the same incubator, thus
helping to ensure environmental stability during the culture period. It was demonstrated that reducing door
opening from six to four times over a 6-day period on a small-box incubator resulted in significantly improved
blastocyst formation (53% vs. 43%) and “good” quality blastocysts (60% vs. 51%), although no differences in
Day 3 embryo quality, implantation, or clinical pregnancy rates were observed [62]. Thus, IVF cases should
be evenly distributed between all available incubators to avoid overuse or excessive openings of an individual
incubator.
A specific number of incubators required within the IVF lab is difficult to recommend but is obviously based
on incubator type and capacity and caseload and is influenced by cost. A minimum of two culture incubators is
required, and the optimal number will be based on the weekly caseload of patients, rather than on the annual
patient number. One can appreciate that if two laboratories both perform 300 cases annually, but one batches
these patients into 2 weeks each month, then the laboratory that does not batch patients will likely require more
incubators if they wish to keep the same incubator workflow (same number of patients per incubator, same num-
ber of openings and closings) as the lab that does not batch.
Regardless of how many incubators are present within a laboratory, some regular method of efficiency
should be assessed to determine proper functions and management (Appendix 3E and Figure 3E.1). Tracking
how many patients are placed into each incubator on a monthly basis is useful to monitor workflow and prevent
overcrowding. Another useful indicator to monitor efficacy is to track embryo development and pregnancy per
incubator. A useful threshold is >50% blastocyst conversion; this threshold will vary from patient to patient, but
an overall average of >50% across all patients is a QC good indicator. Blastocyst quality should also be consid-
ered, although quality will vary based on individual grading criteria. Individual thresholds should be set and
monitored on a monthly basis to track performance. These values can then be compared between incubators to
validate proper management and functioning. If variation in the patient population is problematic, examination
of the better prognosis patients, such as younger patients or the donor population, may be more insightful in
determining whether a true problem exists in the culture system within a specific incubator or whether prob-
lems lie in the population itself.
APPENDIX 3E: DAILY INCUBATOR QC FORM

..........Limits............
Incubators 5.5%–6.5% CO2
4–6% O2
Temp. 36.5°C–37.2°C
Month:
Daily –#1 DAILY Weekly Monthly
Day Tech. CO2 O2 Temp H2O in pans Hygrometer Comments/actions
Change H2O & Oil
Inc Display Measure Inc Display Measure Inc Display Therm (√) (>80)
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
# of patients:
% Blastocyst formation:
FIGURE 3E.1 Sample recording sheet for daily or monthly incubator QC. One sheet can be placed on each incubator.
A “comments” section can be used to record weekly pH values or indicate corrective actions taken during regular QC pro-
cedures. The number of patients, blastocyst conversion rates, or both may be tabulated at the end of each month for each
incubator to aid in QC.
Conclusions
Proper QC is essential to optimize function of the IVF lab and to maximize reproductive outcomes. Although
purchase of high-quality materials and equipment can aid in improving culture conditions, proper oversight
and implementation are crucial. Despite having the same equipment and media in place, two laboratories can
have dramatically different outcomes because several variables within the culture system are controlled directly
within the individual lab. Examples include pH, osmolality, air quality, and incubator workflow. However,
another variable that needs to be examined includes variation between technicians to ensure consistency in lab
practices and approaches. This not only applies to procedures such as embryo grading, Intracytoplasmic sperm
injection (ICSI), and catheter loading but also to how daily QC readings are taken [47], how QC assays are per-
formed, and how media and dishes are prepared. Subtle deviations may introduce an environmental stressor into
the culture system or miss a slight variation that could affect outcome.
REFERENCES
1. Morbeck DE. Importance of supply integrity for in vitro fertilization and embryo culture. Semin Reprod Med
2012; 30(3): 182–190.
2. Lee BE, Boone WR, Brackelsberg PO and Carmichael RA. Development of screening systems for evaluation of
materials used in mammalian embryo transfer. Theriogenology 1988; 30(3): 605–612.
3. Scott LF, Sundaram SG and Smith S. The relevance and use of mouse embryo bioassays for quality control in an
assisted reproductive technology program. Fertil Steril 1993; 60(3): 559–568.
4. Esterhuizen AD, Bosman E, Botes AD, Groenewald OA, Giesteira MV, Labuschagne GP, et al. A comparative
study on the diagnostic sensitivity of rodent sperm and embryos in the detection of endotoxin in Earle’s balanced
salt solution. J Assist Reprod Genet 1994; 11(1): 38–42.
5. Rinehart JS, Bavister BD and Gerrity M. Quality control in the in vitro fertilization laboratory: Comparison of
bioassay systems for water quality. J In Vitro Fert Embryo Transf 1988; 5(6): 335–342.
6. van den Bergh M, Baszo I, Biramane J, Bertrand E, Devreker F and Englert Y. Quality control in IVF with mouse
bioassays: A four years’ experience. J Assist Reprod Genet 1996; 13(9): 733–738.
7. Gardner DK, Reed L, Linck D, Sheehan C and Lane M. Quality control in human in vitro fertilization. Semin
Reprod Med 2005; 23(4): 319–324.
8. Punt-van der Zalm JP, Hendriks JC, Westphal JR, Kremer JA, Teerenstra S and Wetzels AM. Toxicity testing
of human assisted reproduction devices using the mouse embryo assay. Reprod Biomed Online 2009; 18(4):
529–535.
9. Morimoto Y, Hayashi E, Ohno T, Kawata A, Horikoshi Y and Kanzaki H. Quality control of human IVF/ICSI
program using endotoxin measurement and sperm survival test. Hum Cell 1997; 10(4): 271–276.
10. Fleetham JA, Pattinson HA and Mortimer D. The mouse embryo culture system: Improving the sensitivity for use
as a quality control assay for human in vitro fertilization. Fertil Steril 1993; 59(1): 192–196.
11. Davidson A, Vermesh M, Lobo RA and Paulson RJ. Mouse embryo culture as quality control for human in vitro
fertilization: The one-cell versus the two-cell model. Fertil Steril 1988; 49(3): 516–521.
12. Khan Z, Wolff HS, Fredrickson JR, Walker DL, Daftary GS and Morbeck DE. Mouse strain and quality control
testing: Improved sensitivity of the mouse embryo assay with embryos from outbred mice. Fertil Steril 2013; 99:
847–854.e2.
13. Wolff HS, Fredrickson JR, Walker DL and Morbeck DE. Advances in quality control: Mouse embryo morphoki-
netics are sensitive markers of in vitro stress. Hum Reprod 2013; 28(7): 1776–1782.
14. Morbeck DE, Khan Z, Barnidge DR and Walker DL. Washing mineral oil reduces contaminants and embryotox-
icity. Fertil Steril 2010; 94(7): 2747–2752.
15. Thomas T. Culture systems: Air quality. Methods Mol Biol 2012; 912: 313–324.
16. Sharmin R and Ray MB. Application of ultraviolet light-emitting diode photocatalysis to remove volatile organic
compounds from indoor air. J Air Waste Manag Assoc 2012; 62(9): 1032–1039.
17. Chapuis Y, Klvana D, Guy C and Kirchnerova J. Photocatalytic oxidation of volatile organic compounds using
fluorescent visible light. J Air Waste Manag Assoc 2002; 52(7): 845–854.
18. Hall J, Gilligan A, Schimmel T, Cecchi M and Cohen J. The origin, effects and control of air pollution in labora-
tories used for human embryo culture. Hum Reprod 1998; 13(Suppl 4): 146–155.
19. Cohen J, Gilligan A and Willadsen S. Culture and quality control of embryos. Hum Reprod 1998; 13(Suppl 3):
137–144; discussion 145–137.
20. Cohen J, Gilligan A, Esposito W, Schimmel T and Dale B. Ambient air and its potential effects on conception in
vitro. Hum Reprod 1997; 12(8): 1742–1749.
21. Boone WR, Johnson JE, Locke AJ, Crane MM, 4th and Price TM. Control of air quality in an assisted reproduc-
tive technology laboratory. Fertil Steril 1999; 71(1): 150–154.
22. Khoudja RY, Xu Y, Li T and Zhou C. Better IVF outcomes following improvements in laboratory air quality.
J Assist Reprod Genet 2013; 30(1): 69–76.
23. Merton JS, Vermeulen ZL, Otter T, Mullaart E, de Ruigh L and Hasler JF. Carbon-activated gas filtration during
in vitro culture increased pregnancy rate following transfer of in vitro-produced bovine embryos. Theriogenology
2007; 67(7): 1233–1238.
24. Guris J, Janssens R and Vercammen M. Air quality managment. In: Practial Manual of In Vitro Fertilization (eds.
Nagy Z, Varghese A and Argarwal A). New York: Springer, 2012, pp. 17–27.
25. Higdon HL, 3rd, Blackhurst DW and Boone WR. Incubator management in an assisted reproductive technology
laboratory. Fertil Steril 2008; 89(3): 703–710.
26. Schimmel T, Gilligan A, Garrisi G, Espositi B, Cecchi M, Dale B, et al. Removal of volatile organic compounds
from incbuators used for gamete and embryo culture. Fertil Steril 1997; 68(Suppl 1): s165.
27. Mayer J, Nechchiri F, Weedon V, Jones E, Kalin H, Ochniger S, et al. Prospective randomized crossover analysis
of the impact of an IVF incbuator air filtration system (coda, genx) on clinical pregnancy rates. Fertil Steril 1999;
72(Suppl 1): s42.
28. Baltz JM and Tartia AP. Cell volume regulation in oocytes and early embryos: Connecting physiology to success-
ful culture media. Hum Reprod Update 2010; 16(2): 166–176.
29. Hammer MA and Baltz JM. Beta-alanine but not taurine can function as an organic osmolyte in preimplantation
mouse embryos cultured from fertilized eggs. Mol Reprod Dev 2003; 66(2): 153–161.
30. Richards T, Wang F, Liu L and Baltz JM. Rescue of postcompaction-stage mouse embryo development from
hypertonicity by amino acid transporter substrates that may function as organic osmolytes. Biol Reprod 2010;
82(4): 769–777.
31. Moravek M, Fisseha S and Swain JE. Dipeptide forms of glycine support mouse preimplantation embryo devel-
opment in vitro and provide protection against high media osmolality. J Assist Reprod Genet 2012; 29(3):
283–290.
32. Swain JE, Cabrera L, Xu X and Smith GD. Microdrop preparation factors influence culture-media osmolality,
which can impair mouse embryo preimplantation development. Reprod Biomed Online 2012; 24(2): 142–147.
33. Anas MK, Hammer MA, Lever M, Stanton JA and Baltz JM. The organic osmolytes betaine and proline
are transported by a shared system in early preimplantation mouse embryos. J Cell Physiol 2007; 210(1):
266–277.
34. Dawson KM and Baltz JM. Organic osmolytes and embryos: Substrates of the Gly and beta transport systems
protect mouse zygotes against the effects of raised osmolarity. Biol Reprod 1997; 56(6): 1550–1558.
35. Dawson KM, Collins JL and Baltz JM. Osmolarity-dependent glycine accumulation indicates a role for glycine
as an organic osmolyte in early preimplantation mouse embryos. Biol Reprod 1998; 59(2): 225–232.
36. Hammer MA and Baltz JM. Betaine is a highly effective organic osmolyte but does not appear to be transported
by established organic osmolyte transporters in mouse embryos. Mol Reprod Dev 2002; 62(2): 195–202.
37. Hammer MA, Kolajova M, Leveille M, Claman P and Baltz JM. Glycine transport by single human and mouse
embryos. Hum Reprod 2000; 15(2): 419–426.
38. Lane M, Mitchell M, Cashman KS, Feil D, Wakefield S and Zander-Fox DL. To QC or not to QC: The key to a
consistent laboratory? Reprod Fertil Dev 2008; 20(1): 23–32.
39. Swain JE. Optimizing the culture environment in the IVF laboratory: Impact of pH and buffer capacity on gamete
and embryo quality. Reprod Biomed Online 2010; 21(1): 6–16.
40. Swain JE. Is there an optimal pH for culture media used in clinical IVF? Hum Reprod Update 2012; 18(3):
333–339.
41. Swain JE and Pool TB. New pH-buffering system for media utilized during gamete and embryo manipulations for
assisted reproduction. Reprod Biomed Online 2009; 18(6): 799–810.
42. Lane M, Lyons EA and Bavister BD. Cryopreservation reduces the ability of hamster 2-cell embryos to regulate
intracellular pH. Hum Reprod 2000; 15(2): 389–394.
43. Phillips KP and Baltz JM. Intracellular pH regulation by HCO3-/Cl- exchange is activated during early mouse
zygote development. Dev Biol 1999; 208(2): 392–405.
44. Phillips KP, Petrunewich MA, Collins JL and Baltz JM. The intracellular pH-regulatory HCO3-/Cl- exchanger in
the mouse oocyte is inactivated during first meiotic metaphase and reactivated after egg activation via the MAP
kinase pathway. Mol Biol Cell 2002; 13(11): 3800–3810.
45. Hamamah S, Magnoux E, Royere D, Barthelemy C, Dacheux JL and Gatti JL. Internal pH of human spermato-
zoa: Effect of ions, human follicular fluid and progesterone. Mol Hum Reprod 1996; 2(4): 219–224.
46. Johnson JE, Boone WR, Lee ST and Blackhurst DW. Using Fyrite to monitor incubator carbon dioxide levels.
J Assist Reprod Genet 1995; 12(2): 113–117.
47. Boone WR, Johnson JE, Lee ST, Detry MA and Blackhurst DW. Sources of variation in measurements of carbon
dioxide levels inside incubators. J Assist Reprod Genet 1996; 13(7): 606–608.
48. Sun XF, Wang WH and Keefe DL. Overheating is detrimental to meiotic spindles within in vitro matured human
oocytes. Zygote 2004; 12(1): 65–70.
49. Wang WH, Meng L, Hackett RJ, Oldenbourg R and Keefe DL. Rigorous thermal control during intracytoplasmic
sperm injection stabilizes the meiotic spindle and improves fertilization and pregnancy rates. Fertil Steril 2002;
77(6): 1274–1277.
50. Wang WH, Meng L, Hackett RJ, Odenbourg R and Keefe DL. Limited recovery of meiotic spindles in living
human oocytes after cooling-rewarming observed using polarized light microscopy. Hum Reprod 2001; 16(11):
2374–2378.
51. Leese HJ, Baumann CG, Brison DR, McEvoy TG and Sturmey RG. Metabolism of the viable mammalian
embryo: Quietness revisited. Mol Hum Reprod 2008; 14(12): 667–672.
52. Dewdney, A. K., 200% of Nothing: An Eye Opening Tour Through the Twists and Turns of Math Abuse and
Innumeracy. New York: Wiley, 1993.
53. Elert G et al., Temperature of a healthy human body (body temperature), The Physics Factbook, http://www.
hypertextbook.com/facts.
54. Grinsted J, Kjer JJ, Blendstrup K and Pedersen JF. Is low temperature of the follicular fluid prior to ovulation
necessary for normal oocyte development? Fertil Steril 1985; 43(1): 34–39.
55. David A, Vilensky A and Nathan H. [Temperature changes in different parts of the rabbit oviduct. Preliminary
report]. Harefuah 1971; 80(4): 180–182.
56. Hunter RH, Bogh IB, Einer-Jensen N, Muller S and Greve T. Pre-ovulatory graafian follicles are cooler than
neighbouring stroma in pig ovaries. Hum Reprod 2000; 15(2): 273–283.
57. Hunter RH, Grondahl C, Greve T and Schmidt M. Graafian follicles are cooler than neighbouring ovarian tissues
and deep rectal temperatures. Hum Reprod 1997; 12(1): 95–100.
58. Hunter RH and Nichol R. A preovulatory temperature gradient between the isthmus and ampulla of pig oviducts
during the phase of sperm storage. J Reprod Fertil 1986; 77(2): 599–606.
59. Hunter RH. Temperature gradients in female reproductive tissues. Reprod Biomed Online 2012; 24(4): 377–380.
60. Hong K, Forman E, Lee H, Ferry K, Treff N and Scott R. Optimizing the temperature for embyo culture in
IVF: A randomized controlled trial (RCT) comparing standard culture temperature of 37C to the reduced more
physiologic temperature of 36C. Fertil Steril 2012; 98(3): s167.
61. Cooke S, Tyler JP and Driscoll G. Objective assessments of temperature maintenance using in vitro culture
techniques. J Assist Reprod Genet 2002; 19(8): 368–375.
62. Zhang JQ, Li XL, Peng Y, Guo X, Heng BC and Tong GQ. Reduction in exposure of human embryos outside the
incubator enhances embryo quality and blastulation rate. Reprod Biomed Online 2010; 20(4): 510–515.
4
Morphological Selection of Gametes and Embryos: Sperm
Pierre Vanderzwalmen, Magnus Bach, Olivier Gaspard, Bernard Lejeune,
Anton Neyer, Françoise Puissant, Maximilian Schuff, Astrid Stecher,
Sabine Vanderzwalmen, Barbara Wirleitner, and Nicolas H. Zech
Importance of Spermatozoa Morphology in Fertility

Intracytoplasmic sperm injection (ICSI) introduced more than 20 years ago was a tremendously helpful tool to
overcome the infertility of couples when conventional in vitro fertilization (IVF), partial zona dissection, or sub-
zonal sperm injection treatments had failed [1]. The injection of a single spermatozoon into the ooplasm provided
an apt answer to severe male infertility diagnosis indicated by low sperm count, poor sperm motility, or both, or
to infertility due to morphology deficiency. Interestingly, since the introduction of ICSI, less attention has been
devoted to the sperm’s morphology in itself. In addition, it is even more remarkable that after the introduction of
ICSI, even though human spermatozoa exhibit a wide range of shapes, several studies found no correlation between
the injection of sperm with normal or abnormal morphology and ICSI outcomes [2,3]. However, such observations
were most probably biased by the selection performed by the embryologist who tried to select the best “normal-
looking” motile spermatozoa before ICSI, which does not always reflect the quality of the whole semen population.
The assessment of sperm morphology by Kruger’s strict criteria is routinely applied and widely accepted as
the best method of prediction for male fertility potential and highlights the concept that sperm morphology is a
very important parameter in the analysis of the whole population of a semen sample [4]. Of all semen parameters,
sperm morphology turns out to be the best predictor of a man’s fertilizing potential [5].
But the situation is completely different when we move from the diagnostic level toward the laboratory prac-
tice of ICSI where we have to select only one spermatozoon for injection.
Although there is still a controversy as to whether morphological defects of spermatozoa even have an effect
on fertilization or subsequent embryonic development [6], it becomes more and more accepted that abnormally
shaped spermatozoa from patients diagnosed with terato- and asthenozoospermia have a significantly increased
frequency of aneuploidy, a higher DNA fragmentation index (DFI), and an increased rate of mitochondrial
dysfunction [7]. The importance of morphologically normal sperm selection is reinforced when facing the repro-
ductive outcomes in terms of fertilization, embryo development, pregnancy rates, and abortion rates when the
oocyte injections can be done only with abnormally shaped spermatozoa, that is, sperm with elongated, tapered,
or amorphous heads; broken necks; or cytoplasmic droplets [8].
In view of ICSI bypassing the natural barriers of reproduction, it seems reasonable to develop
optimized sperm selection techniques.
With the implementation of restrictive laws regulating the number of embryos for transfer, methods of gamete
and embryo selection are of paramount importance. If we keep in mind that ICSI bypasses the natural barriers of
reproduction and that an abnormal spermatozoon bears the danger of transferring putative negative effects on to
the offspring, it might be reasonable to develop optimized selection techniques. Now, for almost a decade, there-
fore, more attention has been directed towards physiological selection methods based on the biochemical ability of
the spermatozoa to bind either to solid hyaluronic acid or to zona pellucida before intracytoplasmic injection [9].
59
In addition, the importance of sperm morphology is being studied [8], with several novel microscopic
approaches, such as differential interference contrast microscopy [10], digital holographic microscopy [11], and
atomic force microscopy [12], currently being used or in development to allow a more detailed observation of the
different parts of the sperm head (nucleus and acrosome), midpiece, and tail.
Bartoov et al. [10] introduced, 10 years ago, an innovative, noninvasive technique for a more precise mor-
phological evaluation of motile spermatozoa. The so-called motile-sperm organelle-morphology examination
(MSOME) changed the perception of how a spermatozoon suitable for injection should appear. In fact, MSOME
permits not only simple observation in terms of a sperm’s size and shape but also a detailed examination in real
time of the subtle subcellular morphology and abnormalities, such as nuclear defects. The MSOME technique
allowed a stricter discrimination of spermatozoa, with normal nuclei defined by an oval shape with a smooth
configuration, including a normal nuclear content [10,13].
With the replacement of the standard bright-field or the Hoffman modulation contrast (HMC) optics by the
Nomarski differential interference contrast (DIC) optics, a better three-dimensional view of the sperm head
became available. Bright-field or HMC can reveal major abnormalities of the spermatozoon’s aberrations in head
shape and at the level of the midpiece and tail. With Nomarski interference contrast, a more in-depth evaluation
of the quality of semen that provides substantial information about the normalcy of the head, neck, and midpiece
is possible. In consequence, this method of spermatozoa evaluation and selection for ICSI indication generated
the intracytoplasmic morphologically selected sperm injection (IMSI) technique [14,15].
The aim of the first part of this chapter is to give the reasons to select spermatozoa free of nuclear and neck–
midpiece defects or those from patients suffering from globozoospermia syndrome. In addition, a brief overview
of the putative nature of nuclear vacuoles according to the present status of knowledge and their consequences
on embryo development is presented. In the second part of this chapter, we depict the different spermatozoa
classification systems and the technical procedures to select spermatozoa so as not to impair oocyte quality.
Tangible Arguments to Select Sperm Free of Defects
One decade after the first application of MSOME and IMSI, can we still consider these as
useful techniques?
Sperm Head Vacuoles: An Additional Sperm Defect That Can Be Better

Highlighted with Nomarski Optics
The meticulous approach in sperm selection applying the Nomarski DIC optics allows the detection of subtle
sperm head nuclear abnormalities that were designated as vacuoles by Bartoov et al. [10] (Figure 4.1a). These
nuclear defects are otherwise not evident at a 400× magnification with HMC microscopy [15–17] (Figure 4.1b).
In fact, we had to wait for the introduction of Nomarski optics to be concerned about the presence of large and
small vacuoles and then consider them as potential defects. Before the introduction of MSOME and IMSI, there
was to our knowledge no report of the presence of vacuoles and analysis of their location, origin, structure, and
effect on reproductive outcome after the conventional ICSI procedure.
Spermatozoa might contain heterogeneous vacuoles varying in number, size, and content. To simplify this
matter, we often only refer to the term “vacuole” in any circumstance. But is vacuole the appropriate terminology?
From a biological point of view, vacuoles are vesicles surrounded by a membrane. However, this structural
abnormality that we airily term vacuole might more likely represent a kind of depression in the nuclear part of the
head. Some of these so-called vacuoles are in fact deep, such as crater or hollow. Watanabe et al. [18] suggested
that vacuoles are hollows where in the plasma membrane falls into the nucleus, and Westbrook et al. [19] even
talked about nuclear crater formation in the sperm head. Toshimori [20] differentiates between large v acuoles
with amorphous substances or membranous structures inside, and small vacuolous patterns without any
structures inside.
Sperm 61
LNV ?
LNV
?
SV
LNV ?
SV
? LNV
? LNV
(a) (b) (c)
FIGURE 4.1 Observation of spermatozoa with HMC and DIC microscopy (Nomarski). (a) Nomarski at 6000× magnification.
(b) Conventional HMC at 6000× magnification. (c) Conventional HMC at 600× after adjustment of the polarizer. SV, small vacuole;
LNV, large nuclear vacuole.
Boitrelle et al. [12,21] reported that the plasma membrane was sunken, but intact, both large and small vac-
uoles were identified as an abnormal, “thumbprint”-like nuclear concavity covered by acrosomal and plasma
membranes.
Reasons to Select Vacuole-Free Sperm

Pathological Character of Nuclear Vacuoles
One crucial question to investigate concerns the significance of vacuoles. Vacuoles are like liquid bubbles that
can be seen in the sperm’s head, where the nucleus is located, and that have been correlated with DNA anoma-
lies. Considering the publications of Berkovitz et al. [15], Cayli et al. [22], and Hazout et al. [23], it is most
likely that vacuoles reflect molecular defects that are responsible for anomalies of sperm chromatin packaging
and abnormal chromatin remodeling during sperm maturation; such defects, in turn, may render spermatozoa
more vulnerable to DNA damage. According to different studies, the integrity of the chromatin is related to the
presence or absence of vacuoles in the head of spermatozoa and the loss of chromatin compaction renders the
exposed DNA more prone to reactive oxygen species [24–26].
Several DNA and chromatin staining assays, including aniline blue, chromomycin A3, and acridine orange,
were applied to assess more precisely the integrity of the DNA in vacuolated spermatozoa. Interestingly, several
studies found highly statistically significant differences in the DNA integrity between spermatozoa with or
without vacuoles [27].
According to the growing body of literature on this subject, it is now more and more accepted that the pres-
ence of large nuclear vacuoles (LNVs) is correlated with failures of chromatin condensation [7,28,29]. It is
reported in selected teratozoospermic populations that sperm vacuoles were exclusively of nuclear origin and
preferentially located toward the anterior part of the sperm head [30]. Chromatin condensation defects were the
main alterations observed in spermatozoa with large vacuoles [7,12,21,28,30–32]. The chromatin in sperm with
large vacuoles was atypically decondensed, showing a high degree of immaturity.
Indeed, chromatin condensation is a crucial step in protecting the paternal genome during the transit from
the male to the oocyte before fertilization (epididymal transit) [33]. Rousseaux et al. [34] demonstrated a new
keystone in DNA compaction in humans and murines. They postulated that histones are replaced by transi-
tional proteins called bromodomains before protamination takes place. Structural abnormalities of the nucleus
include incomplete or impaired chromatin condensation and nuclear vacuoles and inclusions. Karyolytic changes
or the presence of large intranuclear lacunae or vacuoles are the morphological manifestations of underlying
biochemical alterations.
Early embryogenesis is a critical time for epigenetic regulation, and these epigenetic modulation processes are
sensitive to environmental factors [35,36]. Epigenetic patterns are usually faithfully maintained during development.
However, this maintenance sometimes fails, resulting in the disturbance of epigenetic p rocesses representing the
basis of numerous human disorders. It is known that before histone replacement by protamines, the nucleosomes are
destabilized by hyperacetylation, resulting in rise in the DNA methylation levels [37,38]. These potential epigenetic
mechanisms could be implied in chromatin condensation failures [39].
Poor chromatin condensation may expose the spermatozoon’s nuclear DNA to damage (e.g., DNA fragmenta-
tion) during its journey through both the male and female genital tracts [40,41], and large vacuoles were found
to be associated with DNA fragmentation [7,28]. However, other studies failed to establish a strong link between
large vacuoles and DNA fragmentation [12,18,30,32].
This could explain the differences in terms of DNA fragmentation between different men. A disorder in
spermiogenesis may result in uncondensed and vulnerable chromatin, and according to exposure to the level of
oxidative stress conditions, differences in DNA fragmentation can be noticed [35].
Vacuoles and Reproductive Outcome

The importance of selecting normal spermatozoa becomes obvious when comparing the reproductive outcomes
in terms of fertilization, embryo development, and pregnancy and abortion rates when oocyte injections are done
with morphologically normal sperm and spermatozoa exhibiting different subcellular defects.
Selection of sperm devoid of sperm head vacuoles has multiple benefits.
Vacuoles and Embryo Development

The size and the number of sperm nuclear vacuoles, most accurately identified under Nomarski optics, negatively
affect blastocyst development. In successive studies [42–45], the existence of LNVs together or not with abnor-
mal shape was shown to reduce the percentage of good-quality blastocysts after culture until Day 5. Following
the outcome of each embryo after injection of spermatozoa, it was clearly demonstrated that the use of spermato-
zoa with no vacuoles or fewer than two small vacuoles can be associated with significantly higher blastocyst rates
compared with injecting spermatozoa with more than two small vacuoles or one large vacuole with or without
abnormal shape. Such observations reinforce previous studies suggesting early and late paternal effects on initial
embryonic development [46–48].
As mentioned, it seems that nuclear vacuoles are related to sperm DNA damage and might therefore negatively
affect human embryo development [47]. In addition, it is well known that the integrity of sperm chromatin plays
a key role in embryo development [25]. DNA helices in defective chromatin remodeling during spermiogenesis
would be more vulnerable to physical and chemical stress such as reactive oxygen species [26]. Subsequent
attack by reactive oxidative species may cause sperm’s DNA fragmentation and might even affect later blastocyst
development [26,49] and pregnancy outcomes in a negative manner [47,50,51].
Vacuoles and Pregnancy and Miscarriage Rates

Berkovitz et al. [52] were the first to carry out a more specific analysis on the impact of sperm cells with
normal nuclear shape but with large vacuoles on two matched IMSI groups of 28 patients each. Spermatozoa
with strictly defined normal nuclear shape but large vacuoles were selected for injection and compared with a
control group that included normal nuclear shape spermatozoa lacking vacuoles. No difference in the fertiliza-
tion and early embryo development up to Day 3 was reported. However, injection of spermatozoa with strictly
normal nuclear shape but large vacuoles appeared to significantly reduce pregnancy outcomes (18% vs. 50%)
and seemed to be associated with early miscarriage (80% vs. 7%).
Other studies showed that selection of normally shaped spermatozoa with a vacuole-free head was positively
associated with pregnancy and lower abortion rates after Day 3 embryo transfers in couples with previous and
repeated implantation failures [13,14,42,52,53] and in patients with an elevated degree of DNA-fragmented
spermatozoa [23].
Knez et al. [44] showed that there was no significant difference in the pregnancy rates after IMSI and ICSI
procedures and blastocyst transfer. However, after ICSI, more pregnancies were terminated by spontaneous
Sperm 63
abortion, whereas after IMSI there were no spontaneous abortions. One explanation could be that the IMSI pro-
cedure permits selection of spermatozoa without defects and thus provides more “healthy” blastocysts without
chromosomal abnormalities, possibly in spite of very comparable development and morphology to ICSI-derived
blastocysts.
Vacuoles, Health of Babies Born, and Incidence of Malformations

Still, there are concerns about the long-term safety of injecting spermatozoa carrying vacuoles. We have to be
cautious, especially in the light of Aitken’s work [54] on the putative negative effects of sperm DNA fragmenta-
tion for the next generation. Depending on the level of sperm nuclear DNA fragmentation, oocytes may partially
repair fragmented DNA, producing blastocysts capable of implanting and producing live offspring. However, the
incomplete repair may lead to long-term pathologies. The data of Fernández-Gonzalez et al. [55] gained on the
basis of a mouse model indicate that the use of DNA-fragmented spermatozoa in ICSI can generate effects such
as aberrant growth, premature aging, abnormal behavior, and tumors derived from the mesenchymal lineage that
emerge only in later life.
To date, there have not been sufficient numbers of published studies concerning the health of children born
after ICSI to draw any firm conclusions about the long-term safety of this procedure. However, it is important to
emphasize that animal data are absolutely unequivocal on this point and clearly indicate that DNA damage in the
male germ line is potentially hazardous for the embryo and therefore for the resulting offspring [56]. According
to Cassuto [39], sperm nucleus morphological normalcy, assessed at high magnification, could decrease the
prevalence of major fetal malformations in ICSI children.
Midpiece Selection
MSOME permits not only the observation of nuclear vacuole defects but also other abnormalities that may result
in infertility. Ugajin et al. [57] showed that they were able to clearly differentiate between a straight-shaped and
tapering-shaped midpiece (Figure 4.2). The tapering-shaped midpiece is related to abnormal centrosomal func-
tion as a consequence of aberrant microtubule organization [57]. According to Van Blerkom and Davis [58], the
centriole is, after the nucleus, the most important sperm organelle for initiation of the intraooplasmic fertilization
(a) (b) (c)
FIGURE 4.2 Tapering-shaped (a), intermediate-shaped (b), and straight-shaped (c) midpiece.

process, being responsible for the formation of the sperm aster. The centrosomes play a crucial role in fertiliza-
tion [59], and their dysfunction may cause fertilization failure due to lack of a sperm aster [60,61]. The injection
of selected sperm with a morphologically straight midpiece by IMSI may result in choosing sperm with func-
tional centrosomes, thereby positively influencing fertilization rates and embryo development after ICSI [57].
MSOME Selection in Globozoospermia to Circumvent Artificial Activation

Globozoospermia is a male-factor infertility involving several abnormalities of sperm remodeling during sper-
miogenesis that can lead to formation defects or premature elimination of acrosomal structures. Fertilization
capacity after ICSI is low [62], mainly due to a lack of sperm-specific phospholipase C (PLCζ) [63] that is
responsible for inducing Ca2+ oscillations essential for oocyte activation.
Presently, 21 births have been reported after ICSI followed by artificial Ca2+ oscillations induction via the use
of artificial oocyte activation protocols by exposing oocytes to certain chemical agents (ionophore) [64–67].
Another approach proposed by Kashir et al. [68] and Gatimel et al. [67] is to apply MSOME to maximize the
efficacy of ICSI and IMSI. MSOME increases the efficacy of fertilization by selecting spermatozoa with superior
oocyte activation ability. Sermondade et al. [69] selected spermatozoa that exhibited a small acrosomal bud that
was not visible using conventional sperm selection methodology. Gatimel et al. [67] identified spermatozoa with
some sparse oval forms, revealing the presence of Golgi residues. These studies showed that some globozoosper-
mic spermatozoa possess an acrosomal bud that contains a sufficient amount of PLCζ that is not significantly
different from fertile controls and that correlates to successful oocyte activation and fertilization without the
need for artificial oocyte activation with exogenous chemical agents.
Evaluation of Sperm Head Shape

At high magnification (6300×), Utsuno et al. [70] decomposed sperm head shapes of motile spermatozoa into
four quantitative parameters: ellipticity, anteroposterior symmetry, lateral symmetry, and angularity. Stepwise
forward multiple logistic regression analysis showed a statistically significant increase of the percentage of DNA
fragmentation in spermatozoa with abnormal ellipticity, angularity, and LNVs. This may ensure the advantage
of morphological assessment of spermatozoa at high magnification.
IMSI in 2014: Still a Debate

As mentioned, there is a real benefit to select morphological spermatozoa free of vacuoles. But, for different
reasons, there is undeniable evident skepticism about this method of spermatozoa selection. Controversial pub-
lications led to the hesitation to apply IMSI. What are the main indications for IMSI? How can we assess the
benefit of IMSI?
Superiority of IMSI versus ICSI—But for Which Indications?

When should we propose IMSI instead of ICSI? In principle, all patients can benefit from using IMSI; however, it is
especially useful after several previous implantation failures, in cases of severe teratozoospermia, for patients hav-
ing experienced an unexplained abortion, and definitely in cases with no blastocysts of or low blastocyst rate(s) in
previous IVF attempts. However, the superiority of IMSI over ICSI is still a matter of debate. To date, randomized
and well-powered studies to confirm a benefit of IMSI are limited, and they even depict conflicting results [71].
Controversial conclusions have been drawn, especially in terms of fertilization, top-quality embryo rates, and
pregnancy rates between IMSI and ICSI. Some studies have shown that IMSI improves reproductive outcomes
in cases of male-factor infertility and previous failed ICSI attempts in terms of implantation and clinical preg-
nancy rates compared with conventional ICSI [14,16,23,42,44,72–77]. Alternatively, IMSI and conventional ICSI
seemed to provide comparable laboratory and clinical results when an unselected infertile population was evalu-
ated [78,79] or when IMSI was applied as the first treatment option [80].
Sperm 65
Full Benefit of IMSI Is Attained with Blastocyst Culture

Even if the superiority of IMSI over ICSI is still a matter of debate, we have to recognize that the type of sper-
matozoa selected for injection influences the outcome in terms of embryo development, pregnancy, miscarriage,
and malformation.
One of the main benefits of IMSI is the improvement in embryo quality and the higher rate of blastocysts
obtained per cycle when morphologically good-quality spermatozoa are selected [42–45].
The type of spermatozoa selected for injection influences the outcome in terms of embryo
development, pregnancy, miscarriage, and malformation.
The full benefit of a better spermatozoa selection is highlighted if we are able to produce one more blastocyst
per cycle. In fact, more blastocysts provide a higher chance for the patient to become pregnant in successive
vitrified embryo transfer cycle(s), thus increasing the cumulative pregnancy rate. Moreover, when using IMSI,
the abortion ratios are reduced by 50%, increasing as a consequence the full-term gestation possibilities.
Also, prolonged embryo culture to the blastocyst stage (5 days) can serve as a strong diagnostic tool, reflecting
indications of male and female infertility and yielding useful information regarding the implantation potential
of the human embryo [44].
Explaining the Discrepancies between Studies

Different conclusions may be drawn to explain the absence of differences observed by some studies. For example,
for several studies, embryo transfer is performed on Day 2 or Day 3. It is fair to mention that the selection of
embryos with the higher developmental potential and implantation capacity is almost impossible at this stage,
even with new technologies such as the time lapse. In fact, embryo selection on Day 2 or Day 3 completely
neglects the paternal effect that becomes important only after embryonic genome activation (EGA) on Day 3.
It was previously shown that such “late paternal effect” negatively influences preimplantation embryo develop-
ment and clinical outcomes without any detectable impairment in zygote development, such as cleaving speed
or embryo quality at this stage [48,72].
Another possible explanation for these conflicting results could be the way in which conventional ICSI is
performed in an individual laboratory, and how experienced the embryologists are in performing the ICSI proce-
dure and in particular the selection of spermatozoa for injection. Collecting sperm using a 20× or 40× objective
is quite common. However, at this low-magnification, morphological defects of the sperm head can be hardly
detected [81].
But with a well-aligned microscope equipped with 40× Hoffman contrast optics and optimal adjustment of
both the optical beam and the polarizer, detection of sperm head alterations is possible. In addition, with a
1.5-fold increase in the magnification, it is practicable to observe LNV at 600× magnification (Figure 4.1c).
However a more accurate and simple detection of small vacuoles is attained with Nomarski optics.
From our experience, we realized after some months of application of MSOME-IMSI that it was possible to
observe nuclear defect even with a classical objective. When we started to analyze in a sibling study the rate
of blastocysts after IMSI or ICSI, we observed a significant difference of almost 25% in favor of IMSI. With
increasing experience and being aware of the presence of a vacuole, the probability to select, with the conven-
tional ICSI microscope, spermatozoa free of a vacuole was increased. This has as a consequence an increase in
the rate of blastocysts after ICSI and attenuation in the difference between both techniques.
The conclusions of such observations are that regardless of potential indications of IMSI, we have to select
the best spermatozoa possible and exclude those carrying nuclear defects for all male infertility patients.
Of course, not all spermatozoa with a nuclear vacuole will have a negative impact on the developmental rate,
but if morphologically normal spermatozoa are present in the suspension, it is mandatory to try to select them
independently of the technique that is available.
Thus, we have to do all we can to select the best spermatozoa. There are absolutely no indications to select
bad-quality spermatozoa if good spermatozoa are present in the prepared semen sample. Are there still
indications where improved sperm selection before fertilization is not necessary or low-magnification micros-
copy using HMC is more than enough? Most probably this is not the case.
Technical Aspects
Several reservations concerning technical aspects serve as arguments not to apply this method of spermatozoa
selection:
1. Too sophisticated system? Too complicated to perform.

2. Lack of standardized sperm selection criteria according to MSOME (classification).
3. Time-consuming aspect during spermatozoa selection.
4. Prolonged selection time of male gametes at the expense of oocyte aging.
a. Problems in managing assisted reproductive technology (ART) laboratory
b. High cost
The aim of our work is first to use this technology in a user-friendly way and second to classify the semen and
provide in real time a spermocytogram (MSOME), and for the IMSI technique to select spermatozoa in a manner
that does not impair oocyte quality.
Materials
Basic Materials
Observation at high magnification (MSOME) or sperm selection (before oocyte injection) is performed on an
inverted light microscope equipped with DIC/Nomarski DIC optics, with dry or immersion 63× or 100× mag-
nification objectives lens. In our IVF units, we use a Leica 6000 (Leica Microsystems, Germany) equipped with
63× and 100× DIC objectives.
Dry objectives instead of immersion are more convenient when dishes or slides are moved or replaced for each
oocyte injection. Fast motion or dynamic movements between drops induce air bubble formation between the
objective lens and the dish, preventing the visualization of the semen sample. The use of dry objective permits
an easy handling of the dishes, in particular when a new dish is used for each oocyte injection.
Additional Materials
Using a variable zoom lens (HC Vario C-mount; Leica Microsystems) and subsequent magnification of the
image with a high-definition digital video camera makes it possible to evaluate spermatozoan morphology
on a monitor at magnifications between 6600× and 12,000× in real time or after spermatozoa immobiliza-
tion. Despite all the available equipment and computer software, there is not a high priority to perform such
observation and selection of spermatozoa using high magnification (>6000×) in conjunction with digital
image capture systems to analyze spermatozoa in detail after their capture and visualization on a color
monitor screen.
Practical and technical aspects of sperm selection need to be considered to facilitate the workflow
while performing IMSI.
For IMSI, inverted microscopes are equipped with the classical ICSI equipment (Narashige or Eppendorf
micromanipulators and injectors). Calculation of the total magnification on the screen monitor depends on sev-
eral technical specifications of the objective, the magnification selector of the microscope, the variable zoom
lens, the camera chip diagonal dimension, and the television monitor diagonal dimension.
Sperm 67
MSOME: A Useful Tool for Routine Laboratory Semen Analysis

Morphological Normalcy of Spermatozoa Assessed by MSOME
Based on electron microscopy, and high-magnification optic microscopy, Bartoov et al. [10] defined the
morphological normalcy of motile-sperm nucleus according to the shape and chromatin content. The shape
has to be smooth, symmetric, and oval, with average length and width limits estimated to 4.75 μm ± 0.28 μm and
3.28 μm ± 0.20 μm, respectively (Figure 4.3).
The chromatin mass has to be homogeneous and should contain no extrusions or invaginations, with a
maximum of one vacuole involving <4% of the nuclear area. The acrosome and post-acrosomal lamina are con-
sidered abnormal if absent, partial, or vesiculated. An abaxial neck with the presence of disorders or c ytoplasmic
droplets as well as the presence of broken, short, or double and coiled tail is considered abnormal.
Establishment of an MSOME Sperm Classification

According to Bartoov et al. [10], normal spermatozoa should not contain nuclear vacuole(s) exceeding >4%
of the nuclear area. Saidi et al. [82] and Perdrix et al. [83] classified the relative area of the vacuole into
three groups: (1) <5.9%–6.5%; (2) between 6.5% and 13% or between 5.9% and 12.4%; and (3) >13% or 12.4%,
respectively. For Franco et al. [28], LNVs in spermatozoa were defined by the presence of vacuoles occupying
≥50% of the sperm nuclear area.
Under those circumstances where no normal spermatozoa in the semen sample can be found, the only alter-
native is then to select the morphologically “second-best spermatozoa.” Hence, it is essential to know from the
second-choice spermatozoa with vacuoles, abnormal shape, or both as to which spermatozoa we have to select.
It seemed that the establishment of a classification system of the spermatozoa according to different types of
abnormalities was mandatory for the selection of spermatozoa in a more tangible way and also to analyze their
influence on further outcomes.
To estimate the impact of specific sperm defects on embryo development and further outcomes in an accurate
way, different groups established models of sperm classification according to the normalcy of the shape and
the presence and size of vacuoles [42,43]. For example, Vanderzwalmen et al. [42] classified spermatozoa into
4.75 μm + 2.8 µm
3.28 μm + 0.2 µm
4.0 μm + 5.0 µm
0.78 + 0.18
FIGURE 4.3 Representation of a normal spermatozoon according to Bartoov criteria.

2
3b 3b
3c 3b 1c
2
3b
3c
3a
1b
1b
1a
FIGURE 4.4 Spermatozoa vacuolization classification according to Vanderzwalmen et al. [42], observed with Nomarski optics
63× dry objective. Grade I spermatozoa with normal head shape without vacuole (1a), with one vacuole (1b), and with maximum of
two small vacuoles (1c). Grade II spermatozoa with large nuclear vacuole (LNV) (2). Grade III spermatozoa with abnormal shape
without vacuole (3a), small vacuoles (3b), and LNV (3c).
FIGURE 4.5 Spermatozoa vacuolization pattern. (a) High frequency of grade I spermatozoa (fertilization after conventional
insemination). (b) Severe teratozoospermia (100 spermatozoa grade III).
three grades according to the presence and size of vacuoles: Grade I, normal shape and a maximum of two small
vacuoles; Grade II, normal shape and more than two small vacuoles or at least one large vacuole; Grade III,
abnormal head shapes with or without large vacuoles in conjunction with other abnormalities at the level of the
base (Figures 4.4 and 4.5) [42].
Cassuto et al. [43] established a detailed classification scoring scale ranging between 6 and 0 points according
to the normalcy of the head (2 points if normal), the symmetry of the base (1 point if normal), and the absence
of vacuole (3 points if absent). Three grades of scoring were therefore established: grade 1, high-quality sper-
matozoa with calculated score of 4–6; grade 2, medium-quality spermatozoa with calculated score of 1–3; and
grade 3, low-quality spermatozoa with calculated score of 0.
MSOME for Routine Laboratory Semen Analysis

To assess the usefulness of the evaluation of sperm morphology by MSOME, two studies were undertaken in
a first instance by Bartoov et al. [10,13] that estimated the correlation between MSOME and the World Health
Organization (WHO) routine method. More recently, Oliveira et al. [84] compared the MSOME evaluation with
the Tygerberg classification criteria.
Both works conclude that the MSOME criteria appear to be much more restrictive, presenting significantly
lower sperm normalcy percentages for the semen samples compared with those found after routine analysis by
WHO criteria and the Tygerberg classification. In addition, MSOME represented a much stricter evaluation,
because the use of Nomarski optics enabled the identification of vacuoles that could not be described with the
same accuracy with other methods.
Sperm 69
These studies point toward a benefit in sperm morphology and quality evaluation by including MSOME
among the criteria for routine laboratory semen analysis before ICSI or conventional IVF procedures. A previ-
ous MSOME spermocytogram revealing a high percentage of vacuoles may be judicious to propose directly
IMSI as the best therapy for ICSI candidates. Furthermore the additional information gained by MSOME may
help to avoid fertilization failure in IVF cycles.
Proposed Cutoff Threshold for Further Treatment Decisions

With the aim to define a predictive value of sperm normalcy using MSOME on the outcome of combined IVF-
ICSI, Wittemer et al. [85] undertook a study including 55 couples with previous failure of implantation after
intrauterine insemination (IUI) treatments. In their next attempt, a combination of conventional IVF and ICSI was
proposed for each couple. They concluded that below a threshold of 8% of morphological normal s permatozoa
observed by MSOME, ICSI must be performed instead of conventional IVF to avoid the risk of fertilization failure.
Cassuto (2011, personal communication) observed that if <42% of score 6 spermatozoa (equivalent to our grade I
classification) are present in the semen sample, ICSI-IMSI has to be performed instead of c onventional IVF.
Falagario et al. [86] identified a cutoff of 20, 28% of sperm nuclear vacuolization (SNV) on the total of sperm in
a seminal sample as a physiological threshold. In their study, they observed that patients undergoing ICSI, grouped
according to SNV, showed similar percentages of fertilization and embryo development after Day 2 embryo trans-
fer but that more pregnancies were achieved with a higher implantation rate when SNV was ≤ 20, (28%). They con-
cluded that SNV rate could be introduced as an easy diagnostic evaluation before performing an ICSI–IMSI cycle.
Sperm Preparation for Morphological Observation

For MSOME, observation of the spermatozoa may be done on the native semen or after the sperm washing
procedure on a three-layer gradient of pure sperm (Nidacon, Sweden), as described previously [87]. It should be
stressed that MSOME was applied exclusively to motile spermatozoa that under low-light microscopy magnifica-
tion have a high potential to be selected for ICSI.
How to Perform MSOME

In certain situations, when we have to decide quickly after the oocyte pick-up (OPU) whether IVF or ICSI-IMSI
has to be performed, a rapid assessment of the morphology may be a good decision. If MSOME was part of
a previous exam (e.g., spermocytogram), assessment of the morphology is done in a more sophisticated way with
the microchannel capture technique.
Rapid Morphological Assessment

To receive a rapid qualitative evaluation of the semen sample, a fast morphological assessment is performed
using a microscope coverglass (24 × 60 mm). Elongated drops (~5 μL–10 μL) of 3%–10% polyvinyl pyrrolidone
(PVP) solutions are placed on the coverglass (Figure 4.6a). An aliquot of (1 μL–10 μL) native or washed semen
sample is transferred on the border of one elongated drop (Figure 4.6b). In case of too low sperm concentration
or too low motility, drops of semen sample are placed in the three elongated drops of 3% PVP.
When spermatozoa start to swim, the evaluation of the morphology of motile spermatozoa is done in real time.
According to the initial concentration, 50–100 spermatozoa are observed, classified, and recorded in a file (Figure 4.7).
Morphological Examination Using the Microdrops Capture Channel

In cases where quantitative evaluation of the semen sample is needed for research purposes, elongated drops of
culture medium are placed into a glass-bottomed dish (GWST 5040; WillCo, World Precision Instruments Wells BV,
the Netherlands) and covered with mineral oil (Irvine Scientific, Ireland). The number of elongated drops depends on
the semen quality. With reduced sperm quality (severe oligoasthenozoospermia), more elongated drops are prepared.
After the washing step, the sperm suspension is deposited on the border at one end of the elongated drop.
When motile spermatozoa start to swim along the edge of the elongated drop, sperm capture channels are created
Microscope
cover glass
Elongated PVP drops
(a) (b)
FIGURE 4.6 (a) Preparation of the slide with PVP drops for MSOME. (b) Arrow indicates where the microdrop of semen is deposited.
MSOME
SPZ Class 1 SPZ Class 2 SPZ Class 3

(normal form; no, one, or two vacuoles ≤4%) (normal form, vacuoles >4%) (more abnormalities)
NF Normal Form without vacuoles NFLV Normal Form, one Large Vacuole AFL/SV(n) Abnormal Form, Large + Small Vacuoles
NFSV <4% Normal Form, Small Vacuole <4% NFLV(n) Normal Form, Large Vacuoles AFLV Abnormal Form, one Large Vacuole
NFSV(2) <4% Normal Form, two Small Vacuoles <4% NFL/SV(n) Normal Form, Large + Small Vacuoles AF Abnormal Form without vacuole
FIGURE 4.7 Flow chart reporting the percentages of spermatozoa according to the different vacuolization patterns.
by pulling channels from the drop with a microinjection pipette ICSI pipette (Microtech, Czech Republic) that is
fixed on the micromanipulator [88] (Figure 4.8).
A proper setup of the dish for selecting and capturing sperm facilitates the routine
application of IMSI.
Motile spermatozoa enter the small bays and accumulate at the end, where they stop their progressive
movement due to a restriction of the free moving space.
A total of eight channels per elongated drop are created and examined (Figure 4.8). Because the majority of
the spermatozoa present at the end of the channel show no progressive motility, it is possible to follow them and
take several pictures. Morphological assessment of the spermatozoa may be performed in real time under the
microscope at magnifications between 630× and 1000×.
A subsequent analysis is conducted on the monitor screen after taping the channel using time-lapse recording
(Leica Application Suite magnification optics; Leica). Nevertheless, static sperm imaging only allows evaluation
of the visible part of sperm and might leave some morphological alterations undetected.
A minimum of 100 spermatozoa were analyzed and documented on a specific file (Figure 4.7).
Sperm 71
Channel of culture medium with sperm

suspension
Sperm microcapture channels
Area of
continuous
observation
FIGURE 4.8 Application of the sperm-microcapture channels, where spermatozoa accumulate for MSOME.
Assessment with a Transparent Celluloid Form

Bartoov et al. [13] advised assessing the normalcy of spermatozoa by superimposing a fixed transparent celluloid
form on the motile examined gametes. This fixed, transparent, celluloid form represents a sperm nucleus fitting
the criteria of normalcy of the head (average length, 4.75 μm ± 0.28 μm; average width, 3.28 μm ± 0.20 μm). The
correct sperm size, calculated by the ratio of expected normal sperm size to the actual size, can be visualized on
the monitor screen. The nuclear shape is documented as abnormal if it varies in length or width by 2x standard
deviation from the normal mean axes values [13]. This way of analyzing the morphology is extremely precise.
The disadvantage of this technique is that it is very time-consuming and difficult in practice to compare the
spermatozoa fitting the criteria of normalcy with moving spermatozoa.
IMSI
Sperm Preparation before IMSI
Before IMSI, only washed semen was used after density-gradient centrifugation on one to three layers of pure
sperm (Nidacon, Sweden) [87]. The number of layers depended on the initial sperm concentration.
Two Strategies to Perform IMSI

Three criticisms about IMSI are frequently argued. The first concerns the prolonged period of time for oocytes
out of the incubator during spermatozoa selection. The second criticism is that the prolonged selection time of
male gametes may be at the detriment of oocyte and promote oocyte aging. Finally, some argue that exposing
the spermatozoa for a longer period at 37°C increases the rate of vacuoles. To reduce the time of the oocytes out
of the incubator, two approaches were implemented, as presented below.
First Approach: Sperm Selection and Oocyte Injection on Two Different Microscopes
The spermatozoa are selected using the Nomarski optic, and oocyte injection is performed with a conventional
ICSI microscope. With such a strategy, the oocytes are not present in the dish during selection of the spermato-
zoa. After selection, the dish is incubated for 15–30 min to stabilize the temperature, pH, or both.
In cases of severe teratozoospermia, taking into account that more time for sperm selection is needed, we plan
the selection at least around the time of OPU. With such a policy, oocyte injection can be performed 2–3 hr after
OPU (38–40 hr post–human chorionic gonadotropin [hCG] administration) on the ICSI station, thereby avoiding
oocyte aging. Another advantage of this approach concerns the organization of the laboratory work. In cases of
several IMSI, the IMSI station is occupied only for the selection process and not for the injection phase. Also
with this approach, the IMSI microscope is not occupied for an excessive period, and the oocytes are removed
from the incubator only for the injection step.
Another possibility is to select the spermatozoa in a glass-bottomed dish and place them directly in a conven-
tional ICSI dish.
Preparation of IMSI Dishes Under sterile conditions, several drops are deposited in the glass-bottomed dishes
(Figure 4.9).
Sperm Selection PVP Drops (A) On the left side of the dish, place one to three elongated drops of 7.5%–10%
PVP (~5 μL–10 μL). In this drop, selection of the spermatozoa is performed. The number of drops depends on the
quality of the semen that was assessed in a previous MSOME analysis or on the number of metaphase II (MII)
oocytes, which are needed for injection. The aim is that the spermatozoa that we select stay in the PVP drop for
a period that does not exceed 15 min. Moreover, according to this rule, several elongated drops and even IMSI
dishes should be prepared if many MII oocytes are present.
Sperm Aliquots Drops (B) Under each sperm selection drop (A), a drop of ~10 μL of human tubal fluid
(HTF)-HEPES (IVFonline, Canada) containing 6% human synthetic albumin (HSA) (Irvine Scientific) is
deposited.
Host-Selected Spermatozoa Microdrops (C) Adjacent to the elongated drops of PVP (A), very small drops (<1
μL) (C) of HTF-HEPES containing 6% HSA are deposited with a small stripper pipette. The microdrops will
host the spermatozoa that were selected in drop A until oocyte injection.
Sperm Immobilization Drops (D) A small drop of 10% PVP in which sperm immobilization will take place is
deposited in the upper part of the dish.
D
D
E E
A A A E E A A A E
C E E
C E
C C E
C
B B B C E
C
B B B
FIGURE 4.9 Position of the drops in an IMSI dish (A, D) 7.5%–10% PVP. (B, C, E) Culture medium. (A) Sperm selection PVP
drops. (B) Sperm aliquot drops. (C) Host-selected spermatozoa microdrops. (D) Sperm immobilization drops. (E) Oocytes injection
drops.
Sperm 73
Oocytes Injection Drops (E) The right side of the dish contains five drops of HTF-HEPES containing 6% HSA
in which oocyte injection will take place following the policy of IMSI procedure.
All drops were covered with sterile mineral oil (Irvine Scientific).
Method of Spermatozoa Selection and Injection The entire selection process is performed at room temperature.
At the beginning of IMSI, a small aliquot of washed sperm (1 μL–10 μL according to the semen sample quality)
is transferred into drop B (Figure 4.10a) and a small bridge is formed between drop B and drop A (Figure 4.10b).
The motile spermatozoa swim into the PVP drop and the morphologically normal spermatozoa (or the best second
class) are selected with an ICSI pipette and transferred into a small drop of culture media (drop C) (Figure 4.10c).
They are selected at a magnification of 1000×.
After collecting spermatozoa (if possible, 1.5 times the number of oocytes to inject), the dish is removed from
the IMSI station and placed on a 37°C heating stage for temperature recovery (~30 min).
After this period of incubation at 37°C, the oocytes are placed into the culture media (E), and ICSI is performed
using a conventional Hoffman microscope at 400× magnification (Figure 4.10d). Motile spermatozoa are aspi-
rated from the preselected host drop (C) and immobilized in the PVP drop (D) before injection.
Second Approach: Sperm Selection and Oocyte Injection on the IMSI Station
When the purpose of the study is to follow the outcome of embryo development in relation to the type of injected
spermatozoon (photodocumentation of the injected spermatozoon), a maximum of two oocytes are placed
directly in the IMSI dish (Figure 4.11). A minimum of two dishes are prepared as presented previously, except
that sperm host drops are not present before starting such an approach, the quality of the spermatozoa should be
assessed to be sure that the time spent to select one class I or II spermatozoa will not exceed 2 min.
This procedure is performed at 37°C. At the beginning of the IMSI procedure, a small aliquot of washed sperm
(1 μL–10 μL) is transferred into drop B and a small bridge is formed between drop B and the elongated selection
drop. The motile spermatozoa swim in the PVP drop, and the motile spermatozoa are morphologically selected
at a magnification of 1000× (Figure 4.11a) before immobilization in drop D (Figure 4.11b). Use of a variable
IMSI IMSI
microscope microscope
(a) (b)
IMSI
Hoffman
microscope
microscope
(c)
(d)
FIGURE 4.10 IMSI. (a–c) Description of the different steps for sperm selection on the IMSI microscope followed by (d) oocyte
injection on the Hoffman ICSI microscope.
IMSI IMSI
(a) (b)
FIGURE 4.11 IMSI. Description of the different steps for sperm selection (a) and oocyte injection on the IMSI microscope (b).
zoom lens (HC Vario C-mount; Leica) allows for the reevaluation (after immobilization) of the morphology on
the monitor at magnifications between 6600× and 12,000× and the performance of photodocumentation for a
subsequent analysis. Injections are performed directly after immobilization in the same dish. After IMSI, single
drop culture (in a petri dish or with the time-lapse technology) is performed.
The next two oocytes are placed in a new IMSI dish prewarmed on the heating stage. The previous IMSI dish
is then placed on a heating stage at 37°C until the next injection selection and injection step. This technique is
easy to apply with dry objectives, so that no problem with oil occurs when changing dishes.
Time Spent to Select Spermatozoa

Regarding time allotted to select spermatozoa, the primary intention is to choose normal-shaped spermatozoa
without any vacuoles (grade I) for injection into the oocytes. Depending on the degree of impaired sperm mor-
phology, the mean time required for selecting the best sperm ranged between 2 and 15 min. However, when it
is obvious after 15 min of sperm examination that spermatozoa of a normal morphology cannot be found, the
second-best spermatozoa with the least number of vacuoles, other abnormalities, or a combination is selected
for injection.
In such situations, it might be difficult to decide whether to stop the search for a normal spermatozoon or to
proceed. It might take 15 min and even longer; of course, this also depends on the number of oocytes that have
to be injected and on the number of patients that have to be treated.
Temperature for Sperm Evaluation or Selection

Another crucial question is whether the duration of sperm selection and the temperature might influence vacuole
formation. A study by Peer et al. [89] demonstrated that after 2 hr of incubation at 37°C in culture media, the
incidence of spermatozoa with vacuolated nuclei was significantly higher compared with incubation at 21°C.
Peer and colleagues suggested that prolonged (≥2 hr) sperm manipulations for ART should be performed at
21°C rather than 37°C. Schwarz et al. [90] reported a significant increase in SNV in washed sperm but not in
swim-up sperm. They concluded that the mode of sperm preparation does influence SNV and that vacuolization
is unaffected by temperature in motile sperm isolated by swim-up.
Several experiments were conducted in our centers to analyze the formation of vacuoles in real time. We imple-
mented a time-lapse recording approach on selected Grade I spermatozoa incubated for 24 hr at 37°C. Compared
with the control group, no changes in the morphology of the spermatozoa were observed. No vacuoles appeared.
Sperm 75
Even when the same experiment was conducted on spermatozoa with vacuoles, no changes in the size and shape
of the spermatozoa after 24 hr incubation at 37°C were found [88].
Conclusions
One of the most essential questions is not under which technical conditions the selection of spermatozoa should
be recommended but rather whether we have to consider to select the best spermatozoa and, if possible, exclude
those carrying defects.
For this reason, MSOME and IMSI were developed. MSOME appears to be a powerful method to improve our
understanding of human spermatozoa, and IMSI is now routinely used in ART practices.
However, the clinical use of MSOME and IMSI remains unclear in the fields of male infertility diagnosis
and ARTs. Answers to a lot of questions are pending or unclear for the following issues: (1) the t erminology
of vacuoles, their classification, and their location on the sperm head; (2) vacuole origin and formation;
(3) application of MSOME-IMSI for specific indications, such as teratozoospermia or to a large popula-
tion; (4) the application of IMSI instead of IVF in cases of unexplained infertility; and (5) oocyte repairing
factors. Nevertheless, it is increasingly obvious that large vacuoles reflect a pathological situation, most
probably correlated with sperm chromatin immaturity. In addition, we have to be aware that this technique
is challenging and has to be p erformed under the best working conditions so as to not impair the quality
of oocytes.
The introduction of IMSI made embryologists aware that in times of ICSI the selection of sperm
has to be given proper attention.
The introduction of IMSI has the advantage that embryologists realize that more attention has been given to
sperm selection even in cases of classical ICSI. The application of IMSI leads to more and better quality blasto-
cysts and thus it increases the chance of selecting the proper embryo for transfer with the highest implantation
potential. The full benefit of using MSOME as an additional tool to ICSI procedure manifests when it is per-
formed in combination with Day 5 embryo culture of all fertilized oocytes. Also, knowing the excellent results
obtained with vitrification should not be underestimated.
Even though there are only few reports in the human species on the abnormal outcome generated by sperma-
tozoa carrying vacuoles, a higher and better-resolution technique has to be added as an additional tool for ICSI,
knowing the possible consequence of sperm DNA damage for the offspring. Presently, one study reports higher
de novo malformation after ICSI selection than after IMSI selection [39].
REFERENCES
1. Palermo G, Joris H, Devroey P, Van Steirteghem A. Pregnancies after intracytoplasmic injection of single sperma-
tozoon into an oocyte. Lancet. 1992; 340: 17–18.
2. Nagy ZP, Liu J, Joris H, et al. The result of intracytoplasmic sperm injection is not related to any of the three basic
sperm parameters. Hum. Reprod. 1995; 10: 1123–1129.
3. Svalander P, Jakobsson AH, Forsberg AS, Bengtsson AC, Wikland M. The outcome of intracytoplasmic sperm
injection is unrelated to ‘strict criteria’ sperm morphology. Hum. Reprod. 1996; 11: 1019–1022.
4. Vawda A, Gunby J, Younglai E. Semen parameters as predictors of in-vitro fertilization: The importance of strict
criteria morphology. Hum. Reprod. 1996; 11: 1445–1450.
5. Nikolettos N, Kiipker W, Demirel C, et al. Fertilization potential of spermatozoa with abnormal morphology.
Hum. Reprod. 1999; 14(Suppl 1): 47–70.
6. Host E, Ernst E, Lindenberg S, Smidt-Jensen S. Morphology of spermatozoa used in IVF and ICSI from
oligozoospermic men. Reprod. Biomed. Online. 2001; 3: 212–215.
7. Garolla A, Fortini D, Menegazzo M, et al. High power magnification microscopy and functional status analysis
of sperm in the evaluation and selection before ICSI. Reprod. Biomed. Online. 2008; 17: 610–616.
8. De Vos A, Van De Velde H, Joris H, et al. Influence of individual sperm morphology on fertilization, embryo
morphology, and pregnancy outcome of intracytoplasmic sperm injection. Fertil. Steril. 2003; 79: 42–48.
9. Ebner T, Filicori M, Tews G, Parmegiani L. A plea for a more physiological ICSI. Andrologia. 2011; 44: 1–18.
10. Bartoov B, Berkovitz A, Eltes F. Selection of spermatozoa with normal nuclei to improve the pregnancy rate with
intracytoplasmic sperm injection. N. Engl. J. Med. 2001; 345: 1067–1068.
11. Coppola G, Di Caprio G, Wilding M, et al. Digital holographic microscopy for the evaluation of human sperm
structure. Zygote. 2013; 7: 1–9.
12. Boitrelle F, Ferfouri F, Petit JM, et al. Large human sperm vacuoles observed in motile spermatozoa under
high magnification: Nuclear thumbprints linked to failure of chromatin condensation. Hum. Reprod. 2011; 26:
1650–1658.
13. Bartoov B, Berkovitz A, Eltes F, Kogosowski A, Menezo Y, Barak Y. Real-time fine morphology of motile human
sperm cell is associated with IVF-ICSI outcome. J. Androl. 2002; 23: 1–8.
14. Bartoov B, Berkovitz A, Eltes F, et al. Pregnancy rates are higher with intracytoplasmic morphologically selected
sperm injection than with conventional intracytoplasmic injection. Fertil. Steril. 2003; 80: 1413–1419.
15. Berkovitz A, Eltes F, Yaari S, et al. The morphological normalcy of the sperm nucleus and pregnancy rate of
intracytoplasmic injection with morphologically selected sperm. Hum. Reprod. 2005; 20: 185–190.
16. Berkovitz A, Eltes F, Lederman H, et al. How to improve IVF-ICSI outcome by sperm injection. Reprod. Biomed.
Online. 2006; 12: 634–638.
17. Dorphin B, Lefevre M, Gout C, Oger P, Yazbeck C, Rougier N. Fine localization and characterization of some
abnormalities observed in human sperm heads by interferential differential contrast. Hum. Reprod. 2010; 25: P–043.
18. Watanabe S, Tanaka A, Fujii S, Mizunuma H. No relationship between chromosome aberrations and vacuole-like
structures on human sperm head. Hum. Reprod. 2009; 24: i94–i96.
19. Westbrook VA, Diekman AB, Klotz KL, et al. Spermatid-specific expression of the novel X-linked gene product
SPAN-X localized to the nucleus of human spermatozoa. Biol. Reprod. 2000; 63; 469–481.
20. Toshimori K. Dynamics of the Mammalian Sperm Head. Advances in Anatomy, Embryology and Cell Biology
Volume 204, 2009, pp. 45–51.
21. Boitrelle F, Albert M, Petit J-M, et al. Small human sperm vacuoles observed under high magnification are
pocket-like nuclear concavities linked to chromatin condensation failure. Reprod. Biomed. Online. 2013;

27: 201–211.
22. Cayli S, Jakab A, Ovari L, et al. Biochemical markers of sperm function: Male fertility and sperm selection for
ICSI. Reprod. Biomed. Online. 2003; 7: 462–468.
23. Hazout A, Dumont-Hassan M, Junca AM, Cohen Bacrie P, Tesarik J. High-magnification ICSI overcomes paternal
effect resistant to conventional ICSI. Reprod. Biomed. Online. 2006; 12: 19–25.
24. Gopalkrishnan K, Padwal V, Meherji PK, Gokral JS, Shah R, Juneja HS. Poor quality of sperm as it affects
repeated early pregnancy loss. Arch. Androl. 2000; 45: 111–117.
25. Hammadeh M, Nkemayim D, Georg T, Rosenbaum P, Schmidt W. Sperm morphology and chromatin condensa-
tion before and after semen processing. Arch. Androl. 2000; 44: 221–226.
26. Mahfouz R, Sharma R, Thiyagarajan A, et al. Semen characteristics and sperm DNA fragmentation in infertile
men with low and high levels of seminal reactive oxygen species. Fertil. Steril. 2010; 94: 2141–2146.
27. Perrin A, Huong Nguyen M, Douet-Guilbert N, Morel F, De Braekeleer M. Motile sperm organelle morphology
examination: Where do we stand 12 years later? Expert Rev. Obstet. Gynecol. 2013; 8: 249–260.
28. Franco J, Baruffi R, Mauri A, Petersen C, Oliveira J, Vagnini L. Significance of large nuclear vacuoles in human
spermatozoa: Implications for ICSI. Reprod. Biomed. Online. 2008; 17: 42–45.
29. Franco J, Mauri A, Petersen C, et al. Large nuclear vacuoles are indicative of abnormal chromatin packaging in
human spermatozoa. Int. J. Androl. 2012; 35: 1–6.
30. Perdrix A, Travers A, Chelli MH, et al. Assessment of acrosome and nuclear abnormalities in human spermatozoa
with large vacuoles. Hum. Reprod. 2011; 26: 47–58.
31. Boitrelle F, Ferfouri F, Salama S, et al. Nature of sperm-head vacuoles and results and indications of IMSI (intra-
cytoplasmic morphologically selected sperm injection). Andrologie. 2011; 21: 234–239.
32. Cassuto NG, Hazout A, Hammoud I, et al. Correlation between DNA defect and sperm-head morphology. Reprod.
Biomed. Online. 2012; 24: 211–218.
33. Ward WS. Function of sperm chromatin structural elements in fertilization and development. Mol. Hum. Reprod.
2010; 16: 30–36.
Sperm 77
34. Rousseaux S, Petosa C, Müller CW, Müller CW, Khochbin S. A novel key element in post-meiotic male genome
reprogramming. Med. Sci. 2010; 26: 130–132.
35. Aitken RJ, De luliis GN. Origins and consequences of DNA damage in male germ cells. Reprod. Biomed. Online.
2007; 14: 727–733.
36. Koppers AJ, De Iuliis GN, Finnie JM, McLaughlin EA, Aitken RJ. Significance of mitochondrial reactive oxygen
species in the generation of oxidative stress in spermatozoa. J. Clin. Endocrinol. Metab. 2008; 93: 3199–3207.
37. Palermo GD, Neri QV, Takeuchi T, Squires J, Moy F, Rosenwaks Z. Genetic and epigenetic characteristics of
ICSI children. Reprod. Biomed. Online. 2008; 17: 820–833.
38. Miller D, Brinkworth M, Iles D. Paternal DNA packaging in spermatozoa: More than the sum of its parts? DNA,
histones, protamines and epigenetics. Reproduction. 2010; 139: 287–301.
39. Cassuto G. Low birth defects by deselecting abnormal spermatozoa before ICSI. Reprod. Biomed. Online. 2014;
28: 47–53.
40. Aitken RJ, Koppers AJ. Apoptosis and DNA damage in human spermatozoa. Asian J. Androl. 2011; 13: 36–42.
41. Oliva R, de Mateo S. Medical implications of sperm nuclear quality. In: Epigenetics and Human Reproduction
(eds. Rousseaux S, Khochbin S). Berlin: Springer-Verlag, 2011, pp. 45–55.
42. Vanderzwalmen P, Hiemer A, Rubner P, et al. Blastocyst development after sperm selection at high magnification
is associated with size and number of nuclear vacuoles. Reprod. Biomed. Online. 2008; 17: 617–627.
43. Cassuto NG, Bouret D, Plouchart JM, et al. A new real-time morphology classification for human spermatozoa:
A link for fertilization and improved embryo quality. Fertil. Steril. 2009; 92: 1616–1625.
44. Knez K, Zorn B, Tomazevic T, Vrtacnik-Bokal E, Virant-Klun I. The IMSI procedure improves poor embryo
development in the same infertile couples with poor semen quality: A comparative prospective randomized study.
Reprod. Biol. Endocrinol. 2011; 9: 123–130.
45. Knez K, Tomazevic T, Vrtacnik-Bokal E, Virant-Klun I. Developmental dynamics of IMSI-derived embryos:
A time-lapse prospective study. Reprod. Biomed. Online. 2013; 27: 161–171.
46. Shoukir Y, Chardonnens D, Campana A, Sakkas D. Blastocyst development from supernumerary embryos after
intracytoplasmic sperm injection: A paternal influence. Hum. Reprod. 1998; 13: 1632–1637.
47. Tesarik J, Greco E, Mendoza C. Late but not early paternal effect on human embryo development is related to
sperm DNA fragmentation. Hum. Reprod. 2004; 19: 611–615.
48. Tesarik J. Paternal effects on cell division in the preimplantation embryo. Reprod. Biomed. Online. 2005; 10:
370–375.
49. Seli E, Gardner DK, Schoolcraft WB, Moffatt O, Sakkas D. Extent of nuclear DNA damage in ejaculated
spermatozoa impacts on blastocyst development after in vitro fertilization. Fertil. Steril. 2004; 82: 378–383.
50. Larson KL, DeJonge CJ, Barnes AM, Jost LK, Evenson DP. Sperm chromatin structure assay parameters as pre-
dictors of failed pregnancy following assisted reproductive techniques. Hum. Reprod. 2000; 15: 1717–1722.
51. Nasr-Esfahani MH, Salehi M, Razavi S, et al. Effect of sperm DNA damage and sperm protamine deficiency on
fertilization and embryo development post-ICSI. Reprod. Biomed. Online. 2005; 11: 198–205.
52. Berkovitz A, Eltes F, Ellenbogen A, Peer S, Feldberg D, Bartoov B. Does the presence of nuclear vacuoles in
human sperm selected for ICSI affect pregnancy outcome? Hum. Reprod. 2006; 21: 1787–1790.
53. Junca AM, Cohen-Bacrie P, Hazout A. Improvement of fertilisation and pregnancy rate after intracytoplasmic fine
morphology selected sperm injection. Fertil. Steril. 2004; 82(Suppl 2): S173.
54. Aitken R. Origins and consequences of DNA damage in male germ cells. Reprod. Biomed. Online. 2007;
14: 727–733.
55. Fernandez-Gonzalez R, Nuno Moreira P, Perez-Crespo M, et al. Long-term effects of mouse intracytoplasmic
sperm injection with DNA-fragmented sperm on health and behavior of adult offspring. Biol. Reprod. 2008;
78: 761–772.
56. Berkovitz A, Eltes F, Paul M. The chance of having a healthy normal child following intracytoplasmic
Morphologically-selected sperm injection (IMSI) treatment is higher compared to conventional IVF-ICSI treat-
ment. Fertil. Steril. 2007; 88: S20.
57. Ugajin T, Terada Y, Hasgawa H, Nabeshima H, Suzuki K, Yaegashi N. The shape of the sperm midpiece in
intracytoplasmic morphologically selected sperm injection relates sperm centrosomal function. J. Assit. Reprod.
Genet. 2010; 27: 75–81.
58. Van Blerkom J, Davis P. Evolution of the sperm aster after microinjection of isolated human sperm centrosomes
into meiotically mature human oocytes. Hum. Reprod. 1995; 10: 2179–2182.
59. Schatten H, Sun QY. The role of centrosomes in mammalian fertilization and its significance for ICSI. Mol. Hum.
Reprod. 2009; 15: 531–538.
60. Rawe V, Olmedo S, Nodar F, Doncel G, Acosta A, Vitullo A. Cytoskeletal organization defects and abortive acti-
vation in human oocytes after IVF and ICSI failure. Mol. Hum. Reprod. 2000; 6: 510–516.
61. Rawe V, Terada Y, Nakamura S, Chillik C, Olmedo S, Chemes H. A pathology of the sperm centriole responsible
for defective sperm aster formation, sygamy and cleavage. Hum. Reprod. 2002; 17: 2344–2349.
62. Dam AH, Feenstra I, Westphal JR, Ramos L, van Golde RJ, Kremer JA. Globozoospermia revisited. Hum. Reprod.
Update. 2007; 13: 63–75.
63. Cox L, Larman MG, Saunders CM, Hashimoto K, Swann K, Lai FA. Sperm phospholipase Czeta from humans
and cynomolgus monkeys triggers Ca2+ oscillations, activation and development of mouse oocytes. Reproduction.
2002; 124: 611–623.
64. Kashir J, Heindryckx B, Jones C, De Sutter P, Parrington J, Coward K. Oocyte activation, phospholipase C zeta
and human infertility. Hum. Reprod. Update. 2010; 16: 690–703.
65. Montag M, Köster M, van der Ven K, Bohlen U, van der Ven H. The benefit of artificial oocyte activation is depen-
dent on the fertilization rate in a previous treatment cycle. Reprod. Biomed. Online. 2012; 24(5): 521–526.
66. Stecher A, Bach M, Neyer A, Vanderzwalmen P, Zintz M, Zech NH. Case report: Live birth following ICSI with
non-vital frozen-thawed testicular sperm and oocyte activation with calcium ionophore. J. Assist. Reprod. Genet.
2011; 28: 411–414.
67. Gatimel N, Léandri D, Foliguet B, Bujan L, Parinaud J. Sperm cephalic vacuoles: New arguments for their non
acrosomal origin in two cases of total globozoospermia. Andrology. 2013; 1: 52–56.
68. Kashir J, Sermondade N, Sifer C, et al. Motile sperm organelle morphology evaluation-selected globozoospermic
human sperm with an acrosomal bud exhibits novel patterns and higher levels of phospholipase C zeta. Hum.
Reprod. 2012; 27(11): 3150–3160.
69. Sermondade N, Hafhouf E, Dupont C, et al. Successful childbirth after intracytoplasmic morphologically selected
sperm injection without assisted oocyte activation in a patient with globozoospermia. Hum. Reprod. 2011; 26:
2944–2949.
70. Utsuno H, Oka K, Yamamoto A, Shiozawa T. Evaluation of sperm head shape at high magnification revealed
correlation of sperm DNA fragmentation with aberrant head ellipticity and angularity. Fertil. Steril. 2013;
99: 1573–1580.
71. Perdrix A, Rives N. Motile sperm organelle morphology examination (MSOME) and sperm head vacuoles: State
of the art in 2013. Hum. Reprod. Update. 2013; 19: 527–541.
72. Greco E, Scarselli F, Fabozzi G, et al. Sperm vacuoles negatively affect outcomes in intracytoplasmic morpho-
logically selected sperm injection in terms of pregnancy, implantation, and live-birth rates. Fertil. Steril. 2013;
100: 379–385.
73. Antinori M, Licata E, Dani G, et al. Intracytoplasmic morphologically selected sperm injection: A prospective
randomized trial. Reprod. Biomed. Online. 2008; 16: 835–841.
74. Nadalini M, Tarozzi N, Distratis V, Scaravelli G, Borini A. Impact of intracytoplasmic morphologically selected
sperm injection on assisted reproduction outcome. Reprod. Biomed. Online. 2009; 19: 45–55.
75. Balaban B, Yakin K, Alatas C, Oktem O, Isiklar A, Urman B. Clinical outcome of intracytoplasmic injection
of spermatozoa morphologically selected under high magnification: A prospective randomized study. Reprod.
Biomed. Online. 2011; 22: 472–476.
76. Setti SA, Ferreira RC, Paes de Almeida Ferreira Braga D, de Cássia Sávio Figueira R, Iaconelli A, Jr, Borges E,
Jr. Intracytoplasmic sperm injection outcome versus intracytoplasmic morphologically selected sperm injection
outcome: A meta-analysis. Reprod. Biomed. Online. 2010; 21: 450–455.
77. Delaroche L, Yazbeck C, Gout C, Kahn V, Oger P, Rougier N. Intracytoplasmic morphologically selected sperm
injection (IMSI) after repeated IVF or ICSI failures: A prospective comparative study. Eur. J. Obstet. Gynecol.
Reprod. Biol. 2013; 167: 76–80.
78. De Vos A, Van de Velde H, Bocken G, et al. Does intracytoplasmic morphologically selected sperm injection
improve embryo development? A randomized sibling-oocyte study. Hum. Reprod. 2013; 28: 617–626.
79. Mauri A, Petersen C, Oliveira J, Massaro F, Baruffi R, Franco JG, Jr. Comparison of day 2 embryo quality
after conventional ICSI versus intracytoplasmic morphologically selected sperm injection (IMSI) using sibling
oocytes. Eur. J. Obstet. Gynecol. Reprod. Biol. 2010; 150: 42–46.
Sperm 79
80. Leandri RD, Gachet A, Pfeffer J, et al. Is intracytoplasmic morphologically selected sperm injection (IMSI) ben-
eficial in the first ART cycle? A multicentric randomized controlled trial. Andrology. 2013; 5: 692–697.
81. Montag M, Toth B, Strowitzki T. Sperm selection in ART. J. Reproduktionsmed. Endokrinol. 2012; 9:
485–489.
82. Saidi R, Rives N, Gruel E, Mazurier S, Mousset-Simeon N, Mace B. New classification of spermatocytogram at
high magnification. Med. Reprod. Gyn. Endo. 2008; 10: 315–324.
83. Perdrix A, Saïdi R, Ménard JF, et al. Relationship between conventional sperm parameters and motile sperm
organelle morphology examination (MSOME). Int. J. Androl. 2012; 35: 491–498.
84. Oliveira JB, Massaro FC, Mauri AL, et al. Motile sperm organelle morphology examination is stricter than
Tygerberg criteria. Reprod. Biomed. Online. 2009; 18: 320–326.
85. Wittemer C, Pujol A, Boughali H, et al. The impact of high-magnification evaluation of sperm on ART o utcome.
Hum. Reprod. 2006; 22(Suppl 1): i59–i60.
86. Falagario D, Brucculeri A, Depalo R, Trerotoli P, Cittadini E, Ruvolo G. Sperm head vacuolization affects clinical
outcome in ICSI cycle. A proposal of a cut-off value. J. Assist. Reprod. Genet. 2012; 29: 1281–1287.
87. Van der Zwalmen P, Bertin-Segal G, Geerts L, Debauche C, Schoysman R. Sperm morphology and IVF
pregnancy rate: Comparison between Percoll gradient centrifugation and swim-up procedures. Hum. Reprod.
1991; 6: 581–588.
88. Neyer A, Vanderzwalmen P, Bach M, Stecher A, Spitzer D, Zech N. Sperm head vacuoles are not affected by
in-vitro conditions, as analysed by a system of sperm-micro capture channels. Reprod. Biomed. Online. 2013;
26: 368–377.
89. Peer S, Eltes F, Berkovitz A, Yehuda R, Itsykson P, Bartoov B. Is fine morphology of the human sperm nuclei
affected by in vitro incubation at 37C? Fertil. Steril. 2007; 88: 1589–1594.
90. Schwarz C, Köster M, van der Ven K, Montag M. Temperature-induced sperm nuclear vacuolization is dependent
on sperm preparation. Andrologia. 2012; 44(Suppl 1): 126–129.
5
Morphological Selection of Gametes and Embryos: Oocyte
Introduction
Assessment of oocyte morphology and determination of its correlation with quality and viability and with the
clinical outcome is a difficult task, as the underlying mechanisms that change its appearance are multifactorial
and complex. Optimal oocyte morphology (Figure 5.1) is defined as an oocyte with spherical structure enclosed
by a uniform zona pellucida (ZP), with a uniform translucent cytoplasm free of inclusions and a size-appropriate
polar body (Pb) [1,2]. However, metaphase II (MII)-stage oocytes retrieved from patients after ovarian stimula-
tion are known to show significant morphological variations that may affect the developmental competence and
implantation potential of the derived embryo.
More than half of the oocytes collected can contain at least one morphological abnormality, and this abnor-
mality may be correlated with the asynchrony between nuclear and cytoplasmic maturation of the MII oocyte,
playing an important role in its viability and in the clinical outcome. Morphological variations of the oocytes
may also result from other intrinsic factors, such as age and genetic defects, or extrinsic factors, such as stimula-
tion protocols, culture conditions, and nutrition. Conflicting results have been published regarding the effect of
morphological variations of the oocyte on embryo development and implantation.
This chapter reviews the correlation of morphological abnormalities of the MII oocyte and the clinical out-
come, the effect on genetic disorders, the predictive value of specific abnormalities, and whether any of these
parameters can be used in all the scoring systems applied in in vitro fertilization (IVF) laboratories.
Morphological abnormalities of the oocyte are discussed under two different subgroups: cytoplasmic abnor-
malities and extracytoplasmic abnormalities [1–4].
Cytoplasmic Abnormalities
Cytoplasmic abnormalities of MII oocytes include different types and degrees of cytoplasmic granulations
(slightly diffused or excessive whole/centrally located granulation) and appearance of refractile bodies, smooth
endoplasmic reticulum clusters (sERCs), or vacuolization in the ooplasm. Detection of cytoplasmic variations
was first termed as cytoplasmic dysmorphism by Van Blerkom and Henry in 1992 [5] and since then has been
used as a selection method to assess the viability and implantation potential of the derived embryos.
Morphological Appearance of the Cytoplasm

Although the appearance of cytoplasm was considered a potential predictive factor for the success of clinical
outcome, different definitions and grouping of multiple morphological features in various published studies
make a comparative analysis difficult. The terms that were used in the literature are dark cytoplasm [6–8], dark
cytoplasm–granular cytoplasm [9], dark cytoplasm with slight granulation [10], dark granular appearance of the
cytoplasm [11], and diffused cytoplasmic granularity [12]. The high subjectivity of the definition of these types
of granulations in various laboratories provides limited predictive value for the clinical outcome. Despite the fact
81
FIGURE 5.1 Normal mature oocyte. FIGURE 5.2 Oocyte with dense cytoplasmic granularity.
that the majority of the clinical trials examining the effect of dark granular cytoplasm as an individual feature
showed no detrimental effect on the viability of the derived embryo [6,9–11], and even may be associated with
higher fertilization when compared with the group of oocytes with total absence of granularity [13], contro-
versial results were also published [7,8,12]. Variations published in these articles may be correlated with the
subjectivity of the definition, because diffused slight cytoplasmic granularity, also called as dark cytoplasm, can
be ill defined and could differ by different modulation of the optical path in phase contrast microscopies in vari-
ous laboratories. Homogeneity of the cytoplasm is expected (Figure 5.1); however, the biological significance of
nonhomogeneity is unknown, and based on current evidence, it may represent only variability between oocytes
rather than a dysmorphism of developmental significance [1].
Centrally Located Granulation of the Cytoplasm

Condensed granulation that is centrally located within the cytoplasm with a clear border (Figure 5.2) is unlike
various degrees of diffused granulation described above because it is easily distinguishable with a significantly
darker appearance than normal cytoplasm that could be clearly visible by any modulation type of the optical
path in different phase contrast microscopies. It had first been defined by Serhal et al. [14] with a detrimental
effect on the outcome of intracytoplasmic sperm injection (ICSI), and was later named centrally located granular
cytoplasm (CLCG) by Kahraman et al. [15]. It was indicated that CLCG is a rare morphological feature of the
oocyte that is diagnosed as a large, dark, spongy, granular area in the cytoplasm and that the severity is based on
both the diameter of granular area and the depth of the lesion.
Even though fertilization rate and embryo quality were not affected in the study by Kahraman et al. [15], poor
ongoing pregnancy chance correlated with high aneuploidy (52.3%), and abortion rates (54.5%) were obtained by
the transfer of embryos derived from oocytes with severe CLCG. A study by Meriano et al. [16] defined CLCG
oocytes as organelle clustering and determined that this certain abnormality is the only repetitive dysmorphism
in consecutive cycles and is a negative predictor of pregnancy and implantation rates in intracytoplasmic injection
cycles. Even though more research is needed to define the subcellular and molecular mechanisms of organelle
clustering, hypoxia of the follicle was shown to be correlated with oocytes of such poor developmental compe-
tence [17]. It has also been demonstrated that MII oocytes that exhibited severe cytoplasmic disorganization had a
lower intracytoplasmic pH and ATP content as well as increased incidence of aneuploidy and chromosomal scat-
tering [18,19], findings that were later confirmed by Kahraman et al. [15]. Yakin et al. [20] had also shown that
the embryos that were derived from oocytes with severe cytoplasmic abnormalities where CLCG oocytes were
included had a higher rate of aneuploidy (60.0%) compared with a group of embryos derived from oocytes with nor-
mal morphological appearance (41.9%). However, the results were not statistically different, most likely based on
Oocyte 83
the insufficient number of embryos included in the comparison. The same group demonstrated that cryopreserved
Day 3 cleavage-stage embryos derived from oocytes with severe cytoplasmic abnormalities where CLCG oocytes
consisted the majority of the experimental group had a significantly lower cryosurvival rate. Even if these embryos
survived, they had lower rates of blastocyst formation, and none of the blastocysts obtained were of good quality or
were able to successfully complete the hatching process [10]. Based on the evidence proof, it is important to inform
patients about such morphological defects of the MII oocyte, and the reduced implantation outcome and increased
risk of aneuploidy with the resultant embryos [1].
Refractile Bodies
Refractile bodies (Figures 5.3 and 5.4) are cytoplasmic inclusions that can be dark incorporations, fragments,
spots, dense granules, lipid droplets, and lipofuscin. Transmission electron microscopy studies and Schmorl
staining have shown that refractile bodies >5 μm in diameter showed the conventional morphology of lipofuscin
inclusions that consisted of a mixture of lipids and dense granule materials [21]. Lipofuscin bodies in human
oocytes can be detected throughout meiotic maturation (GV, germinal vesicle; MI, metaphase I; and MII), a situ-
ation that is different from that of other cytoplasmic abnormalities in humans, such as sERCs that appear only
in mature MII-stage oocytes.
The average diameter of a recognizable refractile body under bright-field microscopy is approximately
10 μm [22]. In the majority of published articles, cytoplasmic abnormalities are examined jointly, whereas data
on the individual predictive value of refractile bodies on clinical outcome are limited. According to some investi-
gations, cytoplasmic inclusions did not appear to affect fertilization, embryo quality, and implantation rates [5,8].
Rienzi et al. [12] also showed that refractile bodies do not detrimentally affect fertilization and normal pronu-
clear morphology rates. In contrast, Xia [23] and Otsuki et al. [21] reported decreased fertilization and embryo
development. It is most likely that controversial results might be correlated to the factors that are still unknown,
and one possible confounding factor could be the differing diameters of refractile bodies. Only one study in the
literature has examined precisely the relationship between the sizes of the refractile bodies and the developmen-
tal competence of oocytes, and this study found that lipofuscin inclusions were associated with reduced fertiliza-
tion and unfavorable blastocyst development only when their diameter is >5 μm. This study also showed that the
size of refractile bodies is not correlated with the age of the woman, or with different stimulation protocols, and
that the embryo developmental outcome was not significantly affected by stimulation regimes [21]. According
to the outcome found by Otsuki et al. [21], the aging of oocytes during inactive phases of oogenesis may not be
involved with lipofuscinogenesis; instead, the accumulation of lipofuscin may occur during the growth phase
FIGURE 5.3 Presumably telophase I oocyte with dominant FIGURE 5.4 Oocyte with central refractile body and granule
refractile body (15 μm). in the perivitelline space.
of the oocytes when dominant follicles are being recruited into the preovulatory pathway. The occurrence of
large lipofuscin bodies in normal aging may also be related to conditions of the developing ovarian follicles,
such as perifollicular blood circulation and follicular fluid composition. Other explanations may be related to
oxidative stress [24], proteolytic degradation [25], or lipid metabolism as a source of energy s upply [26]. Further
research is needed to investigate whether any of these possibilities are involved in refractile bodies that are
mainly correlated with lipofuscinogenesis in human oocytes.
Vacuoles
Vacuoles are membrane-bound cytoplasmic inclusions filled with fluid (Figure 5.5) that is virtually identical
with perivitelline space (PVS) liquid [27]. Their sizes and numbers may vary, and it is assumed that vacuoles
arise either spontaneously [27] or by fusion of preexisting vesicles derived from smooth endoplasmic reticulum
(sER), Golgi apparatus, or both [28].
The incidence of vacuoles in MII oocytes varies from 3.1% [11] to 12.4% [29]. However multiple vacuoliza-
tion is a less likely seen phenomenon, with approximately 1%–1.5% [6,7,30]. De Sutter et al. [6] reported a
severely reduced fertilization rate in vacuolated oocytes (40%) compared with oocytes without any vacuoles.
Ebner et al. [30] also reported a significantly decreased fertilization with oocytes containing single vacuoles
(51.6%) and multiple vacuoles (43.8%) compared with oocytes without vacuoles (65.3%). Rienzi et al. [12]
demonstrated significantly reduced fertilization rate for vacuolated oocytes; however, pronuclear morphology
and embryo quality were not detrimentally affected. Only one study [30] had a subgroup analysis examining
the effect of the size of the vacuoles on fertilization rates and found that the group of oocytes that fertilized
normally contained vacuoles with diameters of <9.8 μm ± 3.7 μm, a value that was significantly smaller than
the diameter of vacuoles that the unfertilized oocytes contained (17.6 μm ± 9.0 μm). A cutoff value of 14 μm
for vacuole diameter was noted, above which fertilization did not occur. This study showed that a larger vacuole
or multiple vacuoles may cause a much more detrimental effect to the oocyte than a small vacuole, because a
larger portion of the cytoskeleton (e.g., microtubules) cannot function as it is supposed to function. Van Blerkom
et al. [27] also suggested that large vacuoles might displace the MII spindle from its polar position and, in turn,
result in fertilization failure, cleavage abnormalities, an abnormal cytokinesis pattern, or various combina-
tions of these effects. Even though the presence of few small vacuoles (5 μm–10 μm in diameter) that are fluid
filled but transparent are unlikely to be of biological consequence, observation of large vacuoles >14 μm in
diameter should be noted [1]. Besides the deficiency on fertilization rates, it has also been shown that blasto-
cyst formation, good quality, and hatching blastocyst rates can significantly decrease after ICSI of vacuolated
oocytes. The percentage of aneuploid embryos can also be affected by the use of vacuolated oocytes (41.9%)
FIGURE 5.5 Vacuolized MII oocyte.

Oocyte 85
compared with embryos that are derived from oocytes with normal morphology (60.0%) [20]. Vacuolization
in MII oocytes can also decrease cryosurvival rates and subsequent embryonic development of the derived
cryopreserved embryos [10].
sER Clusters
The presence of sERCs in the cytoplasm of MII oocytes is one of the most important cytoplasmic defects and
demands careful examination (Figure 5.6). A correlation between the presence of sERCs in MII oocytes and the
clinical outcome was first published by Otsuki et al. [31]. They reported that in approximately 10% of the cycles,
cytoplasmic localization of translucent vacuoles similarly sized as pronuclei exists at the MII stage of human
oocytes after the denuding procedure for ICSI. This incidence was 5% for the study by Mateizel et al. [32],
whereas it was 7% for the study by Braga et al. [33]. However, it is most likely that the number of oocytes with
sERCs can be underestimated, because it has been shown by transmission electron microscopic analysis that
there are at least three forms of sERCs: large (18 μm); medium (10 μm–17 μm), which can be classified by light
microscopy; and small (2 μm–9 μm), which are not visible under the conditions used in clinical embryology
laboratories for examination. sERCs can easily be distinguished from fluid-filled vacuoles because they are
not separated from the rest of the ooplasmic volume by a membrane, and are seen as translucent vacuoles.
Even though the mechanism responsible for sERCs is still unknown, there are some human and animal stud-
ies assuming that it could be correlated to some functional and structural alterations of the sER during oocyte
maturation, such as an increase in the sensitivity of the IP3 receptor for Ca²+ [34], increased storage of Ca²+
that is released during oscillation [35], changes in the structure from a sheet-like form to a spherical form in
starfish oocytes [36], and distribution of the sER in mouse oocytes [37]. In human oocytes, the localization of
mobilizable Ca²+ was detected in the small vesicles beneath the plasma membrane of sER. Otsuki et al. [31]
compared the clinical outcome of patients with oocytes containing sERCs and patients with retrieved oocytes
without sERCs and examined whether any confounding parameters, such as stimulation methods and hormonal
levels, can affect the outcome. Fertilization rate and embryo quality were not detrimentally affected; however,
significantly lower clinical pregnancy and implantation and significantly higher biochemical pregnancy were
observed for sERC-positive cycles. Due to the limited number of samples tested, no significant differences
were found between the study groups when stimulation protocols were compared. However, the number of
sERC-positive oocytes obtained by the short protocol was about three times larger than that by the long pro-
tocol. Serum estradiol levels on the day of hCG administration were significantly higher in sERC-positive
cycles. This study had clearly shown that the viability of an embryo is significantly reduced with the presence
of sERCs. Even though the embryo is derived from an oocyte without any clusters, its implantation potential
FIGURE 5.6 Oocyte with an aggregation of the smooth endoplasmic reticulum.

is detrimentally affected if the oocyte is from the cohort of oocytes where at least one oocyte is with clusters.
Besides the viability of the derived embryos, the most important issue with the presence of sERCs had been the
neonatal safety based on evidence proof in the literature. In the Otsuki study [31], only one pregnancy derived
from gametes with sER defect has been reported at which the baby was diagnosed with Beckwith–Wiedemann
syndrome.
Ebner et al. [38] showed that the occurrence of sERCs is significantly related with longer duration and higher
dosage of the stimulation. Fertilization and blastocyst formation rate were significantly lower for oocytes with
sERCs compared with oocytes without sERCs. Take-home baby rate was significantly lower in the group
with sERCs and miscarriage rate was significantly higher in the same group of patients. Pregnancies in women
with affected gametes had a significantly higher incidence of obstetric problems. Birth weight of babies born in
the group with sERCs was significantly lower, and there were two unexplained neonatal deaths reported in the
group with affected gametes, whereas there were no deaths reported in the group without sERCs. Malformation
rate was similar in both groups, with one case of diaphragmatic hernia reported in the group with sERCs.
Similar findings reported by Ebner [38] and Otsuki et al. [31] support the idea that this phenomenon is the
manifestation of an intrinsic oocyte defect caused by a suboptimal ovarian stimulation [27], and perhaps as a
result of o verstimulation. Following other studies reporting multiple malformations [39] and ventricular septal
defects [39] after the transfer of embryos originating from oocytes with sERCs, it was strongly recommended
that oocytes with this feature should not be inseminated, and even that the sibling oocytes should be carefully
examined [1]. However, a study [32] in which 394 sERC-positive cycles and 6845 s ERC-negative cycles were
retrospectively analyzed led the investigators to further research because this study reported that there was no
difference in the rate of major malformations between sERC-positive cycles (5.3%) and s ERC-negative cycles
82.1%). Three new borns, from single embryo transfer with frozen-thawed embryos originating from sERC-
positive oocytes were delivered and presented no major malformations. Taking into account this controversial
data on neonatal outcome, fate of babies born from oocytes with this feature should be very c arefully evaluated
and reported.
Extracytoplasmic Abnormalities
A variety of extracytoplasmic anomalies exist that in part negatively influence fertilization (consistency and
thickness of the zona, Pb1 decay, and debris within the PVS), blastulation (thickness of zona, Pb1 fragmentation),
and pregnancy (thickness of zona, Pb1 morphology, debris in PVS). Characteristics of the ZP and the PVS are
most probably associated with the health of the developing follicle, for example, its vascularization and oxygen
content. Any disturbance during growth might severely alter oocyte morphology, resulting in a pool of gametes
with different prognoses.
Dysmorphic ZP
As a result of the mutual dependence between somatic cells (e.g., cumulus cells) and the egg, it is likely that
any disturbance negatively affecting the follicle will have a comparable impact on the oocyte itself. Among the
conceivable changes in oocyte performance, it is possible that the secretion or patterning of the ZP from the
secondary follicle onward could be altered or interrupted [40,41]. This could either result in dysmorphism that
can be seen under a light microscope (Figure 5.7) or in more subtle changes of the three-dimensional structure
of the ZP.
Definitely, the most severe form of impaired growth of the ZP is its complete absence. Normally, up to four
zona proteins [42,43] build up the three-dimensional matrix of the outer protective shell. Filaments are con-
structed of repeating zona protein (ZP) 2 and 3 units that are cross-linked by ZP1 [44], thus contributing to the
structural integrity of the ZP. Experiments in mice lacking the ZP1 gene showed that secreting only ZP2 and
ZP3 results in a thinner (Figure 5.8), more loosely organized ZP [45]. In contrast, disruption of ZP2 and ZP3 led
to the absence of the a-cellular coat, resulting in infertility [46].
Oocyte 87
FIGURE 5.7 Extremely thick zona pellucida (>30 μm) at FIGURE 5.8 Immature oocyte showing extremely thin zona
the 6 o’clock position. pellucida.
In humans, a defect in gene expression was shown to cause a failure in glycoprotein matrix, even though
the ovum itself showed intact corona cells [47]. In such rare cases, the ovum fails to fertilize in conventional
IVF. In ICSI, there is a considerable risk of exposing the gametes to mechanical stress during the denudation
process. Studies [47,48] showed that in patients with zona-free eggs, pregnancies can be achieved by simply
leaving the coronal cell layer attached, because it acts as a supporting structure keeping the oocyte in shape
during injection.
From conventional IVF, it is known that thicker zonae (e.g., >20 μm) are associated with lower fertiliza-
tion rates [49]. This has been linked to patient and stimulation parameters [50]. In ICSI, however, a thicker
zona neither interferes with subsequent fertilization nor with implantation because assisted hatching can be
applied.
The multilaminar structure of the ZP can also be analyzed quantitatively using polarized light
m icroscopy [51]. Although variation exists in the thickness of zona layers around individual eggs and
between members of a cohort, it is evident that the inner zona layer is the most dominant part of the zona
[41,51]. It has been reported that the birefringence of the inner zona is directly proportional to its thickness
[41,52,53].
Shen et al. [41] found an almost 30% higher mean light retardance in conception cycles compared with non-
conception cycles, indicating that stimulated cycles may yield oocytes of affected quality. There are two retro-
spective studies that have suggested a relationship between ZP birefringence (inner layer) and preimplantation
development. Montag et al. [53] noted a higher rate of good-quality embryos on Day 3 (but not on Day 2) in
an oocyte group with high zona birefringence (41.7%) compared with a cohort with low birefringence (24.4%).
Others [52] observed a difference in progression to the blastocyst stage. The lower the measured retardance, the
lower the blastocyst formation (Figures 5.9 and 5.10).
Discoloration
Irrespective of the actual thickness of the ZP, ovarian stimulation sometimes generates gametes showing a ZP
that appears dark or brownish under a light microscope (Figure 5.11). Mostly, the egg itself is affected. It is
reported that the presence of a discolored ZP is a common phenomenon, at 9.5%–25.7% [6,9,29,54].
That said, it is not completely clear that dark or brown zonae/oocytes occur for the same reasons. These
oocytes have been termed “brown eggs” because they were found to be dark with a thick ZP, a rather small PVS
(sometimes filled with debris), and granular cytoplasm [11]. Esfandiari et al. [11] prospectively compared the
FIGURE 5.9 Zona imaging of oocyte with optimal zona pel- FIGURE 5.10 Zona imaging of oocyte with heterogeneous
lucida and spindle with lower birefringence (3 o’clock position). zona pellucida but optimal spindle (2 o’clock position).
FIGURE 5.11 Brownish oocyte showing intact first polar body and perivitelline space granularity.
outcome of brown gametes with that of gametes of normal appearance. Although the zona in discolored eggs was
thicker than that of control gametes, in conventional IVF the fertilization rate was similar. The same was true
for the fertilization after ICSI, embryo quality, implantation rate, and clinical pregnancy rate. However, because
of the thick zonae, brown oocytes were subjected to laser-assisted hatching significantly more often than the
control group.
Shape Anomalies
Even if the thickness or color of the ZP is inconspicuous, it is not automatic that the shape of the gametes is
spherical. Indeed, there is evidence that extremely ovoid eggs exist [55]. Such gametes have been shown to be
fertilizable and may lead to the birth of a healthy baby. However, a major problem with these reports is that
the degree of the shape anomaly was not quantified, and rather more imprecise descriptions have been given
(e.g., cucumber shaped).
Oocyte 89
FIGURE 5.12 Spherically shaped oocyte with ovoid zona FIGURE 5.13 Ovoid oocyte with first polar body at 4 o’clock
pellucida. position (out of focus).
Our group successfully measured ovoid oocytes [56] and calculated a roundness index (RI, length divided
by width). Actually, two indices were determined to assess whether the whole oocyte was affected (show-
ing an ovoid ooplasm and zona) or only the ZP was of ovoid shape (with the ooplasm being perfectly round)
(Figures 5.12 and 5.13). Special care was taken to detect splitting of the innermost zona layer that might keep
ooplasm in a round shape (whereas the zona is ovoid). The latter dysmorphism was shown to be associated with
implantation failure [41].
The degree of shape anomaly was neither correlated to fertilization nor embryo quality [57], and thus was in
line with previous data [55]; interestingly, two types of cleavage pattern were observed on Day 2. Either ovoid
gametes cleaved normally like a tetrahedron (a crosswise arrangement of four cells with three blastomeres lying
side by side) or, if the ovoid zona failed to exert its shaping function, they resulted in a rather flat array of four
blastomeres. Because the abnormal pattern reduces the number of cell-to-cell contact points from six to five or
four, compaction and blastulation of the corresponding embryos may be delayed [56,58].
Two possible mechanisms may account for the occurrence of ovoid oocytes. First, mechanical stress during
oocyte puncture, denudation processes, or both could deform the egg. This unwanted occurrence would create
ovoid gametes with both ooplasm and zona being affected. In these artificially damaged gametes, a tendency
toward recovery within a day has been suggested [56]. Thus, for the majority of ovoid ova, it can be assumed that
the deformation is a preexisting anomaly generated during maturation within the follicle.
Perivitelline Space
The size of the PVS is closely related to the maturational phase of the oocyte. Whereas at the GV stage (prophase I)
the expansion of PVS is minimal, it begins to increase after the resumption of meiosis. In detail, at MI, the PVS
can clearly be detected, and after completion of maturation (MII) its full size is reached.
Several studies have noted that up to 50% of all ova show a large PVS [12,23]. In oocytes with a larger PVS
(Figure 5.14), a lower fertilization rate was observed (67%) compared with gametes with a normally sized gap
(85%) [23]. This is more or less in line with the results of an Italian group [12] who showed that a large PVS
is detrimental to fertilization and zygote morphology. Interestingly, patient parameters such as female age and
indication did not seem to influence PVS performance [23], but the ratio of estradiol to testosterone (and to pro-
gesterone) did [57].
Data from in vitro- and in vivo-matured oocytes indicate that a large PVS may be ascribed to overmature
eggs [59,60]. Such eggs have shrunk in relation to the ZP, presenting a large gap between them. A large PVS
FIGURE 5.14 Metaphase II oocyte with large perivitelline FIGURE 5.15 Oocyte with large fragmented first polar body.
space.
would also occur if a larger portion of cytoplasm is extruded together with the haploid chromosomal set during
first Pb formation. This would result in a large first Pb and a large PVS.
First Pb Morphology
For a long time, it was thought that Pb1 extrusion marks the completion of nuclear maturation ending in a MII
oocyte. But by using polarized light microscopy, it has been demonstrated that some oocytes showing a Pb1 were
actually in telophase I (Figure 5.3) and not in MII [61,62]. Otsuki et al. [63] found a chromosome aggregation
phase that occurred not only from GV breakdown to MI but also from telophase I to MII. If ICSI is performed,
although chromosomes are unaligned, it may result in failed fertilization or three pronuclear zygotes due to
abnormal chromosomal segregation.
The impact of Pb1 morphology on outcome is still discussed controversially. Although some Pb1s in humans
remain intact for >20 hr after ovulation (Figures 5.1, 5.6, and 5.14), they generally have a shorter lifetime [64].
Taking this time dependency into consideration, it might be hypothesized that Pb1 morphology provides ade-
quate information on the actual postovulatory age of the corresponding egg [60].
Ebner et al. [54] tried to focus exclusively on the status of the Pb1. Ova showing an intact Pb1 gave rise
to higher rates of implantation and pregnancy [65], probably due to an increase in blastocyst formation [66].
However, these data are still a matter of debate [12,54,66–70].
Apparently, the benefit of selecting oocytes according to the morphology of the Pb1 is somewhat reduced
with increasing time span between ovulation induction and ICSI, because studies with different schedules
could not find a relationship between constitution of the Pb1 and subsequent ICSI outcome [67,69,70]. In these
data sets, the percentage of oocytes with fragmented Pbs (Figure 5.15) was higher [23,67,69,70] than that
reported in the work of Ebner et al. [54] in which Pbs were scored 2 hr after collection. This is in line with
the finding that of all intact Pb1, 13% were already fragmented at a second inspection 3 hr later [70]. For
practical reasons and to minimize the risk of ovarian aging in vitro [33], ICSI should be finished within 42 hr
post-ICSI [71].
Data from Hungary [69] suggest that a large Pb1 is the worst case (Figure 5.16). When large Pb1s were
extruded, embryos with multinucleated blastomeres were significantly (p < 0.001) more frequent (26.7%) than
in all other Pb1 classes (~8%). It has been postulated that the extrusion of an abnormally large Pb1 is due to dis-
location of the meiotic spindle [72]. This would in part explain the observed impact on fertilization and embryo
development [12,54,67].
Oocyte 91
FIGURE 5.16 Oocyte with large first polar body.
Debris in the PVS

Sometimes, it is difficult to distinguish between heavily fragmented Pb1s and debris within the PVS (Figures 5.4
and 5.11). Two hypotheses have been proposed to explain the origin of the latter dysmorphism. One hypothesis
is derived from ultrastructural data indicating the presence of an extracellular matrix comprised of granules and
filaments in the space between oolemma and ZP because the matrix is identical to that found between cumulus
cells and the corona radiata [73,74].
The second hypothesis is based on the existence of coronal cell processes passing the ZP and reaching the
egg early in maturation. It is suggested that after withdrawal of these processes, some remnants remain within
the PVS [75].
The findings of Hassan-Ali et al. [76] support the latter theory because they found a close relationship between
the frequency of PVS granularity and maturation. In detail, they never detected debris in prophase I eggs, but
they found debris in 4% of the MI and in 34.3% of MII gametes. They were also able to show that the presence of
PVS granules was dependent on gonadotropin dose. If <30 ampoules were used to stimulate the patient, 17.4% of
the eggs were positive for this anomaly compared with 45.4% in high-dose patients (>45 ampoules). Fertilization
rate, cleavage rate, and embryo quality were found to be unaffected by the presence of coarse granules in the
PVS [76,77]; however, rates of implantation and pregnancy seem to be affected [77], because transfer of embryos
derived from PVS granule-free oocytes increased implantation rate by 5% and pregnancy rate by 21%.
Conclusions
The endpoint for evaluating morphological abnormalities of MII oocytes is to be able to correlate them with
oocyte quality and viability and in correlation increase the overall efficiency of human-assisted reproduction in
terms of clinical success and safety of offspring. High heterogeneity of the published material and subjectivity
of the evaluation of morphological deviations of MII oocytes may provide only limited take-home messages on
the predictive value of outcome parameters for successful results. A meta-analysis [78] examining the clinical
results of 40 relevant articles on previously described extracytoplasmic and cytoplasmic abnormalities showed
that there was no clear tendency to a general increase in predictive value of morphological features and that
these contradicting findings underline the importance of more intensive and coordinated research that could
lead to objective criteria with better predictive value to determine the viability of derived embryos. Rapidly
developing continuous research on new biomarkers of oocyte quality may perhaps be used in addition or as
FIGURE 5.17 Diploid giant MII oocyte showing two first FIGURE 5.18 Diploid prophase I oocyte showing two germi-
polar bodies. nal vesicles.
an alternative to morphological assessment in the future; however, current, limited, predictive value of mor-
phology should not be underestimated considering that it still remains the sole method of choice for selection
until a more effective technology can substitute for it in routine practice in various clinical IVF laboratories
worldwide. Beyond the predictive value of oocyte morphology, it must not be forgotten that information linking
dysmorphism with genetic disorders is of great value and scarce because these disorders are directly correlated
with the health of the offspring in ART applications [79]. According to a common hypothesis [5], the major-
ity of extracytoplasmic anomalies occur late in maturation because they are associated with fertilization and
developmental failure rather than with aneuploidy (e.g., giant eggs). However, evidence-based data clearly dem-
onstrate that some specific severe cytoplasmic defects are correlated with chromosomal aneuploidy and genetic
disorders as described above. It is obvious that some anomalies, for example, the so-called giant eggs [80,81]
with an almost double-sized diameter (Figures 5.17 and 5.18), show a diploid chromosomal set that contributes
to digynic triploidy. Other studies [82] analyzed embryos genetically according to their Pb classes. No correla-
tion was observed between Pb shape and genetic constitution; however, the only Pb group bearing a theoretical
risk of chromosomal disorder, considering the larger volume of ooplasm in large polar bodies (Figure 5.16),
was not analyzed.
REFERENCES
1. ALPHA Scientists in Reproductive Medicine, ESHRE Special Interest Group of Embryology. Istanbul consensus
workshop on embryo assessment: Proceedings of an expert meeting. Reprod Biomed Online 2011;22:632–646.
2. Rienzi L, Balaban B, Ebner T, Mandelbaum J. The oocyte. Hum Reprod 2012;27:i2–i21.
3. Balaban B, Urman B. Effect of oocyte morphology on embryo development and implantation. Reprod Biomed
Online 2006;12:608–615.
4. Ebner T. Is oocyte morphology prognostic of embryo developmental potential after ICSI? Reprod Biomed Online
2006;12:507–512.
5. Van Blerkom J, Henry G. Oocyte dysmorphism and aneuploidy in meiotically mature human oocytes after o varian
stimulation. Hum Reprod 1992;7:379–390.
6. De Sutter P, Dozortsev D, Qian C, et al. Oocyte morphology does not correlate with fertilization rate and embryo
quality after intracytoplasmic sperm injection. Hum Reprod 1996;11:595–597.
7. Loutradis D, Drakakis P, Kallianidis K, et al. Oocyte morphology correlated with embryo quality and pregnancy
rates after intracytoplasmic sperm injection. Fertil Steril 1999;72:240–244.
8. Ten J, Mendiola J, Vioque J, de Juan J, Bernabeu R. Donor oocyte dysmorphisms and their influence on fertiliza-
tion and embryo quality. Reprod Biomed Online 2007;14:40–48.
Oocyte 93
9. Balaban B, Urman B, Sertac A, et al. Oocyte morphology does not affect fertilization rate, embryo quality and
implantation rate after intracytoplasmic sperm injection. Reprod Biomed Online 1998;13:3431–3433.
10. Balaban B, Ata B, Isiklar A, et al. Severe cytoplasmic abnormalities of the oocyte decrease cryosurvival and
subsequent embryonic development of cryopreserved embryos. Hum Reprod 2008;23:1778–1785.
11. Esfandiari N, Burjaq H, Gotlieb L, et al. Brown oocytes: Implications for assisted reproductive technology. Fertil
Steril 2006;86:1522–1525.
12. Rienzi L, Ubaldi FM, Lacobelli M, et al. Significance of metaphase II human oocyte morphology on ICSI out-
come. Fertil Steril 2008;90:1692–1700.
13. Wilding M, Di ML, D’Andretti S, et al. An oocyte score for use in assisted reproduction. J Assist Reprod Biomed
Online 2007;24:350–358.
14. Serhal PF, Ranieri DM, Kinis A, et al. Oocyte morphology predicts outcome of intracytoplasmic sperm i njection.
Hum Reprod 1997;12:1267–1270.
15. Kahraman S, Yakin K, Donmez E, et al. Relationship between granular cytoplasm of oocytes and pregnancy
outcome following intracytoplasmic sperm injection. Hum Reprod 2000;15:2390–2393.
16. Meriano JM, Alexis J, Visram–Zaver S, et al. Tracking of oocyte dysmorphisms for ICSI patients may prove
relvant to the outcome in subsequent patient cycles. Hum Reprod 2001;16:2118–2123.
17. Van Blerkom J, Antczak M, Schrader R. The developmental potential of the human oocyte is related to the
dissolved oxygen content of follicular fluid: Association with vascular endothelial growth factor levels and
perifollicular blood flow characteristics. Hum Reprod 1997;12:1047–1055.
18. Van Blerkom J, Davis PW, Lee J. ATP content of human oocytes and developmental potential and outcome after
in vitro fertilization and embryo transfer. Hum Reprod 1995;10:1047–1055.
19. Van Blerkom J. The influence of intrinsic and extrinsic factors on the developmental potential and chromosomal
normality of the human oocyte. J Soc Gynecol Invest 1996;3:3–11.
20. Yakin K, Balaban B, Isiklar A, et al. Oocyte dysmorphism is not associated with aneuploidy in the developing
embryo. Fertil Steril 2006;88:811–816.
21. Otsuki J, Nagai Y, Chiba K. Lipofuscin bodies in human oocytes as an indicator of oocyte quality. J Assist Reprod
Genet 2007;24:263–270.
22. Veeck LL. Atlas of the Human Oocyte and Early Conceptus. Baltimore: Williams & Wilkins, 1991, pp. 121–166.
23. Xia P. Intracytoplasmic sperm injection. Correlation of oocyte grade based on polar body, perivitelline space and
cytoplasmic inclusions with fertilization rate and embryo quality. Hum Reprod 1997;12:1750–1755.
24. Marzabadi MR, Sohal RS, Brunk UT. Mechanisms of lipofuscinogenesis: Effect of the inhibition of lysosomal
proteinases and lipases under varying concentrations of ambient oxygen in cultured rat neonatal myocardial cells.
APMIS 1991;99:416–426.
25. Iyy GO, Roopsingh R, Kanai S, et al. Leupeptin causes an accumulation of lipofuscin–like substances and other
signs of aging in kidneys of young rats: Further evidence for the protease inhibitor model of aging. Ann N Y Acad
Sci 1996;786:12–23.
26. Mitchell M, Armstrong DT, Robker RL, et al. Adipokines: Implications for female fertility and obesity.
Reproduction 2005;130:583–597.
27. Van Blerkom J. Occurrence and developmental consequences of aberrant cellular organization in meiotically
mature human oocytes after exogenous ovarian hyperstimulation. J Electron Microsc Tech 1990;16:324–346.
28. El Shafie M, Sousa M, Windt ML, et al. Ultrastructure of human oocytes: A transmission electron microscopic
view. In: An Atlas of the Ultrastructure of Human Oocytes. A Guide for Assisted Reproduction. New York:
Partheon Publishing, 2000, pp. 151–171.
29. Alikani M, Palermo G, Adler A, et al. Intracytoplasmic sperm injection in dysmorphic human oocytes. Zygote
1995;3:283–288.
30. Ebner T, Moser M, Sommergruber M, et al. Occurrence and developmental consequences of vacuoles throughout
preimplantation development. Fertil Steril 2005;83:1635–1640.
31. Otsuki J, Okada A, Morimoto K, et al. The relationship between pregnancy outcome and smooth endoplasmic
reticulum clusters in MII human oocytes. Hum Reprod 2004;7:1591–1597.
32. Mateizel I, Van Landuyt L, Tournaye H, et al. Deliveries of normal healthy babies from embryos originating from
oocytes showing the presence of smooth endoplasmic reticulum clusters. Hum Reprod 2013.
33. Braga D, Setti A, Cassia R, et al. Influence of oocyte dysmorphisms on blastocyst formation. Fertil Steril
2013;100:748–754.
34. Goud PT, Goud AP, Van Oostveldt P, et al. Presence and dynamic redistribution of type I inositol
1,4,5–trisphosphate receptors in human oocytes and embryos during in-vitro maturation, fertilization and
early cleavage divisions. Mol Hum Reprod 1999;5:441–451.
35. Carroll J, Jones KT, Whittingham DG. Ca²+ release and the development of Ca²+ release mechanisms during
oocyte maturation: A prelude to fertilization. Rev Reprod 1996;1:137–143.
36. Jaffe LA, Tesaraki M. Structural changes in the endoplasmic reticulum of starfish oocytes during meiotic
maturation and fertilization. Dev Biol 1994;164:579–587.
37. Mehlmann LM, Terasaki M, Jaffe LA, et al. Reorganization of the endoplasmic reticulum during meiotic
maturation of the mouse oocyte. Dev Biol 1995;170:607–615.
38. Ebner T. Prognosis of oocytes showing aggregation of smooth endoplasmic reticulum. Reprod Biomed Online
2008;16:113–118.
39. Akarsu C, Caglar G, Vicdan K, et al. Smooth endoplasmic reticulum aggregations in all retrieved oocytes causing
recurrent multiple anomalies: Case report. Fertil Steril 2009;92:1496e1–1496e3.
40. Epifano O, Liang LF, Familari M, et al. Coordinate expression of the three zona pellucida genes during mouse
oogenesis. Development 1995;121:1947–1956.
41. Shen Y, Stalf T, Mehnert C, et al. High magnitude of light retardation by the zona pellucida is associated with
conception cycles. Hum Reprod 2005;20:1596–1606.
42. Hughes DC, Barratt CLR. Identification of the true human orthologue of the mouse ZP1 gene: Evidence for
greater complexity in the mammalian zona pellucida? Biochem Biophys Acta 1999;1447:303–306.
43. Lefiévre L, Conner SJ, Salpekar A, et al. Four zona pellucida glycoproteins are expressed in the human. Hum
Reprod 2004;19:1580–1586.
44. Wassarman PM. Zona pellucida glycoproteins. Ann Rev Biochem 1998;57:415–442.
45. Rankin T, Talbot P, Lee E, et al. Abnormal zonae pellucida in mice lacking ZP1 results in early embryonic loss.
Development 1999;126:3847–3855.
46. Liu C, Litscher ES, Mortillo S, et al. Targeted disruption of the mZP3 genes results in production of eggs lacking
the zona pellucida and infertility in female mice. Proc Natl Acad Sci USA 1996;93:5431–5436.
47. Stanger JD, Stevenson K, Lakmaker A, et al. Pregnancy following fertilization of zona-free, coronal cell intact
human ova. Hum Reprod 2001;16:164–167.
48. Ding J, Rana N, Dmowski WP. Intracytoplasmic sperm injection into zona-free human oocytes results in normal
fertilization and blastocyst development. Hum Reprod 1999;14:476–478.
49. Bertrand E, Van den Bergh M, Englert Y. Does zona pellucida thickness influence the fertilization rate? Hum
Reprod 1995;10:1189–1193.
50. Loret de Mola JR, Garside WT, Bussi J, et al. Analysis of the human zona pellucida during culture: Correlation
with diagnosis and the preovulatory hormonal environment. J Assist Reprod Genet 1997;14:332–336.
51. Pelletier C, Keefe D, Trimarchi JR. Noninvasive polarized light microscopy quantitatively distinguishes the mul-
tilaminar structure of the zona pellucida of living human eggs and embryos. Fertil Steril 2004;81:850–856.
52. Rama Raju GA, Prakash GJ, Krishna KM, Madan K. Meiotic spindle and zona pellucida characteristics as
predictors of embryonic development: A preliminary study using PolScope imaging. Reprod Biomed Online
2007;14:166–174.
53. Montag M, Schimming T, Zhou C. Validation of an automatic scoring system for prognostic qualitative zona
imaging in human oocytes. Hum Reprod 2007;22(Suppl 1):i11.
54. Ebner T, Yaman C, Moser M, et al. Prognostic value of first polar body morphology on fertilization rate and
embryo quality in intracytoplasmic sperm injection. Hum Reprod 2000;15:427–430.
55. Esfandiari N, Ryan EAJ, Gotlieb L, Casper RF. Successful pregnancy following transfer of embryos from oocytes
with abnormal zona pellucida and cytoplasm morphology. Reprod Biomed Online 2005;11:620–623.
56. Ebner T, Shebl O, Moser M, et al. Developmental fate of ovoid oocytes. Hum Reprod 2008;23:62–66.
57. Xia P, Younglai EV. Relationship between steroid concentrations in ovarian follicular fluid and oocyte
morphology in patients undergoing intracytoplasmic sperm injection (ICSI) treatment. J Reprod Fertil
2000;118:229–233.
58. Suzuki H, Togashi M, Adachi J, et al. Developmental ability of zona-free mouse embryos is influenced by cell
association at the 4-cell stage. Biol Reprod 1995;53:78–83.
59. Mikkelsen AL, Lindenberg S. Morphology of in-vitro matured oocytes: Impact on fertility potential and embryo
quality. Hum Reprod 2001;16:1714–1718.
Oocyte 95
60. Miao YL, Kikuchi K, Sun QY. Oocyte aging: Cellular and molecular changes, developmental potential and rever-
sal possibility. Hum Reprod Update 2009;15:573–585.
61. Eichenlaub–Ritter U, Shen Y, Tinneberg HR. Manipulation of the oocyte: Possible damage to the spindle appara-
tus. Reprod Biomed Online 2002;5:117–124.
62. Montag M, Schimming T, Van der Ven H. Spindle imaging in human oocytes: The impact of the meiotic cell
cycle. Reprod Biomed Online 2006;12:442–446.
63. Otsuki J, Nagai Y. A phase of chromosome aggregation during meiosis in human oocytes. Reprod Biomed Online
2007;15:191–197.
64. Ortiz ME, Lucero P, Croxatto HB. Postovulatory aging of human ova: II—Spontaneous division of the first polar
body. Gamete Res 1983;7:269–276.
65. Ebner T, Moser M, Yaman C, et al. Elective transfer of embryos selected on the basis of first polar body morphol-
ogy is associated with increased rates of implantation and pregnancy. Fertil Steril 1999;72:599–603.
66. Ebner T, Moser M, Sommergruber M, et al. First polar body morphology and blastocyst formation rate in ICSI
patients. Hum Reprod 2002;17:2415–2418.
67. De Santis L, Cino I, Rabellotti R, et al. Polar body morphology and spindle imaging as predictors of oocyte qual-
ity. Reprod Biomed Online 2005;11:36–42.
68. Younis JS, Radin O, Mirsky N, et al. First polar body and nuclear precursor body morphology is related to the
ovarian reserve of infertile women. Reprod Biomed Online 2008;16:851–858.
69. Fancsovits P, Tothne Z, Murber A, et al. Correlation between first polar body morphology and further embryo
development. Acta Biol Hung 2006;57:331–338.
70. Ciotti PM, Notarangelo L, Morselli–Labate AM, et al. First polar body morphology before ICSI is not related to
embryo quality or pregnancy rate. Hum Reprod 2004;19:2334–2339.
71. Van de Velde H, De Vos A, Joris H, et al. Effect of timing of oocyte denudation and microinjection on survival,
fertilization and embryo quality after intracytoplasmic sperm injection. Hum Reprod 1998;13:3160–3164.
72. Verlhac MH, Lefebvre C, Guillaud P, et al. Asymmetric division in mouse oocytes: With or without MOS. Curr
Biol 2000;10:1303–1306.
73. Dandekar P, Talbot P. Perivitelline space of mammalian oocytes: Extracellular matrix of unfertilized oocytes and
formation of a cortical granule envelope following fertilization. Mol Reprod Dev 1992;31:135–143.
74. Dandekar P, Aggeler J, Talbot P. Structure, distribution and composition of the extracellular matrix of human
oocytes and cumulus masses. Hum Reprod 1992;7:391–398.
75. Sathanathan H. Ultrastructure of the human egg. Hum Cell 1997;10:21–38.
76. Hassan-Ali H, Hisham-Saleh A, El-Gezeiry D, et al. Perivitelline space granularity: A sign of human menopausal
gonadotropin overdose in intracytoplasmic sperm injection. Hum Reprod 1998;13:3425–3430.
77. Farhi J, Nahum H, Weissman A, et al. Coarse granulation in the perivitelline space and IVF–ICSI outcome.
J Assist Reprod Genet 2002;19:545–549.
78. Rienzi L, Gajta G, Ubaldi F. Predictive value of oocyte morphology in human IVF: A systematic review of the
literature. Hum Reprod Update 2011;17:34–45.
79. Sa R, Cunha M, Silva J, et al. Ultrastructure of tubular smooth endoplasmic reticulum aggregates in human meta-
phase II oocytes and clinical implications. Fertil Steril 2011;96:143–149.
80. Rosenbusch B, Schneider M, Gläser B, et al. Cytogenetic analysis of giant oocytes and zygotes to assess their
relevance for the development of digynic triploidy. Hum Reprod 2002;17:2388–2393.
81. Balakier H, Bouman D, Sojecki A, et al. Morphological and cytogenetic analysis of human giant oocytes and
giant embryos. Hum Reprod 2002;17:2394–2401.
82. Verlinsky Y, Lerner S, Illkevitch N, et al. Is there any predictive value of first polar body morphology for embryo
genotype or developmental potential? Reprod Biomed Online 2003;7:336–341.
6
Morphological Selection of Gametes
and Embryos: 2PN/Zygote
Introduction
The question of the start of life is as old as life itself and nobody can really answer it.
Life is the search from nothing for something.

— Christian Morgenstern
The pronuclear (PN)-stage embryo is the first stage of life where the genetic material of both female and male
is visible as pronuclei and when fertilization is initiated. Although it is possible to judge the morphology of the
oocyte and the spermatozoa by itself, the PN stage is the first event that allows for morphology rating of the new
life by various small details that are indicative for the future of the embryo.
The basis of all is small. “Omnia rerum principia parva sunt.”
— Marcus Tullius Cicero (106–43 BCE)
There are numerous possibilities for noninvasive markers to evaluate the developmental potential of an
embryo, but studies on transcriptomics [1], proteomics [2], and metabolomics [3] are still largely experimental.
Thus, light microscopic morphological examination of the cells is still the most common routine in the labora-
tory for examining development from the oocyte to the embryo.
The challenge is that in the in vitro fertilization (IFV) program, only about 10% of the collected oocytes
have the potential to develop into an embryo that can implant [4,5]. About 50%–70% [6] of the oocytes are
aneuploid [7], and about 70% of the embryos, 44 hr after fertilization, present aneuploid blastomeres [8]. With
reference to the success or failure of an assisted reproductive technique (ART) therapy, selection based on micro-
scopic evaluation of morphology is essentially still the preferred method and essential.
After ovulation or ovum pick-up in therapy, the human oocyte has to be arrested in the second meiotic division
(metaphase-II), usually characterized by the presence of a first polar body. The entry or the injection of the sperm
into the oocyte initiates the biochemical activation of the oocyte by a sperm-specific protein called phospholi-
pase C zeta [9,10] and leads to a change in the membrane potential, a rise of the intracellular Ca2+ level [10], and
the second meiotic division. The second polar body containing the chromatids from one haploid chromosome
set is extruded, and the female pronucleus is formed (Figure 6.1). During this process, the ooplasm rotates in a
periodical way, and in parallel the sperm chromatin decondensates. The sperm cell also delivers the centriole
that has a leading role in further development and control of microtubules that are important for the symmetry of
the developing embryo [11]. These microtubules pull the haploid female pronucleus toward the male pronucleus.
Both pronuclei finally migrate to the center of the cell and align [12–14]. Microtubules also mediate the organi-
zation of a mitochondrial clustering to the center of the cell in the area around the pronuclei. This could be the
result of the different metabolic needs of the new life. The mitochondria are where ATP generation takes place,
thus they are essential for further embryo development [13,15].
97
FIGURE 6.1 Time-lapse sequence showing the course of polar body extrusion. (a) 2.3 hr post-ICSI (pI). (b) 3.6 hr pI. (c) 6.7 hr pI.
First appearance of pronuclei in the periphery. (d) 11.3 hr pI. Centralization of the pronuclei.
G1 S G2 M
Interphase Division
Time 0 hr Time 24 hr
FIGURE 6.2 Cell cycle of an eukaryotic cell is divided into four stages. (From Alberts B et al., Molekularbiologie der Zelle,
Zweite Auflage VCH Verlagsgesellschaft, Weinheim, 1990.)
The G1 phase starts approximately 2–3 hr after sperm entry, and pronuclei appear after 4–6 hr. This process
is finished 18–22 hr after sperm entry or injection. Figure 6.2 [16] illustrates this cell cycle of an eukaryotic
cell. A routine microscopic morphological examination [17–19] of the pronuclei is performed 16–18 hr after
sperm entry. The judgment is done with reference to the number, size, and the symmetry of the pronuclei.
Nuclear precursor bodies are formed in the pronuclei and constitute the nucleoli. Nucleoli are where pre-rRNA
synthesis takes place. The symmetry and synchrony of the nuclear precursor bodies in the pronuclei can be used
to evaluate the potential of the developing embryo [20–22].
2PN/Zygote 99
FIGURE 6.3 The pronuclear membrane breakdown (a) and early cleavage (b).
Anomalies in the cytoplasm of the PN-stage oocyte, such as vacuoles and refractile bodies, can negatively
influence development of the embryo [23–25]. The appearance of a peripheral cytoplasmic translucency is seen
in the majority of PN-stage oocytes and is discussed in the literature as another potential marker [14,26,27].
These criteria are most commonly used to evaluate the PN-stage oocyte.
The identical reduplication of the haploid chromosome set of each pronuclei takes place in the S phase within
6 hr after the G1 phase, followed by the onset of PN membrane breakdown and syngamy. With the fusion of the
male and female genetic material, the fertilization cascade is finished, and embryo development starts by cellular
division cycles. Figure 6.3 demonstrates this fusion and the start of the cellular division cycle.
The PN-stage judgment comprises (1) number of pronuclei; (2) size of pronuclei; (3) number, size, and distribu-
tion of nucleoli; (4) cytoplasmic halo; and (5) recommendations.
Number of Pronuclei
The occurrence of the pronuclei signals the initiation of the fertilization cascade. The number of pronuclei is an
important indicator for aneuploidy. A normal fertilized cell presents with two centrally positioned, juxtaposed
pronuclei.
A regular fertilized oocyte shows, in general, two pronuclei.
Existence and Number of Pronuclei

One day after oocyte retrieval (16–18 hr after sperm injection), it is possible to examine the pronuclei. Not all cells
exhibit two pronuclei; there is the possibility of the appearance of only one or three and more pronuclei (Figure 6.4).
The judgment of the number of pronuclei is not always simple due to the three-dimensional spherical struc-
ture, with pronuclei being located somewhere inside this structure. To determine exactly the number and posi-
tion of pronuclei, it is important to use a suitable optical mode, such as the Hoffmann modulation contrast optic,
at a sufficiently high magnification (e.g., 40× objective). A simple stereomicroscope does not give sufficient res-
olution. Sometimes, it is very important to rotate the oocyte to identify pronuclei that are positioned above one
another. Figure 6.5 illustrates this problem, where two pronuclei are positioned exactly above each other so that
it looks like one PN cell. The same problem could appear by looking at a cell with three pronuclei (Figure 6.6).
Depending on the orientation, such a cell could be judged as normal when looked at just once without turning.
Oocyte with one Oocyte with two Oocyte with three

pronucleus pronuclei pronuclei
FIGURE 6.4 Oocyte with one, two, and three pronuclei.
FIGURE 6.5 One cell seen in different positions, illustrating the difficulty in identifying the number of pronuclei. (a) This could
be a cell with one pronucleus. (b-e) Rotation reveals the presence and position of two pronuclei.
Oocytes with One Pronucleus

An oocyte exhibiting 1PN has to be judged in a different way. Time-lapse sequences can show the develop-
ment from PN appearance up to fusion. This allows identification of whether pronuclei appear in asynchrony
or whether one pronucleus disappears earlier than the other pronucleus or whether pronuclei fuse prematurely.
Also, cells that develop from the beginning with only one pronucleus can clearly be identified. Looking only
once may not enable a correct judgment, and the potential of 1PN-stage oocytes depends on the fertilization
method that determines, for example, the risk of aneuploidy.
2PN/Zygote 101
FIGURE 6.6 One cell in different positions to identify correctly the number of pronuclei. (a) This could be a cell with two
pronuclei. (b-e) The same cell rotated to identify clearly three pronuclei.
In Vitro Fertilization
Oocytes with just one pronuclei (1PN stage) that develop after insemination of the cumulus-oocyte complex are
not always aneuploid. Plachot et al. [28,29] showed that between 46% and 69% of these cells are haploid and that
between 13% and 29% are diploid. Staessen et al. [30] analyzed even more cells (80.3%) to be diploid and found
only 12.5% to be diploid. The asynchronous development of the cell can be a reason for 1PN-stage cells (early
appearance or disappearance of one pronucleus) but parthenogenetic activation also could be a reason. Another
possibility is the entry of a sperm that did not decondense or two pronuclei that fused. Balakier et al. [31] reported
after fluorescent in situ hybridization (FISH) analyses that 50% of 1PN-stage oocytes were diploid. These results
lead, in the routine work, to the question how to handle 1PN-stage oocytes after IVF when no other cells are
available. A solution is to discuss with the patient the possible risk of aneuploidy of 1PN-stage oocytes after IVF.
If the patient agrees to it, it is possible to have an embryo transfer. Staessen et al. [32] reported repeated birth
from healthy children after transfer of 1PN-stage oocytes from IVF.
Intracytoplasmic Sperm Injection

The appearance of 1PN-stage cells after intracytoplasmic sperm injection (ICSI) is different from that of cells
after IVF. Sultan et al. [33] could show that only 9.5% of these cells were diploid in comparison with 61.9% after
IVF. Literature reveals that 1PN-stage oocytes after ICSI can be diploid but that the chance is just between 5.3%
and 27.9% [30,34,35]. The risk of chromosomal anomalies (aneuploidy) in cells with just one pronucleus after
ICSI is clearly higher than after IVF, leading to the recommendation not to transfer or freeze these cells.
Use of PN-stage oocytes with just ONE pronucleus

IVF ICSI
Diploid, 13%–80.5% Diploid, 5.3%–27.9%
When no other cell is available, further culture and transfer Further culture and transfer
can be performed after informing the patient about the possible risks not recommended
No cryopreservation No cryopreservation
FIGURE 6.7 Cells with irregular fertilization or development with three and four pronuclei. (a) Oocyte with three pronuclei.
(b) Oocyte with four pronuclei.
Three or More Pronuclei in an Oocyte

The appearance of three or more pronuclei in one oocyte (Figure 6.7) usually indicates an underlying error that
has occurred during the developmental process. These cells are at high risk of being polyploid.
Three PN-stage cells have three haploid chromosome sets in comparison with the regular case with two
haploid chromosome sets. If the redundant chromosome set originates from the maternal side, it is called digyny;
if it is from the paternal side, it is called diandry. A triploid cell does normally not result in a pregnancy or
miscarriage. In case of a pregnancy, the child dies after birth [36]. In IVF, the reason for diandry can be due to
entry of two sperms into the oocyte [37]. The entry of a diploid sperm is very unlikely but possible. A possible
explanation for digyny is failure of extrusion of the second polar body, and this is the most common reason for
the presence of three pronuclei after ICSI [38].
PN-stage oocytes with three pronuclei or more after IVF or ICSI should be discarded and not be
used for transfer or cryopreservation.
PN-Stage Cells with Two Pronuclei

PN Size and Position
After the entry or the injection of the sperm into the oocyte, the fertilization cascade is initiated. Within the
next 2.5–4.5 hr, the second polar body is discharged and the PN formation starts. The sperm chromatin is very
tightly packed with protamines, and these have to be exchanged with histones that are part of the decondensation
process. At the same time, the maternal pronucleus is formed; it is slightly smaller than the paternal pronucleus.
Both pronuclei grow in size during their development and move together in the center of the cell (Figure 6.8).
The position is very important because the first cleavage furrow goes through the PN axis [39].
In most cases, variation in this sequential event shows a lower developmental potential. When one pronucleus
is clearly smaller, as shown in Figure 6.9, the corresponding cells have a poor prognosis for initiating a success-
ful pregnancy [17,40].
2PN/Zygote 103
FIGURE 6.8 Pronuclear (PN)-stage oocyte with two regular FIGURE 6.9 Pronuclear stage with uneven size of pronuclei.
pronuclei centrally positioned and of equal size (paternal pro-
nucleus slightly larger).
FIGURE 6.10 Pronuclei stage oocyte with two pronuclei not aligned in the center.
The pronuclei should be located side by side in the center of the cell. If this is not the case and if pronuclei are
not aligned, as shown in Figure 6.10, further development is usually slow and irregular.
PN stage with two pronuclei that are not of the same size or that are not aligned at the center of
the oocyte show a reduced development potential.
A reason for the low development potential could be the asymmetry in the cell. The first cleavage furrow
is generally formed in the PN plane, and if pronuclei are located toward the edge of the oocyte, this can lead
to problems and can be a potential reason for poor development [41]. In lower order animals, such as insects,
worms, and reptiles, polarity is a well-established fact. In mammals, it is discussed whether it is asymmetry
or polarity. Invertebrate and vertebrate oocytes present with a pronounced asymmetry in the position of the
Centrally Peripherally
positioned pronuclei positioned pronuclei
FIGURE 6.11 Location of the pronuclei in the oocyte: centrally or peripherally.
organelle distribution. If this is a manifestation of structural and molecular events and is invariable as well
as irreplaceable, it is called polarity. The fact that it is possible to remove a blastomere from a human embryo
shows that the polarity is not as strict as in lower order animals. Nevertheless, although in the human PN-stage
asymmetry occurs, nonalignment of the pronuclei or asymmetry of the nucleolar precursor bodies is not a
good sign.
However, it is very important to remember that PN scoring and PN appearance and disappearance is a
time-dependent development and only time-lapse imaging can really see the dynamic structure of the develop-
ment [42]. This dynamic can be the reason for the different opinions in the literature when pronuclei are located
in the periphery of the cell, as demonstrated in Figure 6.11. Ebner et al. [27,43] showed a lower developmental
potential for these cells. The relatively rare phenomenon of peripheral pronuclei could be a reason why it is
not often found in the literature. Garello et al. [44] reported that this phenomenon is associated with failure or
retarded embryo development, but his observations were based on only 19 cells. The problem of peripheral pro-
nuclei is that the first cleavage furrow goes through the PN axis [39], and when the PN are peripherally located,
this can result in an abnormal cleavage pattern, eventually leading to uneven two-celled embryos. Nevertheless,
we also observed live birth after transfer of embryos derived from such oocytes. The prognostic value with
regard to biological processes is not necessarily 100% correct. Still, it is important to identify the cells with the
best statistical chances for achieving a pregnancy.
Number and Distribution of Nuclear Precursor Bodies (Nucleoli)

During the development of the pronuclei, the nucleolar precursor bodies appear. These are regions where rRNA
synthesis and processing take place. They bind at repeated rDNA sequences of certain chromosomes and are
responsible for the active transcription, processing, and protein packaging of ribosomal RNAs [45]. It is possible
to determine the number and distribution of the nucleolar precursor bodies in pronuclei using Hoffmann contrast
optics at 400× magnification. The underlying symmetry gives information about the development potential of
the cell. Symmetry and balance are very important. The Alpha and ESHRE consensus paper (2012; [20,21]) has
tried to define and classify PN scoring based on nucleolar precursor bodies. The consensus paper defined a good
prognosis, when the nuclear precursor bodies show a symmetrical picture (range 1), but they mention that there
are more specialized systems, such as Z-scoring [39]. Nonsymmetrical or other arrangements, including periph-
erally sited pronuclei, are rated as range 2, and cells with pronuclei with absent or just a single nuclear precursor
body are rated abnormal.
2PN/Zygote 105
The Alpha and ESHRE paper is a consensus and reflects the diversity in this topic in the literature. In routine work,
it is often a problem that we just look once at a cell and we do not look at the development. This static observation
can be the reason why different results from the same scoring system are reported. It has to be mentioned that some
studies found no correlation between PN scoring and the developmental potential of the resulting embryo [46–51].
Nucleoli appear and disappear in a time-dependent way [52], and it is a good example how fast and symmetric
development can be. This means that nucleolar patterns do rapidly change in a highly time-dependent manner in
most oocytes during the alignment with the cleavage furrow between the two pronuclei and the gradual reduction
of the number of nucleoli [53]. This can also been seen in time-lapse monitoring [42]. With this knowledge, it is
obvious that the time of entry of the sperm into the oocyte and the maturational state itself play an important role.
Nonsymmetrical formation of nucleolar precursor bodies reflects problems in the oocyte, but it does not necessarily
mean that the cell cannot solve this problem. For the biologist, it is important to record and assess the right signals
and symmetries to be able to compare the cells of one patient to identify those that are best suited for embryo transfer.
With ICSI, we know the exact time of entry of the sperm. Early development after IVF is in most cases slower
because the sperm themselves have to find the way into the oocyte. In routine work, this is estimated to cause a
delay in the range of 2 hr.
In our opinion, the Z-scoring model proposed by Scott et al. [14,17,39] is an easy scoring system that covers
most of the other systems and variations presented in the literature [27,54–61]. According to Scott’s work, the
symmetric forms Z1 and Z2 show a high potential for initiating a pregnancy. It is very similar to the scoring
system presented in the Alpha and ESHRE consensus paper (2011) [20,21].
PN Scoring According to Scott et al. [17,62]

Z1: Both pronuclei with equal numbers of nuclear precursor bodies (nucleoli) aligned at the PN junction
Z2: Both pronuclei with equal numbers of nuclear precursor bodies (nucleoli) scattered symmetrically
Z3: Both pronuclei with inequality of numbers or alignment of nuclear precursor bodies (nucleoli)
Z4: Both pronuclei of different size or not aligned in a central position in the cell
PN Scoring According to the Alpha and ESHRE Consensus Meeting 2011

Range 1: Symmetrical Equivalent to Z1 and Z2
Range 2: Nonsymmetrical Other arrangements, including peripherally sited pronuclei
Range 3: Abnormal Pronuclei with 0 or 1 nuclear precursor body
Examples of these scoring systems are shown in Figure 6.12 [14,17,20,21,62].

The appearance and development of the nucleolar precursor bodies are time-dependent events and show how
far the cell has progressed in the cell cycle. Symmetries are important in the development of cells, and the sym-
metric view of the Z1 and Z2 scores characterizes a PN stage with high potential. Figure 6.13 shows the time-
dependent formation of the nucleoli in the pronuclei. For selection of the cells for transfer, a symmetric formation
and pronounced progression in the cell cycle warrant a positive choice.
Timing of the development of the PN stage is influenced by the culture medium and by the culture conditions.
This can be another reason for explaining the reported differences in the literature. However, for the decision of
which cell has the greatest potential, it is important to compare all cells from one patient in a given cycle to others
so as to choose the best cells. Asymmetric behavior of the cells can lead to a change in the stimulation regimes
or the external conditions in the next cycle to reach better results.
Symmetry and Polarity

In nonmammalian species, the animal and the vegetal pole can be distinguished [44,63]. In the develop-
ment of the mammalian cells, symmetries and polarization play an important role [41], but this role is
not clearly understood yet. This is also reflected in the PN score, but it looks like symmetry and polarity
are essential in allvdevelopmental steps. It starts in the oocyte and continues through fertilization events
until embryo development. Symmetry and polarity of the orientation of the polar body can play a role in
embryo quality [44]. Polar bodies that are not situated next to each other and those that display a large
angle between them are suggested to be of poorer quality [64]. A reason for this reduced quality could be
a suboptimal orientation of the pronucleus that leads to cytoplasmic turbulence or uneven cleavage and
fragmentation [22]. However, at least the first polar body is not affixed to the oolemma, hence timing can
play an important role.
Presence of the So-Called Halo

The outer coat of the oocyte, the zona pellucida, is transparent. This is the reason why one can see the pronuclei
as well as intracytoplasmic structures such as vacuoles and inclusion bodies. A cytoplasmic halo is formed when
mitochondria and cytoplasmic components are pulled by microtubules from the periphery to the more central
region [14]. Because of this pulling, the periphery gets cleared of granular structures, forming the so-called
halo [65,66], as shown in Figure 6.14.
(a)
(b)
FIGURE 6.12 Scoring systems for pronuclear (PN) stages. (a) PN stage with Rang 1 (Z1) score (symmetrical). (b) PN stage with
Rang 1 (Z2) score (symmetrical).
2PN/Zygote 107
(c)
FIGURE 6.12 (Continued) Scoring systems for pronuclear (PN) stages. (c) PN stage with Rang 2 (Z3) score (nonsymmetrical).
(d)
FIGURE 6.12 (Continued) Scoring systems for pronuclear (PN) stages. (d) PN stage with Rang 3 (abnormal, 0 or just 1 nuclear
precursor body).
The phenomenon of a subplasmalemmal zone was first mentioned by Payne et al. [67]. Different reports exist
about the importance and significance of this region in relation to the potential of the corresponding PN stage.
Most of the cells (about 67.7%–88.7%) display a halo but in a different specificity [47,65,66]. We know that it
appears because of the transport from mitochondria and cell toward the pronuclei [15], but the importance and
the meaning of symmetrical and polar halos are not yet understood [14,47]. Ebner et al. [65] evaluated the halo
as a positive sign, whereas Salumets et al. [47] found no influence and Zollner et al. [61] differentiated between
normal halo as positive and concentric or extreme halos as negative. Our own study [68,69] found no influence
by just looking to see whether there is a halo or not, but it could be that the halo also appears and disappears and
it is just a momentum picture in the development of the cell. This could be the reason why the question toward
the potential of the halo cannot be simply answered as yes or no. Time-lapse imaging should enable to gain more
information in this regard.
2PN/Zygote 109
(e)
FIGURE 6.12 (Continued) Scoring systems for pronuclear (PN) stages. (e) PN stage with (Z4) score. PN of different sizes are
not aligned in a central position in the cell. (Modified from Scott L, Smith S, Hum Reprod 13, 1003–1013, 1998; Scott L et al., Hum
Reprod 15, 2394–2403, 2003; Scott L et al., Hum Reprod 22; 230–240, 2007; Alpha Scientists in Reproductive Medicine, ESHRE
Special Interest Group of Embryology, Reprod Biomed Online 22, 2011, 632–646; Alpha Scientists in Reproductive Medicine,
ESHRE Special Interest Group of Embryology, Hum Reprod 2011, 1–14.)
(a) (b) (c) (d)
FIGURE 6.13 Time-dependent progression of the orientation of the nucleoli in the pronuclei of the pronuclear stage from (a) dis-
persed to (b) moving towards centre to ( c) alignment at side of pronuclear contact to (d) fusion of nucleoli and further alignment.
FIGURE 6.14 Occurrence of the halo effect.
REFERENCES
1. Devreker F. Uptake and Release of Glucose, Pyruvate and Lactate. In: ESHRE Campus Non-Invasive Embryo
Quality Assessment. ed. ESHRE, Grimbergen, Belgium. 2005, pp. 77–91.
2. Brewis IA. Proteomics in reproductive research: The potential importance of proteomics to research in reproduc-
tion. Hum Reprod 1999; 14: 2927–2929.
3. Kere J, Hofmann B. Comprehensive molecular analysis of oocytes and embryos. Hum Reprod 2011; 26
(Suppl 1): i56.
4. Borini A, Lagalla C, Sciajno R, Distratis V, Bonu MA, Cattoli M, et al. Artificial reproductive technology achieve-
ments for optimizing embryo quality. Ann N Y Acad Sci 2004; 1034: 252–261.
5. Coticchio G, Sereni E, Serrao L, Mazzone S, Iadarola I, Borini A.What criteria for the definition of oocyte qual-
ity? Ann N Y Acad Sci. 2004;1034:132–144.
6. Geraedts J, Montag M, Magli C, Repping S, Handyside A, Staessen C, et al. Polar body array CGH for prediction
of the status of the corresponding oocyte. Part I: Clinical results. Hum Reprod 2011; 26: 3173–3180.
7. Van Blerkom J. Epigenetic influences on oocyte development competence: Perifollicular vascularity and intrafol-
licular oxygen. J Assist Reprod Genet 1998; 15: 226–234.
8. Ziebe S, Lundin K, Loft A, Bergh C, Nyboe Andersen A, Selleskog U, et al. FISH analysis for chromosomes
13, 16, 18, 21, 22, X and Y in all blastomeres of IVF pre-embryos from 144 randomly selected donated human
oocytes and impact on pre-embryo morphology. Hum Reprod 2003; 18: 2575–2581.
9. Saunders CM. PLC zeta: A sperm specific trigger of Ca2+ oscillations in eggs and embryo development.
Development 2002; 129: 3533–3544.
10. Heytens E, Parrington J, Coward K, Young C, Lambrecht S, Yoon S-Y, et al. Reduced amounts and abnormal
forms of phospholipase C zeta (PLCζ) in spermatozoa from infertile men. Hum Reprod 2009; 24: 2417–2428.
11. Palermo G, Munné S, Cohen J. The human zygote inherits its mitotic potential from the male gamete. Fertil Steril
1994; 9: 1220–1225.
12. Ebner T, Moser M, Tews G. Die Bedeutung der Morphologie der Oozyte hinsichtlich ihres weiteren
Entwicklungspotentials [The significance of oocyte morphology regarding it’s further developmental potential].
J Reprod Endocrinol 2006; 1: 17–23.
13. Van Blerkom J, Davis P, Merriam J, Sinclair J. Nuclear and cytoplasmic dynamics of sperm penetration, pronu-
clear formation and microtubule organization during fertilization and early preimplantation development in the
human. Hum Reprod Update 1995; 1: 429–461.
14. Scott L, Smith S. The successful use of pronuclear embryo transfers the day following oocyte retrieval. Hum
Reprod 1998; 13: 1003–1013.
15. Van Blerkom J, Davis P, Alexander S. Differential mitochondrial distribution in human pronuclear embryos leads
to disproportionate inheritance between blastomeres: Relationship to microtubular organization, ATP content and
competence. Hum Reprod 2000; 15: 2621–2633.
2PN/Zygote 111
16. Alberts B, Bray D, Lewis J, Raff M, Roberts K, Watson J. Molekularbiologie der Zelle [Molecular Biology
of the Cell. Weinheim: 2nd edition, VCH Verlagsgesellschaft, 1990]. Weinheim: Zweite Auflage VCH
Verlagsgesellschaft, 1990.
17. Scott L, Alvero R, Leondiris M, Miler B. The morphology of human pronuclear embryos is positively related to
blastocyst development and implantation. Hum Reprod 2000; 15: 2394–2403.
18. Hill GA, Freeman M, Bastias MC, Rogers BJ, Herbert CM, Osteen KG, et al. The influence of oocyte maturity
and embryo quality on pregnancy rate in a program for in vitro fertilization-embryo transfer. Fertil Steril 1989;
52: 801–806.
19. Steer C, Mills C, Tan S, Campbell S, Edwards R. The cumulative embryo score: A predictive embryo scoring
technique to select the optimal number of embryos to transfer in an in-vitro fertilization and embryo transfer
programme. Hum Reprod 1992; 7: 117–119.
20. Alpha Scientists in Reproductive Medicine, ESHRE Special Interest Group Embryology. Istanbul con-
sensus workshop on embryo assessment: Proceedings of an expert meeting. Reprod Biomed Online 2011;
22: 632–646.
21. Alpha Scientists in Reproductive Medicine, ESHRE Special Interest Group of Embryology. The Istanbul consen-
sus workshop on embryo assessment: Proceedings of an expert meeting. Hum Reprod 2011; 26: 1270–1283.
22. Papale L, Fiorentino A, Montag M, Tomasi G. The zygote. Hum Reprod 2012; 27 (Suppl 1): i22–i4.
23. Kahraman S, Yain K, Dönmez E, Samli H, Bahce M, Cengiz G, et al. Relationship between granular cytoplasm of
oocytes and pregnancy outcome following intracytoplasmic sperm injection. Hum Reprod 2000; 15: 2390–2393.
24. Ebner T, Moser M, Sommergruber M, Gaiswinkler U, Shebl O, Jesacher K, et al. Occurrence and developmental
consequences of vacuoles throughout preimplantation development. Fertil Steril 2005; 83: 1635–1640.
25. Balaban B, Urman B. Effect of oocyte morphology on embryo development and implantation. Reprod Biomed
Online 2006; 12: 608–615.
26. Sadler T. Medizinische Embryologie. New York: Georg Thieme Verlag Stuttgart, 1998, pp. 12–17.
27. Ebner T, Moser M, Sommergruber M, Tews G. Selection based on morphological assessment of oocytes
and embryos at different stages of preimplantation development: A review. Hum Reprod Update 2003;
9 (3): 251–262.
28. Plachot M, Mandelbaum J, Junca A-M, de Grouchy J, Salat-Baroux J, Cohen J. Cytogenetic analysis and devel-
opmental capacity of normal and abnormal embryos after IVF. Hum Reprod 1989; 4 (Suppl 8): 99–103.
29. Plachot M, Crozet N. Fertilization abnormalities in human in vitro fertilization. Hum Reprod 1992; 7 (Suppl 1):
89–94.
30. Staessen C, Van Steirteghem AC. The chromosomal constitution of embryos developing from abnormally fertil-
ized oocytes after intracytoplasmic sperm injection and conventional in-vitro fertilization. Hum Reprod 1997; 12:
321–327.
31. Balakier H, Squire J, Casper RF. Characterization of abnormal one pronuclear human oocytes by morphology,
cytogenetics and in-situ hybridization. Hum Reprod 1993; 8: 402–408.
32. Staessen C, Janssenswillen C, Van den Abbeel E, Devroey P, Van Steirteghem AC. Avoidance of triplet pregnan-
cies by elective transfer of two good quality embryos. Hum Reprod 1993; 8: 1650–1653.
33. Sultan KM, Munné S, Palermo GD, Alikani M, Cohen J. Chromosomal status of uni-pronuclear human zygotes
following in-vitro fertilization and intracytoplasmic sperm injection. Hum Reprod 1995; 10: 132–136.
34. Staessen C, Janssenswillen C, Devroey P, Van Steirteghem AC. Cytogenetic and morphological observations of
single pronucleated human oocytes after in-vitro fertilization. Hum Reprod 1993; 8: 221–223.
35. Plachot M, Selva J, Wolf JP, Bastit P, de Mouzon J. Consequences of oocyte dysmorphy on the fertilization rate
and embryo development after intracytoplasmic sperm injection. A prospective multicenter study. Gynecol Obstet
Fertil 2002; 10: 772–779.
36. Baraitser M, Winter RM. Missbildungs Syndrome. Bern: Hans Huber, 2001, pp. 10–11.
37. Rosenbusch B, Schneider M, Sterzik K. The chromosomal constitution of multipronuclear zygotes resulting from
in-vitro fertilization. Hum Reprod 1997; 12: 2257–2262.
38. Grossmann M, Calafell JM, Brandy N, Vanrell JA, Rubio C, Pellicer A, et al. Origin of tripronucleate zygotes
after intracytoplasmic sperm injection. Hum Reprod 1997; 12: 2762–2765.
39. Scott L. Pronuclear scoring as a predictor of embryo development. Reprod Biomed Online 2003; 6: 201–214.
40. Scott L. Morphological correlates of oocyte and embryo competence-identification. Hum Fertility 2002; 5: 206–214.
41. Scott L. Oocyte and embryo polarity. Semin Reprod Med 2001; 18: 171–183.
42. Montag M, Liebenthron J, Köster M. Which morphological scoring system is relevant in human embryo develop-
ment? Placenta 2011; 32: S252–S256.
43. Beuchat A, Thévenaz P, Unser M, Ebner T, Senn A, Urner F, et al. Quantitative morphometrical characterization
of human pronuclear zygotes. Hum Reprod 2008; 23 (9): 1983–1992.
44. Garello C, Baker H, Rai J, Montgomery S, Wilson P, Kennedy CR, et al. Pronuclear orientation, polar body place-
ment, and embryo quality after intracytoplasmic sperm injection and in-vitro fertilization: Further evidence for
polarity in human oocytes. Hum Reprod 1999; 14: 2588–2595.
45. Tesarik J, Kopecny V. Assembly of the nucleolar precursor bodies in human male pronuclei is correlated with
early RNA synthetic activity. Exp Cell Res 1990; 191: 153–156.
46. Payne JF, Raburn DJ, Couchman GM, Price TM, Jamison MG, Walmer DK. Relationship between pre-embryo
pronuclear morphology (zygote score) and standard day 2 or 3 embryo morphology with regard to assisted repro-
ductive technique outcomes. Fertil Steril 2005; 84: 900–909.
47. Salumets A, Hyden-Granskog C, Suikkari A, Tiitinen A, Tuuri T. The predictive value of pronuclear morphology
of zygotes in the assessment of human embryo quality. Hum Reprod 2001; 16: 2177–2181.
48. Weitzmann VN, Schnee-Riesz J, Benadiva C, Nulsen J, Siano L, Maier D. Predictive value of embryo grading for
embryos with known outcomes. Fertil Steril 2010; 93: 658–662.
49. James A, Hennessy S, Reggio B, Wiemer K, Larsen F, Cohen J. The limited importance of pronuclear scoring of
human zygotes. Hum Reprod 2006; 21: 1599–1604.
50. Nicoli A, Valli B, Di Girolamo R, Di Tommaso B, Gallinelli A, La Sala GB. Limited importance of pre-embryo
pronuclear morphology (zygote score) in assisted reproduction outcome in the absence of embryo cryopreserva-
tion. Fertil Steril 2007; 88: 1167–1173.
51. Brezinova J, Oborna I, Svobodova M, Fingerova H. Evaluation of day one embryo quality and IVF-outcome –
a comparison of two scoring systems. Reprod Biol Endocrinol 2009; 7: 9.
52. Tesarik J, Kopecny V. Development of human male pronucleus: Ultrastructure and timing. Gamete Res 1989; 24:
135–149.
53. Plachot M. Fertilization. Hum Reprod 2000; 15 (Suppl 4): 19–30.
54. Balaban B, Urman B, Isiklar A, Alatas C, Aksoy S, Mercan R, et al. The effect of pronuclear morphology on
embryo quality parameters and blastocyst transfer outcome. Hum Reprod 2001; 16: 2357–2361.
55. Montag M, van der Ven H, German Pronuclear Morphology Study Group. Evaluation of pronuclear morphology
as the only selection criterion for further embryo culture and transfer: Results of a prospective multicentre study.
Hum Reprod 2001; 11: 2384–2389.
56. Nagy Z, Dozortsev D, Diamond M, Rienz L, Ubaldi F, Abdelmassih R, et al. Pronuclear morphology evaluation
with subsequent evaluation of embryo morphology significantly increases implantation rates. Fertil Steril 2003;
80: 67–74.
57. Senn A, Urner F, Chanson A, Primi MP, Wirthner DW, Germont M. Morphological scoring of human pronuclear
zygotes for prediction of pregnancy outcome. Hum Reprod 2006; 21: 234–239.
58. Tesarik J, Greco E. The probability of abnormal preimplantation development can be predicted by a single static
observation on pronuclear stage morphology. Hum Reprod 1999; 14: 1318–1323.
59. Tesarik J, Junca A, Hazout A, Aubriot F, Nathan C, Cohen-Bacrie P, et al. Embryos with high implantation poten-
tial after intracytoplasmic sperm injection can be recognized by a simple, non-invasive examination of pronuclear
morphology. Hum Reprod 2000; 15: 1396–1399.
60. Wittmer C, Bettahar-Lebuge K, Ohl J, Rongieres C, Nisand I, Gerlinger P. Zygote evaluation: An efficient tool for
embryo selection. Hum Reprod 2000; 15: 2591–2597.
61. Zollner U, Zollner KP, Hartl G, Dietl J, Steck T. The use of a detailed zygote score after IVF/ICSI to obtain good
quality blastocysts: The German experience. Hum Reprod 2002; 17: 1327–1333.
62. Scott L, Finn A, O’Leary T, McLellan S, Hill J. Morphologic parameters of early cleavage-stage embryos that
correlate with fetal development and delivery: Prospective and applied data for increased pregnancy rates. Hum
Reprod 2007; 22 (1): 230–240.
63. Edwards RG, Beard HK. Oocyte polarity and cell determination in early mammalian embryos. Mol Hum Reprod
1997; 3: 863–905.
64. Gianaroli L, Magli MC, Ferraretti AP, Fortini D, Grieco N. Pronuclear morphology and chromosomal abnormali-
ties as scoring criteria for embryo selection. Fertil Steril 2003; 80: 341–349.
2PN/Zygote 113
65. Ebner T, Moser M, Sommergruber M, Gaiswinkler U, Wiesinger R, Puchner M, et al. Presence, but not type
or degree of extension, of a cytoplasmic halo has a significant influence on preimplantation development and
implantation behaviour. Hum Reprod 2003; 18: 2406–2412.
66. Stalf T, Herrero J, Mehnert C, Manolopoulos K, Lenhard A, Gips H. Influence of polarization effects and pronu-
clei on embryo quality and implantation in an IVF program. J Assist Reprod Genet 2002; 19: 355–362.
67. Payne D, Flaherty SP, Barry MF, Mathews CD. Preliminary observations on polar body extrusion and pronuclear
form score as a prognostic tool in IVF treatment. Hum Reprod 1997; 12: 705–708.
68. Greuner M. Untersuchung morphologischer Kriterien der Gameten und Vorkernstadien zur Erzielung einer
Schwangerschaft [Investigation of morphological criteria of gametes and PN-stage oocytes for achieving
a pregnancy]. Dissertation. Fakultät Charite, Universitätsmedizin Berlin, 2008.
69. Greuner M, Winkler S, Maxrath B, Montag M, Schmiady H. Charakterisierung des morphologischen
Entwicklungspotenzials von der Oozyte bis zum Embryo [Characterization of the developmental potential from
the oocyte to the embryo based on morphology assessment]. J Reproduktionmed Endokrinol 2012; 1: 13–19.
7
Morphological Selection of Gametes and Embryos: Embryo
Introduction
Fertilization occurs in vivo in the ampullary-isthmic region of the fallopian tube. In the subsequent days, the
zygote undergoes successive cleavage divisions while traversing the fallopian tube, until 2–3 days after fertiliza-
tion, and the embryo enters the uterus at the morula-blastocyst stage of development [1]. Until this century when
quality commercial in vitro fertilization (IVF) media designed specifically to meet the needs of the growing
embryo were introduced into IVF practice, embryos were traditionally transferred to the patient’s uterus at the
cleavage stage of development on Day 2 or Day 3 of in vitro culture at the four- to eight-cell stage of development.
Cleavage-stage embryos range from the two-cell stage to the morula stage when the embryo consists of more
than eight cells and each blastomere is so closely juxtaposed so as to give the appearance of no individual inter-
nal cellular boundaries as a result of the formation of tight junctions between individual blastomeres. This light
microscopic appearance at the morula stage is known as compaction. During cleavage, the embryo does not
increase in size but undergoes successive reduction in blastomere size as cell numbers increase with each cleav-
age division. For the first 2 days of development, the embryo is under the control of stored maternal message
inherited from the oocyte and it is not until the four- to eight-cell transition that the embryo is under the control
of an activated embryonic genome [2]. Many embryos undergo developmental arrest between the four- to eight-
cell stages at the time the embryonic genome becomes activated, and this embryonic block can be exacerbated
by poor in vitro culture conditions.
It is imperative that critical morphological selection parameters are used to improve the chances
of a successful pregnancy outcome.
Transferring embryos at the cleavage stage of development has the distinct disadvantage that most of the original
zygotes undergo cleavage and are theoretically available for selection for transfer. It is therefore imperative that
critical morphological selection parameters are used to improve the chances of a successful pregnancy outcome. A
distinct advantage of transferring embryos at the cleavage stage of development, particularly on Day 2 at the four-
cell stage, is that cryopreservation is very successful with pregnancy outcomes after transfer of thawed embryos
equivalent to those achieved after transfer of fresh embryos, provided only good morphological quality embryos are
selected for cryopreservation and there is no loss of blastomeres during the cryopreservation-thawing process [3].
Morphological criteria remain the gold standard for selection of embryos for transfer, and they have acquired
greater significance as the global trend moves toward a single embryo transfer policy. Many morphological
criteria and scoring systems have been proposed for the selection of the “best” cleavage-stage embryo for trans-
fer. Embryos are scored based on cleavage kinetics, cell numbers and size of blastomeres, spatial orientation
of blastomeres, pattern and extent of fragmentation, nuclear status, cytoplasmic anomalies, and in the case of
morula-stage embryos the degree of compaction [4–25].
This chapter illustrates the various morphological characteristics of the two-cell to morula-stage embryo that
have been used to select the best embryo for transfer and then links the characteristics to implantation potential,
where known.
115
Cleavage Kinetics and Cell Numbers

It was recognized early in the history of IVF that the kinetics of cleavage, that is, the number of cells at a specific
point in time postinsemination, was linked to implantation outcome [4]. Faster cleaving embryos were more
likely to implant than slower cleaving embryos, with the exception of embryos showing very rapid cleavage [4].
Rapidly dividing embryos can result from tripronucleate zygotes that cleave from one to three cells and then to
six cells within two cleavage divisions [26]. Furthermore, it has been demonstrated that embryos showing normal
cleavage kinetics are more likely to be euploid than slower cleaving embryos and embryos showing very rapid
cleavage kinetics [27–29].
The time of the first cleavage division to the two-cell stage has been linked to subsequent embryo quality
and implantation rates [30–38]; however, some have claimed that this relationship holds true only for embryos
generated in gonadotropin-releasing hormone (GnRH) agonist cycles and not for GnRH antagonist cycles
[39,40].
It has recently been agreed that the timing for the observation of optimal cleavage rates that have been linked to
the highest implantation potential should be two cells at 26 ± 1 hr postinjection or 28 ± 1 hr postinsemination
(Figure 7.1a), four cells at 44 ± 1 hr (Figure 7.1b), eight cells at 68 ± 1 hr (Figure 7.1c), and morula at 92 ± 2 hr
(Figure 7.1d) [24]. Transfer of four-cell embryos on Day 2 results in higher pregnancy rates than transfer of
embryos with lower or higher cell numbers on Day 2 [22,41,42].
FIGURE 7.1 Examples of embryos showing normal cleavage kinetics and normal cytokinesis on Day 1 (a), Day 2 (b), Day 3 (c), and
Day 4 (d). Original magnification, 300×.
Embryo 117
Similarly, transfer of embryos with eight cells on Day 3 is associated with a higher pregnancy rate [12,43],
and the transfer of slow-cleaving embryos on Day 3, with five or fewer cells, has been associated with early
pregnancy loss [44]. It must be remembered that this information is based on nearly four decades of single light
microscopic observations of embryo development on Days 1–4. The introduction of continuous observation
through time-lapse microscopy may further refine our knowledge in this area in the coming decade, particularly
the optimal timing between successive mitotic divisions.
Transfer of embryos on Day 3 with eight cells is associated with a higher pregnancy rate, and trans-
fer of slow-cleaving embryos with five or fewer cells has been associated with early pregnancy loss.
Size of Blastomeres
Cell number is related to implantation outcome; so is cell size. Regular cytokinesis resulting in blastomeres
of even size has been related to higher implantation rates when observed as early as the first cleavage divi-
sion [45,46]. Subsequent cleavage divisions should also produce daughter cells of equivalent size, and poorer
outcomes are associated with embryos whose daughter cells vary in diameter by more than one-third [6].
Regularity in blastomere size on Day 2 has been shown to correlate with higher implantation rates [8,9,22,47].
Irregular cytokinesis may also be associated with irregular nucleokinesis because multinucleation and
chromosomal abnormalities have been observed at a higher frequency in embryos with blastomeres of uneven
size [47,48].
Time-lapse imaging has demonstrated that embryos showing synchronous division on Day 2 with a time
difference between the second and third mitosis (three cell to four cell) of no greater than 1 hr have a greater
potential to develop to blastocyst and implant [49,50].
Irregular cytokinesis must be distinguished from asynchronous cleavage divisions, both of which result in
embryos with blastomeres of uneven size. Embryos that undergo synchronous division should result in two
cells, four cells, and eight cells after the first, second, and third cleavage divisions (or the first, third, and sev-
enth mitosis), respectively (Figure 7.1). If there has also been regular cytokinesis, then all blastomeres in these
embryos should be expected to be of the same size.
Regular cytokinesis results in blastomeres of even size that are related to higher implantation
rates when observed as early as the first cleavage division.
Embryos that undergo asynchronous divisions result in three cells, fives cells, six cells, and seven cells on
Day 2 and Day 3 (Figure 7.2). These embryos would therefore be expected to have a difference in cell size result-
ing from the cell that failed to undergo synchronous cleavage. A diagrammatic representation of normal and
abnormal cell sizes for embryos that have undergone synchronous division, that is, two cell, four cell, and eight
cell, are depicted in Figure 7.3, and asynchronous division, that is, three cell, five cell, six cell, and seven cell,
are depicted in Figure 7.4.
Spatial Orientation of Blastomeres

There is polarization in the distribution of organelles, proteins, and molecular message within the oocyte [51–53].
Therefore, mitotic division results in differential inheritance of organelles and molecules in the developing
embryo. This spatial arrangement of organelles and molecules can be further regulated by the execution of tan-
gential cleavage planes in the second cleavage division (Figure 7.5) [54]. The first cleavage plane in mammalian
embryos is meridional with the axis of division running from the position of the polar bodies (the animal pole)
to the opposite pole (the vegetal pole), resulting in two equal-sized daughter cells with each inheriting similar
FIGURE 7.2 Examples of embryos showing asynchronous division but normal cytokinesis. (a) Three-cell embryo, one large
blastomere and two smaller blastomeres. (b) Five-cell embryo, three large blastomeres and two smaller blastomeres. (c) Six-cell
embryo, two large blastomeres and four evenly sized smaller blastomeres. (d) Seven-cell embryo, one large blastomere and six
evenly sized smaller blastomeres. Original magnification, 300×.
distributions of animal and vegetal cytoplasm. The second cleavage plane is meridional, like the first cleavage
plane, but the third cleavage plane is equatorial [54]. The resultant four-cell embryo has a tetrahedral appearance
with a differential in distribution of animal and vegetal cytoplasm (Figure 7.5) [55].
Disturbances in these defined cleavage planes result in embryos with a nontetrahedral appearance, such as the
clover-shaped embryo (Figure 7.6), and the resultant disturbance in distribution of animal and vegetal cytoplasm
would theoretically have consequences on subsequent viability. Approximately 3% of embryos show a nontetra-
hedral appearance at the four-cell stage, and these embryos have decreased development to the blastocyst stage
and a reduced implantation rate compared with embryos arising from a tetrahedral arrangement of cells at the
four-cell stage [56].
Pattern and Extent of Fragmentation

Embryo fragmentation has long been included along with blastomere size and number as an important morpho-
logical criterion linked to embryo viability. Fragments are defined as anuclear, membrane-bound cytoplasmic
structures found in the extracellular spaces between blastomeres or between the blastomeres and the zona pel-
lucida in the perivitelline space. The extent of fragmentation is most often expressed as a percentage of the total
volume of the embryo. In very fragmented embryo, it is often difficult to assess large anucleate fragments from
Embryo 119
Synchronous cleavage, Synchronous cleavage,

regular cytokinesis irregular cytokinesis
(a) (b)
(c) (d) (e)
(f) (g)
FIGURE 7.3 Diagrammatic representation of embryos on Days 1–3 that have undergone synchronous division with regular cytoki-
nesis or irregular cytokinesis. (a) Two-cell embryo showing regular cytokinesis. (b) Two-cell embryo showing irregular cytokinesis.
(c) Four-cell embryo showing regular cytokinesis. (d) Four-cell embryo showing irregular cytokinesis in one of the two originating
blastomeres. (e) Four-cell embryo showing irregular cytokinesis in both of the originating blastomeres. (f) Eight-cell embryo show-
ing regular cytokinesis. (g) Eight-cell embryo showing irregular cytokinesis in at least one of the originating blastomeres.
blastomeres, and it has been suggested that for Day 2 embryo fragments are defined as those <45 μm in diameter
and for Day 3 embryos fragments are defined as those <40 μm in diameter [57], based on the observation that
cytoplasmic structures of these sizes do not contain DNA. In addition, it must be remembered that when perform-
ing isolated assessments, the degree of fragmentation may appear to decrease from one observation to the next.
This is because fragments or cytoplasmic blebbing often forms around the time of cleavage, and these cytoplas-
mic blebs or fragments are often transient structures that disappear by resorption or lysis with time [58,59].
Implantation and pregnancy rates have been reported to be low when embryos with a high degree of fragmen-
tation are transferred on Day 2 [8,9] or Day 3 [43], particularly when the volume of fragmentation exceeds 25%
of the total embryo volume. This observation would equate to the observation that thawed Day 2 embryos have
a similar implantation potential to fresh embryos unless they suffer blastomere loss, that is, one cell of a four-
cell embryo is equivalent to a 25% volumetric loss [3]. Regular embryos with a minor degree of fragmentation
(<10%) however have similar implantation potential to embryos with no fragmentation [9,10]. The consensus
document published by ALPHA Scientists in Reproductive Medicine and ESHRE Special Interest Group of
Embryology suggests that fragmentation should be classified as mild (<10%; Figure 7.7a), moderate (10%–25%;
Figure 7.7b), and severe (>25%; Figure 7.7c) [24].
(a) (b) (c) (d)
(e) (f) (g) (h)
(i) (j) (k) (l)
FIGURE 7.4 Diagrammatic representation of embryos on Days 2 and 3 that have undergone asynchronous division with regular
cytokinesis or irregular cytokinesis. (a) Three-cell embryo showing regular cytokinesis in one of the two originating blastomeres.
(b) Three-cell embryo showing irregular cytokinesis in one of the two originating blastomeres. (c) Three-cell embryo showing three
equally sized blastomeres that is sometimes the result of the first mitotic division of a tripronucleate zygote. (d) Five-cell embryo
showing regular cytokinesis in one of the four originating blastomeres. (e) Five-cell embryo showing irregular cytokinesis in one
of the originating blastomeres. (f) Five-cell embryo showing irregular cytokinesis resulting in blastomeres of even size. (g) Six-cell
embryo showing regular cytokinesis in two of the four originating blastomeres. (h) Six-cell embryo showing irregular cytokinesis
in two of the four originating blastomeres. (i) Six-cell embryo showing irregular cytokinesis resulting in six blastomeres of even
size. (j) Seven-cell embryo showing regular cytokinesis in three of the four originating blastomeres. (k) Seven-cell embryo showing
irregular cytokinesis in one of the four originating blastomeres. (l) Seven-cell embryo showing irregular cytokinesis resulting in
seven blastomeres of even size.
When the volume of fragmentation exceeds 25% of the total embryo volume, implantation and
pregnancy rates have been reported to be low.
The reduction in implantation and pregnancy potential observed after the transfer of early cleavage stage
embryos with a high degree of fragmentation (>25%) is likely to be due to the finding that these embryos, when
cultured to the blastocyst stage, have a reduction in cell numbers in both the trophectoderm and inner cell mass.
In contrast, embryos with mild-to-moderate fragmentation have a reduction in cell numbers in the trophectoderm
only [60]. In addition, the rate of aneuploidy has been observed to increase with the degree of fragmentation [61–63].
Embryo 121
(a) First mitotic division—meridional (b) Second mitotic division—meridional (c) Third mitotic division—equatorial
FIGURE 7.5 Diagrammatic representation of the first, second, and third mitotic divisions. (a) The first mitotic division is meridi-
onal; (b) the second mitotic division in one of the two originating blastomeres is meridional; and (c) the third mitotic division in
the remaining blastomere is equatorial, resulting in a four-cell embryo with mixed inheritance patterns of organelles, proteins, and
molecular message.
FIGURE 7.6 Examples of four-cell embryos showing different spatial distribution of blastomeres. (a) A four-cell embryo with
tetrahedral arrangement of blastomeres expected as a result of the meridional and equatorial divisions. (b) A four-cell embryo show-
ing a planar arrangement of blastomeres resulting in a clover-shaped embryo that may be reflective of disturbances in the normal
cleavage patterns. Original magnification, 300×.
It has been suggested that the pattern of fragmentation is just as important as the extent of fragmentation
to viability [10,53]. When the extent of fragments is similar but the fragments are scattered throughout the
embryo rather than localized in one place in the perivitelline space, the implantation rate is compromised [10].
It has also been observed that there is a tendency toward a higher proportion of chromosomal errors in embryos
with scattered fragments compared with embryos with localized fragments [62]. However, because time-lapse
observations have shown that the position of fragments is dynamic, with fragments moving around and some-
times disappearing within the embryo, the consensus document published by ALPHA Scientists in Reproductive
FIGURE 7.7 Examples of four-cell embryos showing different degrees of fragmentation. (a) Four-cell embryo with mild
fragmentation, that is, <10% fragments. (b) Four-cell embryo with moderate fragmentation, that is, 10%–25% fragments. (c) Four-
cell embryo with severe fragmentation, that is, >25% fragments. Original magnification, 300×.
Medicine and ESHRE Special Interest Group of Embryology concluded that no weight should be given to the
spatial positioning of fragments in morphology scores [24].
Nuclear Status
It is not uncommon to observe a nucleus in one or more blastomeres during development, and this nucleation is
dependent on the stage of the cell cycle at the time of observation, with the nucleus being visible only during inter-
phase (Figure 7.8a). Because embryos are usually observed at a single discrete time point on any day of develop-
ment, the incidence of multinucleation is probably underreported. In around one-third of all embryos, more than
one nucleus can be observed within a single blastomere (Figure 7.8b) [64]. Multinucleation (two or more nuclei per
blastomere) can be observed in embryos on Days 1–3; however, it is often more difficult to observe the nuclear status
in embryos on Day 3 due to the smaller cell sizes [64]. The nuclei in a multinucleated blastomere may be of normal
size (Figure 7.8b) or much smaller (micronuclei) (Figure 7.8c). Multinucleation is believed to arise through karyoki-
nesis in the absence of cytokinesis; partial fragmentation of nuclei; or errors in chromosome segregation, packaging
at mitosis, or both [65,66]. As a result, multinucleated embryos are associated with a high degree of chromosomal
errors [47,62,63,66].
Embryo quality has been reported to be linked with nuclear status, with four-cell embryos on Day 2 and eight-
cell embryos on Day 3 having a lower incidence of multinucleation than other cell numbers on these days [64].
Uneven cleavage has also been reported to be associated with a higher rate of multinucleation [47]. Transfer of
FIGURE 7.8 Examples of two-cell embryos showing different nucleation status. (a) Two-cell embryo showing a single nucleus in
each blastomere (red arrows). (b) Two-cell embryo showing two nuclei in one blastomere (red arrows). (c) Two-cell embryo showing
micronuclei in one blastomere (red circle delineates the micronuclei). Original magnification, 300×.
Embryo 123
embryos with multinucleated blastomeres has been shown to result in lower implantation and pregnancy rates
[47,64,67–70] and higher abortion rates [42].
It is generally agreed that embryos that have been identified, at some point in their development, to contain
a multinucleated blastomere should not be selected for transfer if an alternative embryo is available [24].
However, it has been demonstrated that it is possible for binucleated cells on Day 1 to cleave to normal cells on
Day 2 [66], and live births have been achieved, if less frequently, from the transfer of multinucleated embryos.
Cytoplasmic Anomalies
The cytoplasm of embryos is normally homogeneous in appearance and is normally pale and clear or slightly
granular in appearance except perhaps around the time of cleavage, both before and immediately after, when
significant organelle movement within the cytoplasm may result in a more granular appearance that may or may
not be h omogeneous at the time of observation. Several dysmorphisms have been identified in the cytoplasm
of oocytes and cleavage-stage embryos and include cytoplasmic pitting, vacuoles, refractile bodies, plaques of
smooth endoplasmic reticulum (sER), and the severe clustering of organelles that results in large areas, usually
subcortical, of clear or “halo” cytoplasm [71]. With the possible exception of plaques of sER and severe cluster-
ing or centralization of organelles, there has been very little literature published on the impact of cytoplasmic
dysmorphism on implantation outcome.
Cytoplasmic pitting is characterized by numerous tiny pits in the cytoplasm, resulting in a very granular
appearance that may be associated with organelle reorganization, as mentioned above, before or immediately
after cleavage. Studies have failed to identify any association between cytoplasmic pitting and implantation
potential [14,72], but one study reported an increased incidence of early loss of gestational sacs after transfer of
embryos showing cytoplasmic pitting [73].
Vacuoles are membrane-bound cytoplasmic inclusions filled with fluid [74]. Vacuoles can vary in size, num-
ber, and time of appearance during embryo development (see Chapter 5) [75]. Three different types of vacuoles
can be identified: vacuoles arising within the oocyte, vacuoles artificially created during the intracytoplasmic
sperm injection procedure, and vacuoles arising de novo during development [75]. Vacuolation de novo can
occur throughout development but peaks at Day 4 of development when it is associated with developmental
arrest [75]. It was observed that the later the vacuoles arose in the cytoplasm of the developing embryo, the
greater the detrimental impact on blastocyst formation [75]. It has also been agreed that small vacuoles (<10 μm
in diameter) have little consequence to development but that large vacuoles (>14 μm in diameter) can interfere
with cleavage planes and result in reduction in development to the blastocyst stage [24].
Refractile bodies observed under bright-field microscopy are refractile due to the composition of lipid material
and dense granules [76]. The lipid material has been demonstrated to be lipofuscin [77]. The presence of large
refractile bodies (>5 μm in diameter) not only has an impact on fertilization but also results in significantly lower
blastocyst developmental rates [77].
Plaques of sER can occasionally be identified in the cytoplasm of oocytes [78] and early cleavage-stage
embryos [79]. Although similar in appearance, the localized accumulation of sER can be clearly differentiated
from fluid-filled vacuoles at the light microscope level because they are translucent (see Chapter 5). The plaques
can vary in size and can be as large as a pronucleus in diameter [78]. It has been reported that plaques of sER can
perturb calcium signaling events and can have significant developmental consequences [24]. Once an oocyte that
contains sER plaques is fertilized, there appears to be no impact on d evelopment to Day 4, but a significant reduc-
tion in the numbers of embryos developing to the blastocyst stage is noted [80]. Pregnancy outcomes after transfer
of embryos from patients who produced oocytes with sER plaques are associated with a lower chance of success-
ful pregnancy [78,80] regardless of whether the transferred embryo was derived from an oocyte containing an
sER plaque. In addition, reports of fetal abnormalities [81] and in one case the birth of a child with the Beckwith–
Wiedemann imprinting disorder [78] have been associated with patients who produce oocytes with sER plaques.
Occasionally, severe central clustering of organelles can be observed in one or more blastomeres of a cleavage-
stage embryo, leaving a clear subcortical region or halo. It has been suggested that these blastomeres are destined
to degenerate and that the resultant embryo has a reduced implantation potential [13,24].
Compaction
Up to Day 3 of development, each blastomere is a discrete entity, clearly distinguishable from the next blasto-
mere. Between Day 3 and Day 4, tight junctions begin to form between neighboring blastomeres, and it becomes
more difficult to distinguish one cell from the next. This process is known as compaction, and a completely
compacted embryo is known as a morula. An ideal morula on Day 4 of development should contain between
16 and 32 cells, that is, it has undergone the fourth cleavage division or both the fourth and the fifth cleavage
division [18] (Figures 7.1d and 7.9a). However, embryos can be observed on Day 4 that have initiated but not
completed the compaction process, that is, partially compacted embryos (Figure 7.9b) or have failed to undergo
the compaction process (Figure 7.9c).
The compaction process should begin on Day 3 at the eight-cell stage; however, compaction may be observed
earlier in embryos with fewer than eight cells and this does not appear to have a negative impact on implanta-
tion potential [14]. A delay in the compaction process resulting in failure of, or partial compaction, on Day 4
may reflect a slight retardation in developmental kinetics, and given more time, these embryos may progress to
complete compaction [18]. Alternatively, failure of, or partial compaction, in some embryos may be indicative of
blastomeres with limited to no developmental potential, resulting in their exclusion from the compaction process
and ultimately the formation of small compacted embryos with limited ability to form an inner cell mass of any
appreciable size and viability when development progresses [18].
FIGURE 7.9 Examples of Day 4 embryos. (a) Fully compacted morula. (b) Partially compacted embryo. (c) Twelve-cell embryo
showing little sign of compaction. (d) Cavitating morula. Original magnification, 300×.
Embryo 125
Regardless of the reason for the incomplete or failed compaction, it is evident that this morphological f eature is
reflective of a reduction in viability that is most significant when compaction is completely absent. The consensus
is that if more than half of the embryo is not involved in the compaction process by 92 ± 2 hr, these embryos
have a reduced implantation potential [18,24]. Partially compacted embryos on Day 4 (at least half of the embryo
is involved in the compaction process) have a similar implantation potential to fully compacted embryos, pro-
vided the embryo has entered the fourth cleavage division by Day 4 [23]; this finding concurs with our own
observations (Vaxevanoglou, unpublished data).
Incomplete (more than half of the embryo is not involved in the compaction process by 92 ± 2 hr)
or completely absent compaction is reflective of reduced viability, resulting in a reduced
implantation potential.
Occasionally, advanced developmental kinetics can be observed on Day 4, with early signs of cavitation within the
morula, the first step toward blastocyst formation (Figure 7.9d) [15,23,82]. These early cavitating embryos are associ-
ated with a very high implantation potential. Advanced developmental kinetics on Day 4 seems to confer an advan-
tage to viability rather than being detrimental, as has been reported for early cleavage-stage embryos [8,9] where
advanced developmental kinetics is more often associated with an increase in chromosomal a bnormalities [27].
Conclusions
Morphology remains the gold standard for selecting the most viable embryo for transfer, and this is true for all
stages of development. Genetic analysis of a Day 3 embryo or blastocyst-stage embryo can provide the addi-
tional advantage of selecting a chromosomally normal embryo, but biopsy for genetic analysis is not possible for
embryos before Day 3.
Embryologists have reached consensus on the morphologic criteria that have the greatest predictive value in
discriminating between viable and nonviable embryos on Days 2–4 (Tables 7.1 and 7.2) [24]:
1. The kinetics of development is probably the single most important predictor of normal develop-
ment; therefore, it is essential that observations of embryo morphology are performed at fixed times
as described in the consensus document. It is likely that in the next decade, continuous time-lapse
observation of embryo morphology that is now available within incubators will reveal the ideal time
intervals between successive cleavage divisions and further refine the predictive value of cleavage
kinetics.
TABLE 7.1
Consensus Scoring System for Day 2 and Day 3 Embryos in Addition to
the Ideal Cell Numbers at 44 ± 1 hr and 68 ± 1 hr Postinsemination
Grade Rating Description
1 Good • <10% fragmentation
• Stage-specific cell size
• No multinucleation
2 Fair • 10%–25% fragmentation
• Stage-specific cell size for majority of cells
• No multinucleation
3 Poor • Severe fragmentation (>25%)
• Cell sizes are not stage specific
• Evidence of multinucleation
Source: ALPHA Scientists in Reproductive Medicine, ESHRE Special Interest
Group of Embryology, Hum Reprod 26, 1270–1283, 2011.
TABLE 7.2
Consensus Scoring System for Day 4 Embryos
Grade Rating Description
1 Good • Entered into fourth round of cleavage, i.e., >8 cells
• Evidence of compaction that involves virtually all the embryo
volume
2 Fair • Entered into fourth round of cleavage, i.e., >8 cells
• Compaction involves the majority of the embryo volume
3 Poor • Disproportionate compaction involving less than half of
the embryo with more than half of the embryo remaining as
discrete blastomeres
Source: ALPHA Scientists in Reproductive Medicine, ESHRE Special Interest Group
of Embryology, Hum Reprod 26, 1270–1283, 2011.
2. Blastomere size should be stage appropriate, that is, reflecting normal cytokinesis.
3. Fragmentation volume but not pattern was agreed to affect viability, particularly when it equated to
approximately 25% or more of the embryo volume.
4. Multinucleation is best observed on Day 2, most likely due to the association of a high incidence of
chromosomal error with multinucleation.
5. Day 4 embryos should have entered the fourth round of cleavage and should be compacted or compacting.
6. It must be remembered, however, that despite decades of linking embryo morphology to pregnancy
potential, the predictive value of any one morphologic criterion remains low. The predictive value for
pregnancy potential can be improved by combining several criteria and can be further improved by
combining the observations over successive days of development (see Chapters 5, 6, and 8). However,
morphological criteria improve the chances of selecting the best embryo within a cohort for transfer,
but the chances of pregnancy remains a statistical probability rather than a certainty.
REFERENCES
1. Croxatto HB, Diaz S, Fuentaealba B, et al. Studies on the duration of egg transport in the human oviduct. I.
The time interval between ovulation and egg recovery from the uterus in normal women. Fertil Steril 1972; 23:
447–458.
2. Braude P, Bolton V, Moore S. Human gene expression first occurs between the four- and eight-cell stages of
preimplantation development. Nature 1988; 332: 459–461.
3. Edgar DE, Bourne H, Speirs AL, et al. A quantitative analysis of the impact of cryopreservation on the implanta-
tion potential of human early cleavage stage embryos. Hum Reprod 2000; 15: 175–179.
4. Cummins JM, Breen TM, Harrison KL, et al. A formula for scoring human embryo growth rates in in vitro fer-
tilization: Its value in predicting pregnancy and in comparison with visual estimates of embryo quality. J In Vitro
Fert Embryo Transf 1986; 3: 284–295.
5. Claman P, Armant DR, Seibel MM, et al. The impact of embryo quality and quantity on implantation and the
establishment of viable pregnancies. J In Vitro Fert Embryo Transf 1987; 4: 218–222.
6. Puissant F, Van Rysselberge M, Barlow P, et al. Embryo scoring as a prognostic tool in IVF treatment. Hum
Reprod 1987; 2: 705–708.
7. Steer CV, Mills CL, Tan SL, et al. The cumulative embryo score: A predictive embryo scoring technique to select
the optimal number of embryos to transfer in an in-vitro fertilization and embryo transfer programme. Hum
Reprod 1992; 7: 117–119.
8. Giorgetti C, Terriou P, Auquier P, et al. Embryo score to predict implantation after in-vitro fertilization: Based on
957 single embryo transfers. Hum Reprod 1995; 10: 2427–2431.
9. Ziebe S, Petersen K, Lindenberg S, et al. Embryo morphology or cleavage stage: How to select the best embryos
for transfer after in vitro fertilization. Hum Reprod 1997; 12: 1545–1549.
Embryo 127
10. Alikani M, Cohen J, Tomkin G, et al. Human embryo fragmentation in vitro and its implications for pregnancy
and implantation. Fertil Steril 1999; 71: 836–842.
11. Gerris J, De Neubourg D, Mangelschots K, et al. Prevention of twin pregnancy after in vitro fertilization or intra-
cytoplasmic sperm injection based on strict embryo criteria: A prospective randomized clinical trial. Hum Reprod
1999; 14: 2581–2587.
12. Van Royen E, Mangelschots K, De Neubourg D, et al. Characterization of a top quality embryo, a step towards
single-embryo transfer. Hum Reprod 1999; 14: 2345–2349.
13. Veeck LL. Preembryo grading and degree of cytoplasmic fragmentation. In: An Atlas of Human Gametes and
Conceptuses: An Illustrated Reference for Assisted Reproductive Technology. New York: Parthenon Publishing,
1999, pp. 46–51.
14. Desai NN, Goldstein J, Rowland DY, et al. Morphological evaluation of human embryos and derivation of
an embryo quality scoring system specific for day 3 embryos: A preliminary study. Hum Reprod 2000; 15:
2190–2196.
15. Huisman GJ, Fauser BC, Eijkemans MJ, et al. Implantation rates after in vitro fertilization and transfer of a maxi-
mum of two embryos that have undergone three to five days of culture. Fertil Steril 2000; 73: 117–122.
16. Fisch JD, Rodriguez H, Ross R, et al. The Graduated Embryo Score (GES) predicts blastocyst formation and
pregnancy rate from cleavage-stage embryos. Hum Reprod 2001; 16: 1970–1975.
17. De Placido G, Wilding M, Strina I, et al. High outcome predictability after IVF using a combined score for zygote
and embryo morphology and growth rate. Hum Reprod 2002; 17: 2402–2409.
18. Tao J, Tamis R, Fink K, et al. The neglected morula/compact stage embryo transfer. Hum Reprod 2002; 17:
1513–1518.
19. Ebner T, Moser M, Sommergruber M, et al. Selection based on morphological assessment of oocytes and embryos
at different stages of preimplantation development: A review. Hum Reprod Update 2003; 9: 251–262.
20. Baczkowski T, Kurzawa R, Głabowski W. Methods of embryo scoring in in vitro fertilization. Reprod Biol 2004;
4: 5–22.
21. Rienzi L, Ubaldi F, Iacobelli M, et al. Significance of morphological attributes of the early embryo. Reprod
Biomed Online 2005; 10: 669–681.
22. Holte J, Berglund L, Milton K, et al. Construction of an evidence-based integrated morphology cleavage embryo
score for implantation potential of embryos scored and transferred on day 2 after oocyte retrieval. Hum Reprod
2007; 22: 548–557.
23. Feil D, Henshaw RC, Lane M. Day 4 embryo selection is equal to Day 5 using a new embryo scoring system
validated in single embryo transfers. Hum Reprod 2008; 23: 1505–1510.
24. ALPHA Scientists in Reproductive Medicine, ESHRE Special Interest Group of Embryology. The Istanbul con-
sensus workshop on embryo assessment: Proceedings of an expert meeting. Hum Reprod 2011; 26: 1270–1283.
25. Prados FJ, Debrock S, Lemmen JG, et al. The cleavage stage embryo. In: Atlas of Human Embryology: From
Oocytes to Preimplantation Embryos (eds. Magli MC, Jones GM, Lundin K, van den Abbeel E). Hum Reprod
2012; 27(Suppl 1): i22–i49.
26. Kola I, Trounson A, Dawson G, et al. Tripronuclear human oocytes: Altered cleavage patterns and subsequent
karyotypic analysis of embryos. Biol Reprod 1987; 37: 395–401.
27. Magli MC, Gianaroli L, Munné S, et al. Incidence of chromosomal abnormalities from a morphologically normal
cohort of embryos in poor-prognosis patients. J Assist Reprod Genet 1998; 15: 297–301.
28. Munné S. Chromosome abnormalities and their relationship to morphology and development of human embryos.
Reprod Biomed Online 2006; 12: 234–253.
29. Finn A, Scott L, O’leary T, et al. Sequential embryo scoring as a predictor of aneuploidy in poor-prognosis
patients. Reprod Biomed Online 2010; 21: 381–390.
30. Shoukir Y, Campana A, Farley T, et al. Early cleavage of in-vitro fertilized human embryos to the 2-cell stage: A
novel indicator of embryo quality and viability. Hum Reprod 1997; 12: 1531–1536.
31. Sakkas D, Shoukir Y, Chardonnens D, et al. Early cleavage of human embryos to the two-cell stage after intracy-
toplasmic sperm injection as an indicator of embryo viability. Hum Reprod 1998; 13: 182–187.
32. Bos-Mikich A, Mattos AL, Ferrari AN. Early cleavage of human embryos: An effective method for predicting
successful IVF/ICSI outcome. Hum Reprod 2001; 16: 2658–2661.
33. Lundin K, Bergh C, Hardarson T. Early embryo cleavage is a strong indicator of embryo quality in human IVF.
Hum Reprod 2001; 16: 2652–2657.
34. Fenwick J, Platteau P, Murdoch AP, et al. Time from insemination to first cleavage predicts developmental com-
petence of human preimplantation embryos in vitro. Hum Reprod 2002; 17: 407–412.
35. Salumets A, Hydén-Granskog C, Mäkinen S, et al. Early cleavage predicts the viability of human embryos in
elective single embryo transfer procedures. Hum Reprod 2003; 18: 821–825.
36. Hammoud I, Vialard F, Casasnovas P, et al. How viable are zygotes in which the PN are still intact at 25 hours?
Impact on the choice of embryo for transfer. Fertil Steril 2008; 90: 551–556.
37. Fu J, Wang XJ, Wang YW, et al. The influence of early cleavage on embryo developmental potential and IVF/ICSI
outcome. J Assist Reprod Genet 2009; 26: 437–441.
38. Lee MJ, Lee RK, Lin MH, et al. Cleavage speed and implantation potential of early-cleavage embryos in IVF or
ICSI cycles. J Assist Reprod Genet 2012; 29: 745–750.
39. Yang WJ, Hwu YM, Lee RK, et al. Early cleavage does not predict treatment outcome following the use of GnRH
antagonists in women older than 35. Fertil Steril 2007; 88: 1573–1578.
40. Yang WJ, Hwu YM, Lee RK, et al. Early-cleavage is a reliable predictor for embryo implantation in the GnRH
agonist protocols but not in the GnRH antagonist protocols. Reprod Biol Endocrinol 2009; 7: 20.
41. Thurin A, Hardarson T, Hausken J, et al. Predictors of ongoing implantation in IVF in a good prognosis group of
patients. Hum Reprod 2005; 20: 1876–1880.
42. Scott L, Finn A, O’Leary T, et al. Morphologic parameters of early cleavage-stage embryos that correlate with
fetal development and delivery: Prospective and applied data for increased pregnancy rates. Hum Reprod 2007;
22: 230–240.
43. Racowsky C, Stern JE, Gibbons WE, et al. National collection of embryo morphology data into Society for
Assisted Reproductive Technology Clinic Outcomes Reporting System: Associations among day 3 cell number,
fragmentation and blastomere asymmetry, and live birth rate. Fertil Steril 2011; 95: 1985–1989.
44. Hourvitz A, Lerner-Geva L, Elizur SE, et al. Role of embryo quality in predicting early pregnancy loss following
assisted reproductive technology. Reprod Biomed Online 2006; 13: 504–509.
45. Ciray HN, Karagenç L, Ulug U, et al. Early cleavage morphology affects the quality and implantation potential
of day 3 embryos. Fertil Steril 2006; 85: 358–365.
46. Hesters L, Prisant N, Fanchin R, et al. Impact of early cleaved zygote morphology on embryo development and
in vitro fertilization-embryo transfer outcome: A prospective study. Fertil Steril 2008; 89: 1677–1684.
47. Hardarson T, Hanson C, Sjögren A, et al. Human embryos with unevenly sized blastomeres have lower
pregnancy and implantation rates: Indications for aneuploidy and multinucleation. Hum Reprod 2001;
16: 313–318.
48. Magli MC, Gianaroli L, Ferraretti AP. Chromosomal abnormalities in embryos. Mol Cell Endocrinol 2001; 183:
29–34.
49. Wong CC, Loewke KE, Bossert NL, et al. Non-invasive imaging of human embryos before embryonic genome
activation predicts development to the blastocyst stage. Nat Biotechnol 2010; 28: 1115–1121.
50. Meseguer M, Herrero J, Tejera A, et al. The use of morphokinetics as a predictor of embryo implantation. Hum
Reprod 2011; 26: 2658–2671.
51. Antczak M, Van Blerkom J. Oocyte influence on early development: The regulatory proteins leptin and STAT 3
are polarized in mouse and human oocytes and differentially distributed within the cells of the preimplantation
stage embryo. Mol Hum Reprod 1997; 3: 1067–1086.
52. Edwards RG, Beard HK. Oocyte polarity and cell determination in early mammalian embryos. Mol Hum Reprod
1997; 3: 863–905.
53. Antczak M, Van Blerkom J. Temporal and spatial aspects of fragmentation in early human embryos: Possible
effects on developmental competence and association with the differential elimination of regulatory proteins from
polarized domains. Hum Reprod 1999; 14: 429–447.
54. Gulyas BJ. A reexamination of cleavage patterns in eutherian mammalian eggs: Rotation of blastomere pairs
during second cleavage of the rabbit. J Exp Zool 1975; 193: 235–248.
55. Edwards RG, Hansis C. Initial differentiation of blastomeres in 4-cell human embryos and its significance for
early embryogenesis and implantation. Reprod Biomed Online 2005; 11: 206–218.
56. Ebner T, Maurer M, Shebl O, et al. Planar embryos have poor prognosis in terms of blastocyst formation and
implantation. Reprod Biomed Online 2012; 25: 267–272.
57. Johansson M, Hardarson T, Lundin K. There is a cutoff limit in diameter between a blastomere and a small
anucleate fragment. J Assist Reprod Genet 2003; 20: 309–313.
Embryo 129
58. Hardarson T, Löfman C, Coull G, et al. Internalization of cellular fragments in a human embryo: Time-lapse
recordings. Reprod Biomed Online 2002; 5: 36–38.
59. Van Blerkom J, Davis P, Alexander S. A microscopic and biochemical study of fragmentation phenotypes in
stage-appropriate human embryos. Hum Reprod 2001; 16: 719–729.
60. Hardy K, Stark J, Winston RML. Maintenance of the inner cell mass in human blastocysts from fragmented
embryos. Biol Reprod 2003; 68: 1165–1169.
61. Ziebe S, Lundin K, Loft A, et al. FISH analysis for chromosomes 13, 16, 18, 21, 22, X and Y in all blastomeres
of IVF pre-embryos from 144 randomly selected donated human oocytes and impact on pre-embryo morphology.
Hum Reprod 2003; 18: 2575–2581.
62. Magli MC, Gianaroli L, Ferraretti AP, et al. Embryo morphology and development are dependent on the chromo-
somal complement. Fertil Steril 2007; 87: 534–541.
63. Munné S, Chen S, Colls P, et al. Maternal age, morphology, development and chromosome abnormalities in over
6000 cleavage-stage embryos. Reprod Biomed Online 2007; 14: 628–634.
64. Van Royen E, Mangelschots K, Vercruyssen M, et al. Multinucleation in cleavage stage embryos. Hum Reprod
2003; 18: 1062–1069.
65. Pickering SJ, Taylor A, Johnson MH, et al. An analysis of multinucleated blastomere formation in human
embryos. Hum Reprod 1995; 10: 1912–1922.
66. Staessen C, Van Steirteghem A. The genetic constitution of multinuclear blastomeres and their derivative daugh-
ter blastomeres. Hum Reprod 1998; 13: 1625–1631.
67. Jackson KV, Ginsburg ES, Hornstein MD, et al. Multinucleation in normally fertilized embryos is associ-
ated with an accelerated ovulation induction response and lower implantation and pregnancy rates in in vitro
fertilization-embryo transfer cycles. Fertil Steril 1998; 70: 60–66.
68. Pelinck MJ, De Vos M, Dekens M, et al. Embryos cultured in vitro with multinucleated blastomeres have poor
implantation potential in human in-vitro fertilization and intracytoplasmic sperm injection. Hum Reprod 1998;
13: 960–963.
69. Moriwaki T, Suganuma N, Hayakawa M, et al. Embryo evaluation by analysing blastomere nuclei. Hum Reprod
2004; 19: 152–156.
70. Saldeen P, Sundström P. Nuclear status of four-cell preembryos predicts implantation potential in in vitro
fertilization treatment cycles. Fertil Steril 2005; 84: 584–589.
71. Van Blerkom J, Henry G. Oocyte dysmorphism and aneuploidy in meiotically mature human oocytes after o varian
stimulation. Hum Reprod 1992; 7: 379–390.
72. Rienzi L, Ubaldi F, Minasi MG, et al. Blastomere cytoplasmic granularity is unrelated to developmental potential
of day 3 human embryos. J Assist Reprod Genet 2003; 20: 314–317.
73. Ebner T, Tews G, Sommergruber M, et al. Cytoplasmic pitting has a negative influence on implantation outcome.
J Assist Reprod Genet 2005; 22: 239–344.
74. Van Blerkom J. Occurrence and developmental consequences of aberrant cellular organization in mei-
otically mature human oocytes after exogenous ovarian hyperstimulation. J Electron Microsc Tech 1990;
16: 324–346.
preimplantation development. Fertil Steril 2005; 83: 1635–1640.
76. Veeck LL. Abnormal morphology of the human oocyte and conceptus. In: An Atlas of Human Gametes and
Conceptuses: An Illustrated Reference for Assisted Reproductive Technology. New York: Parthenon Publishing,
1999, pp. 121–166.
77. Otsuki J, Nagai Y, Chiba K. Lipofuscin bodies in human oocytes as an indicator of oocyte quality. J Assist Reprod
Genet 2007; 24: 263–270.
78. Otsuki J, Okada A, Morimoto K, et al. The relationship between pregnancy outcome and smooth endoplasmic
reticulum clusters in MII human oocytes. Hum Reprod 2004; 19: 1591–1597.
79. Makabe S, Naguro T, Nottola SA, et al. Ultrastructural dynamic features of in vitro fertilization in humans. Ital
J Anat Embryol 2001; 106(2 Suppl 2): 11–20.
80. Ebner T, Moser M, Shebl O, et al. Prognosis of oocytes showing aggregation of smooth endoplasmic reticulum.
81. Akarsu C, Cağlar G, Vicdan K, et al. Smooth endoplasmic reticulum aggregations in all retrieved oocytes causing
recurrent multiple anomalies: Case report. Fertil Steril 2009; 92: 1496e1–1496e3.
8
Morphological Selection of Gametes
and Embryos: Blastocyst
Thomas Ebner
Introduction
Morula or blastocyst stage marks the switch from a cell cluster of individual blastomeres to a relatively smooth
mass with indistinguishable cell outlines capable of actively regulating its internal environment. Thus, any pro-
longation of in vitro culture will allow for a more accurate prediction of developmental capacity. Other potential
advantages of blastocyst culture include synchronization between developmental stage of the embryo and the
uterine milieu, reduction of multiple pregnancy rate, or indirect selection of chromosomal anomalies.
Prolonged culture to blastocyst stage (Days 4–6 of preimplantation development) is certainly not a new
approach. Some 20 years ago, acceptable blastocyst formation rates as high as 40% were already reported [1];
however, associated pregnancy data indicate that blastocyst development does not automatically correspond
to implantation potential. Obviously, it is important to differentiate between the ability of a particular culture
system (e.g., culture medium, embryo density, type of incubator, oxygen tension) to support blastocyst formation
in vitro and the potential of the said system to give rise to a viable blastocyst [2].
Change from a cleavage stage to a blastocyst transfer program requires a proper adaption strategy.
If a clinical embryologist needs to address the problem of changing a cleavage-stage transfer program into a
blastocyst program, it is recommended to start working with supernumerary embryos first. Once a successful
Day 2 or Day 3 transfer has been performed, further culture of sibling embryos may indicate whether a given
culture system is capable of creating blastocysts at all. It should be kept in mind that the quality of the leftover
embryos should be comparable with that of embryos transferred to allow for proper prediction of culture system
quality. Thus, good-prognosis patients with sufficient embryos in abundance (e.g., more than five, to seven)
should preferably be considered for training purposes.
In cases where a regular blastocyst formation rate of 40%–50% is reached and the quality of the developed
blastocysts appears appropriate, one could proceed to transfer (and/or cryopreserve) them. However, if the oppo-
site scenario is encountered, every effort needs to be made to increase the rate of blastulation.
Old-fashioned methods such as coculture with Vero have become rather obsolete because the improved
understanding of both the physiological changes in oviduct and uterus and the different metabolic needs of the
cleavage-stage and the blastocyst-stage embryos led to the development of a new generation of sequential and
global culture media. If a change of media supplier is a nonissue, it is good to know that almost all major players
provide simple as well as complex media. However, if the decision is made to persevere with feeder cells, in situ
coculture with attached cumulus cells (as the result of an incomplete denudation process) that turned out to be
advantageous in terms of blastocyst formation [3] would be a more commonly used technique.
Apart from somatic cells, stimulatory effects may also derive from neighboring embryos. Thus, grouping
embryos in culture (in contrast to single culture) would facilitate accumulation of autotrophic factors that in turn
have been found to be beneficial for blastulation [4]. Obviously, this is a balancing act because optimal embryo
density, for example, the maximum number of embryos in a given volume of culture medium, is uncertain, as is
131
the actual negative impact of decomposition products such as ammonium. To be on the safe side for blastocyst
culture, most clinical embryologists tend to change the culture medium on Day 3, regardless of the type (global
or sequential) of medium used.
Usually, the above mentioned minor changes in culture conditions should result in acceptable survival rates
to Day 5. Only in the event of little avail would major changes of the laboratory setup need to be considered,
provided that physical premises (e.g., pH, temperature) are fulfilled. One such adaption would be the use of low-
ered incubator oxygen tension (approximately 5% instead of 20%), a tension that has been reported to support
development to blastocyst stage and live birth rate [5,6].
Low oxygen tension in the incubator has been reported to support development to blastocyst
stage and live birth rate.
Assuming that a sufficient number of blastocysts of acceptable quality will finally be available, adequate
scoring of the grown concepti is of particular importance for selecting the best candidates for transfer. To avoid
comparing apples and oranges in in vitro fertilization (IVF)-intracytoplasmic sperm injection (ICSI), timing
of fertilization and embryo development is crucial. In this respect, an international consensus was reached [7]
emphasizing that on Day 4, the first possible day of blastocyst formation, morphological scoring should be done
at 92 ± 2 hr postinsemination. Typically, on the fifth day of in vitro culture (116 ± 2 hr), preimplantation develop-
ment should culminate in the formation of the blastocyst. Once fully developed, human blastocysts consist of
two different cell types: an outer layer of trophectoderm (TE) responsible for the accumulation of fluid in the
blastocyst cavity and specialized for implantation and an inner cell mass (ICM) forming all three germ layers
of the fetus.
Scoring of Blastocysts
Establishment of blastocyst transfer programs as a matter of routine came along with two key requirements: the
need of an efficient cryopreservation program and an easily applicable blastocyst scoring system. In the early
years of blastocyst grading, particular attention was focused on developmental stage, for example, degree of
blastocyst expansion [8,9]. A more recent scoring system [10] took additional morphological parameters into
consideration, namely, morphology of the ICM and TE. To summarize, according to the degree of expansion,
blastocysts were scored using numbers in ascending order ranging from Grade 1 (Figure 8.1 [1]; blastocyst cavity
less than half of the volume of the embryo) to Grade 6 (completely hatched blastocyst). Beginning with the full
blastocyst stage (Figure 8.2; Grade 3), additional assessment of ICM and TE could be done (based on cell number
and cohesion) to predict developmental competence.
For reasons of accuracy, the more detailed Gardner approach [10] gained acceptance and allowed for reducing
the number of transferred blastocysts without limiting pregnancy rate [11]. A randomized prospective e valuation
compared the two scoring systems [12]. Although similar numbers of blastocysts were transferred in c omparable
patient cohorts, the Gardner score turned out to be superior to the Dokras score in terms of implantation
(37.6% vs. 25.0%) and multiple pregnancy (38.6% vs. 17.1%).
Blastocysts of different qualities can be grown per patient, making an optimized blastocyst
scoring system indispensable.
Blastocyst Cell Number

A factor in common to both scoring systems is the emphasis on blastocyst expansion (e.g., the size of the
blastocoel). Expansion of blastocyst on Day 5 or Day 6 ranges from retarded morula to expanded or even hatch-
ing blastocyst stage (Figure 8.3). Although most embryologists rely on visual judgment rather than m
easuring
Blastocyst 133
FIGURE 8.1 Early blastocyst with blastocoel taking up FIGURE 8.2 Full IVF blastocyst with optimal cell lineages
less than half of the volume. Fragments are extruded into the (3AA).
perivitelline space.
FIGURE 8.3 Optimal blastocyst showing hatching site close to the inner cell mass (5AA).
blastocyst diameter, Shapiro et al. [13] accurately measured blastocysts before transfer and found that the diam-
eter of a transferred blastocyst was the most significant variable in predicting clinical pregnancy.
In this context, a high variability in cell numbers has been observed. The mitotic activity is considered to be
a reliable indicator of blastocyst viability and developmental capacity [14]; however, to count the actual number
of nuclei in a blastocyst, its cells have to be fixed.
Although it has been suggested [15] that noninvasive assessment of the total cell number (TCN) is possible
under good microscopical optics, the majority of studies on TCN were performed using stained cells of spare
blastocysts donated to research (often of reduced quality) and thus probably not representing the actual cell
number of healthy blastocysts.
On Day 5, cell number of blastocysts ranged from 42 to 58 [1,16–18]. However, using sequential media, the rate
of mitosis was found to be increased, for example, from 64 to 111 cells on Day 5 [19] and, on average, 167 cells
on Day 6 [20]. It can be summarized that a full human blastocyst at Day 5 of development should exceed 60 cells
and at least have its cell number doubled on Day 6.
Predictably, any process that severely reduces cytoplasmic volume of the embryo will cause a dramatic loss of
cells at the blastocyst stage, if at all this stage is reached. Indeed, one report [21] describes that blastomere loss
after cryopreservation resulted in significantly lower blastocyst cell numbers on Day 6 (n = 45) compared with
blastocysts derived from fully intact cleavage-stage embryos after thawing (n = 58). Extensive fragmentation at
earlier stages showed the same detrimental effect on TCN on Day 6 [22], for example, a significant decrease in
cell count from 69 (embryos without fragmentation) to 29 (>25% fragmentation). Interestingly, for minimal and
moderate levels of fragmentation, the reduction in cell number was largely confined to the TE, whereas a steady
number of ICM cells were maintained. This finding suggests a homeostatic regulation of the lineage that gives
rise to the fetus [22].
Cell Lineages
Hardy et al. [1] realized certain differences in the growth rate of both cell types. In general, mitotic rate of the
TE is approximately 1.5 times higher than that of the ICM; however, compared with some other mammals, the
overall proportion of the ICM is approximately 33% of all cells on Day 5 and 51% on Day 6 [1]. The striking
peak on Day 6, with half of all cells in the blastocyst being part of the ICM, may be the result of increase in ICM
growth rate between Day 5 and Day 6, a time when the number of TE cells is more or less unchanged. Because
there is widespread cell death of even morphologically normal cells in both cell l ineages [1], it is s uspected that
the preservation of cell number within cell lineages is regulated by apoptotic phenomena [23].
Inner Cell Mass

The health of a blastocyst is not only strongly dependent on the overall cell number [24] but also on the proper
formation of both cell lineages. According to the scoring system of Gardner and Schoolcraft [10], the embryo-
blast is considered to be optimal (Grade A) if the ICM shows a tight package of numerous cells (Figures 8.2
through 8.4). Any reduction in number and contact affected the quality of this cell lineage; thus, loosely grouped
cell accumulations are scored Grade B (Figures 8.5 through 8.7), whereas absence (Figures 8.8 through 8.10)
or presence of only few cells randomly distributed within the cavity of the blastocyst are classified as Grade C.
Quantitative grading of the ICM emphasized the importance of its size and shape [25]. In contrast to blastocyst
expansion and TE cell number, ICM size was significantly related to implantation. Blastocysts showing an ICM
<3800 μm2 showed lower implantation rates (18%) compared with blastocysts with an ICM >4500 μm2 (45%).
FIGURE 8.4 Full blastocyst with optimal inner cell mass and FIGURE 8.5 Suboptimal full blastocyst (3BC) developed
impaired trophectoderm (3AB). after conventional IVF. Trophectoderm is not cohesive between
6 o’clock and 2 o’clock positions. Inner cell mass consists of
few cells only.
Blastocyst 135
FIGURE 8.6 Full blastocyst with optimal inner cell mass but FIGURE 8.7 Expanded blastocyst with both cell lineages
bad quality TE (3AC). consisting of few cells (4BB).
FIGURE 8.8 Hatching (5 o’clock position) blastocyst without FIGURE 8.9 Ovoid full blastocyst with blastomeres extruded
inner cell mass and with reduced trophectoderm (5CB). into perivitelline space (11 o’clock position) and blastocoel (3CB).
It has to be clarified that these studies [25] measured blastocysts of different sizes, for example, ranging from
full to expanded stage. Recent data indicate that the size of the embryoblast is closely related to the degree of
expansion [26]. This seems to be associated with a more peripheral location of the cell mass within the blas-
tocyst cavity as the blastocyst expands, an increased cohesion within ICM cells, or both. The latter is further
supported by the observation that at full blastocyst stage, the number of ICM cells can still be estimated, while
at expanded stage, the number cannot be identified accurately. Taking these data into consideration, it was not
surprising that Richter et al. [25] noticed a reduction in ICM size of Day 6 blastocysts compared with Day 5
blastocysts.
In addition, the above mentioned study [25] evaluated the possible influence of ICM shape on o utcome by
introducing the roundness index (RI), an index that represents the length-to-width proportion of the

ICM (Figure 8.11). Extreme RI values of <1.04 (almost round) and >1.20 (too oval) had a worse prognosis
(implantation rates of 7% and 33%, respectively) compared with those with intermediate RI. Implantation rates
were highest for embryos with both optimal ICM size and shape.
FIGURE 8.10 Expanded blastocyst of poor quality (4CB). FIGURE 8.11 Expanded blastocyst of good quality (4AA)
Blastomere extruded into blastocoel should not be mixed up with ovoid inner cell mass (roundness index 2.0).
with inner cell mass.
FIGURE 8.12 Expanded IVF blastocyst (4AA) with necrotic FIGURE 8.13 Full blastocyst with both cell lineages con-
area in trophectoderm (8 o’clock position). Two cytoplasmic sisting of few cells (3BB).
processes bridge the blastocoel.
Striving to replace blastocyst with large ICM, embryologists should not forget that disproportionately o versized
ICMs, for example, with apoptotic processes not working properly [22], could cause problems in maintaining
healthy central cells because of the increased distance over which nutrients and oxygen have to diffuse or they
could contribute to large-offspring syndrome [27].
Trophectoderm
TE can be classified in the same way as ICM [10]. The outer layer is considered to be optimal if it consists of
numerous sickle-shaped cells (Figures 8.2, 8.11, and 8.12), forming a cohesive epithelium (Grade A). If number
and cohesion of these cells are reduced or shows several gaps, the TE is scored B (Figures 8.4, 8.8, 8.10, 8.13,
and 8.14). The worst-case scenario (Grade C) would be a TE consisting of very few larger cells with a low num-
ber of tight junctions (Figures 8.5 and 8.6).
Blastocyst 137
FIGURE 8.14 Expanded blastocyst of ovoid shape (4AB).
Recent studies suggest that of all blastocyst parameters, TE is the only significant independent
predictor of live birth outcome.
Kovacic et al. [28] noted a 36% implantation rate in blastocysts showing an impaired trophoblast (e.g., flattened
cells; no sickle-shaped cells; no junctions between TE cells; cells with granulation, pigmentation, vacuolization,
or a combination), and other studies supported these findings [13]. Some studies [13,28] could not find any
difference between the mean number of TE cells and the occurrence of a clinical pregnancy. One should keep
in mind that they evaluated the number of TE cells in one single focal plane only, which might be an over- or
underestimate of the actual quality of the TE. However, it can be hypothesized that any preliminary reduction in
TE cell number could easily be compensated by the accelerated growth rate from Day 6 onward [1].
This is in contrast to the recent findings [29] that suggest that a decrease in the quality of TE had an almost
linear relationship to a reduced rate of live births. In detail, all three morphological parameters had a significant
effect on live birth; however, once adjusted for known significant confounders (e.g., female age, number of good-
quality embryos, follicle-stimulating hormone [FSH] dosage), it was shown that TE was the only significant
independent predictor of live birth outcome. In another retrospective study by the same working group [30] data
from fresh single blastocyst transfers could be confirmed analyzing vitrified/warmed single blastocyst transfers.
Others reported [31] that TE morphology is significantly related to the rates of ongoing pregnancy and miscar-
riage. Interestingly, the only data prospectively collected in a large multicenter trial revealed that it is rather the
degree of expansion and hatching than the quality of both cell lineages that predicts outcome [32].
Bad-Quality Blastocysts
Only a limited number of papers analyzed the fate of bad-quality blastocysts [28,33]. This group of blastocysts with
poor prognosis is usually associated with lower cell numbers and a higher degree of chromosomal d isorders [33,34].
Bad-quality blastocysts consist of numerous different morphological subtypes, such as b lastocysts with excessive
fragmentation, excluded blastomeres, and necrotic cells, as well as trophoblast vesicles. Trophoblast vesicles are
characterized by the absence of the ICM [35] and a rudimentary TE with only a minor number of cells (Figure 8.15).
Transfer of trophoblast vesicles resulted in a disastrous live birth rate of approximately 7% [28].
The fate of all the other inferior blastocyst variations, although they may show acceptable ICMs or TEs, will also
be compromised by excluded fragments, blastomeres, or both (Figures 8.1 and 8.16) [1]. These cellular remnants are
accompanied by reduced cell numbers on Day 5 [22] and an impaired hatching process [36]. Live birth rate (approx-
imately 17%) in these bad-quality blastocysts were found to be slightly higher than that of trophoblast vesicles [28].
FIGURE 8.15 Pseudoblastocyst presumably consisting of <12 FIGURE 8.16 Early blastocyst showing exclusion of two
cells. blastomeres.
FIGURE 8.17 Transition of early to full blastocyst stage. The FIGURE 8.18 Expanded IVF blastocyst (4AA) with inner cell
expansion of the blastocoel is more than half of the volume, and mass showing a large vacuole.
intended location of the inner cell mass can be estimated.
Transfer of blastocysts showing necrotic foci in one of the cell lineages (Figure 8.17) [2,3] is a common
phenomenon (Figure 8.12). A study that assessed the actual location of such degenerative areas indicated that
outcome was worse if the ICM was affected (23% live births) and slightly better if only the TE was affected
(32.8%). Consequently, ICM compactness and multicellularity contributed more to vital implantation than TE
cohesiveness [28].
Vacuolization of blastocysts could have a detrimental effect on developmental capacity as well (Figure 8.18). It
has been reported that approximately one-third of Day 5 blastocysts showed vacuoles [37]. It appears that elimi-
nation of vacuoles from the ICM is a common scenario [37] because the majority of vacuoles could be located
in the TE [37,38], indicating that embryos might develop strategies to minimize a n egative impact of vacuoliza-
tion on implantation behavior. Reports of pregnancies achieved after transfer of vacuolated blastocysts are rare;
however, prognosis was much better if vacuolization was restricted to the TE [37].
Another important morphological characteristic is the presence of cytoplasmic extensions (Figure 8.12)
bridging the blastocyst cavity at the full or later stages (Grades 3–6). These processes are commonly present
in half of the junctional TE cells spanning the boundary between the polar and the mural region of TE and are
Blastocyst 139
directed toward the surface of the ICM [39]. These cytoplasmic strings are thought to be related to the flow of
cells from the polar to the mural TE; consequently, they tend to withdraw as the cells reach their final location.
Interestingly, variations in both shape (from broad triangles to string-like projections) and length (some fail
to reach the ICM surface) have been observed [39]. Persistence of cytoplasmic processes up to hatching stage
possibly marks blastocysts of developmental lability.
Interestingly, blastocysts of ovoid shape (Figures 8.9 and 8.14) seem to have the same genetic risk and implan-
tation potential as their round counterparts. The only striking finding is a delayed growth in these blastocysts
deriving from ovoid oocytes due to a reduced number of cell–cell contacts at four-cell stage [40].
Spontaneous Hatching
Regardless of its respective quality, every blastocyst surviving until expanding stage is forced to escape from its
zona pellucida (ZP) to not face total arrest of development. In vitro spontaneous hatching of the human embryo
is rather supported by the tremendous increase of internal pressure caused by both a gradual accumulation of
fluid and preferential proliferation of TE. The degree to which calcium-activated K+ channels are involved in the
process of hatching is currently under discussion [41].
In the absence of the uterine milieu, the hatching process usually starts with small vesicles protruding through
the ZP. It should be kept in mind that this blebbing does not necessarily indicate the precise location of subsequent
hatching [38]. Once a small opening has been created, the TE starts to herniate and, governed by TE projections,
a larger opening is created by mechanical forces. Additional help may come from blastocyst “breathing” [38],
a sequence of collapses and reexpansions considered to assist final extrusion from the r uptured ZP (Figure 8.19).
In humans, hatching occurs at various regions of the ZP. Although some studies suggest that blastocysts
show hatching sites mainly close to the ICM [35], Sathananthan et al. [36] postulated that most of the blasto-
cysts hatch from the abembryonic pole. Considering the proportions within a blastocyst, the likelihood of blas-
tocysts hatching from the smaller embryonic site is much lower than the chance of herniating near the rather
extensive mural TE. A recent study [42] supports the latter theory because out of all hatching blastocysts (Grade 5),
<40% showed a zona breach close to the embryonic pole.
It seems that human blastocysts have a developmental benefit if they hatch adjacent to the ICM, irrespective of
whether it is focused on fresh [42] or thawed blastocysts [43]. Because this area corresponds to the cells (syncy-
tiotrophoblast) that will later drive invasion into the endometrium, theoretically, hatching close to the ICM would
accelerate contact between those TE cells that are supposed to draw the b lastocyst into the uterine wall and the
endometrium. This mutual interaction between blastocyst and uterus may be impaired or delayed if herniation
takes place opposite the ICM, if hatching difficulties occur, or both.
FIGURE 8.19 Hatching (7 o’clock position) blastocyst of optimal quality (5AA) during collapsing phase (breathing).
Conclusions
Blastocyst culture as the method of choice for all patients is a balancing act and could result in inacceptable rates
of cycle cancellation. However, it should be kept in mind that prolonged culture to the blastocyst stage might be
associated with negative scenarios.
First, an almost threefold incidence of monozygotic twinning (5.6% vs. 2%) was observed in blastocyst
transfers compared with cleavage-stage transfers [44]. Others identified blastocyst culture as the major risk in
terms of monozygotic twinning, whereas embryo freezing, type of stimulation protocol used, ICSI fertilization,
or zona removal did not influence its incidence [45]. The other suggested risk with blastocyst culture is a theoreti-
cal change in methylation patterns, resulting in affected birth weights. A couple of studies found a significantly
lower birth weight in children born from blastocyst transfer compared with cleavage-stage transfer [46,47].
These results have not been unchallenged [48]. Interestingly, the opposite was indicated when the birth weights
of children were analyzed according to the length of embryo culture [49]. With increasing time in culture from
Day 2 to Day 6, an increase in the proportion of large for gestational age (LGA) babies was found.
However, one prerequisite for optimizing blastocyst culture, transfer, and cryopreservation is to develop
a better understanding of morphological criteria to adequately predict implantation potential of a certain
blastocyst.
Identifying the blastocyst with the highest implantation potential paves the way to single
embryo transfer.
Usually, several blastocysts of different qualities can be grown per patient, making an optimized blastocyst
s coring system indispensable. With the current trend toward a significant reduction of the number of embryos
considered for transfer, every effort should be made to filter out the blastocyst with the highest implantation
potential.
Thus, it is recommended to weigh the morphological criteria at the blastocyst stage as follows: quality of the
ICM, quality of the TE, expansion (e.g., blebbing out of the zona at the embryonic pole), and few anomalies
(excluded blastomeres, vacuoles, necrotic foci, cytoplasmic strings). There is increasing evidence that the impact
of the TE on live birth is at least the one of the ICM. In order to identify the actual potential of TE, multilayer
analysis is recommended. If semiautomatic grading of human blastocysts [50] is the way to proceed may be
questioned.
REFERENCES
1. Hardy K, Handyside AH, Winston RM. The human blastocyst: Cell number, death and allocation during late
preimplantation development in vitro. Development 1989; 107: 597–604.
2. Gardner DK, Lane M. Culture and selection of viable blastocysts: A feasible proposition for human IVF? Hum
Reprod Update 1997; 3: 367–382.
3. Ebner T, Moser M, Sommergruber M, et al. Incomplete denudation of oocytes prior to ICSI enhances embryo
quality and blastocyst development. Hum Reprod 2006; 21: 2972–2977.
4. Ebner T, Shebl O, Moser M, et al. Group culture of human zygotes is superior to individual culture in terms of
blastulation, implantation and life birth. Reprod Biomed Online 2010; 21: 762–768.
5. Kovacic B, Vlaisavljević V. Influence of atmospheric versus reduced oxygen concentration on development of
human blastocysts in vitro: A prospective study on sibling oocytes. Reprod Biomed Online 2008; 17: 229–236.
6. Meintjes M, Chantilis SJ, Douglas JD, et al. A controlled randomized trial evaluating the effect of lowered incuba-
tor oxygen tension on live births in a predominantly blastocyst transfer program. Hum Reprod 2009; 24: 300–307.
7. Alpha Scientists in Reproductive Medicine. Istanbul consensus workshop on embryo assessment: Proceedings of
an expert meeting. Reprod Biomed Online 2011; 32: 632–646.
8. Dokras A, Sargent IL, Ross C, et al. The human blastocyst: Its morphology and HCG secretion in vitro. Hum
Reprod 1991; 6: 1143–1151.
Blastocyst 141
9. Dokras A, Sargent IL, Barlow DH. Human blastocyst grading: An indicator of developmental potential. Hum
Reprod 1993; 8: 2119–2127.
10. Gardner DK, Schoolcraft WB. In vitro culture of human blastocysts. In: Towards Reproductive Certainity:
Infertility and Genetics Beyond 1999 (eds. Jansen R, Mortimer D). Carnforth: Parthenon Press, 1999, pp. 378–388.
11. Gardner DK, Lane M, Stevens J, et al. Blastocyst score affects implantation and pregnancy outcome: Towards a
single blastocyst transfer. Fertil Steril 2000; 73: 1155–1158.
12. Balaban B, Yakin K, Urman B. Randomized comparison of two different blastocyst grading systems. Fertil Steril
2006; 85: 559–563.
13. Shapiro BS, Daneshmand ST, Garner FC, et al. Large blastocyst diameter, early blastulation, and low preovula-
tory serum progesterone are dominant predictors of clinical pregnancy in fresh autologous cycles. Fertil Steril
2008; 90: 302–309.
14. Brinsko SP, Ball BA, Miller PG, et al. In vitro development of day 2 embryos obtained from young, fertile mares
and aged, subfertile mares. J Reprod Fertil 1994; 102: 371–378.
15. Bongso A. Defining blastocyst quality. In: Handbook on Blastocyst Culture (ed. Bonso A). Singapore: Sydney
Press Indusprint, 1999, pp. 62–79.
16. Winston NJ, Braude PR, Pickering SJ, et al. The incidence of abnormal morphology and nucleocytoplasmic ratios
in 2–, 3– and 5–day human pre-embryos. Hum Reprod 1991; 6: 17–24.
17. Van Blerkom J. Development of human embryos to the hatched blastocyst stage in the presence or absence of a
monolayer of Vero cells. Hum Reprod 1993; 8: 1525–1539.
18. Vlad M, Walker D, Kennedy RC. Nuclei number in human embryos co-cultured with human ampullary cells.
Hum Reprod 1996; 11: 1678–1686.
19. Dumoulin JCM, Derhaag JG, Bras M, et al. Growth rate of human preimplantation embryos is sex dependent after
ICSI but not after IVF. Hum Reprod 2005; 20: 484–491.
20. Fong CY, Bongso A. Comparison of human blastulation rates and total cell number in sequential culture media
with and without co-culture. Hum Reprod 1998; 14: 774–781.
21. Archer J, Gook DA, Edgar DH. Blastocyst formation and cell numbers in human frozen-thawed embryos
following extended culture. Hum Reprod 2003; 18: 1669–1673.
22. Hardy K, Stark J, Winston RML. Maintenance of the inner cell mass in human blastocysts from fragmented
embryos. Biol Reprod 2003; 68: 1165–1169.
23. Hardy K, Stark J. Mathematical models of the balance between apoptosis and proliferation. Apoptosis 2002; 7:
373–381.
24. Matsuura K, Hayashi N, Takiue C, et al. Blastocyst quality scoring based on morphologically grading correlates
with cell number. Fertil Steril 2010; 94: 1135–1137.
25. Richter KS, Harris DC, Daneshmand ST, et al. Quantitative grading of human blastocyst: Optimal inner cell mass
size and shape. Fertil Steril 2001; 76: 1157–1167.
26. Ebner T, Vanderzwalmen P, Shebl O, et al. Morphological aspects of human blastocysts and the impact of
vitrification. J Reprod Med Endocrinol 2011; 8: 13–20.
27. Leese HJ, Donnay I, Thompson JG. Human assisted conception: A cautionary tale. Lessons from domestic
animals. Hum Reprod 1998; 13 (Suppl 4): 184–202.
28. Kovacic B, Vlaisavljevic V, Reljic M, et al. Developmental capacity of different morphological types of day 5
human morulae and blastocysts. Reprod Biomed Online 2004; 8: 687–694.
29. Ahlström A, Westin C, Reismer E, et al. Trophectoderm morphology: An important parameter for predicting live
birth after single blastocyst transfer. Hum Reprod 2011; 27: 89–96.
30. Ahlström A, Westin C, Reismer E, et al. Prediction of live birth in frozen-thawed single blastocyst transfer cycles
by pre-freeze and post-thaw morphology. Hum Reprod 2013; 28: 1199–1209.
31. Honnma H, Baba T, Sasaki M, et al. Trophectoderm morphology significantly affects the rates of ongoing preg-
nancy and miscarriage in frozen-thawed single-blastocyst transfer cycle in vitro fertilization. Fertil Steril 2012;
98: 361–367.
32. Van den Abbeel E, Balaban B, Ziebe S, et al. Association between blastocyst morphology and outcome of single-
blastocyst transfer. Reprod Biomed Online 2013; 27: 353–361.
33. Hardarson T, Caisander G, Sjögren A, et al. A morphological and chromosomal study of blastocysts developing
from morphologically suboptimal human pre-embryos compared with control blastocysts. Hum Reprod 2003; 18:
399–407.
34. Alfarawati S, Fragouli E, Colls P, et al. The relationship between blastocyst morphology, chromosomal
abnormality, and embryo gender. Fertil Steril 2011; 95: 520–524.
35. Ménézo YJR, Kaufmann R, Veiga A, et al. A mini-atlas of the human blastocyst in vitro. Zygote 1999; 7: 61–65.
36. Sathananthan H, Menezes J, Gunasheela S. Mechanisms of human blastocyst hatching in vitro. Reprod Biomed
Online 2003; 7: 228–234.
preimplantation development. Fertil Steril 2005; 83: 1635–1640.
38. Veeck LL, Zaninovic N. Human blastocysts in vitro. In: An Atlas of Human Blastocysts (eds. Veeck L, Zaninovic N).
London: Parthenon Publishing, 2003, pp. 99–137.
39. Gardner RL. Flow of cells from polar to mural trophectoderm is polarized in the mouse blastocyst. Hum Reprod
2000; 15: 694–701.
40. Ebner T, Shebl O, Moser M, et al. Developmental fate of ovoid oocytes. Hum Reprod 2008; 23: 62–66.
41. Lu YC, Ding GL, Yang J, et al. Small-conductance calcium-activated K (+) channels 3 (SK3) regulate blastocyst
hatching by control of intracellular calcium concentration. Hum Reprod 2012; 27: 1421–1430.
42. Ebner T, Gruber I, Moser M. Location of herniation predicts implantation behaviour of hatching blastocysts.
J Turk Ger Gynecol Assoc 2007; 8: 184–188.
43. Miyata H, Matsubayashi H, Fukutomi N, et al. Relevance of the site of assisted hatching in thawed human
blastocysts: A preliminary report. Fertil Steril 2010; 94: 2444–2447.
44. Milki AA, Jun SH, Hinckley MD, et al. Incidence of monozygotic twinning with blastocyst transfer compared to
cleavage-stage transfer. Fertil Steril 2003; 79: 503–506.
45. Kawachiya S, Bodri D, Shimada N, et al. Blastocyst culture is associated with an elevated incidence of
monozygotic twinning after single embryo transfer. Fertil Steril 2011; 95: 2140–2142.
46. Källén B, Finnström O, Lindam A, et al. Blastocyst versus cleavage stage transfer in in vitro fertilization:
Differences in neonatal outcome? Fertil Steril 2010; 94: 1680–1683.
47. Nakashima A, Araki R, Tani H, et al. Implications of assisted reproductive technologies on term singleton birth
weight: An analysis of 25,777 children in the national assisted reproduction registry of Japan. Fertil Steril 2013;
99: 450–455.
48. Fernando D, Halliday JL, Breheny S, et al. Outcomes of singleton births after blastocyst versus nonblastocyst
transfer in assisted reproductive technology. Fertil Steril 2012; 97: 579–584.
49. Mäkinen S, Söderström-Anttila V, Vainio J, et al. Does long in vitro culture promote large for gestational age
babies? Hum Reprod 2013; 28: 828–834.
50. Santos Filho E, Noble JA, Poli M, et al. A method for semi-automatic grading of human blastocyst microscope
images. Hum Reprod 2012; 27: 2641–2548.
9
Noninvasive Techniques: Gamete Selection—Sperm
Introduction
The ultimate aim of any sperm selection method is to provide the best-quality sperm possible so as to maximize
the outcome of whatever assisted reproductive technology (ART) procedures are to be undertaken. For artificial
insemination and in vitro fertilization (IVF), the main requirement is the provision of a sample enriched with
progressive motile sperm, because these techniques are dependent on the ability of sperm to find and penetrate
the oocyte by themselves. To this end, several preparatory techniques have been developed, the most used of
which have been detailed in the preceding chapter.
With the advent of intracytoplasmic sperm injection (ICSI), many of the elements of the aforementioned
techniques were circumvented; consequently, the onus for sperm selection shifted from motility to other
parameters more influential for ICSI itself, namely, sperm morphology, content, function, and nuclear DNA
integrity. The purpose of this chapter is not to list every technique that has been developed or trialed but
rather to present the most common clinically used of these methods to describe their underlying principle,
detail the procedure, and discuss the advantages and disadvantages of each. As an adjunct, an exciting new
use of a relatively old technique that holds the potential of noninvasive and nondestructive sperm selection is
also presented.
Microscopy Beyond Morphology

Since Antoni van Leeuwenhoek first trained his microscope on the secretion “born from the seed of the genitals
of animals” [1], the microscope has been central to the evaluation of semen. In the ensuing centuries, a multi-
tude of publications have focused on the microscopic description of sperm and their qualities, with morphology
being the predominant criterion considered by embryologists for their selection of sperm to be used in ICSI.
The importance of sperm morphology and its associations with ARTs is dealt in depth in Chapter 4; here we
describe a feature beyond morphological “normalcy.”
Polarization Microscopy
Background
In contrast to the evaluation of oocytes with polarizing light, where it has been used to locate the spindle and
assess the thickness of the zona pellucida, the use of polarizing light to appraise sperm quality is a relatively
new development. The principle behind such measurements is that the nucleoprotein filaments together with the
arrangement and orientation of the acrosome affect the birefringence, that is, the refractive index based on polar-
ization and direction of propagated light, in such a way that it is indicative of the sperm’s health and maturity [2].
Because these aspects are associated with nuclear (n)DNA integrity, birefringent appearance has been suggested
as a further selection criterion of sperm, and as such many studies have incorporated polarized light microscopy
into their conventional ICSI system [2–6].
143
BOX 9.1 APPLYING POLARIZATION MICROSCOPY FOR SPERM SELECTION
Overview
A. Preparation
B. Examination
C. Selection
Instrumentation and Materials

• ICSI system normally used (e.g., inverted microscope, Hoffman contrast, motorized
micromanipulators)
• Polarizing and analyzing lenses (20× or 63×)
• Polyvinylpyrrolidone solution (PVP) (7%)
• Mineral oil
Procedure
A. Preparation
A.1. Obtain the sperm suspension by the method routinely used (e.g., swim-up, density gradient
centrifugation).
A.2. Dilute the resulting suspension to 3 × 106/mL.
A.3. Place 1 μL of sperm suspension in 10 μL drops of PVP covered with oil.
A.4. Incubate at room temperature for 30 min [5].
B. Birefringence examination
B.1. Observe on the inverted microscope equipped with polarizing and analyzing lenses.
B.2. Identify the birefringent sperm.
B.3. Observe whether the birefringence is displayed in the complete head or only partially in
the posta–crosomal region.
C. Sperm selection
C.1. Select sperm with micromanipulators that
−− Are birefringent
−− Show partial birefringence (post–acrosomal region) because these are reacted and
should be better for ICSI [4]
−− Do not have vacuoles
Pros and Cons

Pros Cons
• Can be performed on living sperm • Expensive equipment
• Using a 63× objective, vacuoles can also be distinguished in both • Only a limited number of groups use and few studies
birefringent and nonbirefringent sperm [4], thereby providing performed on the method
more information for selection • Further validation of the method’s clinical performance
• Acrosome-reacted sperm, reported to be beneficial for ICSI [4], is still needed
can be identified
• Identification of birefringent sperm is relatively easy and does not
add significantly to workload
Gamete Selection—Sperm 145
Sperm Binding
Selection based on the ability or inability to bind various substances presents an attractive option because, in
theory, the process does not interfere with structure, affect viability, or endanger the genetic contents of the
sperm. The type of separation that is achievable is dependent on the properties of the binding substrate, such
as the preference for mature sperm (hyaluronic acid [HA] [7]), apoptotic cells (Annexin V, Yo Pro 1), and an
affinity for disrupted DNA (oligopeptide p53 [8]). A further advantage of the approach is that the underlying
principle allows for its adaptation to several sorting techniques (e.g., magnet-activated cell sorting [MACS], flow
cytometry, and glass wool filtration).
Magnet-Activated Cell Sorting

Background
Regardless of the binding substance used, the underlying aspects of MACS are similar, that is superparamag-
netic microbeads that have been conjugated with the binding substrate are incubated with the sperm to allow for
binding, and then the resulting amalgams are placed into a magnetic field that entraps the beads that have sperm
bound to them. The sperm that do not bind and therefore are not restricted in their passage can be collected and
are thus available for use (Figure 9.1).
MACS sperm selection before ART has been reported to improve pregnancy rates and to be safe and
reliable [9], although the extent of the latter is not yet clear. The efficacy of the technique has reportedly been
improved when it is used in combination with sperm preparation methods (e.g., density gradient), in that it
achieves better sperm yields in terms of apoptosis, motility, vitality, and DNA integrity [10–12].
FIGURE 9.1 Schematic of the MACS system. A sperm suspension is incubated with Annexin V-conjugated microbeads and loaded
onto the separation column that is exposed to a magnetic field. Apoptotic sperm that have externalized phosphatidylserine on the
cell surface are bound to the microbeads (red) and thus remain in the column, leaving nonapoptotic sperm (green) to pass through
and be collected.
BOX 9.2 APPLYING MACS

Although the following description is based on separation using Annexin V [9], as stated above, the basis
of the protocol is similar for other binding substrates.
Overview
A. Setup
B. Labeling
C. Separation

• MiniMACS™ Starting kit (e.g., Miltenyi Biotec GmbH, Germany)
This kit contains
• Separator
• Multistand
• Columns
• Microbeads
• Degassed buffer (e.g., phosphate-buffered saline [PBS] supplemented with EDTA and bovine
serum albumin [BSA]; see MACS Miltenyi Biotec GmbH product datasheet)
Procedure
A. Setup
A.1. Adapt the MiniMACS separator to the MultiStand.
A.2. Insert column to the front of the separator (column wings to the front).
A.3. Place a collection tube under the column.
B. Cell magnetic labeling
B.1. Obtain sperm suspension (e.g., by density-gradient preparation).
B.2. Prepare microbead suspension.
Incubate sperm suspension with 100 μL of superparamagnetic Annexin V-conjugated micro-
beads for 15 min at room temperature.
C. Separation
C.1. Load the labeled suspension onto the column.
C.2. Wash three times with degassed buffer by adding 500 μL each time (waiting until the
reservoir is empty before adding the buffer).
C.3. Collect nonlabeled (i.e., nonapoptotic) fraction passing through the column.
C.4. Remove the column from the magnetic field.
C.5. Place the column on another collection tube.
C.6. Add 1 mL of buffer to the column.
C.7. Flush out the labeled (apoptotic) fraction retained in the column by using the plunger
provided in the kit.
Pros and Cons

Pros Cons
• The apoptotic cell fraction can be separated • Other cell types cannot be distinguished (e.g., immature germ cells, leukocytes)a
• It can be performed on living sperm • Not suitable for samples with low sperm concentration
a Therefore, it has been suggested to be used together with density-gradient sperm preparation [11].
Fluorescence-Activated Cell Sorting

Background
Sorting by flow cytometry is a well-established technique dating back to the late 1960s, and it is based upon
the detection of a fluorescent signal generated by a stain that has been incorporated into or onto a cell. As with
MACS, numerous stains indicative of various properties (e.g., acridine orange, propidium iodide [nDNA status],
Yo Pro 1 [apoptosis], Hoechst 33342 [vitality]) have been used either singularly or in combination in FACS [13].
Similarly to magnetic separation, the fundamental aspects of FACS are irrespective of the staining or
binding substance.
BOX 9.3 APPLYING FACS

The following description is based on the separation strategy of Ribeiro et al. [13] using Yo Pro 1, a protocol
that is adaptable for any of the other staining substrates.
Overview
A. Setup
B. Staining
C. Separation

• Flow cytometer (in the cited paper, the instrument used was an influx flow cytometer (BD
Biosciences) equipped with a solid-state laser (100 mW) for excitation at 355 nm, a blue laser
(200 mW) for excitation at 488 nm, and a red laser (120 mW) for excitation at 640 nm. Analyses
were performed using Sortware software from the same manufacturer)
• Yo Pro 1
• IVF medium or PBS
• BSA
• Penicillin and streptomycin
Procedure
A. Setup
A.1. Prepare sperm suspension (e.g., swim-up).
A.2. Centrifuge an aliquot of 0.3–1 × 106 spermatozoa for 10 min at 460g.
A.3. Remove supernatant resuspend pellet in 1 mL of IVF medium or PBS.
B. Staining
B.1. Add 1 μL of YO-PRO solution (final concentration, 0.2 μmol/L) to the sperm suspension.a
Incubate at 37°C and 5% CO2 for 15 min in the dark.
B.2. Filter the stained suspension with Celltrics 100 mm filters (Partec).
C. Separation
C.1. The settings used by Ribeiro et al. [13] for sorting were 100 μm nozzle; sample pressure,
19.5–20.5 psi; and event rate, 1000–9000 events per second.
C.2. Sterile PBS containing 0.1% BSA, 50 U/mL penicillin, and 50 μL/mL streptomycin was
used as sheath fluid.
C.3. Sorted samples are collected in polystyrene round-bottomed tubes containing 1 mL of
IVF medium.
a If more than one stain is to be used, it is added here.
Pros and Cons

Pros Cons
• Large numbers of sperm can be sorted in a reasonable time • Not suitable for samples with low sperm concentration
• Objective selection • Requires expensive equipment
• More than one stain can be used simultaneously • Exclusion of the stain from the sorted sperm not proven
• Benefit to ART outcome as yet not established
Selection Using Micromanipulators

Two further procedures have been developed that use the binding ability of sperm themselves.
HA Binding
A major component of the cumulus for which only mature sperm have a receptor, HA is the basis of two
commercially available separation methods: SpermSlow™ and the PICSI® system (both from Origio a/s,
Denmark) (see Video 9.1). The rationale behind both approaches is that because only mature sperm have
intact nDNA, the fraction that is bound to the acid must consist of sperm with high nDNA integrity. In practi-
cal terms, the HA solution in SpermSlow acts as an immobilizing agent, slowing down mature sperm so that
they can be collected, whereas the PICSI system involves a dish precoated with a solid-state HA formulation;
addition of the sample; and after an incubation step, the collection of the bound sperm using a micropippete.
Although there is some evidence for the efficacy of the approach [7], at present its true clinical value is yet to
be established [14].
Zona Pellucida Binding

Collection of sperm bound to the zona pellucida [15] is a more “physiological” use of sperm–oocyte binding,
because it does not involve any direct intervention or treatment and is closer to “natural” conditions. However,
access to and availability of suitable zonae are serious obstacles to the routine use of the method, as is the lack
of verification of the technique’s clinical worth.
VIDEO 9.1 Binding of spermatozoa to solid HA in dish. (Video provided by Dr. Markus Montag, ilabcomm GmbH, Germany.
Accessible on the “Downloads/Updates” tab at: http://www.crcpress.com/product/isbn/9781842145470.)
Electric Charge
Due to their concentrated, and relative to their size, large DNA content, sperm possess a high negative electric
charge. Because this charge varies depending on chromosomal complement, it was initially used as a means of
sperm sex selection [16]. More recently, variations in electrical charge have been shown to also be associated
with chromatin compaction and DNA integrity [17]; as a consequence, methods have been developed that use
these differences to separate and sort sperm.
Zeta Potential
The zeta (ζ) or electrokinetic potential is defined as the electric potential in the slip plane between a sperm
membrane and its surroundings. Mature sperm possess a ζ between –16 and –20 mV [18] whereby they are
able to adhere to positively charged substances such as the walls of plastic or glass tubes. The inference
being that this adhering ability allows for the quick separation of sperm with intact (i.e., possessing high
ζ values) from fragmented DNA (i.e., lower ζ values). The validation and use of this method have thus far
been limited [19].
Electrophoresis
Background
The isolation of sperm based on their electrophoretic movement is based on the premise that the higher
the electronegativity, the higher the sperm quality in terms of morphology, capacitation ability, and
DNA integrity [20]. Marketed commercially as MicroFlow® or SpermSep® CS10 (Nusep Ltd., Australia),
the system consists of a cartridge divided into two chambers that allow for the directional movement
of cells when exposed to an electric potential. The chamber is divided by a 5 μm polycarbonate mem-
brane that allows for the crossing of sperm but excludes larger contaminating cells (i.e., leukocytes and
germ cells). Although the true efficacy of the system is yet to be established, a full-term pregnancy has
been reported using electrophoretically isolated sperm from a male having high numbers of sperm with
DNA damage [21].
BOX 9.4 APPLYING ELECTROPHORESIS FOR SPERM SELECTION
Overview
A. Setup
B. Separation
Materials
• SpermSep CS10 (NuSep Ltd.) [22]
• Electrophoresis buffer
Procedure
A. Setup
A.1. Prepare the electrophoresis buffer 10 mM HEPES, 30 mM NaCl, and 0.2 M sucrose
(osmolarity, 310 mOsm kg−1; pH 7.4, adjusted with 2 M KOH)
A.2. Autoclave the separation cartridge.
A.3. Sterilize the electrophoresis buffer with a 0.22 μm filter.
A.4. Insert the separation cartridge into housing on the upper part of the SpermSep CS10.
A.5. Pipette 400 μL of electrophoresis buffer into the separation chamber.
A.6. Fill the buffer reservoir with 80 mL of electrophoresis buffer and place the reservoir
housing into the front of the instrument.
A.7. Run the buffer pump for 1 min before the sperm separation.
B. Separation
B.1. Make sure that the separation unit is sealed.
B.2. Load 400 μL of the semen sample into the corresponding chamber using a sterile pipette.
B.3. Leave to equilibrate for 5 min.
B.4. Start the electrophoresis (constant current of 75 mA (18–21V) for 5 min).
B.5. After the separation, the sperm preparation is recovered from the separation chamber of
the cartridge with an elongated sterile tip.
N O T E: If separated in the appropriate medium, the retrieved sperm can be used directly for IVF or ICSI.
For intrauterine insemination (IUI), the separated suspension could be used if prostaglandins in seminal
plasma have been removed or reduced to insignificant levels [22].
Pros and Cons

Pros Cons
• Taking only 5 min, the separation run is very rapid in comparison • Recovery rates are low (approximately 20% [23])
with other methods • May not be suitable for severe oligozoospermic samples
• No centrifugation needed, thus no oxidative stress caused by the • Equipment and procedure outside of normal andrology
physical forces of centrifugation laboratory practice [19]
• Not associated with increased reactive oxygen species (ROS) or • Not clinically verified or validated
oxidative damage
Future Perspectives
The rapid advances in miniaturization (e.g., labs on a chip), computerization, and optics have made a ccessible to
biomedicine, instruments that were once the sole preserve of physics and chemistry researchers. Although not
yet routine, these new methods provide opportunities for diagnosis and treatment that were previously unimagi-
nable and possess aptitudes that could make the dependable and safe assessment, grading, and selection of live
gametes a reality. One such technique that has already been introduced to reproductive investigations and holds
the promise of possible selection of sperm for ART is Raman microspectroscopy.
Raman Microspectroscopy
Background
Raman spectroscopy is a relatively old method (i.e., the fundamental principle behind the technique was dis-
covered in 1928) that has been recently “rediscovered” and used in biomedicine both as a research tool and
as a diagnostic tool. Based on the inelastic scattering of light (i.e., the Raman effect), the process provides a
“molecular fingerprint” of materials that when coupled with confocal microscopy allows for a detailed profiling
of the chemical components of cells. Already used in numerous fields of medicine [24,25], the method has been
shown to noninvasively and nondestructively analyze whole organs; tissue sections; and (more importantly for
reproductive medicine) living cells. Preliminary work [26–29] has shown the power of this method to identify,
classify, and localize nDNA damage in a single sperm, information that opens the way for the next step, the
selection and use of this sperm for ART (Figure 9.2).
FIGURE 9.2 Raman spectroscopy for sperm. (a) Averaged Raman spectra from sperm (blue) and after ultraviolet (UV) irradia-
tion (red). Arrow indicates main spectral feature in damaged sperm (peak at 1047 cm−1). Spectra have been normalized, baseline-
corrected, and smoothed. (b) Single-point Raman spectra are acquired from the post–acrosomal region of the sperm head. The red
dot represents the position on which the laser is focused.
BOX 9.5 APPLYING RAMAN SPECTROSCOPY FOR SELECTING SPERM

The following method is based on the protocols optimized in Mallidis et al. [28] and Sanchez et al. [29]
for the assessment of air-dried sperm using the LabRAM Aramis system (Horiba Scientific, France). The
settings and procedure are similar to those that would be necessary for the evaluation of living sperm.
Overview
A. Sample preparation
B. Focus localization
C. Spectral acquisition
D. Spectral analysis
Materials
• Raman microspectroscopy system
• Suprasil slides
• PBS
Procedure
A. Preparation
A.1. Obtain the sperm suspension to be analyzed.
A.2. Wash in PBS (5 min, 600g).
A.3. Smear 10 μL of the suspension onto a Suprasil slide.
A.4. Let air-dry overnight.
B. Focus localization
Normally, the visual focus does not correspond to the actual focus. Therefore, a Z-profile
should be done to find the position in the Z-axis where the best possible signal can be obtained.
B.1. Focus the sample.
B.2. Select the cell to analyze.
B.3. Center it under the laser position.

B.4. Set that position to zero.
B.5. Make a quick map in the Z-axis, taking several spectra from ~3 μm below and up to
~3 μm above the current position.
B.6. Take note of the position with the best spectrum in terms of intensity and resolution.
C. Spectral acquisition
C.1. Settings (the following are optimized for the LabRAM Aramis and may differ depending
on the system used):
−− Laser: He-Ne 632.8 nm
−− Filter: 100%
−− Hole: 100 μm
−− Slit: 100 μm
−− Spectrometer: 1200 cm−1
−− Grating: 600 grooves per millimeter
−− Objective: 100×
−− Acquisition time: 2 × 30 s (2 × 5 for Z-maps)
C.2. Move the stage to the position selected after the Z-map.
C.3. Change the acquisition time to 2 × 30 s.
Take the spectrum.
D. Spectral analysis
D.1. Data processing:
−− Baseline correction
−− Normalization (e.g., to the maximum peak within the 600–1800 cm−1 spectral range)
−− Set the lowest intensity to zero
N o t e: The procedures are performed using the software included as part of the microspectroscopic
system. For example, in the LabRAM Aramis, the software provided is LabSpec5.
D.2. Depending on the information of interest (e.g., DNA damage), select a band to analyze
(e.g., ~1095 cm−1).
D.3. Select another band that remains stable in all measurements.
D.4. Extract from the software the peak intensity values corresponding to the selected peaks
and calculate the ratios between them.
Pros and Cons

Pros Cons
• Allows for profiling of specific sperm • Equipment is expensive
• Provides detailed information on chemical composition • Requires specialist knowledge
• Not restricted to nDNA assessment • Results are complex
• Relatively quick and easy • Needs validation and verification of clinical worth
Conclusions
The long-held dream of clinicians and scientists in reproductive medicine has been, and remains, the a ssessment,
selection, and use of gametes that will provide not only good fertilization rates but also optimal embryo quality,
implantation, and pregnancy rates, all of which ultimately lead to the birth of healthy c hildren. In this endeavor,
as noted in the preceding sections, numerous techniques have been developed to select the best possible sperm
for ARTs. Although many of the described methods may be considered novelties and lack the robust verification
and validation essential for routine clinical use, they nonetheless constitute a great improvement in the existing
TABLE 9.1
Overview of Different Technologies Used for Sperm Selection
Technique Emerging Used Diagnostic Therapeutic Studies
Polarization microscopy X X X 2–4
MACS X X X 9, 30
FACS X X 13
PICSI X X X 31
HA medium X X X 32
Zona pellucida binding X X X 15, 33, 34
Zeta potential X X X 35
Electrophoresis X X X 21, 21, 23
Raman microspectroscopy X X 26–29
Note: HA, hyaluronic acid.
selection process that is solely dependent on the subjective choice of the embryologists. The main disadvantage
however is that although most of the techniques provide information on which sperm quality can be better clas-
sified, the practical aspects of the procedures either destroy or alter sperm in ways that render them unusable for
ARTs (Table 9.1). Be that as it may, until the full potential of upcoming techniques based on advanced technol-
ogy has been realized, the procedures described in this chapter, regardless of their limitations, represent the best
options presently available to the ART clinic.
REFERENCES
1. Lewenhoeck A. De natis E Semine Genitali Animalculis. Phil Trans 1677.
2. Ghosh S, Chattopadhyay R, Bose G, Ganesh A, Das S, Chakravarty BN. Selection of birefringent spermatozoa
under Polscope: Effect on intracytoplasmic sperm injection outcome. Andrologia 2012;44(Suppl 1):734–738.
3. Gianaroli L, Magli MC, Collodel G, Moretti E, Ferraretti AP, Baccetti B. Sperm head’s birefringence: A new
criterion for sperm selection. Fertil Steril 2008;90(1):104–112.
4. Gianaroli L, Magli MC, Ferraretti AP, Crippa A, Lappi M, Capitani S, et al. Birefringence characteristics in
sperm heads allow for the selection of reacted spermatozoa for intracytoplasmic sperm injection. Fertil Steril
2010;93(3):807–813.
5. Petersen CG, Vagnini LD, Mauri AL, Massaro FC, Cavagna M, Baruffi RL, et al. Relationship between DNA
damage and sperm head birefringence. Reprod Biomed Online 2011;22(6):583–589.
6. Collodel G, Iacoponi F, Mazzi L, Terzuoli G, Pascarelli NA, Moretti E. Light, polarizing, and transmission elec-
tron microscopy: Three methods for the evaluation of sperm quality. Syst Biol Reprod Med 2013;59(1):27–33.
7. Yagci A, Murk W, Stronk J, Huszar G. Spermatozoa bound to solid state hyaluronic acid show chromatin structure
with high DNA chain integrity: An acridine orange fluorescence study. J Androl 2010;31(6):566–572.
8. Enciso M, Pieczenik G, Cohen J, Wells D. Development of a novel synthetic oligopeptide for the detection of
DNA damage in human spermatozoa. Hum Reprod 2012;27:2254–2266.
9. Dirican EK, Ozgun OD, Akarsu S, Akin KO, Ercan O, Ugurlu M, et al. Clinical outcome of magnetic acti-
vated cell sorting of non-apoptotic spermatozoa before density gradient centrifugation for assisted reproduction.
J Assist Reprod Genet 2008;25(8):375–381.
10. Said TM, Agarwal A, Zborowski M, Grunewald S, Glander HJ, Paasch U. Utility of magnetic cell separation as a
molecular sperm preparation technique. J Androl 2008;29(2):134–142.
11. Said TM, Grunewald S, Paasch U, Glander HJ, Baumann T, Kriegel C, et al. Advantage of combining magnetic
cell separation with sperm preparation techniques. Reprod Biomed Online 2005;10(6):740–746.
12. Said T, Agarwal A, Grunewald S, Rasch M, Baumann T, Kriegel C, et al. Selection of nonapoptotic sper-
matozoa as a new tool for enhancing assisted reproduction outcomes: An in vitro model. Biol Reprod
2006;74(3):530–537.
13. Ribeiro SC, Sartorius G, Pletscher F, de Geyter M, Zhang H, de Geyter C. Isolation of spermatozoa with low
levels of fragmented DNA with the use of flow cytometry and sorting. Fertil Steril 2013;100:686–694.
14. Nijs M, Creemers E, Cox A, Janssen M, Vanheusden E, Van der Elst J, et al. Relationship between hyaluronic acid
binding assay and outcome in ART: A pilot study. Andrologia 2010;42(5):291–296.
15. Liu F, Qiu Y, Zou Y, Deng ZH, Yang H, Liu de Y. Use of zona pellucida-bound sperm for intracytoplasmic sperm
injection produces higher embryo quality and implantation than conventional intracytoplasmic sperm injection.
Fertil Steril 2011;95(2):815–818.
16. Engelmann U, Krassnigg F, Schatz H, Schill WB. Separation of human X and Y spermatozoa by free-flow
electrophoresis. Gamete Res 1988;19(2):151–160.
17. Chan PJ, Jacobson JD, Corselli JU, Patton WC. A simple zeta method for sperm selection based on membrane
charge. Fertil Steril 2006;85(2):481–486.
18. Ishijima SA, Okuno M, Mohri H. Zeta potential of human X- and Y-bearing sperm. Int J Androl 1991;14(5):340–347.
19. Said TM, Land JA. Effects of advanced selection methods on sperm quality and ART outcome: A systematic
review. Hum Reprod Update 2011;17(6):719–733.
20. Ainsworth C, Nixon B, Aitken RJ. Development of a novel electrophoretic system for the isolation of human
spermatozoa. Hum Reprod 2005;20(8):2261–2270.
21. Ainsworth C, Nixon B, Jansen RP, Aitken RJ. First recorded pregnancy and normal birth after ICSI using electro-
phoretically isolated spermatozoa. Hum Reprod 2007;22(1):197–200.
22. Fleming S, Aitken J. Electrophoretic sperm separation. In: Practical Manual of In Vitro Fertilization: Advanced
Methods and Novel Devices (eds. Nagy ZP, Varghese AC, Agarwal A, Springer Publishing, New York). 2012, p. 259.
23. Aitken RJ, Hanson AR, Kuczera L. Electrophoretic sperm isolation: Optimization of electrophoresis conditions
and impact on oxidative stress. Hum Reprod 2011;26(8):1955–1964.
24. Caspers PJ, Lucassen GW, Puppels GJ. Combined in vivo confocal Raman spectroscopy and confocal micros-
copy of human skin. Biophys J 2003;85(1):572–580.
25. Downes A, Mouras R, Elfick A. Optical spectroscopy for noninvasive monitoring of stem cell differentiation.
J Biomed Biotechnol 2010;2010:101864.
26. Huser T, Orme CA, Hollars CW, Corzett MH, Balhorn R. Raman spectroscopy of DNA packaging in individual
human sperm cells distinguishes normal from abnormal cells. J Biophotonics 2009;2(5):322–332.
27. Meister K, Schmidt DA, Brundermann E, Havenith M. Confocal Raman microspectroscopy as an analytical tool
to assess the mitochondrial status in human spermatozoa. Analyst 2010;135(6):1370–1374.
28. Mallidis C, Wistuba J, Bleisteiner B, Damm OS, Gross P, Wubbeling F, et al. In situ visualization of damaged
DNA in human sperm by Raman microspectroscopy. Hum Reprod 2011;26(7):1641–1649.
29. Sanchez V, Redmann K, Wistuba J, Wubbeling F, Burger M, Oldenhof H, et al. Oxidative DNA damage in human
sperm can be detected by Raman microspectroscopy. Fertil Steril 2012;98(5):1124–1129.e1–3.
30. Rawe VY, Boudri HU, Alvarez Sedo C, Carro M, Papier S, Nodar F. Healthy baby born after reduction of sperm
DNA fragmentation using cell sorting before ICSI. Reprod Biomed Online 2010;20(3):320–323.
31. Nasr-Esfahani MH, Razavi S, Vahdati AA, Fathi F, Tavalaee M. Evaluation of sperm selection procedure based
on hyaluronic acid binding ability on ICSI outcome. J Assist Reprod Genet 2008;25(5):197–203.
32. Parmegiani L, Cognigni GE, Bernardi S, Troilo E, Ciampaglia W, Filicori M. “Physiologic ICSI”: Hyaluronic
acid (HA) favors selection of spermatozoa without DNA fragmentation and with normal nucleus, resulting in
improvement of embryo quality. Fertil Steril 2010;93(2):598–604.
33. Paes Almeida Ferreira de Braga D, Iaconelli A, Jr, Cassia Savio de Figueira R, Madaschi C, Semiao-Francisco L,
Borges E, Jr. Outcome of ICSI using zona pellucida-bound spermatozoa and conventionally selected spermato-
zoa. Reprod Biomed Online 2009;19(6):802–807.
34. Black M, Liu de Y, Bourne H, Baker HW. Comparison of outcomes of conventional intracytoplasmic sperm injec-
tion and intracytoplasmic sperm injection using sperm bound to the zona pellucida of immature oocytes. Fertil
Steril 2010;93(2):672–674.
35. Kheirollahi-Kouhestani M, Razavi S, Tavalaee M, Deemeh MR, Mardani M, Moshtaghian J, et al. Selection
of sperm based on combined density gradient and Zeta method may improve ICSI outcome. Hum Reprod
2009;24(10):2409–2416.
10
Noninvasive Techniques: Gamete Selection—Oocyte
Introduction
Despite the recent advances in assisted reproductive techniques (ARTs), the embryo wastage remains dramatically
high. In fact, it has been demonstrated that only a few of the embryos among those produced in vitro is actually
able to implant, giving rise to pregnancy and live birth. As a consequence, the majority of ART-started cycles
results in a negative outcome [1,2]. To increase the chances of success, many centers perform multiple embryo
transfer, clearly at the expense of a higher incidence of multiple gestations. The ability to identify and select the
embryos with the most developmental potential is considered one major goal of contemporary in vitro fertilization
(IVF) worldwide, aiming for single embryo transfer (SET) without affecting the overall pregnancy rate.
Assessment of oocyte quality has gained increased consideration, leading to a blossoming of

the literature.
To date, the morphological evaluation of cleavage-stage embryos and blastocysts remains the gold standard
in ARTs, and it is routinely applied in IVF laboratories. However, because embryo viability seems to reflect the
intrinsic developmental potential of the originating gametes, in the past few years, the assessment of oocyte qual-
ity has gained increased consideration, leading to a blossoming of the literature. In this regard, several morpho-
logical features have been proposed as earlier markers predictive of developmental competence and implantation.
Nevertheless, the contradictory results obtained so far underline the importance of further research to reach a
consensus as well as the parallel development of novel noninvasive tests for the assessment of oocyte quality [3].
Polarized Light Microscopy

To date, the first-line oocyte quality assessment relies on the morphological classification of the cumulus–
corona–oocyte complex (CCOC) and, above all, on the correct identification of mature denuded eggs. This latter
analysis is surely a primary goal for the choice of appropriate oocytes for insemination purposes, yet it is not
devoid of difficulties and interpretational misunderstandings.
Usually, nuclear maturity is simply assessed by light microscopy through the visualization of the first polar
body (IPB) extruded in the perivitelline space (PVS): oocytes displaying the IPB are presumably at the meta-
phase II (MII) stage (Figure 10.1) in which the homologous chromosomes are aligned at the spindle equatorial
plate during meiosis II. It is well recognized that in stimulated cycles, 85% of the retrieved oocytes expose the
IPB and are thus classified as MII, whereas 10% show an intracytoplasmic structure called the “germinal vesi-
cle” (GV), characteristic of the prophase of meiosis I (PI) (Figures 10.2 and 10.3), and the last 5% lack both IPB
and GV. These latter oocytes are generally classified as metaphase I (MI) [4], although they may have undergone
GV breakdown but not yet progressed to the true MI stage (Figure 10.4).
However, it is now clear that the sole use of light microscopic analysis is inadequate, even for the seemingly simple
classification of oocytes on the basis of nuclear maturity. For example, it has been demonstrated that some oocytes
morphologically classified as MII are really immature, despite the presence of IPB in the PVS, being at the early
155
FIGURE 10.1 Denuded MII oocyte. An intact IPB is clearly FIGURE 10.2 Denuded GV oocyte with a centrally placed
visible in the PVS (400× magnification). nucleus and a prominent single nucleolus (200× magnification).
FIGURE 10.3 Denuded GV oocytes with eccentrically located FIGURE 10.4 Denuded MI oocytes without any visible IPB
nuclei and prominent nucleoli (200× magnification). The periph- in the typically narrow PVS (200× magnification).
eral location of nuclei indicates that GVBD is probably imminent.
telophase phase of meiosis I (TI). In these cases, additional information can be obtained by the visualization of the
meiotic spindle (MS). This highly ordered microtubular structure is involved in the chromosome segregation pro-
cess; thus, it is crucial for the correct completion of meiosis and subsequent fertilization. It is possible to use its ability
to shift the plane of polarized light inducing a retardance (ability that makes it “birefringent”) for the noninvasive
spotting through polarized light microscopy combined with software for image processing (Figure 10.5) [5].
MS identification contributes to both qualitative and quantitative data. Importantly, the presence of the MS
permits us to better assess nuclear maturity. For example, in the case of early telophase I the MS is clearly
interposed between the IPB and the oocyte (Figure 10.6). About 80%–90% of MII present MS [6,7]; its absence
has been related to lower fertilization rates and in vitro embryo development, although there are still insuf-
ficient data to demonstrate its correlation with an impairment of clinical pregnancy and implantation rates [8].
Even if the presence of the MS is reassuring, it should be noted that it seems to “disappear” in late telophase I,
reforming only 40–60 min later [9,10]. Moreover, the microtubules are highly sensitive to both chemical and
physical stresses, and suboptimal culture conditions can account for MS disappearance [5]. Finally, the MS may
Gamete Selection—Oocyte 157
FIGURE 10.5 MII oocyte observed at polarized light micros- FIGURE 10.6 Telophase I oocyte observed at polarized light
copy (400× magnification). The birefringent MS is visible just microscopy (400× magnification). The MS is visible at the
below the extruded IPB at about the 6 o’clock position. 6 o’clock position. It is interposed between the extruded IPB
and the ooplasm, indicating that the first meiotic division is not
yet concluded.
be artifactually not visualized if the oocyte is not correctly orientated during the analysis [6,11], and it has been
shown that its visualization is more probable if the time elapsed from human chorionic gonadotropin (hCG)
administration is at least 38 hr [7]. This finding may be explained by the fact that it is likely that more oocytes
are still in prometaphase II at time intervals closer to hCG administration [10].
Suboptimal culture conditions and oocyte handling can account for disappearance of the MS
because microtubules are highly sensitive to both chemical and physical stresses.
The MS is not only fundamental for the chromosome segregation process but also is a key organelle in the
creation of the IPB and the second polar body (IIPB). Its position at the very periphery of the cell [12] determines
the true animal pole and the plane of the first cleavage [6] and thus the polar body (PB) extrusion site. However,
the MS is not always perfectly aligned with the extruded PB (Figure 10.7). The IPB dislocation may be due to the
manipulation occurring during the denuding process. It has been demonstrated that only displacements >90° are
correlated with lower fertilization rates [11]. In fact, it should be hypothesized that in these cases the manipula-
tion has been strong enough to possibly cause irreversible damage to the oocytes. Moreover, a displaced IPB
may be a false indicator of the presence of the MS, which can be touched and injured by the micropipette during
intracytoplasmic sperm injection (ICSI) [5,13].
Finally, it has been postulated that the degree of birefringence (directly proportional to the molecular orga-
nization of the structure and the density of microtubules) [14] may correlate with in vitro and clinical results
[15–18], but data are still controversial [19].
Polarization microscopy allows detection of MSs as well as substructural features of

the zona pellucida.
Just to complete the picture, it is noteworthy to mention that polarized light microscopy may also be applied
to the analysis of other egg features considered important for oocyte selection and that usually are evaluated
solely through light microscopy, such as zona pellucida (ZP) characterization [20]. The ZP is a trilaminar glyco
protein structure crucial in the protection of oocytes and embryos and in the fertilization process. Each ZP
layer is characterized by different molecular arrangements and thereby exhibits different birefringence patterns;
FIGURE 10.7 MII oocyte observed at polarized light microscopy (400× magnification). The MS is visible at the 9 o’clock position,
and it is clearly dislocated about 90° from the IPB (placed at the 6 o’clock position).
FIGURE 10.8 MII oocytes observed at polarized light microscopy (250× magnification) using automated zona scoring. The zona
score is shown in the lower left corner and denotes oocytes with a good (a), medium (b), and bad (c) zona score. The score is depen-
dent on the uniformity and the intensity of the birefringence of the inner zona layer. (c) also shows birefringence in the outer zona
layer, which is a clear sign of an impaired zona formation during follicular development. (Images courtesy to Jana Liebenthron,
University of Bonn, Germany.)
in particular, the inner and outer layers appear birefringent due to the highly organized orientation of the
filaments [21]. The darkness, thickness, and retardance of the inner layer have been suggested by many studies
to be markers of oocyte competence and prognostic of IVF success. Even if the data are not uniform, it seems
that a regular and strong zona birefringence characterizes oocytes with better developmental potential [10].
Examples of different birefringence patterns of the ZP are shown in Figure 10.8, showing oocytes with a good
(Figure 10.8a), medium (Figure 10.8b), and bad (Figure 10.8c) zona score. These measurements were performed
with a polarization microscope and a build-in algorithm that enables automatic zona scoring [10].
The “Omics” Technologies

Most recently, the developing field of genomic, transcriptomic, proteomic, and metabolomic (the so-called
“omics” technologies) seems to promise a completely new way of assessing oocyte (and embryo) quality and
competence. In fact, apart from morphology and polarized light microscopy assessment, these approaches offer
promissory and powerful tools in the noninvasive investigation of the differences, at a molecular level, among
eggs that are only apparently similar to each other.
The Genomic Analysis

The genomic analysis relies on the conceivable premise that there should exist genetic determinants for oocyte
(and embryo) viability, such as an appropriate chromosome number and DNA integrity. The early identification
and selection of oocytes with a correct chromosomal status should help in reducing the production and transfer
of aneuploid embryos, thereby potentially improving IVF and clinical results. For many years, the method of
choice for aneuploidy screening was fluorescence in situ hybridization (FISH), but FISH has been replaced by
the more recent comparative genomic hybridization (CGH) technique that permits analysis of all the chromo-
somes throughout their length [22,23].
Whatever the technique of choice, it is essential to underline that the assessment of the chromosomal status of a
cell requires undoubtedly its destruction; thus, it is considered a highly invasive procedure. In the context of repro-
ductive medicine, the investigation of an oocyte DNA content renders it completely useless for subsequent ART
purposes. The cytogenetic study of the IPB should overcome this limitation, because any chromosomal gain or loss
in the PB is indicative of a reciprocal anomaly in the oocyte. Nevertheless, this approach does not take into account
eventual errors occurring during meiosis II, after sperm penetration, nor the paternal effect. Even if the additional
biopsy and study of the IIPB may add useful information, it is clear that (1) the nonimmediacy of the genetic results
and (2) the extensive study of both IPB and IIPB are incompatible with early oocyte selection [22,23].
The Transcriptomic Approach

DNA content is not the whole story. In fact, the phenotype is made up by the conjunctive effect of the genomic
constitution, epigenetic and environmental, acting on gene expression, protein synthesis, and metabolism [23].
The analysis of gene expression may provide useful insight into the way cells work and respond to stimuli
under different conditions. The term “transcriptome” refers to the total amount of RNA in a cell, destined to
protein synthesis as well as to RNA storage, and it comprises mRNA, rRNA, tRNA, and miRNA. Thus, the
“transcriptomic” analysis analyzes the RNA content, both quantitatively and qualitatively [23,24].
Transcriptomic analysis can be performed on the oocyte itself, at the obvious expense of rendering it use-
less for subsequent IVF purposes once again, or more interestingly, on granulosa cells (GCs) or cumulus cells
(CCs) [22–25]. During preantral-to-antral follicle transition, GCs commence differentiation into mural GCs (lin-
ing the follicle walls) and CCs, closely adherent to the oocyte. It is well recognized that GCs are major players
in the follicular differentiation process that result in the creation of an optimal environment for the oocyte matu-
ration and growth [26]. Moreover, CCs establish with the enclosed oocyte a complex cross-talk that promotes
folliculogenesis as well as the acquisition of egg competence [27,28]. As a consequence, it is conceivable that the
study of GC and CC functions may provide valuable information about oocyte quality and its ability to fertilize
and develop into a viable implanting embryo: any observed abnormality in GCs’ and CCs’ molecular patterns
may be either the cause or the consequence of an altered oocyte development [29]. Because in IVF treatments,
after ovulation induction and ovum retrieval, GCs and CCs are usually discarded, they offer a valid, completely
noninvasive alternative to standard morphological assessment and selection of eggs.
The transcriptomic analysis permits us to study the expression of individual defined genes (candidate-
gene approach) as well as to characterize the transcriptomic profile. In the first option, the technology
of choice is represented by quantitative reverse transcription-polymerase chain reaction (qRT-PCR), also
known as real-time RT-PCR; as for the transcriptomic profiling, the microarrays lead the scene, result-
ing in the generation of a list of genes differentially expressed under different physiologic, pathologic, or
experimental conditions [24]. With a combination of these approaches, it is hoped we can soon catego-
rize the pattern of genes differentially expressed in GCs and CCs that surround healthy oocytes versus
not-viable oocytes. To date, the genes already studied are involved in different cellular pathways and pro-
cesses, such as the establishment of bidirectional c ommunication between CCs and an oocyte [30], cumulus
expansion and mucidification [29–35], regulation of apoptosis [29,33,36–38], protection from oxidative
stress and response to hypoxia [33,39,40], cell cycle/proliferation and DNA damage response [29,33,38], and
lipid and carbohydrate metabolism [30,33,36,41].
Despite the results obtained so far being extremely intriguing and encouraging, the data are not uniform and
need to be confirmed, validated, and tested in randomized trials to reach a consensus. One of the major problems
facing the attempts to associate differentially expressed genes to known predictable outcomes is the difficulty
in determining the threshold value for physiologic inter- and intrasample variances. Moreover, the studies cur-
rently available differ in size of study population (generally low), the technology used (characterized by different
sensitivity and specificity values), and the referred outcomes (oocyte maturity, oocyte quality, oocyte aneuploidy,
fertilization rate, cleavage and early cleavage rate, cleavage embryo quality, blastocyst rate, blastocyst quality,
pregnancy rate, and implantation rate). Thus, the large amount of data collected seems to emphasize the notion
that the true endpoint for every investigation should be pregnancy or, even better, live birth rate. In fact, although
some identified factors may be considered prognostic for certain egg and embryo features, they may not be asso-
ciated with other coupled characteristics, even if they are likewise correlated to further developmental potential.
Moreover, it is clearly not coherent to propose novel early markers for oocyte competence prognostic for standard
parameters already known to be inadequate predictors of IVF outcomes. Finally, when multiple embryo transfer
is performed, data analysis is complicated by the fact that it is not possible to track the developmental fate of each
embryo [24,25].
The omics technologies seem to promise a completely new way of assessing oocyte quality
and competence.
The Proteomic and Metabolomic Frontier

Unfortunately, transcriptomic analysis does not reveal the real functional molecular status of a cell, because
it does not take into account the possible posttranslational regulation of gene expression resulting in protein
modification, degradation, or sequestration. It is believed that the protein content of a cell may be more clearly
linked to the phenotype, thus bearing more useful information concerning cell viability [22,23]. The quali-
tative and quantitative protein profiling in clinical specimens is called “proteome,” and “proteomic” is the
technology referred to proteome investigation. The ground-breaking advent of mass spectrometry promises to
solve the technical problems related to conventional gel electrophoresis analysis and radiolabeling. However,
the difficulty of the technique, the high costs, and the very little knowledge related to human (or even mam-
mal) protein content renders this approach far from routine clinical applicability and more intensive search is
needed [23].
The proteomic analysis should be carried on the oocyte itself, again at the expense of losing the precious
gamete, or on the surrounding CCs and GCs. Unfortunately, very few articles have been published on this
subject [42,43], and even a partial pattern profiling is lacking. The follicular fluid (FF) content has attracted
much more interest from the scientific community, maybe due to the relatively easier way of obtaining the
starting material and the supposed simpler protein pattern depicted in comparison with somatic cells [23].
As for CC and GC analysis, FF investigation oriented to oocyte quality determination takes an advantage
from the interconnection between the egg and the provided microenvironment in which it happens to grow
and mature. Indeed, recent publications tried to examine FF protein content to identify novel markers of IVF
success [43–45].
Another frontier in omics technology for noninvasive assessment of the oocyte is represented by metabolo-
mics, namely, the investigation of the complete content of small nonproteinaceous molecules in a given bio-
logical fluid, resulting from the action of different proteins and gene expression pathways [46,47]. In contrast to
genomics and proteomics, metabolomics provides the immediate and punctual functional status of a biological
system [46,47]. The contribution of metabolomics in oocyte selection is currently under investigation, both in FF
and culture medium. In particular, it seems that the consumption of glucose and oxygen may be an interesting
prognostic factor related to oocyte competence and IVF success [47–50].
Follicular Vascularity and Oxygenation

Another supposed critical factor affecting the maturation and growth of a competent oocyte, able to develop
into a viable embryo, is the extent of perifollicular vascularity (PFV) and thus follicular oxygenation. Pulsed
and power Doppler ultrasonography have been successfully used to measure the blood flow around maturing
follicles in stimulated cycles, thus allowing us to determine differences among follicles apparently similar at
a conventional ultrasound analysis [51–63]. According to the percentage of follicular circumference present-
ing visible blood flow, different PFV grades have been described: Grade 1, <25%; Grade 2, <50%; Grade 3,
<75%; and Grade 4, >75%. Conventionally, Grade 1–2 follicles are referred as poorly vascularized, whereas
Grade 3–4 follicles are highly vascularized [52]. Contradictory results failed to find any definite correlation
between the extent of PFV and follicular maturity (expressed as follicular size, oocyte yield, and MII retrieval
rate) as well as oocyte morphological assessment, fertilization rate, embryo quality, and cleavage status
[51–54,56,57,59,62,64,65]. Nevertheless, tracking the developmental fate of oocytes and embryos derived from
follicles with high-grade versus low-grade PFV showed that there seems to be a positive correlation between
high PFV and pregnancy rates and live birth [52,54,57,58,60,61,64,65], even if some studies did not confirm
this trend [62,63].
PFV could reflect the efforts to abate hypoxia increasing the intrafollicular dissolved oxygen concentration
via gaseous diffusion as the antrum expands. It has been proposed that poorly vascularized and hypoxic follicles
can eventually lead to higher oocyte aneuploidy rates by means of reduced ATP level and decreased intracellular
pH (pHi). In fact, such an altered metabolic pattern seems to affect the organization and structural stability of
the spindle, thus favoring chromosome displacement and segregation disorders that affect the embryos’ further
ability to develop and implant [55,56,66,67]. This finding is consistent with the results obtained by the cytoge-
netic analysis of noninseminated or unfertilized MII derived by highly vascularized versus poorly vascularized
follicles [52,54].
To investigate the cellular basis for the apparent altered oocyte metabolism in cases of low PFV, some studies
examined the structure and function of mitochondria [55,56], revealing that they were heterogeneously distributed
in the cytoplasm of the majority of oocytes from poorly vascularized hypoxic follicles. As a consequence, some
ooplasmic regions resulted in reduced metabolic activity. It has been speculated that this nonhomogeneous mito-
chondrial distribution may progress to asymmetric inheritance by the daughter cells after cleavage, causing
embryo developmental arrest and demise. Nevertheless, the exact role of mitochondria remains to be elucidated.
Differences in PFV and follicular oxygenation could run in parallel with the production of angiogenic
growth factors by the GCs. A correlation has been proposed between the grade of PFV and the presence of
angiogenic growth factors in the FF. In particular, the role of vascular endothelial growth factor (VEGF)
has been quite extensively studied because it is a strong promoter of vascularization expressed in human fol-
licles by both cumulus and granulosa-lutein cells. Some studies suggested that the measurement of VEGF
FF concentration could be used as an index of PFV and hypoxia [56,65,68,69], but contradictory results were
obtained. Moreover, this is a time-consuming and expensive technique, and it has not yet been introduced in
clinical practice.
Conclusions
Early selection of oocytes is considered a major goal of contemporary IVF worldwide, allowing the identifica-
tion of the most competent gametes to inseminate. In turn, this would help in reducing the number of embryos
produced in vitro and progress to SET. Unfortunately, standard morphological evaluation is not precise, and a
consensus is still lacking. However, new noninvasive tools for oocyte selection are gaining increased interest
from the scientific community, from the more classic polarized light microscopy analysis to the evaluation of
PFV and, to conclude, the ground-breaking omics technology. The results obtained so far are really intrigu-
ing and encouraging, but it would be wise to raise some concerns. For polarized light microscopy analysis, the
contradictory results underline the need of more intense study to reach a consensus. As to omics and, in part,
PFV evaluation, the high costs, the difficulty of the techniques, and the time required for testing are limiting
their routine applicability. Moreover, even if these approaches are all considered “noninvasive,” we still need
more evidence for the safety of the techniques (i.e., the possible effect of additional time required for each oocyte
outside the incubator or unindicated removal of CCs and, as a consequence, ICSI performance). Finally, prospec-
tive randomized studies are required to determine their predictive power, alone or in combination with other
factors, so that further efforts enrich our current knowledge [24].
REFERENCES
1. Kovalevsky G, Patrizio P. High rates of embryo wastage with use of assisted reproductive technology: A look at
the trends between 1995 and 2001 in the United States. Fertil Steril 2005; 84: 325–330.
2. Bromer JG, Seli E. Assessment of embryo viability in assisted reproductive technologies: Shortcomings of cur-
rent approaches and the emerging role of metabolomics. Curr Opin Obstet Gynecol 2008; 20: 234–241.
3. Rienzi L, Vajta G, Ubaldi F. Predictive value of oocyte morphology in human IVF: A systematic review of the
literature. Hum Reprod Update 2011; 17: 34–45.
4. Rienzi L, Ubaldi F. Oocyte retrieval and selection. In: Textbook of Assisted Reproductive Technologies. Laboratory
and Clinical Perspectives, 3rd ed. (eds. Gardner DK, Weissman A, Howles CM, Shoham Z). London, UK:
Informa Healthcare; 2009, pp. 85–101.
5. Oldenbourg R. Polarized light microscopy of spindles. Methods Cell Biol 1999; 61: 175–208.
6. Cooke S, Tyler JP, Driscoll GL. Meiotic spindle location and identification and its effect on embryonic cleavage
plane and early development. Hum Reprod 2003; 18: 2397–2405.
7. Cohen Y, Malcov M, Schwartz T, et al. Spindle imaging: A new marker for optimal timing of ICSI? Hum Reprod
2004; 19: 649–654.
8. Petersen C. Relationship between visualization of meiotic spindle in human oocytes and ICSI outcomes: A meta-
analysis. Reprod Biomed Online 2009; 18: 235–243.
9. Montag M, Schimming T, Van der Ven H. Spindle imaging in human oocytes: The impact of the meiotic cell
cycle. Reprod Biomed Online 2006; 12: 442–446.
10. Montag M, Köster M, Van der Ven K, et al. Gamete competence assessment by polarizing optics in assisted repro-
duction. Hum Reprod Update 2011; 17: 654–666.
11. Rienzi L, Ubaldi F, Martinez F, et al. Relationship between meiotic spindle location with regard to the polar body
position and oocyte developmental potential after ICSI. Hum Reprod 2003; 18: 1289–1293.
12. Maro B, Verlhac MH. Polar body formation: New rules for asymmetric divisions. Nat Cell Biol 2002; 4:
E281–E283.
13. Silva CP, Kommineni K, Oldenbourg R, et al. The first polar body does not predict accurately the location of the
metaphase II meiotic spindle in mammalian oocytes. Fertil Steril 1999; 71: 719–721.
14. Sato H, Ellis GW, Inoue S. Microtubular origin of meiotic spindle from birifrangence. Demonstration of the
applicability of Wiener’s equation. J Cell Biol 1975; 67: 501–517.
15. Trimarchi JR, Karin RA, Keefe DL. Average spindle retardance observed using the polscope predicts cell number
in day 3 embryos. Fertil Steril 2004; 82 (Suppl 2): S268.
16. Shen Y, Stalf T, De Santis L, et al. Light retardance by human oocyte spindle is positively related to pronuclear
score after ICSI. Reprod Biomed Online 2006; 6: 737–751.
17. Rama Raju GA, Prakash GJ, Krishna KM, et al. Meiotic spindle and zona pellucida characteristics as predic-
tors of embryonic development: A preliminary study using Polscope imaging. Reprod Biomed Online 2007; 14:
166–174.
18. Kilani S, Cooke S, Kan A, et al. Are there non-invasive markers in human oocytes that can predict pregnancy
outcome? Reprod Biomed Online 2009; 18: 674–680.
19. De Santis L, Cino I, Rabellotti E, et al. Polar body morphology and spindle imaging as predictors of oocyte qual-
ity. Reprod Biomed Online 2005; 11: 36–42.
20. Ebner T, Balaban B, Moser M, et al. Automatic user-independent zona pellucida imaging at the oocyte stage
allows for the prediction of preimplantation development. Fertil Steril 2010; 94: 913–920.
21. Pellettier C, Keefe DL, Trimarchi JR. Non-invasive polarized light microscopy quantitatively distinguishes
the multilaminar structure of the zona pellucida of living human eggs and embryos. Fertil Steril 2004; 81:
850–856.
22. Patrizio P, Fragouli E, Bianchi V, et al. Molecular methods for selection of the ideal oocyte. Reprod Biomed
Online 2007; 15: 346–353.
23. Seli E, Robert C, Sirard MA. OMICS in assisted reproduction: Possibilities and pitfalls. Mol Hum Reprod 2010;
16: 513–530.
24. Huang Z, Wells D. The human oocyte and cumulus cells relationship: New insights from the cumulus cell tran-
scriptome. Mol Hum Reprod 2010; 16: 715–725.
25. Uyar A, Torrealday S, Seli E. Cumulus and granulosa cell markers of oocyte and embryo quality. Fertil Steril
2013; 99: 979–997.
26. Adashi EY. Endocrinology of the ovary. Hum Reprod 1994; 9: 815–827.
27. Albertini DF, Combelles CM, Benecchi E, et al. Cellular basis for paracrine regulation of ovarian follicle develop-
ment. Reproduction 2001; 121: 647–653.
28. Gilchrist RB, Lane M, Thompson JG. Oocyte-secreted factors: Regulators of cumulus cell function and oocyte
quality. Hum Reprod Update 2008; 14: 159–177.
29. Zhang X, Jafari N, Barnes RB, et al. Studies of gene expression in human cumulus cells indicate pentraxin 3 as a
possible marker for oocyte quality. Fertil Steril 2005; 83 (Suppl 1): 1169–1179.
30. Feuerstein P, Cadoret V, Dalbies-Tran R, et al. Gene expression in human cumulus cells: One approach to oocyte
competence. Hum Reprod 2007; 22: 3069–3077.
31. McKenzie LJ, Pangas SA, Carson SA, et al. Human cumulus granulosa cell gene expression: A predictor of fer-
tilization and embryo selection in women undergoing IVF. Hum Reprod 2004; 19: 2869–2874.
32. Cillo F, Brevini TAL, Antonini S, et al. Association between human oocyte developmental competence and
expression levels of some cumulus genes. Reproduction 2007; 134: 645–650.
33. Van Montfoort APA, Geraedts JPM, Dumoulin JCM, et al. Differential gene expression in cumulus cells as a
prognostic indicator of embryo viability: A microarray analysis. Mol Hum Reprod 2008; 14: 157–168.
34. Anderson RA, Sciorio R, Kinnell H, et al. Cumulus gene expression as a predictor of human oocyte fertilisation,
embryo development and competence to establish a pregnancy. Reproduction 2009; 138: 629–637.
35. Gebhardt KM, Feil DK, Dunning KR, et al. Human cumulus cell gene expression as a biomarker of pregnancy
outcome after single embryo transfer. Fertil Steril 2011; 96: 47–52.e2.
36. Assou S, Haouzi D, Mahmoud K, et al. A non-invasive test for assessing embryo potential by gene expression
profiles of human cumulus cells: A proof of concept study. Mol Hum Reprod 2008; 14: 711–719.
37. Maraldi T, Riccio M, Sena P, et al. MATER protein as substrate of PKC epsilon in human cumulus cells. Mol Hum
Reprod 2009; 15: 499–506.
38. Fragouli E, Wells D, Iager AE, et al. Alteration of gene expression in human cumulus cells as a potential indicator
of oocyte aneuploidy. Hum Reprod 2012; 27: 2559–2568.
39. Matos L, Stevenson D, Gomes F, et al. Superoxide dismutase expression in human cumulus oophorus cells. Mol
Hum Reprod 2009; 15: 411–419.
40. Ito M, Muraki M, Takahashi Y, et al. Glutathione S-transferase theta 1 expressed in granulosa cells as a biomarker
for oocyte quality in age-related infertility. Fertil Steril 2009; 90: 1026–1035.
41. Hamel M, Dufort I, Robert C, et al. Identification of differentially expressed markers in human follicular cells
associated with competent oocytes. Hum Reprod 2008; 23: 1118–1127.
42. Wang N, Zhang P, Guo X, et al. Comparative proteome profile of immature rat ovary during primordial follicle
assembly and development. Proteomics 2009; 9: 3425–3434.
43. Atiomo WU, Khalid S, Ziauddin A, et al. Framework for a system approach to proteomic biomarker profiling in
polycystic ovary syndrome. Expert Rev Proteomics 2009; 6: 469–499.
44. Estes SJ, Ye B, Qiu W, et al. A proteomic analysis of IVF follicular fluid in women <or=32 years old. Fertil Steril
2009; 92: 1569–1578.
45. Jarkovska K, Martinkova J, Liskova L, et al. Proteome mining of human follicular fluid reveals a crucial role
of complement cascade and key biological pathways in women undergoing in vitro fertilization. J Proteome
Res 2010; 5: 1289–1301.
46. Nagy ZP, Sakkas D, Behr B. Symposium: Innovative techniques in human embryo viability assessment. Non-
invasive assessment of embryo viability by metabolomic profiling of culture media (‘metabolomics’). Reprod
Biomed Online 2008; 17: 502–507.
47. Revelli A, Delle Piane L, Casano S, et al. Follicular fluid content and oocyte quality: From single biochemical
markers to metabolomics. Reprod Biol Endocrinol 2009; 7: 40.
48. Roberts R, Stark J, Iatropoulou A, et al. Energy substrate metabolism of mouse cumulus-oocyte complexes:
Response to follicle-stimulating hormone is mediated by the phosphatidylinositol 3–kinase pathway and is asso-
ciated with oocyte maturation. Biol Reprod 2004; 71: 199–209.
49. Preis KA, Seidel G, Jr, Gardner DK. Metabolic markers of developmental competence for in-vitro matured mouse
oocytes. Reproduction 2005; 130: 475–483.
50. Scott L, Berntsen J, Davies D, et al. Human oocyte respiration rate measurement- potential to improve oocyte and
embryo selection? Reprod Biomed Online 2008; 17: 461–469.
51. Nargund G, Bourne T, Doyle P, et al. Association between ultrasound indices of follicular blood flow, oocyte
recovery, and preimplantation embryo quality. Hum Reprod 1996; 11: 109–113.
52. Chui DKC, Pugh ND, Walker SM, et al. Follicular vascularity – the predictive value of transvaginal power
Doppler ultrasonography in an in-vitro fertilization programme: A preliminary study. Hum Reprod 1997; 12:
191–196.
53. Van Blerkom J, Antczak M, Schrader R. The developmental potential of the human oocyte is related to the dis-
solved oxygen content of follicular fluid: Association with vascular endothelial growth factor levels and perifol-
licular blood flow characteristics. Hum Reprod 1997; 12: 1047–1055.
54. Van Blerkom J. Can the developmental competence of early human embryos be predicted effectively in the clini-
cal IVF laboratory? Hum Reprod 1997; 12: 1610–1614.
55. Van Blerkom J. Epigenetic influences on oocyte developmental competence: Perifollicular vascularity and intra-
follicular oxygen. J Assist Reprod Genet 1998; 15: 226–234.
56. Van Blerkom J. Intrafollicular influences on human oocyte developmental competence: Perifollicular vascularity,
oocyte metabolism and mitochondrial function. Hum Reprod 2000; 15: 173–188.
57. Bhal PS, Pugh ND, Chui DK, et al. The use of transvaginal power Doppler ultrasonography to evaluate the rela-
tionship between perifollicular vascularity and outcome in in-vitro fertilization treatment cycles. Hum Reprod
1999; 14: 939–945.
58. Bhal PS, Pugh ND, Gregory L, et al. Perifollicular vascularity as a potential variable affecting outcome in stimu-
lated intrauterine insemination treatment cycles: A study using transvaginal power Doppler. Hum Reprod 2001;
16: 1682–1689.
59. Huey S, Abuhamad A, Barroso G, et al. Perifollicular blood flow Doppler indices, but not follicular pO2, pCO2,
or pH, predict oocyte developmental competence in in vitro fertilization. Fertil Steril 1999; 72: 707–712.
60. Borini A, Maccolini A, Tallarini A, et al. Perifollicular vascularity and its relationship with oocyte maturity and
IVF outcome. Ann N Y Acad Sci 2001; 943: 64–67.
61. Borini A, Tallarini A, Maccolini A, et al. Perifollicular vascularity monitoring and scoring: A clinical tool for
selecting the best oocyte. Eur J Obstet Gynecol Reprod Biol 2004; 115 (Suppl): S102–S105.
62. Palomba S, Russo T, Falbo A, et al. Clinical use of the perifollicular vascularity assessment in IVF cycles: A pilot
study. Hum Reprod 2006; 21: 1055–1061.
63. Palomba S, Russo T, Orio F, et al. Perifollicular vascularity assessment for selecting the best oocytes for in vitro
fertilization programs in older patients. Fertil Steril 2008; 90: 1305–1309.
64. Robson SJ, Barry M, Norman RJ. Power Doppler assessment of follicle vascularity at the time of oocyte retrieval
in in vitro fertilization cycles. Fertil Steril 2008; 90: 2179–2182.
65. Monteleone P, Artini PG, Simi G, et al. Follicular fluid VEGF levels correlate with perifollicular blood flow in
normoresponder patients undergoing IVF. J Assist Reprod Genet 2008; 25: 183–186.
66. Gaulden M. The enigma of Down syndrome and other trisomic conditions. Mutat Res 1992; 269: 69–88.
67. Van Blerkom J, Davis P, Lee J. ATP content of human oocytes and developmental potential and outcome after
in vitro fertilization and embryo transfer. Hum Reprod 1995; 10: 415–424.
68. Barroso G, Barrionuevo M, Rao P, et al. Vascular endothelial growth factor, nitric oxide, and leptin follicular fluid
levels correlate negatively with embryo quality in IVF patients. Fertil Steril 1999; 72: 1024–1026.
69. Malamitsi-Puchner A, Sarandakou A, Baka SG, et al. Concentrations of angiogenic factors in follicular fluid and
oocyte-cumulus complex culture medium from women undergoing in vitro fertilization: Association with oocyte
maturity and fertilization. Fertil Steril 2001; 76: 98–101.
11
Noninvasive Techniques: Embryo Selection
by Oxygen Respiration
Introduction
Assisted human reproduction is characterized by high variability in results regarding pregnancy and birth rates.
In fact, in a cohort of fertilized oocytes from the same stimulation cycle, the developmental rate varies drasti-
cally, even though produced at the same time and under similar conditions [1]. This variability results in large
differences in implantation potential of human embryos developed in vitro, despite similarities in observable
parameters such as embryo developmental rates and morphology. Classic studies suggested that morphologic
embryo quality is correlated with implantation rates, being the best predictor for the treatment outcome [2–4].
Therefore, there is no doubt that embryo morphology (determined by the number, size, and shape of blastomeres;
the proportion of fragments; and the presence of multinucleated blastomeres) has some predictive value on devel-
opment and implantation potential [5]. However, it is important to highlight that sometimes this conventional
evaluation is not enough to get a successful pregnancy, and seemingly good morphology embryos do not always
lead to implantation and birth. This could be explained by the lack of endometrial receptivity and the fact that
many embryos cease to develop before implantation. For this reason, several noninvasive techniques have been
proposed that may be able to detect alterations of culture environment surrounding oocytes and embryos to gain
supplemental information regarding embryo viability [6–8].
Some noninvasive methods such as metabolomic profiling are still at the initial stages of development,
awaiting verification in daily laboratory practice [9], but they could be applied in the future with further
improvements [10,11]. One of these noninvasive methods, namely oxygen consumption (OC), show promise
as a good indicator of overall metabolic activity [12] and may be a valuable parameter for evaluating embryo
quality [13,14]. OC is directly related with the capacity of an embryo to produce ATP via oxidative phosphoryla-
tion, a process that uses 30% and 60%–70% of the oxygen consumed by the embryo at the early cleavage and
blastocyst stages, respectively [15].
Cellular energy (ATP) is produced by glycolysis (anaerobic respiration) and oxidative phosphorylation (aerobic
respiration). The energy created by the first pathway does not involve O2 and produces a low amount of energy,
owing to the incomplete metabolism of glucose (∆G 47 kcal/mol glucose). The energy released from the aerobic
process (second pathway) is higher (686 kcal/mol glucose) than with the anaerobic mechanism, and it is h arnessed in
the Krebs cycle coupled to the mitochondrial electron transport chain during oxidative phosphorylation [1,16–20].
Therefore, ATP production could be estimated by measuring OC, because oxidative phosphorylation is the major
producer of ATP in mammalian early embryos [21–23], and measuring the OC pattern provides a good estimation
of embryo metabolism.
The purpose of this chapter is to describe new and potentially useful aspects for the evaluation of the embryo
implantation potential, focusing on OC as a tentative method for embryo selection. In the first section of this chapter,
we evaluate the influence of different stimulation protocols on the oocyte OC as well as the influence of different
oocyte dysmorphisms in respiration rates. The next step was to study how the fertilization process might be affected
by oxygen uptake. In the second part of the chapter, we focus on embryos, measuring the OC during 3 days of
culture until embryo transfer and evaluating its correlation with embryo development and reproductive outcome.
165
OC Measurements
Respiration or OC measurements need a special microsensor that is provided by the EmbryoScope version C
(Unisense FertiliTech, Denmark) incubator as presented in Figure 11.1.
This incubator has controlled temperature and gas composition and an optional microsensor, facilitating
automatic measurements of the OC for each individual oocyte or embryo based on a steady-state respirometer
principle (Figure 11.1). OC rate measurements are performed by a Clark-type electrochemical O2 sensor, and
the time taken for O2 measurements is 60 s per oocyte or embryo. Sensor calibration is automatic and is part of
each measurement cycle. Calibration is performed by first submerging the microsensor in 0.1 M alkaline sodium
ascorbate (0% O2) and then in the medium inside the beaker (20% O2).
The sensitivity and reproducibility of the technique enable accurate measurement of OC rates down to 0.05 nL/hr
(0.6 × 10 –15 fmol/s). The reproducibility analysis indicates a standard deviation of <0.01 nL/hr (0.083 fmol/s).
The time between measurements is 60 min for four slides, for a 15 min cycle possible with a single slide.
The procedure is performed as follows: the oocyte or the embryo is placed at the bottom of a microwell (0.5 mm
in diameter, 1.5 mm in depth) in the embryo slide containing a culture medium droplet of 25 μL of Quinn Advantage
Cleavage medium (QACM) covered with an overlay of mineral oil to prevent evaporation. As the oocyte or embryo
consumes O2, the concentration at the bottom of the well is reduced, establishing a linear O2 concentration gradient
down through the well. This gradient is quantified by the O2 sensor, and the OC rate can be estimated from the mea-
sured OC gradient within the well containing the sample, according to Fick’s first law of diffusion. Thus, multiply-
ing the gradient by the O2 diffusion coefficient (3.37 × 10 –5 cm2/s at 37ºC and 9% salinity) and the cross-sectional
area of the well (0.25 mm2). The calculated OC rate for each sample is expressed in femtomoles of O2 consumed
per second, using O2 solubility in the medium of 200.4 mmol/L at 37ºC and 9% salinity. The narrowness of the
well prevents convection from taking place at a level that would influence the accuracy of the diffusion calculations.
OC by the oocyte or embryo reduces the concentration of O2 at the bottom of the well, thereby
establishing a linear measurable O2 concentration gradient.
Oxygen Oxygen
microsensor concentration
Depth
∆X
∆C
Glass well ∆C
Respiration = –D*A* ∆X
FIGURE 11.1 Schematic overview of incubator with a microsensor that can be used to measure the oxygen consumption through
the gradient created by oocyte uptake, as explained in the text.
Embryo Selection by Oxygen Respiration 167
The injection-molded EmbryoSlide is impermeable to gaseous exchange, and the O2 microelectrode is reca-
librated automatically at the onset of each new measurement cycle. The interval between consecutive OC rate
measurements for the same oocyte or embryo is 20 min.
Also, the incubator includes an imaging system that uses a low-intensity red light (650 nm) from a single light-
emitting diode (LED) with short illumination times for image acquisition of 30 ms per image. The optics consists
of a modified Hoffmann contrast with a 20× objective (Leica Place), providing optimal light sensitivity and
resolution for the red wavelength. The digital images are collected by a highly sensitive charge-coupled device
(CCD) camera (1280 × 1024 pixels) with a resolution of 3 pixels per μm, monochrome, 8-bit. The time between
acquisitions is 15 min for four slides, and a 4 min cycle is possible for a single slide. The consequence is an image
acquisition time per well including focusing that gives rise to a video recorded by a camera.
Oocyte Respiration
The contemporary classification of oocytes is routinely based solely on a qualitative evaluation of morphological
criteria. For oocytes, polar body morphology [24]; cytoplasm appearance [25]; zona pellucida thickness, appear-
ance, and birefringence [26,27]; and the position or shape of the spindle [28,29] have all been shown to correlate
with viability. The predictive value of these individual parameters is still unresolved but is generally presumed to
be limited. Animal experiments have shown that accurate measurement of different aspects of oocyte metabolic
activity can predict embryo developmental potential [6–8].
The mitochondrial respiration component has been correlated with oocyte quality and subsequent embryo
development [30–32], and a direct correlation between mitochondrial distribution and the subsequent develop-
ment of mammalian embryos has been found in several species [30].
In the oocyte, mitochondria are probably the major energy-producing system (ATP production), many studies
mention that low mitochondrial activity and number are associated with premature arrest of the oocyte, fertiliza-
tion failure, and reduced embryo development [33–35].
If abnormal mitochondrial complement can compromise respiration [36–39], measuring O2 uptake in single
oocytes may indicate both defects on mitochondrial load or dysfunction and therefore O2 uptake measurements
might be useful for the selection of oocytes with better viability. To date, the different techniques developed to mea-
sure O2 are unsuitable for clinical use because they are too invasive for embryos, they have low sensitivity, and they
cannot measure OC without affecting embryo viability. A recent study [40] has demonstrated differences in oocyte
quality based on morphology and reproductive outcome analysis for different ovarian stimulation regimens, such as
urinary human menopausal gonadotropin (hMG), highly purified menotropin (HP-hMG), and recombinant follicle-
stimulating hormone (rFSH). However, there is still little information available on metabolic markers of oocyte
quality. One approximation of oocyte quality was done by Scott et al. [41,42]. They worked with human oocytes,
and although the material consisted of immature or nonfertilized oocytes, they observed differences on respiration
rates depending on maternal age or basal FSH concentrations, two factors directly related with oocyte health.
In human oocytes, OC is a quality marker for oocyte competence and is affected by ovarian
stimulation regimens.
In human oocytes, we have demonstrated that OC is a quality marker for human oocyte competence and is
affected by ovarian stimulation regimens (Figure 11.2). Regarding OC and oocyte morphology, we can see that OC
was not affected by the morphology of the oocytes. Oocytes with normal morphology had similar oxygen uptake
compared with other oocyte phenotypes, such as clustered (clust) and granular (Gr), or with oocytes having large
periviteline space (LPS), refractile bodies (RBs), multiple dysmorphism (M), or the first polar body fragmented
(fPB) (Figure 11.3). Finally, the analysis between OC and the fertilization of oocytes showed a direct correlation
between oocyte OC and normal fertilization. The average OC was approximately 10% higher for oocytes, before
intracytoplasmic sperm injection (ICSI), that were subsequently successfully fertilized (i.e., with two visible
pronuclei) compared with oocytes that failed to fertilize (Figure 11.4).
6.0
5.8
5.6
5.4
5.2
*
5.0 5.68
4.8
4.6 4.81 4.97
4.4
4.2
4.0
hMG
hMG+FSH
Ovarian st
imulation FSH
protocol
FIGURE 11.2 OC depending on stimulation protocol. Note the higher uptake when we stimulate the donor only with FSH,
compared with stimulation with hMG or with a combination of FSH and hMG.
343
8.00
159
6.00
Oxygen uptake
4.00
2.00
0.00
N RB LPS Gr Clust M fPB
Morfo_grup-cod
FIGURE 11.3 Box plot graphics of OC depending on oocyte morphology. Normal aspect (N), clustered (clust), granular (Gr), large
periviteline space (LPS), refractile bodies (RB), multiple dysmorphism (M), or the first polar body fragmented (fPB). The levels of
OC were very similar for the seven categories established with regard to oocyte phenotype.
Oocyte respiration before ICSI
5.2
5.1 * p < .02
5.0 N=299
fmol/s
4.9
4.8
4.7 N=49
4.6
4.5
4.4
Unfertilized Fertilized
FIGURE 11.4 Analysis of OC and its influence on the fertilization process. The level of OC was higher for those oocytes
correctly fertilized, whereas the unfertilized oocytes had lower values. (*) denotes a significant difference (p < .05).
Embryo Respiration
Until recently, the only instrument used for embryo evaluation was the inverted light microscope that provided
information on morphological characteristics. Developmental and morphological information gained from
microscopic assessment has been positively associated with in vitro fertilization (IVF) outcomes [3,4,43], includ-
ing pregnancy and implantation rates, but as mentioned, we need new selection methods that allow prediction
of embryo development potential by improving probabilities of achieving pregnancy for infertile couples [44].
The measurement of OC might be a valuable parameter for evaluating embryo quality [13,14]. Because paternal
(sperm-derived) mitochondria do not survive after fertilization [45], the entire mitochondrial content of the
developing fetus is derived from the oocyte. As it has been described, mitochondria are the main oxygen con-
sumers in mammalian oocytes and early embryos [17,18,21,46,47]. Therefore, changes in mitochondrial activity
should be directly reflected in the OC of the developing embryo.
It has been demonstrated that embryos change their metabolic preferences depending on their developmen-
tal stage [48], but to date it has not been described whether OC changes depend on the embryo developmental
stage.
We have studied OC on embryo developmental stages from fertilization to embryo transfer, dividing the
O2 measurements into four groups based on quartiles with a similar number of measurements in each period:
(1) from ICSI through pronucleus (PN) formation (<17.2 hr), (2) from PN formation through first cleav-
age (17.2–35.0 hr), (3) after first cleavage through second and third cleavages (35.1–52.0 hr), and (4) after
52 hr until moment of the embryo transfer (72 hr). The results revealed different levels of OC according
to embryo stage, giving values for each quartile of 5.34 fmol/s (95% CI, 5.31–5.36), 5.17 fmol/s (95% CI,
5.15–5.20), 4.81 fmol/s (95% CI, 4.78–4.84), and 4.59 fmol/s (95% CI, 4.56–4.62; p < .0001), as shown in
Figure 11.5.
Magnusson et al. [49] suggested that human embryos that consumed more oxygen developed into blastocysts
more quickly than those with lower OC rates [49]. Other studies have demonstrated that embryos with appropri-
ate OC have a high ability to develop into blastocysts. Yamanaka et al. [50] observed higher OC in advanced
stages (hatching or hatched group) compared with the arrested or degenerated stages, as well as a higher con-
sumption in fresh blastocysts than in vitrified, warmed blastocysts.
In our experience, the OC of embryos that achieve an ongoing pregnancy is higher compared with those that
do not, showing the biggest differences in the last quartile (>52.1 hr post-ICSI), when the embryo has five or
more cells (Figure 11.6). We observed the same pattern for embryos that implant compared with those that do not
implant (Figure 11.7). Again, the biggest difference was observed in the last quartile. Therefore, results obtained
Embryo oxygen uptake (average) Embryo O2 uptake (transferred embryos)

6.0 *
6.0
5.80 *
5.5 5.52 *
5.5 5.68 5.31 *
5.26 5.28
5.0
5.34
5.0 4.84
5.17
4.79
4.5
p < .001
4.81
4.5
4.59 R = 0.188
4.0
≤17.2 4.0
17.2–35.0 ≤17.2 17.2–35.0 35.0–52.1 >52.2
35.0–52.1
>52.2 p < .001
Hours after ICSI Not pregnant Pregnant
FIGURE 11.5 Time-dependent embryo OC. Averages in FIGURE 11.6 OC averages in each of the four time ranges
each of the four time ranges are presented. A significant linear for transferred embryos. (*) indicates a significant differ-
correlation was found between embryo development and OC ence (p < .001) between transferred embryos that generated
(Pearson correlation index = –0.188; p < .001). an ongoing pregnancy and embryos that did not generate an
ongoing pregnancy at different time ranges.
Embryo O2 uptake (transferred embryos) Embryo O2 uptake (transferred embryos)

Not implanted 6.0
6.5 * Non viable
6.27 * 100 implanted Frozen embryos
6.0 5.84 * Transferred
5.75 * 5.75 5.5
5.5 5.38 5.55
5.0 4.98 *
4.86 5.0
4.5
4.0 4.5
≤17.2 17.2–35.0 35.0–52.1 >52.2
p < .001 ≤17.2 17.2–35.0 35.0–52.1 >52.2
FIGURE 11.7 OC averages from transferred embryos in FIGURE 11.8 OC averages in each of the four time ranges
each of the four time ranges depending on implantation suc- depending on final embryo viability. (*) indicates significant
cess. (*) indicates a significant difference (p < .001) between difference (p < .001) between T/F and NV embryos.
implanted embryos and nonimplanted embryos, with bigger
differences from 52.1 hr post-ICSI.
in both analyses suggest that respiration patterns from 52 hr after ICSI and onward show the strongest correlation
with implantation and ongoing pregnancy success.
Finally, we observed that the final destination of the embryo was well correlated with OC, with differences
becoming more significant in the advanced stages of embryo cleavage. The OC values were very similar for the
three categories nonviable (NV), frozen (F), and transferred (T) during the first hours of development. However,
as cleavages continued, the differences in OC pattern became more pronounced between the better (T or F) and
the poorer quality (NV) embryos, at 4.57 fmol/s for NV, 4.94 fmol/s for F, and 4.96 fmol/s for T from 52.2 hr
and onward. Embryos with similar morphological features presented similar average OC values, displaying a
correlation between quality and OC (Figure 11.8).
Time Lapse and Oxygen Uptake: Cytokinesis and Embryo Respiration

To understand the relationship between metabolism and embryo competence, we investigated the possible cor-
relation between OC and cytokinesis, a key temporal event (cell cleavage). Studies have reported small peaks of
OC (increases of 3%–10%) preceding cell division and lasting until 2 hr. Detection of the timing of the first cleavage
is critical for subsequent evaluation of embryo quality and viability, because early cleavage is considered a significant
predictor of developmental potential and has been associated with higher pregnancy and implantation rates [51–53].
Taking this into account, embryo OC rates were analyzed at time intervals according to cytokinesis timings
(i.e., at the time in which the cleavage takes place or active period and the next measurement after the first division
or inactive period). Initially, we analyzed these measurements in all embryos, for a further comparison between
consumption of T and NV embryos, and finally, among the T embryos, in those that implanted or not.
We obtained a total of 3815 measurements during the cytokinesis (active phase) and 2204 measurements after
this division (passive phase), and we observed a significant increase in OC levels when cytokinesis occurred,
with values of 4.74 fmol/s (95% CI, 4.72–4.76) between cleavages (interphase period) and 5.14 fmol/s (95% CI,
4.72–4.76) during the cytokinesis period (Figure 11.9).
Silent and active cleavage periods show clear differences in OC.
Differences in OC between silent and active cleavage periods changed substantially among the different events
from PN formation to cleavage to eight cells (Figure 11.10). When we analyzed OC between embryos with
good and poor quality, we observed higher levels in the embryos with better quality in both phases ( interphase
and cytokinesis). The T embryos presented higher OC levels when cytokinesis occurred, 5.25 fmol/s (95% CI,
5.23–5.28) at interphase versus 5.43 fmol/s (95% CI, 5.33–5.52) at cytokinesis, c ompared with discarded embryos
(arrested or bad-quality embryos), 5.09 fmol/s (95% CI, 5.01–5.05) versus 5.28 fmol/s (95% CI, 5.21–5.35),
respectively (Figure 11.11).
Among the T embryos, we found that those that implanted presented a higher increase in OC during cytoki-
nesis, 5.79 fmol/s (95% CI, 5.66–5.91) during interphase period versus 6.71 fmol/s (95% CI, 5.85–7.58) during
cleavage period, than the embryos that failed to implant, 5.21 fmol/s (95% CI, 5.06–5.36) versus 5.28 fmol/s
(95% CI, 5.15–5.41), demonstrating the existence of high OC demand in the cell cleavage in embryos with higher
implantation potential (Figure 11.12).
Cleavage oxygen consumption pattern

5.2
5.0
fmol/s
4.8
4.6
4.4
t
ie
Respiration rate
t
Qu
ie
av
Qu
av
cle
le
t
ie
e-
t
t-c
ie
Qu
Pr
Qu
s
Po
FIGURE 11.9 OC during the division timing. Note the peak corresponds to higher energy demand in the starting of second
cell cycle.
Cleavage O2 consumption pattern

6.0
5.8
5.6
5.4
fmol/s
5.2
5.0
4.8
4.6
4.4
13 hr 21 hr 24 hr 30 hr 35 hr 45 hr 52 hr 58 hr 65 hr
4.2
Hours after ICSI
FIGURE 11.10 OC before and after each division. Note that the peak corresponds to higher energy demand in each embryo
cleavage.
5.5
5.4
5.3
5.2
fmol/s
5.1 Nonviable
5.0 Transferred
4.9
4.8
5 hr
13 hr
21 hr
29 hr
37 hr
Time-hours post ICSI
FIGURE 11.11 OC average between embryos with poor or good quality before and after cytokinesis. Note the higher values cor-
responding to transferred embryos (good quality), compared with lower values of nonviable embryos.
Limitations of the Measurement System

Even though we have seen that systems introducing OC measurements can be really useful in improving embryo
selection in the laboratory, such systems do have some as yet not fully resolved limitations. We must take into
account the fact that the study we have presented was developed using a C version of EmbryoScope and not the
currently available medical device. Being then a prototype, several technical problems and limitations were pres-
ent. This device had a capacity for four slides (12 embryos each) at the same time. It is important to highlight that
the O2 microelectrode was washed with different solutions and recalibrated automatically at the onset of each
new measurement cycle (12 embryos). However, the same sensor was placed in different slides (embryos from
different patients in their corresponding slides). Thus, in our study, we could analyze only one patient each time
to avoid the same sensor handling different patients’ media.
7.0
6.8
6.6
6.4
Not implanted
6.2
Implanted
fmol/s
6.0
5.8
5.6
5.4
5.2
5.0
5 hr
13 hr
21 hr
29 hr
37 hr
FIGURE 11.12 OC average between implanted or not implanted embryos before and after cytokinesis. Note the higher peak
corresponding to implanted embryos, compared with lower peak of not implanted embryos (practically nonexistent peak).
Moreover, even though this version included a time-lapse unit, it was not possible to perform OC measure-
ments and image analysis at the same time. Pictures were taken at different timings when OC measurements
were performed. Having the measurements and time-lapse recordings, data analysis was complex. The soft-
ware available did not combine both data, and then sophisticated Excel spreadsheets were mandatory to do
data analysis. Regardless, the OC measurements were done just before and after each division and not during
cytokinesis.
Future Perspectives: An Embryo Selection by OC?

It is important to highlight that even though morphological evaluation remains the primary method of embryo
assessment, it is not always accurate enough and the good appearance of embryos does not always mean preg-
nancy and birth. Embryo selection based on OC could be a new option to improve the assisted reproduction
outcome, where more than half of the cycles fail to succeed.
The results presented here show that the OC rate of T embryos that achieved an ongoing pregnancy was signif-
icantly higher than the OC rate of embryos that did not achieve it. The high peak observed during first division in
implanted embryos could be regarded as an alternative method of embryo selection, based on our observations of
reduced peaks in nonimplanted embryos. The higher OC differences observed in the advanced stages of cleavage
embryos strengthens the hypothesis of embryo selection by OC: respiration patterns from 52 hr after ICSI and
onward showed the strongest correlation with implantation and ongoing pregnancy success. Such differences,
which increased as cleavage continued, could be indicating that progressive deficiencies in mitochondrial func-
tion are reflected in differences in embryo OC.
Regarding embryo viability, the OC values were very similar for all types of embryos during the first hours
of development, but as cleavage continued, the differences in OC patterns became more pronounced between
the better and the poorer quality embryos. These results corroborate the conclusions mentioned above and
suggest that the measurements should be performed almost immediately before the embryo transfer (from
52 hr post-ICSI) if the intention is to perform the embryo selection by combining OC with standard embryo
morphology.
Toward later cleavage stages, differences in OC patterns become more pronounced between
better and poorer quality embryos.
The OC could be considered as an alternative to assess human embryo metabolomic profiling: OC is the
main portion of metabolism in the growing embryo and the literature suggests that the metabolomic profile is a
potential marker of embryo viability in the human model [54,55]. The combination of both methods (metabolism
and time lapse), considered as noninvasive, offers an opportunity to learn more about the developmental com-
petence of the embryo, without legal and ethical problems. Conventional morphological assessment will remain
important, but the intrinsic subjectivity related to evaluation will be compensated by the additional noninvasive
analysis, thereby increasing accuracy.
OC measurement could be applied in the near future in IVF laboratories as an alternative method for embryo
selection based on the literature and our new findings, but prospective randomized control trials are mandatory
to confirm its usefulness for embryo selection with a remarkable increase in the reproductive outcome.
REFERENCES
1. Wilding M, Coppola G, Dale B, Di Matteo L. Mitochondria and human preimplantation embryo development.
Reproduction 2009;137(4):619–624.
2. Terriou P, Sapin C, Giorgetti C, Hans E, Spach JL, Roulier R. Embryo score is a better predictor of pregnancy
than the number of transferred embryos or female age. Fertil Steril 2001;75(3):525–531.
3. Ziebe S, Petersen K, Lindenberg S, Andersen AG, Gabrielsen A, Andersen AN. Embryo morphology
or cleavage stage: How to select the best embryos for transfer after in-vitro fertilization. Hum Reprod
1997;12(7):1545–1549.
4. Van Royen E, Mangelschots K, De Neubourg D, Valkenburg M, Van de Meerssche M, Ryckaert G,
et al. Characterization of a top quality embryo, a step towards single-embryo transfer. Hum Reprod
1999;14(9):2345–2349.
5. Rienzi L, Ubaldi F, Iacobelli M, Romano S, Minasi MG, Ferrero S, et al. Significance of morphological attributes
of the early embryo. Reprod Biomed Online 2005;10(5):669–681.
6. Rieger D, Loskutoff NM. Changes in the metabolism of glucose, pyruvate, glutamine and glycine during matura-
tion of cattle oocytes in vitro. J Reprod Fertil 1994;100(1):257–262.
7. Lane M, Gardner DK. Selection of viable mouse blastocysts prior to transfer using a metabolic criterion. Hum
Reprod 1996;11(9):1975–1978.
8. Houghton FD, Hawkhead JA, Humpherson PG, Hogg JE, Balen AH, Rutherford AJ, et al. Non-invasive amino
acid turnover predicts human embryo developmental capacity. Hum Reprod 2002;17(4):999–1005.
9. Hardarson T, Ahlstrom A, Rogberg L, Botros L, Hillensjo T, Westlander G, et al. Non-invasive metabolomic pro-
filing of Day 2 and 5 embryo culture medium: A prospective randomized trial. Hum Reprod 2012;27(1):89–96.
10. Nagy ZP, Sakkas D, Behr B. Symposium: Innovative techniques in human embryo viability assessment. Non-
invasive assessment of embryo viability by metabolomic profiling of culture media (‘metabolomics’). Reprod
Biomed Online 2008;17(4):502–507.
11. Seli E, Vergouw CG, Morita H, Botros L, Roos P, Lambalk CB, et al. Noninvasive metabolomic profiling as an
adjunct to morphology for noninvasive embryo assessment in women undergoing single embryo transfer. Fertil
Steril 2010;94(2):535–542.
12. Leese HJ. What does an embryo need? Hum Fertil (Camb) 2003;6(4):180–185.
13. Barnett DK, Bavister BD. What is the relationship between the metabolism of preimplantation embryos and their
developmental competence? Mol Reprod Dev 1996;43(1):105–133.
14. Houghton FD, Thompson JG, Kennedy CJ, Leese HJ. Oxygen consumption and energy metabolism of the early
mouse embryo. Mol Reprod Dev 1996;44(4):476–485.
15. Trimarchi JR, Liu L, Porterfield DM, Smith PJ, Keefe DL. Oxidative phosphorylation-dependent and -independent
oxygen consumption by individual preimplantation mouse embryos. Biol Reprod 2000;62(6):1866–1874.
16. Wilding M, Dale B, Marino M, di Matteo L, Alviggi C, Pisaturo ML, et al. Mitochondrial aggregation patterns
and activity in human oocytes and preimplantation embryos. Hum Reprod 2001;16(5):909–917.
17. Van Blerkom J. Intrafollicular influences on human oocyte developmental competence: Perifollicular vascularity,
oocyte metabolism and mitochondrial function. Hum Reprod 2000;15(Suppl 2):173–188.
18. Van Blerkom J, Davis PW, Lee J. ATP content of human oocytes and developmental potential and outcome after
in-vitro fertilization and embryo transfer. Hum Reprod 1995;10(2):415–424.
19. Pozzan T, Magalhaes P, Rizzuto R. The comeback of mitochondria to calcium signalling. Cell Calcium
2000;28(5–6):279–283.
20. Wilding M, Di Matteo L, Dale B. The maternal age effect: A hypothesis based on oxidative phosphorylation.
Zygote 2005;13(4):317–323.
21. Houghton FD, Leese HJ. Metabolism and developmental competence of the preimplantation embryo. Eur J
Obstet Gynecol Reprod Biol 2004;115(Suppl 1):S92–S96.
22. Shoubridge EA, Wai T. Mitochondrial DNA and the mammalian oocyte. Curr Top Dev Biol 2007;77:87–111.
23. Thompson JG, Partridge RJ, Houghton FD, Cox CI, Leese HJ. Oxygen uptake and carbohydrate metabolism by
in vitro derived bovine embryos. J Reprod Fertil 1996;106(2):299–306.
24. Ebner T, Yaman C, Moser M, Sommergruber M, Feichtinger O, Tews G. Prognostic value of first polar body morphol-
ogy on fertilization rate and embryo quality in intracytoplasmic sperm injection. Hum Reprod 2000;15(2):427–430.
25. Ebner T, Moser M, Tews G. Is oocyte morphology prognostic of embryo developmental potential after ICSI?
Reprod Biomed Online 2006;12(4):507–512.
26. Shen Y, Stalf T, Mehnert C, Eichenlaub-Ritter U, Tinneberg HR. High magnitude of light retardation by the zona
pellucida is associated with conception cycles. Hum Reprod 2005;20(6):1596–1606.
27. Shiku H, Shiraishi T, Ohya H, Matsue T, Abe H, Hoshi H, et al. Oxygen consumption of single bovine embryos
probed by scanning electrochemical microscopy. Anal Chem 2001;73(15):3751–3758.
28. Madaschi C, de Souza Bonetti TC, de Almeida Ferreira Braga DP, Pasqualotto FF, Iaconelli A, Jr, Borges E, Jr.
Spindle imaging: A marker for embryo development and implantation. Fertil Steril 2008;90(1):194–198.
29. Madaschi C, Aoki T, de Almeida Ferreira Braga DP, de Cassia Savio Figueira R, Semiao Francisco L, Iaconelli A,
Jr, et al. Zona pellucida birefringence score and meiotic spindle visualization in relation to embryo development
and ICSI outcomes. Reprod Biomed Online 2009;18(5):681–686.
30. Bavister BD, Squirrell JM. Mitochondrial distribution and function in oocytes and early embryos. Hum Reprod
2000;15(Suppl 2):189–198.
31. Squirrell JM, Lane M, Bavister BD. Altering intracellular pH disrupts development and cellular organization in
preimplantation hamster embryos. Biol Reprod 2001;64(6):1845–1854.
32. Squirrell JM, Schramm RD, Paprocki AM, Wokosin DL, Bavister BD. Imaging mitochondrial organization in
living primate oocytes and embryos using multiphoton microscopy. Microsc Microanal 2003;9(3):190–201.
33. Van Blerkom J. Mitochondria in human oogenesis and preimplantation embryogenesis: Engines of metabolism,
ionic regulation and developmental competence. Reproduction 2004;128(3):269–280.
34. Van Blerkom J. Mitochondria in early mammalian development. Semin Cell Dev Biol 2009;20(3):354–364.
35. Dumollard R, Duchen M, Carroll J. The role of mitochondrial function in the oocyte and embryo. Curr Top Dev
Biol 2007;77:21–49.
36. May-Panloup P, Chretien MF, Jacques C, Vasseur C, Malthiery Y, Reynier P. Low oocyte mitochondrial DNA
content in ovarian insufficiency. Hum Reprod 2005;20(3):593–597.
37. May-Panloup P, Chretien MF, Malthiery Y, Reynier P. Mitochondrial DNA in the oocyte and the developing
embryo. Curr Top Dev Biol 2007;77:51–83.
38. Reynier P, May-Panloup P, Chretien MF, Morgan CJ, Jean M, Savagner F, et al. Mitochondrial DNA content
affects the fertilizability of human oocytes. Mol Hum Reprod 2001;7(5):425–429.
39. Muller-Hocker J, Schafer S, Weis S, Munscher C, Strowitzki T. Morphological-cytochemical and molecu-
lar genetic analyses of mitochondria in isolated human oocytes in the reproductive age. Mol Hum Reprod
1996;2(12):951–958.
40. Melo M, Bellver J, Garrido N, Meseguer M, Pellicer A, Remohi J. A prospective, randomized, controlled trial
comparing three different gonadotropin regimens in oocyte donors: Ovarian response, in vitro fertilization out-
come, and analysis of cost minimization. Fertil Steril 2010;94(3):958–964.
41. Scott L, Berntsen J, Davies D, Gundersen J, Hill J, Ramsing N. Symposium: Innovative techniques in human
embryo viability assessment. Human oocyte respiration-rate measurement—potential to improve oocyte and
embryo selection? Reprod Biomed Online 2008;17(4):461–469.
42. Scott L, Finn A, O’Leary T, McLellan S, Hill J. Morphologic parameters of early cleavage-stage embryos that
correlate with fetal development and delivery: Prospective and applied data for increased pregnancy rates. Hum
Reprod 2007;22(1):230–240.
43. Giorgetti C, Terriou P, Auquier P, Hans E, Spach JL, Salzmann J, et al. Embryo score to predict implantation after
in-vitro fertilization: Based on 957 single embryo transfers. Hum Reprod 1995;10(9):2427–2431.
44. Baczkowski T, Kurzawa R, Glabowski W. Methods of embryo scoring in in vitro fertilization. Reprod Biol
2004;4(1):5–22.
45. Cummins JM. Fertilization and elimination of the paternal mitochondrial genome. Hum Reprod 2000;15(Suppl 2):
92–101.
46. Zweig MH, Campbell G. Receiver-operating characteristic (ROC) plots: A fundamental evaluation tool in clinical
medicine. Clin Chem 1993;39(4):561–577.
47. Lopes AS, Lane M, Thompson JG. Oxygen consumption and ROS production are increased at the time of fertil-
ization and cell cleavage in bovine zygotes. Hum Reprod 2010;25:2762–2773.
48. Botros L, Sakkas D, Seli E. Metabolomics and its application for non-invasive embryo assessment in IVF. Mol
Hum Reprod 2008;14(12):679–690.
49. Magnusson C, Hillensjo T, Hamberger L, Nilsson L. Oxygen consumption by human oocytes and blastocysts
grown in vitro. Hum Reprod 1986;1(3):183–184.
50. Yamanaka M, Hashimoto S, Amo A, Ito-Sasaki T, Abe H, Morimoto Y. Developmental assessment of human
vitrified-warmed blastocysts based on oxygen consumption. Hum Reprod 2011;26(12):3366–3371.
51. Salumets A, Hyden-Granskog C, Suikkari AM, Tiitinen A, Tuuri T. The predictive value of pronuclear morphol-
ogy of zygotes in the assessment of human embryo quality. Hum Reprod 2001;16(10):2177–2181.
52. Salumets A, Hyden-Granskog C, Makinen S, Suikkari AM, Tiitinen A, Tuuri T. Early cleavage predicts the viabil-
ity of human embryos in elective single embryo transfer procedures. Hum Reprod 2003;18(4):821–825.
53. Fancsovits P, Toth L, Takacs ZF, Murber A, Papp Z, Urbancsek J. Early pronuclear breakdown is a good indicator
of embryo quality and viability. Fertil Steril 2005;84(4):881–887.
54. Garrido-Gomez T, Dominguez F, Lopez JA, Camafeita E, Quinonero A, Martinez-Conejero JA, et al. Modeling
human endometrial decidualization from the interaction between proteome and secretome. J Clin Endocrinol
Metab 2011;96(3):706–716.
55. Dominguez F, Pellicer A, Simon C. The human embryo proteome. Reprod Sci 2009;16(2):188–190.
12
Noninvasive Techniques: Embryo Selection
by Time-Lapse Imaging
Alison Campbell
Introduction
The ability to acquire sequential, photographic, time-lapse images of the developing preimplantation human
embryo in vitro has recently provided clinical embryologists with a powerful noninvasive embryo monitor-
ing and selection tool. Time-lapse imaging was used to study fertilization and early human embryo kinetics
more than 15 years ago, but this technology is now available for the routine clinical in vitro fertilization (IVF)
setting [1]. The first live birth after time-lapse imaging of embryos was reported by Pribenszky and colleagues in
2010, and since then time-lapse imaging of embryos has rapidly become a key topic and area of research in the
field of human fertility [2]. This chapter focuses on practical aspects of time-lapse imaging for embryo selection.
It covers some of the key findings that have been reported to date and considers the potential impact this exciting
technology may have on our understanding of embryo development and on clinical IVF outcome.
Standard versus Time-Lapse Methodology in the Embryology Laboratory

Currently, embryo selection methods rely primarily on morphological evaluation of the embryo based on five or
six single, conveniently scheduled observations during their in vitro development. Although these daily observa-
tions are recorded and considered, selection of the embryo(s) for transfer, or cryopreservation, tends to be weighted
toward the morphology of the embryo just before embryo transfer, focusing on cell number and amount of frag-
mentation. Selection is often based on a simple embryo grading scheme or a graduated embryo scoring scheme that
may also take into account several other variables such as static pronuclear morphology [3,4]. These observations
are, in general, not photographic but are recorded by a scribed entry onto a laboratory worksheet, as a database
entry, or both. Although this methodology is evidence based, and now guided by a consensus document, time-lapse
technology has highlighted the limitations of this rather subjective approach to a crucial clinical decision [5,6].
It is logical that a static snap-shot method to accurately study a dynamic process, such as embryo develop-
ment, may not be optimal; however, to date, IVF practitioners have worked within these limitations to study and
improve embryo selection in the interest of their patients.
A static snap-shot method is not accurate to study a dynamic process such as embryo development.
Conventional light microscopical observations, unlike time-lapse imaging, cannot allow the precise timings of
mitoses, or the observation of anomalous cleavage events of the in vitro preimplantation embryo, to be recorded.
Furthermore, without time lapse, transient characteristics such as multinucleation can easily be missed when
embryos are limited to a single daily observation. Many studies have demonstrated the detrimental effect of
multinucleation on embryo implantation, pregnancy, and birth rates [7]. Time-lapse imaging and analysis ensure
that such phenomena are observed and recorded, allowing embryo deselection, where appropriate.
177
Having thousands of time-lapse images available, compared with the usual four to six records of a static obser-
vation with conventional methods, may initially seem daunting to the embryologist responsible for embryo selec-
tion, but, with robust training and quality assurance, confidence and skills develop quickly and the inquiring
mind of the embryologist is opened to this exciting approach.
The wealth of detail that can be observed and information gleaned from such time-lapse images is remark-
able. As an example, even before the first cleavage, and within hours of insemination, polar body extrusion can
be visualized, providing rapid reassurance that fertilization is likely underway. The dynamics and nature of the
proceeding pronuclear formation and morphology now question the extent to which traditional pronuclear grad-
ing can assist in embryo selection.
Throughout the preimplantation embryo’s development, from insemination to uterine transfer, new phenom-
ena, observed using time-lapse imaging and analysis devices, are being defined and observed as the study of
human preimplantation embryology takes on a new direction.
Essential Tools: The Time-Lapse Device

Several clinical time-lapse devices with automatic image capture and software for data collection and analysis
are commercially available for IVF practice. Some use the standard incubator (e.g., Primo Vision, Vitrolife,
Sweden and Eeva, Auxogyn, United States). Such systems use a single microscope objective within a unit placed
inside the incubator chamber to capture images within one culture dish. These systems allow group culture
conditions due to the design of the dishes provided with the devices. Each is modular, allowing multiple devices
to be fitted within one incubator, with the number depending on the size of the incubator. Primo Vision uses
Hoffman contrast integrated optics with green light-emitting diode (LED) (550 nm) illumination and can collect
images through 11 focal planes. The Eeva system uses dark-field image capture, in a single focal plane, and auto-
matic cell division tracking and uses software to analyze early embryo development, providing evidence-based
quantitative data on each embryo’s developmental potential to the blastocyst stage [8]. The Primo Vision system
allows user-defined programming of the software to record morphokinetic variables. It also allows published
or in-house–developed embryo selection algorithms to be used to rank embryos accordingly. Although widely
available, these two systems are not yet cleared for clinical use in the United States.
Selection of a time-lapse system for clinical use is likely to be a task that all IVF centers will be
faced with over the next few years to provide the optimum clinical outcome for their patients.
The most widely used integrated device available and in use worldwide is the EmbryoScope (FertiliTech,
Denmark). Rather than using a conventional incubator, this device in itself is a nonhumidified incubator with inter-
nal circulation of ultraviolet (UV)-light sterilized air through a high-efficiency particulate air (HEPA) and volatile
organic compound filtration system. The EmbryoScope has an in-built red LED (635 nm) illumination camera sys-
tem that arguably allows the most stable and uninterrupted time-lapse imaging and culture. The EmbryoScope can
image up to 72 embryos, through a maximum of nine equidistant focal planes. This system includes embryo selection
modeling software and provides tools for knowledge building through retrospective analysis of embryo developmen-
tal data. A more recently developed device by ESCO Medical (Singapore) is an alternative time-lapse incubation
system with capacity for up to 84 embryos from six patients. This system consists of six individually controlled and
monitored culture compartments and also requires manual annotation of the acquired time-lapse images.
Selection of a time-lapse system for clinical use is likely to be a task that all IVF centers will be faced with over
the next few years to provide the optimum clinical outcome for their patients. It is also likely to be a long-term invest-
ment. When deciding which system, or systems, to introduce, consideration should be given to potential impact of
clinical outcome, opportunities for development and continuous improvement, device specification, focal planes,
image quality and capacity, user friendliness, degree of validation conducted by the supplier and required in-house,
certification and licensing (where required), limitations, space, and cost. Customer support, servicing, and training
should also be taken into account because accessibility to these may be limited by clinic or supplier location.
Embryo Selection by Time-Lapse Imaging 179
Identification and Annotation of Dynamic Embryo Development

When using time lapse, the frequency of image acquisition can take place at variable time intervals set by the
individual laboratory. At present 5, 10, 15, or 20 min intervals are most commonly used. As the technology
advances, these time intervals may be moved closer, to acquire more detailed information, or even varied during
the course of preimplantation embryo development to access more information during certain periods and less
during periods of quiescence or where there is evidence of image capture being of lesser clinical impact.
Time-lapse images played, rewound, and paused sequentially as a video can be assessed automatically or
semiautomatically by software, or manually. Embryologists and researchers can now study the movement (kinet-
ics) and durations of developmental events alongside embryo morphology. The embryo’s developmental patterns
and appearance have been referred to as morphokinetics [9]. To date, several specific morphokinetic variables
have been correlated to embryo viability, and the outcome measures considered have been blastocyst develop-
ment, implantation, and live birth. As data from embryos transferred with a known outcome amasses, the most
predictive morphokinetic variables for viable embryo selection can be identified. It should be noted, of course,
that embryo quality and selection is just one of the several factors influencing IVF treatment success. Factors
such as clinical history, embryo transfer procedure, and endometrial receptivity are also of great importance.
Unless the selected time-lapse system has fully automated image capture and analysis tools, some degree
of user definability, in terms of morphokinetic variables to be recorded, may be required when establishing
a clinical time-lapse service or research program.
Because there may be multiple practitioners in the laboratory involved with the assessment of time-lapse
images, it is recommended that key variables for annotation are defined within the standard operating procedure
and that these variables are routinely recorded. It is crucial that an early decision is made as to whether variables
or morphological features that are not seen in a particular embryo (e.g., compaction or vacuolation) should be
reported as “not seen” or whether not reporting them is sufficient to suggest that they were not observed.
Additional or novel variables can be added at a later date once the practitioners have gained familiarity and
are able to annotate more swiftly. Annotation is the process of interpreting and recording morphokinetic events
on visualizing the time-lapse images and entering this information into the time-lapse device software, where
applicable. The introduction of time lapse should not been seen as a burden but rather as a tool that not only dra-
matically increases flexibility in the embryologists’ working day but also has the potential to train, educate, and
most importantly enhance clinical outcome. Because time-lapse images viewed as a video can be retrospectively
studied and annotated, colleagues can work together to ensure the quality of annotation, discuss queries, and
even go back to historical images to annotate new variables at a later date.
Guidelines and consensus are required for standard time-lapse imaging, and to date, without this agreement,
several alternative definitions have already been used for the same variable and the approach and standard prac-
tice varied from clinic to clinic.
The most commonly used morphokinetic variables are established based on the basic principles of embryol-
ogy and mitosis and include timing of pronuclear appearance and fading, increasing cell numbers (time to two,
three, four, five, six cells, etc.) and times of embryo differentiation to the morula and blastocyst stages. Durations
of mitotic cycles and synchronicity, as used in some of the published dynamic embryo selection algorithms, can
then be calculated from these variables.
Annotation is the process of interpreting and recording morphokinetic events on visualizing

the time-lapse images and entering this information into the time-lapse device software.
If there is a specific research interest, additional “user-defined” variables can be recorded. An example may
be surrounding multinucleation. If the number, size, degree, appearance, fading, and dynamics of nuclei are of
particular interest during embryo development, this information can be recorded. To facilitate downstream data
analysis, it is important that strictly defined terms or phrases are used for these phenomena. An accepted con-
sensus document defining such criteria would assist scientists in data sharing.
Table 12.1 summarizes the commonly used morphokinetic variables and provides basic descriptions and abbre-
viations for them. The abbreviations in bold may be considered core variables, several of which have been referred
to in time-lapse–related clinical publications to date. It is recommended that they are routinely recorded when
using time lapse in a clinical setting. This table is not exhaustive and is included as a guideline for users of time
lapse. It has been adapted from a document, in preparation, by a team of time-lapse experts aiming to encourage
users of time-lapse technologies in IVF to standardize practice for data to be amassed, experiences shared, and
best practice reached. Anomalous and non-time-lapse–dependent variables have not been included but may be
annotated at the embryologist’s discretion and in line with standard operating procedure.
Assurance of Annotation Quality

Ensuring an accurate and objective record of dynamic, and often anomalous, embryo development can be chal-
lenging, whether using automatic detection software, the human eye, or even by committee. To amass data of
high quality and a resource for embryo selection algorithm development to enhance clinical outcome, a quality
assurance system should be introduced early on to ensure that embryologists’ or practitioners’ interpretations
and annotations are objective and consistent. This exercise may be developed in-house or provided by the
time-lapse device supplier. Even where there is a sole annotator, data should be compared over time to ensure
that consistency and objectivity remain.
Several morphokinetic variables are at risk of subjective interpretation. The appearance of pronuclei and
initiation of compaction are just two examples. Thorough training and the use of reference images and audit
should be used to ensure annotation quality.
The Challenges of Annotation

Many of the morphokinetic variables within Table 12.1 (see also Figures 12.1 through 12.4) can readily be iden-
tified on studying the embryo’s time-lapse images, whereas others can cause great discussion and debate. The
current lack of an industry standard makes comparing practice and data difficult. An example of one annotation
challenge may be fragmentation.
Fragmentation is a poorly understood but a very common and dynamic feature of human preimplantation
IVF embryos that has been linked to aneuploidy [10,11]. Time lapse provides an opportunity for the dynamics
of fragmentation to be studied in greater detail and has recently been used in a study by Chavez et al. [12] that
suggested that most fragmented early cleavage-stage embryos were aneuploid. More recently, a preliminary
study by Montgomery et al. [13] has shown an association between the timing and completion of compaction in
fragmented embryos, and blastocyst ploidy.
Fragmented embryos with extended periods of compaction were significantly more likely to give rise to aneu-
ploid blastocysts than embryos that completed compaction within 22 hrs of ICSI. Conventionally, fragmentation
has been reported by an estimation of the proportion of the embryo affected, and ranges are used rather than
specific values [5,6]. To share data and experience with other time-lapse practitioners, surrounding this common
feature of human embryos, when annotating fragmentation, it is recommended that a percentage value and the
number of cells at the time point of recording are used. There is currently no recommendation for the definition
or standard recording of patterning or dynamic movement of fragmentation within the embryo, but there is a
desire for such guidelines to be developed.
Outcome Measures
Successful live birth is the ultimate outcome after IVF treatment, and despite the limitations of any outcome
measure, including live birth, this should be the primary outcome measure when considering the potential of a
developing embryo.
TABLE 12.1
Summary of Morphokinetic Variables and Proposed Definitions
Description
Morphokinetic Variables
Time (t)
t0 IVF or midtime of ICSI/IMSI
tPB2 The second polar body is completely detached from the ooplasm
tnPN Fertilization status is confirmed
(tPN1a) The first pronucleus is first visible
(tPN2a) The second pronucleus is first visible
tPNf All pronuclei have faded (see Figure 12.1)
t2-t9 Two (see Figure 12.2) to nine sequential, distinguished cells are present
tSC The first two cells merge; initiation of compaction observed (see Figure 12.3)
tMx/w Morula is formed or compaction goes no further; “x” corresponds to fully compacted, and “w” corresponds
to partially compacted or cells excluded
tSB The first sign of a cavity is observed as blastulation begins (see Figure 12.4)
tByz Full blastocyst stage is reached; the last frame before the zona pellucida starts to thin; “y” corresponds to
morphology of inner cell mass cells, and “z” corresponds to trophectoderm cells (see Figure 12.4)
tEyz Initiation of expansion is confirmed; the zona pellucida starts to thin
tHNyz Extrusion of cells from the zona pellucida is present
tHDyz Blastocyst is fully hatched from the zona pellucida
Calculated Variables
VP tPNf-tPN1a (period of visible pronuclei)
Cell Cycle
CC1 t2-tPB2
The end of the second meiosis to the formation of two discrete cells
CC2 The time for a two-cell embryo to form a four-cell embryo
The two blastomeres (a and b) can be considered individually
CC2a = t3-t2
CC2b = t4-t2
CC3 The time for a four-cell embryo to form an eight-cell embryo
The four blastomeres can be considered individually
CC3a = t5-t4
CC3b = t6-t4
CC3c = t7-t4
CC3d = t8-t4
Synchronization
S2 The duration of the transition from two sister cells, each dividing to reach the four-cell stage
t4-t3
S3 As above, but from four to eight cells
t8-t5
Duration of Compaction (Morula Stage)

tMx-tSC Full compaction
tMy-tSC Partial compaction
Blastocyst Stage
tHN-tSB Duration of blastulation
Note: Each time point defines the time-lapse frame in which the phenomena described are first observed or recorded.
FIGURE 12.1 Sequential time lapse images of fading pronuclei. Image (c) is tPNf.
FIGURE 12.2 Sequential time lapse image of the first mitosis. Image (c) shows t2.
FIGURE 12.3 Initiation of compaction (tSC). More clearly seen with multiple focal planes. Image (c) shows tSC.
FIGURE 12.4 Initiation of blastulation (tSB). Images (b) and (c) show the increasing cavity and image (d) shows tB.
Due to the relatively recent introduction of time-lapse technology for IVF, the length of the gestational period,
and the time required to acquire and collate obstetric outcome data from patients, alternative and less robust
outcome measures have been used in published time-lapse studies. Blastulation, although of great scientific
interest, has limited clinical value due to the high incidence of blastocysts that fail to implant or are a neuploid [8].
Clinical pregnancy, defined by the presence of a fetal heart on ultrasound scan, has also been used, but due to the
early nature of these reported pregnancies, some pregnancies may be lost before term [14]. The difficulty remains
that failed pregnancies or even failed implantations cannot necessarily always be attributed to the embryo;
however, in terms of outcome measure, live birth arguably remains the most reliable indicator of the viability
potential of the preimplantation embryo for IVF and time-lapse practitioners [15].
KID refers to transferred embryos with a known outcome.
What is commonly referred to as “known implantation data,” or KID, are the morphokinetic data of a specific
and transferred embryo that has a known outcome, with the outcome being either a negative pregnancy test,
a gestational sac or fetal heart on ultrasound scan (at 6–8 weeks’, gestation), or a live birth. Figure 12.5 describes
Single embryo Double embryo

transfer transfer
One embryo One embryo Two embryos Two embryos No KID**

1 × KID 1 × KID 2 × KID 2 × KID
Positive Negative Negative Positive*
Pregnancy
loss × 1
Pregnancy
Pregnancy loss × 2
loss
One live birth No live birth Two live birth No live birth No live birth
1 × KID(LB) 1 × KID(LB) 2 × KID(LB) No KID(LB) 2 × KID(LB)
Positive Negative Positive Negative
FIGURE 12.5 Known implantation data (KID). KID is the morphokinetic data of a specific, transferred embryo that has a known
outcome. *Zygoticity of a twin pregnancy, following double embryo transfer, cannot be ascertained without genetic fingerprinting.
Due to relatively low incidence of dizygotic twinning, practitioners, accepting this limitation, may include these data in KID analy-
ses. **As implantation data cannot be deduced or used following a double embryo transfer resulting in a single implantation or birth,
the KID ratio, or rate, is lower than the implantation rate per embryo transferred, commonly used for IVF data analyses.
how morphokinetic data may be used for analysis after embryo transfer. Data can be compared between embryos
giving positive or negative implantation data (KID+ or KID–, respectively). All data can be used after a single
embryo transfer, or a double embryo transfer with a negative outcome. Using data after multiple embryo transfer
that has resulted in the same number of fetal hearts or babies born may be problematic without the use of genetic
fingerprinting to ascertain the chrorionicity or zygoticity of the pregnancies. However, due to the very low inci-
dence of monozygotic twinning, most analyses could justify the inclusion of these data.
Data Collection and Analysis

Data can be exported from the time-lapse devices for assessment and analysis, or they can be processed using
integrated software tools. Provided the data are quality assured and complete, meaning that all relevant mor-
phokinetic variables have been recorded according to standard policy, where they occurred, and that outcome
of transferred embryos has been updated, there exists a powerful tool for retrospective analysis. The data should
be carefully divided into groups for comparison and statistical analysis. Due to the ranges observed for each
morphokinetic variable, it is recommended that median values are used as opposed to means. This way, extreme
high or low outliers do not introduce a skew.
KID rates (or ratios) can be calculated for each significant variable for exclusion (deselection) and selection
criteria to be identified and then used to develop embryo selection algorithms or models.
KID rates are calculated using the following formula: KID+/KID+ plus KID– × 100%.
Regular data review is recommended to continually improve embryo selection algorithms.
Selection and Deselection Criteria and Algorithms for Embryo Selection

Several time-lapse studies have linked kinetic markers to embryo viability. In a retrospective analysis of
EmbryoScope-acquired time-lapse human embryo data, a significant association was previously demonstrated
between the timing of pronuclear fading and the first three cleavage events, and successful implantation [16].
Another study, looking at early embryo development, showed an inverse relationship between the ability of
embryos to develop to the blastocyst stage and the length of time for zygote division [17].
Time lapse identifies aberrant cleavage patterns.
More recently, a study by Rubio and colleagues [18] focused on the phenomenon of direct, or rapid, cleavage
from a single cell to three cells in <5 hr. This study demonstrated the ability of time lapse to identify aberrant
cleavage divisions and the reduced implantation potential when such embryos were transferred compared with
embryos that did not exhibit this behavior. In a cohort of 1659 transferred embryos, the incidence of this “direct
division” was 13.7%, and the known implantation rate of these embryos was statistically significantly lower than
for embryos with a normal cleavage pattern (1.2% vs. 20.2%, respectively). Figure 12.6 shows a series of time-
lapse images demonstrating this phenomenon within 0.5 hr.
This consecutive series of four time-lapse images, taken at about 10 min intervals, demonstrates the phe-
nomenon of direct cleavage from one to three cells and would not be observed without time lapse and has been
reported to be associated with reduced implantation potential [18].
Several embryo selection models, or algorithms, indicating the added value of using morphokinetic information
when selecting embryos have now been published. The first by Wong and colleagues [8] identified three significant
morphokinetic variables associated with the likelihood of blastulation. These variables and values (in parentheses)
were P1, the duration of the first cytokinesis (within 33 min); P2, the interval between the first and the second cyto-
kineses, that is, the total length of time at the two-cell stage (7.8–14.3 hr); and P3, the time between the second and
third mitoses, that is, the total length of time at the three-cell stage or the synchrony of cleavage of the first two cells
(within 5.8 hr). This work was used to develop the Eeva test (early embryo viability assessment) (Auxogyn).
FIGURE 12.6 Rapid or direct cleavage from one to three cells.
The work of Meseguer et al. [9] used the EmbryoScope and looked beyond blastulation to implantation and
pregnancy rates. By studying 247 transferred embryos with known implantation outcome, optimal ranges for
specific morphokinetic variables were defined. By analyzing multiple preimplantation embryo developmen-
tal milestones and durations, a hierarchical model to classify embryos, according to the most significant
morphokinetic variables, was developed. The timing of the cleavage to five cells (t5) in this study showed the
highest correlation with positive implantation. This model first introduced exclusion criteria. These exclusion
criteria were uneven blastomeres after the first cytokinesis, rapid or direct cleavage from one to three cells,
and multinucleation at the four-cell stage. After the exclusion of embryos fulfilling these criteria, embryos
were classified according to the timings of similar variables proposed by Wong et al. (P2 and P3 above),
referred to as cc2 (within 11.9 hr) and S2 (within 0.76 hr), as well as t5 (48.8–56.6 hr post-intracytoplasmic
sperm injection [ICSI]). Despite there being some similarities in the morphokinetic variables used within
these published models, the timings differ and caution should be exercised when considering applying such
models, developed in a different research or clinical setting, before robust validation is performed
(as d iscussed below). It has been demonstrated that a published embryo selection model from one setting may
not always effectively be transferred to another setting, but it may be useful with some modification, and
after a validation process [19].
Time-lapse–based embryo selection models indicate the value of applied morphokinetics, but care
has to be taken while transferring models from one clinical setting to another.
More recently, an aneuploidy risk classification model has been published [20]. Also, using the EmbryoScope,
20 morphokinetic variables and durations were recorded for blastocysts that then underwent trophectoderm
biopsy and preimplantation genetic screening (PGS) for all chromosomes using array-comparative genomic
TABLE 12.2
Time-Lapse–Derived Model for Classification of Ploidy with Associated Probabilities
of Aneuploidy
Aneuploidy Risk Class Model n Probability of Aneuploidy (%)
Low tB < 122.9 hr and tSB < 96.2 hr 36 37
Medium tB < 122.9 hr and tSB ≥ 96.2 hr 49 69
High tB ≥ 122.9 hr 12 97
All 97 61
hybridization (CGH) or single-nucleotide polymorphism (SNP) array. Two variables significantly correlated with
embryo ploidy were used to develop an algorithm for embryo selection that classified embryos as having a low,
medium, or high risk of aneuploidy. These variables were the start of blastulation (tSB) and the time to reach
the full blastocyst stage (tB). Risk classifications were defined according to the time an embryo reached these
two developmental milestones. Table 12.2 shows the probability of aneuploidy when embryos were partitioned
according to their morphokinetic values for the two significant variables.
This model was later tested retrospectively, by the same researchers, on transferred blastocysts with known
clinical outcome that had not undergone biopsy and PGS. Significant relative increases in positive fetal heart
and live birth rates were demonstrated when an embryo retrospectively classified as low risk was transferred,
compared with the overall rates after blastocyst transfer, indicating the potential clinical applicability of this
noninvasive time-lapse algorithm for embryo selection.
Practitioners working with selection models where embryos are deselected or excluded should consider the
supporting data to help with the decision process regarding the fate of such embryos. Considerations should be
made as to whether the morphokinetic phenomena or timings observed during a particular embryo’s develop-
ment, deeming it deselected for transfer, preclude implantation entirely or are associated with reduced viability
or potential. An embryo identified as having a high risk of aneuploidy or low potential to blastulate by a model,
for example, may be the highest ranking embryo within a cohort and still give the patient a chance of a positive
outcome, albeit small.
Transferability of Embryo Selection Algorithms

Whether such embryo selection algorithms can be directly transferrable between clinics remains to be demon-
strated but based on reports that rates of embryo development can differ according to intrinsic or extrinsic factors,
it is likely that they may not. Either way, it should be preferable for clinics to develop their own models for their
specific patient populations, stimulation regimens, culture conditions, and according to their interpretation of
time-lapse images.
Clinics introducing time-lapse methodologies are advised to do so with strict and standard practice through-
out the process. Until we know the potential impact of subtle deviations from protocol, culture dishes or slides
should be prepared in a standard and precise manner, and time-lapse images, where performed manually, should
be assessed objectively and observations recorded (annotated) in the same way by all practitioners. Only this
will allow centers to collate robust and complete data to develop in-house embryo selection models that can be
fine-tuned as experience and data amasses.
Confounding Factors
There have been several reports on the impact, on embryo morphokinetics, of compounding factors that may be
patient or clinic specific. Examples are gas composition during in vitro culture, age, female body mass index,
and culture media [14,21–23].
A recent study, however, reported that morphokinetic parameters used for embryo selection were not affected
between two different culture media analyzed [24]. Over the next few years, such questions surrounding confounding
factors and those crucially asking whether algorithms for embryo selection are transferable will likely be a key focus
for time-lapse study in the field of IVF. To make the most effective progress in this field, embryologists are urged to
work together, sharing best and common practice to enable this shift in practice to take place most effectively.
Change Control
Due to the possible impact of intrinsic and extrinsic factors on morphokinetic timings, great care should be taken
when a change to practice is being considered or made when using time-lapse systems. The impact of a control-
lable factor such as a change in media or plasticware, for example, on the precise timings of embryo develop-
ment, should be understood and validated before full implementation, particularly if embryo selection models
have been developed under specific conditions. As with all changes made in IVF laboratories, these changes
should be justified, controlled, validated, and evaluated but now with the additional consideration of potential
impact on morphokinetics, when using time-lapse imaging.
A Tool or a Rule? The Role of the Embryologist

As a clinical treatment, we are still in the relatively early stages of clinical implementation of time-lapse
technology. We remain unclear as to whether published algorithms, developed using particular incubation
conditions (such as gas mix and culture media), can be directly transferred between centers using not only
alternative incubation but also potentially differing definitions and operating procedures for recording dynamic
observations of embryo development. Although the expectation is that, eventually, agreement will be reached as
to the most significant indicators of an embryo’s potential to develop through to live birth, the specific timings
may differ according to the variation in, for example, patient history, age, or incubation conditions, and this could
result in clinics having numerous algorithms that can be applied for the purpose of embryo selection, according
to these factors. In time, and with increasing data and experience, the optimal ranges for defined dynamic events
such as the interval between the first and second cytokineses may be further fine-tuned, and additional novel
morphokinetic markers of embryo viability will be identified.
The role of the embryologist remains key, despite the most sophisticated algorithm.
The role of the embryologist remains key. Even with sophisticated morphokinetic selection algorithms, the
embryologist may need to overrule the algorithm should an embryo reaching all of the milestones within the
optimal time ranges also have features considered detrimental, such as smooth endoplasmic reticulum clusters,
large areas of cellular degeneration, or late-onset developmental arrest. The use of bright-field time-lapse devices
provides embryologists with images already familiar to them and allows this facilitative interaction between the
digital and the human eye.
Acknowledgments
I am grateful for permission to use and adapt the table of proposed definitions, currently being drafted for sub-
mission by The Embryo Morphokinetic Consensus Group; Inge Agerholm, Jesus Aguilar, Sandrine Chamayou,
Nadir Ciray, Marga Esbert, Shabana Sayed, and Alison Campbell.
I thank Louise Kellam for assistance with image preparation for this chapter and my CARE Fertility
colleagues.
REFERENCES
1. Payne D, Flaherty SP, Barry MF, et al. Preliminary observations on polar body extrusion and pronuclear forma-
tion in human oocytes using time-lapse video cinematography. Hum Reprod 1997; 12: 532–541.
2. Pribenszky C, Matyas S, Kovacs P, et al. Pregnancy achieved by transfer of a single blastocyst selected by time-
lapse monitoring. Reprod Biomed Online 2010; 21: 533–536.
3. Cutting R, Morroll D, Roberts SA, et al. Elective single embryo transfer guidelines for practice British Fertility
Society and Association of Clinical Embryologists. Hum Fertil 2008; 11: 131–146.
4. Fisch J, Rodriguez H, Ross R, et al. The graduated embryo score predicts blastocyst formation and pregnancy rate
from cleavage stage embryos. Hum Reprod 2001; 16: 1970–1975.
5. Alpha Scientists in Reproductive Medicine, ESHRE Special Interest Group of Embryology. The Istanbul consen-
sus workshop on embryo assessment: Proceedings of an expert meeting. Hum Reprod 2011; 26: 1270–1283.
6. Alpha Scientists in Reproductive Medicine, ESHRE Special Interest Group of Embryology. The Istanbul con-
sensus workshop on embryo assessment: Proceedings of an expert meeting. Reprod Biomed Online 2011; 22:
632–646.
7. Van Royen E, Magelschots K, Vercruyssen M, et al. Multinucleation in cleavage stage embryos. Hum Reprod
2003; 18(5): 1062–1069.
8. Wong CC, Loewke KE, Bossert NL, et al. Non-invasive imaging of human embryos before embryonic genome
activation predicts development to the blastocyst stage. Nat Biotechnol 2010; 28(10): 1115–1121.
9. Meseguer M, Herrero J, Tejera A, et al. The use of morphokinetics as a predictor of embryo implantation. Hum
Reprod 2011; 26(10): 2658–2671.
10. Munne S, Cohen J. Chromosome abnormalities in human embryos. Hum Reprod Update 1998; 4: 842–855.
11. Alikani M, Calderon G, Tomkin G, et al. Cleavage anomalies in early human embryos and survival after pro-
longed culture in vitro. Hum Reprod 2000; 15: 2634–2643.
12. Chavez SL, Loewke KE, Han J, et al. Dynamic blastomere behaviour reflects human embryo ploidy by the four-
cell stage. Nat Commun 2012; 3: 1251.
13. Montgomery S, Duffy S, Bowman N, et al. Does the duration of compaction differ in fragmented cleavage stage
embryos that ultimately become euploid or aneuploidy blastocysts? Is this reflected in the timing of blastulation?
ESHRE, London. Hum Reprod 2013; 28(Suppl 1): i1–i4.
14. Meseguer M, Rubio I, Cruz M, et al. Embryo Incubation and selection in a time-lapse monitoring system
improves pregnancy outcome compared with a standard incubator: A retrospective cohort study. Fertil Steril
2012; 98: 1481–1489.
15. Campbell A, Fishel S, Bowman N, et al. Retrospective analysis of outcomes after IVF using an aneuploidy risk
model derived from time-lapse imaging without PGS. Reprod Biomed Online 2013; 27: 140–146.
16. Herrero J, Alberto T, Ramsing NB, et al. Linking successful implantation with the exact timing of cell division
events obtained by time-lapse system in the EmbryoScope. Fertil Steril 2010; 94(4): S149.
17. Cruz M, Perez-Cano I, Gadea B, et al. Time-lapse video analysis provides a correlation between early embryo
division kinetics and subsequent blastocyst formation and quality. Hum Reprod 2011; 26: P-115.
18. Rubio I, Kuhlmann R, Agerholm I, et al. Limited implantation success of direct-cleaved human zygotes: A time-
lapse study. Fertil Steril 2012; 98: 1458–1463.
19. Best L, Campbell A, Duffy S, et al. Session 57: Does one model fit all? Testing a published embryo selection
algorithm on independent time-lapse data. Hum Reprod 2013; 28: i87–i90.
20. Campbell A, Fishel S, Bowman N, et al. Modelling a risk classification of aneuploidy in human embryos using
non-invasive morphokinetics. Reprod Biomed Online 2013; 26: 477–485.
21. Liebenthron J, Montag M, Koster M, et al. Influence of age and AMH on early embryo development realised by
time-lapse imaging. Hum Reprod 2012; 27: P-135.
22. Bellver J, Mifsud A, Grau N, et al. Similar morphokinetic patterns in embryos derived from obese and normo-
weight infertile women: A time-lapse study. Hum Reprod 2013; 28: 794–800.
23. Ciray HN, Aksoy T, Goktas C, et al. Time-lapse evaluation of human embryo development in single versus
sequential culture media-a sibling oocyte study. J Assist Reprod Genet 2012; 29: 891–900.
24. Basile N, Morbeck D, García-Velasco J, et al. Type of culture media does not affect embryo kinetics: A time-lapse
analysis of sibling oocytes. Hum Reprod 2013; 28: 634–641.
13
Noninvasive Techniques: Embryo Selection by
Transcriptomics, Proteomics, and Metabolomics
Introduction
The number of multiple births has increased >70% since 1980 [1], when approximately 40% of twin births and
80% of triplet and higher order births resulted from infertility therapy [2]. Multiple embryo transfers in in vitro
fertilization (IVF) treatment particularly elevated the frequency of multiple pregnancies in this period. Although
the desired outcome of infertility treatment is the birth of a single healthy infant, more than one embryo has
commonly been transferred in IVF cycles to increase the likelihood of successful outcome. Subsequent multiple
gestations, however, can lead to serious maternal and fetal health problems [3]. The overall cost of healthcare
services provided for multiple pregnancies and resulting preterm births is estimated to exceed the cost of IVF
itself [4–6]. Medical and financial complications associated with IVF multiple pregnancies became much more
important as the prevalence of IVF treatment increased significantly within the past two decades.
To prevent potential negative implications of infertility treatment, regulatory legislation has been enacted in
many countries to set limits on the number of embryos transferred [7]. In the meantime, investigators focused
on selection of the most viable gametes and embryos in an attempt to reduce multiple pregnancies without com-
promising the pregnancy rates. Eventually, the average number of embryos transferred in nondonor IVF cycles
during 2001–2010 has decreased from 3.1 to 2.3 in United States [8] and from 2.2 to 1.7 in United Kingdom [9].
However, elective single embryo transfers still constitute a minor portion of the transfer cycles because current
ability to select the single most developmentally competent embryo is less than perfect.
Embryo Selection: From Morphology to Omics

Soon after the birth of the first IVF baby, controlled ovarian stimulation protocols provided maturation of m ultiple
oocytes in a cycle that, in turn, enabled availability of multiple embryos for transfer. Since then, selection of the
embryo(s) to be transferred has been a critical step affecting the success of the IVF treatment. From the early
times of IVF to the present, embryo assessment strategies have mainly been based on timely morphological
observations of embryos throughout the culture period. Morphology-based embryo selection has practical advan-
tages in the clinical routine and has resulted in significant improvements in pregnancy rates [10]. The efficiency
of morphological assessment has recently been further improved with the introduction of novel technologies that
allow continuous monitoring of the embryo development in vitro and selection of embryos based on additional
dynamic morphological parameters [11,12]. Nevertheless, the precision of morphological criteria remains less
than desired, motivating investigators to pursue additional markers of embryo viability at the molecular level.
The molecular mechanisms underlying distinct biological conditions are mainly characterized by process-
specific gene expression. Genes are expressed through transcription of the genetic code stored in DNA into
RNAs and subsequent translation of messenger RNAs (mRNAs) into proteins. The human genome contains
3 × 109 base pair of nucleotides and approximately 30,000 genes. These genes generate ~200,000 transcripts
and ~1 million proteins. In addition, human metabolome is estimated to have ~3000 metabolites, the small mol-
ecules (<1 kDa) including amino acids, lipids, nucleotides, and signaling molecules. The DNA, RNA, protein,
191
DNA RNA Protein Metabolites
Transcription Translation Metabolism
~3×109 bp nucleotides mRNA, tRNA, rRNA, miRNA Small molecules (<1 kDa)
~1,000,000 proteins
~30,000 genes ~200,000 transcipts ~3000 metabolites
Genomics Transcriptomics Proteomics Metabolomics
FIGURE 13.1 Omics approaches investigate the molecular constitution of biological samples at genomics, transcriptomics,
proteomics, and metabolomics levels.
and metabolite constitutions of organisms are investigated by genomics, transcriptomics, proteomics, and
metabolomics approaches, respectively (Figure 13.1). Omics research basically looks for answers to two ques-
tions: what distinct molecules and how many copies of each molecule exist in a biological sample? Within the
past decade, the omics technologies have been used widely as potential novel embryo assessment tools in assisted
reproduction (for review, see [13]).
The invasive genomic approach to embryo assessment, referred to as preimplantation genetic screening (PGS),
allows the exclusion of aneuploid embryos through genetic profiling of biopsied cells. Although a recent meta-
analysis showed that PGS with fluorescence in situ hybridization (FISH) is associated with lower pregnancy and
live birth rates [14], PGS using more recent comparative genomic hybridization (CGH)-array or single nucleotide
polymorphism (SNP)-array technologies has shown promising results [15,16].
Beyond the oocyte and embryo genome, RNA constitution of follicular somatic cells and protein and metabolite
content of spent embryo culture media serve as potential indicators of embryo viability. This chapter covers an
overview of the techniques and clinical applications of noninvasive transcriptomics, proteomics, and metabolo-
mics approaches to embryo assessment.
Omics research basically looks for answers to two questions: what distinct molecules and how
many copies of each molecule exist in a biological sample?
Embryo Selection by Transcriptomics

The term transcriptome refers to the entire set of RNA molecules in a particular cell or tissue, and includes
mRNAs, transfer RNAs (tRNAs), ribosomal RNAs (rRNAs), microRNAs (miRNAs), and other noncoding
RNAs. Specifically, the relative concentrations of individual mRNAs can be evaluated as an approximation to
expression levels of the corresponding genes. Within the context of IVF, cumulus/granulosa cells transcriptome
has been investigated widely as a noninvasive tool to assess oocyte quality as an indicator of embryo viability
(for review, see [17]).
Folliculogenesis and Oocyte–Somatic Cell Interactions

The oocyte and its surrounding somatic cells grow and develop in a coordinated and mutually dependent
manner [18]. At the initial stage of folliculogenesis, a single layer of flattened granulosa cells surrounds the
perinatally formed primordial follicles. The oocyte in the primordial follicle remains arrested at the prophase
of the first meiotic division until shortly before ovulation. Starting with puberty, selected primordial follicles
grow and differentiate into primary follicles, where granulosa cells surrounding the oocyte become cuboidal.
Embryo Selection by Transcriptomics, Proteomics, and Metabolomics 193
The zona pellucida, a glycoprotein polymer capsule that separates the primary oocyte from the follicular cells,
also d evelops at this stage. As follicles progress to the secondary stage, granulosa cells undergo mitosis, forming
multiple layers around the zona pellucida. Subsequent early antral follicles are characterized by the formation
and growth of fluid-filled cavities (antrum) adjacent to the oocyte.
Primordial, primary, and secondary stages of folliculogenesis occur independently of gonadotropic hormones,
whereas follicular-stimulating hormone (FSH) secreted by the pituitary stimulates the transition from early
antral to antral follicle stage [19]. In the human ovary, antral follicles compete with each other under the influ-
ence of FSH, and the majority of follicles that have fewer FSH receptors stop developing further and undergo
atresia. Eventually, only one dominant follicle becomes the preovulatory follicle. In the preovulatory follicle,
granulosa cells differentiate into two distinct cell types: the mural granulosa cells, lining the antrum and the fol-
licle wall; and cumulus cells, immediately surrounding the oocyte. At this stage, luteinizing hormone (LH) surge
initiates oocyte maturation and cumulus expansion occurs by the synthesis of hyaluronic acid within the cumulus
mass. Synthesized hyaluronic acid is deposited into the extracellular matrix that binds to oocyte and cumulus
cells together. The oocyte resumes meiosis and the ultimate mature cumulus–oocyte complex (COC) contains an
oocyte arrested at the metaphase of the second meiotic division (MII) that is ready for ovulation and fertilization.
During folliculogenesis and oocyte maturation, the oocyte derives most of its substrates for energy metab-
olism and biosynthesis from the surrounding somatic cells [20,21]. The highly specialized cumulus cells
within the COCs have transzonal cytoplasmic processes that penetrate through zona pellucida and form gap
junctions [22]. Cumulus cells communicate with each other and with the oocyte through the gap junctions
that allow bidirectional transfer of metabolites and signaling molecules [23]. The transfer of paracrine fac-
tors between the oocyte and the surrounding cumulus cells is of special importance for oocyte maturation.
Initial observations on oocyte–cumulus cell interaction in relation to embryo viability demonstrated that
oocytes obtained from follicles with impaired cumulus expansion have limited potential for implantation [24].
Consequently, cumulus/granulosa cell transcriptome has been analyzed in an attempt to identify potential bio-
markers of embryo competence.
Technologies for Transcriptomic Analysis of Cumulus and Granulosa Cells

The common initial step in transcriptomics experiments is sample preparation that consists of the stages of cell
and tissue collection; RNA isolation; concentration, purity, and integrity assessment; and reverse transcription
of the RNA to complementary DNA (cDNA) molecules. Once the sample quality is ensured, quantification
of relative mRNA levels is performed using either individual gene expression analysis tools or whole-genome
transcription profiling technologies. This section provides an overview of the sample preparation methods and
analytical techniques used for investigation of relationship between cumulus/granulosa cell transcriptome and
oocyte/embryo competence.
Sample Preparation
Sampling of somatic components of the follicle poses specific challenges due to potential cross-contaminations
or contamination with blood. Mechanical separation of COCs from contaminating cells is easier than that of
granulosa cells. In addition, the effect of regional differences in the cumulus physiology can be minimized using
laser capture technology. Granulosa cells need to be carefully separated from leukocytes because potential
contamination with blood during follicular aspiration may alter the gene expression profile.
RNA must be purified immediately after tissue/cell isolation to prevent potential RNA degradation that would
result in a rapid change in the transcript profile. RNases must be inactivated when the samples need to be stored
before RNA isolation. After RNA isolation, concentration, purity, and integrity of RNA molecules must be con-
firmed using UV spectroscopic analyses [25], gel-based methods [26], or microfluidics technologies [27]. Then,
mRNA molecules are reverse transcribed to cDNA before amplification and quantification. cDNA synthesis
occurs by priming the total RNA with either one of the oligo(dT) random hexamers and gene-specific primers
or a blend of these primers. Priming strategy has been shown to be an important factor affecting the results of
mRNA quantification [28].
qRT-PCR and High-Throughput Transcriptomics

Since the discovery of the thermostable Thermophilus aquaticus (Taq) polymerase [29], polymerase chain
reaction (PCR) has been used as an efficient and practical tool to detect the existence of a particular DNA
(or cDNA) fragment in a biological sample. The PCR method allows exponential amplification of the target
sequence by using a pair of primers, deoxynucleotides, and a polymerase enzyme. Each of the consecutive
amplification cycles in PCR consists of three steps performed at specific temperatures: denaturation of double
strands (~94°C), primer annealing to complementary parts on both strands (~65°C to 55°C), and primer exten-
sion (~72°C). The amount of starting material is doubled at the end of each cycle. After ~25 to 40 cycles, the
resulting PCR product is analyzed on agarose gel containing a fluorescent DNA stain.
Transcriptomics studies make use of PCR after reverse transcription of mRNA molecules to cDNA. The
combined method is called reverse transcription-polymerase chain reaction (RT-PCR) and is widely used for
gene expression analysis. Because agarose gel–based analysis of RT-PCR products do not provide precise
quantitative information, the quantitative reverse transcription-polymerase chain reaction (qRT-PCR), or the
so-called real-time RT-PCR, has been developed for continuous measurement of PCR amplification, allowing
reliable quantification of specific transcripts in a sample [30,31]. Currently, qRT-PCR is commonly used for
quantitative gene e xpression analysis.
To account for potential variations between PCR reactions, gene expression results need to be normalized using
endogenous controls. The most common control genes are beta-actin (β-Actin) or glyceraldehyde 3-phosphate
dehydrogenase (GAPDH). Alternatively, mathematical models such as NormFinder [32] and GeNorm [33] exist that
help with selection of the most suitable normalization genes among a set of candidate genes.
Within the past decade, gene expression research evolved from single-gene analysis to genome-wide
transcription profiling with the advance of microarray technology. A microarray chip consists of single-stranded,
~25 base-length polynucleotide probes that are synthesized in fixed positions in the form of a two-dimensional
array on a solid surface. Each probe is attached to a microscopic spot on the array and corresponds to the base
sequence of a specific mRNA. The large number of probes on a single microarray chip enables simultaneous
expression analysis for all known genes.
The microarray experimental procedure consists of labeling, hybridization, scanning, and data analysis stages
(Figure 13.2). First, total RNA is isolated and then cDNA is synthesized by reverse transcription of mRNA.
Amplified and biotin-labeled cRNA is obtained through in vitro transcription. Then, fragmented cRNAs are
deposited over the array, allowing for hybridization of the labeled targets to the complementary probes. The
amount of hybridization at each probe is expected to be proportional to the level of expression of the gene repre-
sented by that probe. Subsequently, the microarray chip is illuminated by a laser light. Gene expression is then
quantified by means of fluorescence intensity that is captured by the scanners into an image.
Cumulus/granulosa Isolated Reverse transcribed, labeled, Hybridization to Image scanning and Differential
cells RNA and amplified cDNA microarray chip quality control expression analysis
FIGURE 13.2 Major steps of a typical microarray experiment. Total RNA is extracted from the follicular cell samples, and mRNA
is reverse transcribed into cDNA. Amplified and labeled samples are hybridized to a microarray slide, allowing the labeled targets
to bind to their complementary oligonucleotides attached to the microscopic probes. The array is then washed and scanned to obtain
the fluorescent image that is further processed to get the intensity values for differential expression analysis.
Recently, next-generation sequencing technologies have provided an alternative transcriptomics approach

called RNA Sequencing (RNA-Seq) that appears to be a more precise detection and quantification of RNA
transcripts [34,35]. The RNA-Seq procedure starts with the construction of a cDNA library from isolated RNA
molecules. Long RNA molecules, such as mRNAs, need to be fragmented into smaller pieces (by hydrolysis or
nebulization) to be compatible with the sequencing technology. The reverse-transcribed cDNA can also be frag-
mented by DNAse I treatment or sonication. Alternatively, recent direct RNA sequencing technology eliminates
the need for prior conversion of RNA to cDNA [36].
By using the cDNA library constructed, short sequence reads (30–400 bp) are obtained by high-throughput
sequencing of each molecule either from one end (single-end sequencing) or both ends (pair-end sequencing).
The resulting reads are mapped to a reference genome or assembled de novo without a priori knowledge of the
underlying genome. Each sequence read may correspond to a known exon, a splice variant, or a new candidate
gene. The number of sequence reads that map to a particular transcript is expected to be proportional to its
expression level. Hence, read counts are used for quantification of gene expression in RNA-Seq experiments.
Gene expression research evolved from single-gene analysis to genome-wide transcription

rofiling with the advance of microarray technology; however, next-generation sequencing
p
technologies provide an alternative transciptomics approach called RNA sequencing that appears
to be a more precise detection and quantification of RNA transcripts.
Typical RNA-Seq protocols require at least 1 μg of total RNA. However, this amount of starting material
may not be available when studying small tissues such as oocytes. Recently, Antoniou and Taft [37] provided a
protocol, specifically for Illumina, to prepare sequencing libraries from mouse oocytes and suggested that this
protocol can also be used for follicular cells. Unfortunately, to date there are no publications applying RNA-Seq
approach for analysis of follicular cells in the context of assisted reproduction.
Analysis of the data produced by both microarray and RNA-Seq technologies poses significant challenges
because expression levels of thousands of genes are investigated simultaneously in a multistage experimental set-
ting. Considering the underlying technological complexities, several bioinformatics methods have been proposed
to provide reliable and efficient data analysis in transcriptomics studies.
Analysis of Transcriptomics Data

The raw microarray data obtained from the image files needs to be processed before differential expression
analysis. The commonly applied robust multichip average (RMA) preprocessing method consists of background
adjustment, quantile normalization, and probe set summarization stages [38–40]. Preprocessed data are then
analyzed for differential expression. False discovery rate (FDR) corrected statistical analysis is proposed to
prevent excessive Type I error (false-positive rate) during simultaneous testing of the null hypothesis (i.e., the
expression levels are not different between the two experimental conditions) for each gene [41,42]. To conclude
that a gene is differentially expressed, the data analysis should also consider biological significance (fold change
rate) in addition to statistical significance (FDR corrected p-value). Commercial tools such as Partek Genomic
Suite and Bioinformatics toolbox of Matlab exist for differential expression analysis of microarray data.
With regard to RNA-Seq data analysis, the depth of sequencing is a major concern because transcriptome
coverage strongly depends on the sequencing depth. A small amount of sequencing data would be sufficient for
the highly expressed genes where more reads are required for accurate quantification of low abundance targets.
Analysis of RNA-Seq data requires advanced bioinformatics approaches for alignment of the reads to an existing
genome or for assembly of the reads into a new genome. Despite the challenges, RNA-Seq has the advantage of
being able to detect tissue-specific and currently unknown splice variants and polymorphisms.
The results of differential expression analysis of both microarray and RNA-Seq data may (1) reveal no differ-
ence between the populations of interest; (2) identify just a few genes up regulated or down regulated in either
population; or (3) determine hundreds of differentially expressed genes requiring more sophisticated analyses
for biological interpretation, that is, pathway analysis of microarray data.
Clinical Applications of Transcriptomics-Based Embryo Assessment

qRT-PCR and microarray analysis of cumulus and granulosa cell mRNAs have been applied to identify biomark-
ers associated with oocyte and embryo development and cycle outcome in women undergoing IVF. Studies using
qRT-PCR are hypothesis-driven in nature. These studies propose that somatic cells surrounding viable oocytes
have a different expression pattern for genes previously shown to be important for COC development, compared
with cumulus and granulosa cells around nonviable oocytes. Many of these studies commonly investigated
the genes that are up regulated in cumulus or granulosa cells in response to LH stimulation, such as e pidermal
growth factor (EGF)-like factors (amphiregulin [AREG], epiregulin [EREG], and betacellulin [BTC]) and their
downstream regulators (prostaglandin synthase-2 [PTGS2], also termed cyclooxygenase-2 [COX2], tumor necro-
sis factor alpha-induced protein [TNFAIP6], hyaluronan synthase 2 [HAS2]), that are necessary for synthesis
and stabilization of the extracellular matrix by cumulus cells, cumulus expansion, and subsequent ovulation.
Others evaluated genes that mediate cell division and metabolism.
In the first study using qRT-PCR, McKenzie et al. [43] found HAS2, PTGS2, and GREM1 to be elevated
in cumulus cells of oocytes that lead to high-grade embryos on Day 3. Subsequent studies confirmed the
association between GREM1 expression and Day 3 embryo development [44,45] (Table 13.1). More recently,
Gebhardt et al. [46] reported a significantly higher expression of PTGS2 and VCAN in cumulus cells obtained
from oocytes that yielded a live birth.
In 2011, Wathlet et al. [47] used qRT-PCR to analyze cumulus cells for the expression of eight genes:
Syndecan 4 (SDC4), PTGS2, VCAN, activated leukocyte cell adhesion molecule (ALCAM), GREM1, transient
receptor potential cation channel—subfamily M member 7 (TRPM7), Calmodulin 2 (CALM2), and Inositol
1,4,5-trisphosphate 3-kinase A (ITPKA). Day 3 or Day 5 morphology correlated with all genes studied with the
TABLE 13.1
Potential Biomarker Genes of Oocyte and Embryo Competence Identified by Transcriptomic Analysis of Cumulus/
Granulosa Cells
Transcripta Study Technique Result
PTGS2 (COX2) McKenzie et al. (2004) qRT-PCR (Taqman assay) ↑ PTGS2 ∝ fertilization & Day 3 embryo morphology
Feuerstein et al. (2007) qRT-PCR (SYBR Green) ↑ PTGS2 ∝ PB formation
Anderson et al. (2009) qRT-PCR (SYBR Green) ↑ PTGS2 ∝ oocyte maturation
Gebhardt et al. (2011) qRT-PCR (SYBR Green) ↑ PTGS2 ∝ live birth
Wathlet et al. (2011) qRT-PCR (SYBR Green) ↑ PTGS2 ∝ oocyte maturation
GREM1 McKenzie et al. (2004) qRT-PCR (Taqman assay) ↑ GREM1 ∝ fertilization & Day 3 embryo morphology
Cillo et al. (2007) RT-PCR ↑ GREM1 ∝ fertilization & Day 3 embryo morphology
Anderson et al. (2009) qRT-PCR (SYBR Green) ↑ GREM1 ∝ Day 2–3 embryo morphology
HAS2 McKenzie et al. (2004) qRT-PCR (Taqman assay) ↑ HAS2 ∝ oocyte maturation
Cillo et al. (2007) RT-PCR ↑ HAS2 ∝ fertilization & Day 3 embryo morphology
PTX3 Zhang et al. (2005) Microarray ↓ PTX3 ∝ fertilization & embryo quality
Anderson et al. (2009) qRT-PCR (SYBR Green) ↓ PTX3 ∝ cumulus expansion
TRPM7 Wathlet et al. (2011) qRT-PCR (SYBR Green) ↑ TRPM7 ∝ Day 3 embryo morphology
Wathlet et al. (2012) qRT-PCR (SYBR Green) ↑ TRPM7 ∝ Day 3 embryo morphology
VCAN Gebhardt et al. (2011) qRT-PCR (SYBR Green) ↑ VCAN ∝ live birth
Wathlet et al. (2011) qRT-PCR (SYBR Green) ↓ VCAN ∝ oocyte maturation, ↑ VCAN ∝ pregnancy
RGS2 Feuerstein et al. (2012) Microarray ↑ RGS2 ∝ blastocyst development & pregnancy
Hamel et al. (2010) qRT-PCR ↑ RGS2 ∝ pregnancy
Note: Symbols ↑, ↓, ∝ stands for higher level, lower level, and associated with, respectively. COX2, cyclooxygenase-2; GREM1,
gremlin1; HAS2, hyaluronan synthase 2; PTGS2, prostaglandin synthase-2; PTX3, pentraxin 3; RGS2, regulator of G-protein
signaling 2; TRPM7, transient receptor potential cation channel, subfamily M member 7; VCAN, versican.
a Only the transcripts identified by at least two studies have been listed in the table. All studies, except for Hamel et al. (2010),
analyzed cumulus cells. RGS2, identified as a potential biomarker of pregnancy (Hamel et al. 2010), was studied in granulosa cells.
exception of VCAN. Predictive models were developed using multiple parameters, predicting pregnancy with
a sensitivity of 70% and a specificity of 90%. Better cleavage-stage embryo prediction relied on TRPM7 and
ITPKA expression and pregnancy prediction relied on SDC4 and VCAN expression.
The same group of investigators assessed 11 genes in a subsequent study and found no significant difference in
cumulus cell expression of VCAN among the pregnant versus nonpregnant groups, whereas EphrinB2 (EFNB2)
and ITPKA were up regulated and calcium/calmodulin-dependent protein kinase 1D (CAMK1D) showed the
same trend in the pregnant group [48]. EFNB2, CAMK1D, and Stanniocalcin-2 (STC2) were used for the live
birth prediction model.
An increasing number of unbiased studies used the microarray approach for oocyte viability assessment based
on cumulus/granulosa cell transcriptome. In the first such study, Zhang et al. [49] compared pooled cumulus cells
from oocytes that failed to fertilize with those derived from oocytes that developed into eight-cell embryos on
Day 3, and they identified 160 genes differentially expressed between the two groups. They then applied qRT-
PCR to quantify PTX3 and confirmed the association of PTX3 expression with oocyte development, contradic-
tory to the results reported by Cillo et al. [44].
In another microarray study, van Montfoort et al. [50] compared gene expression in cumulus cells from eight
oocytes resulting in early-cleavage embryos and from eight oocytes resulting in non-early-cleavage embryos. In
total, 611 genes were differentially expressed between the groups. Among the genes found to be differentially
expressed, cyclin D2 (CCND2), chemokine (C-X-C motif) receptor 4 (CXCR4), glutathione peroxidase 3 (GPX3),
catenin delta 1 (CTNND1), 7-dehydrocholesterol reductase (DHCR7), disheveled, dsh homolog 3 (DVL3), heat
shock 27 kDa protein 1 (HSBP1), and tripartite motif containing 28 (TRIM28) were reported to be the most
significant genes, confirmed by subsequent qRT-PCR analysis.
Subsequently, Assou et al. [51] identified 630 genes as differentially expressed in cumulus cells from oocytes
that lead to a pregnancy compared with oocytes that did not. Majority of these genes had higher expression
levels in the pregnant group, suggesting transcriptional activation in cumulus cells of viable oocytes. The down
regulation of nuclear factor IB (NFIB) and the up regulation of BCL-like protein 11 (BCL2L11) and phospho-
enolpyruvate carboxykinase 1 (PCK1) in the pregnant group was further confirmed by qRT-PCR in the same
study.
Most recently, Feuerstein et al. [52] collected 197 individual cumulus cell samples from 106 patients undergo-
ing the ICSI procedure and investigated cumulus cell gene expression using 96 microarrays to determine genes
associated with oocyte maturation and subsequent blastocyst development. After microarrays, these researchers
performed a meta-analysis to test identified genes in other data sets available in Gene Expression Omnibus,
and they selected eight genes from the 308 differentially expressed genes for further validation by quantitative
polymerase chain reaction (qPCR). Three of these eight selected genes were validated as potential biomarkers
(perilipin 2 [PLIN2], regulator of G-protein signaling 2 [RGS2], and angiogenin [ANG]). After correction for
additional experimental parameters, including interpatient variability, only the expression level of RGS2 was
confirmed to be related to oocyte developmental competence and clinical pregnancy.
Granulosa cell transcriptome has been analyzed in three consecutive studies by Hamel et al. [53–55] in cor-
relation with pregnancy outcome. In the first study [53], the gene expression of granulosa cells was determined
using both a custom-made cDNA microarray and an Affymetrix GeneChip®. The samples were investigated in
two groups: the follicular cells from oocytes that resulted in a positive pregnancy and the follicular cells from
oocytes resulting in embryos that failed to develop. Among the 115 genes identified as being differentially
expressed, altered expression of 3-beta-hydroxystreoid dehydrogenase (HSD3β1), ferredoxin 1 (FDX1), serine
(or cysteine) proteinase inhibitor clade E member 2 (SERPINE2), cytochrome P450 aromatase (CYP19A1), and
cell division cycle 42 (CDC42) was significantly associated with pregnancy outcome. Building on the findings of
this initial study, they subsequently identified phosphoglycerate kinase 1 (PGK1), RGS2, regulator of G-protein
signaling 3 (RGS3), CDC42 [54], UDP-glucose pyrophosphorylase 2 (UGP2), and pleckstrin homology-like
domain family A member 1 (PHLDA1) [55] as potential follicular markers associated with embryo quality
resulting in a successful pregnancy.
Eventually, transcriptomic analysis of cumulus/granulosa cells enabled identification of various genes as
potential biomarkers of oocyte and embryo competence; the most frequently studied of these genes are listed in
Table 13.1.
Embryo Selection by Proteomics

Proteins are the key factors regulating the cellular functions within an organism. Proteome is the entire set
of proteins expressed by a specific genome under certain environmental conditions at a certain time. Proteomic
profiling is of primary importance to understand the complex nature of the biological processes. Although
transcriptomics technologies are available to provide an estimation of gene expression by quantification of
mRNAs within a sample, mRNA expression level of a particular gene does not always correlate with its protein
expression [56]. Observed differences between mRNA and corresponding protein levels are mainly attributed
to posttranscriptional mechanisms, substantial differences between proteins’ in vivo half-lives, and a significant
amount of experimental errors [57,58]. Recent advances in analytical proteomics techniques and related bioin-
formatics approaches enabled direct quantification of gene expression at the protein level.
Whereas transcriptomics provides an estimation of gene expression by quantification of mRNAs

within a sample, proteomics aims at the actual protein expression of a biological sample.
Proteomics studies may be aimed at (1) profiling proteome content of a biological sample, (2) performing
comparative protein expression analysis, (3) localizing and identifying posttranslational modifications, and
(4) exploring protein–protein interactions [59]. Proteomics technologies could be applied to IVF for noninvasive
assessment of embryonic secretome that includes the proteins produced by the embryos and secreted into the
surrounding culture media [60,61]. Preliminary proteomics studies in this context hypothesized that secretome
profiles of the culture media could potentially correlate with embryo viability [62–64]. Using mass spectrometry
or protein array technologies, these studies reported altered expression of specific proteins associated with blas-
tocyst development or implantation.
Analytical Proteomics Techniques

The basic mass spectrometer consists of an ionization source (matrix-assisted laser desorption ionization
[MALDI], surface-enhanced laser desorption/ionization [SELDI], or electrospray ionization [ESI]), a mass ana-
lyzer (e.g., time-of-flight [TOF], ion trap, Orbitrap), and a mass detector. Mass analyzers are usually used in
tandem (tandem mass spectrometry) to achieve higher degrees of ion separation and identification.
Mass spectrometry–based proteomics analyses are coupled with gel-based or gel-free protein separation tech-
niques and provide high detection specificity without the need for antibodies [65].
In the widely used two-dimensional difference gel electrophoresis (2D-DIGE) method for protein separation,
different protein mixtures are labeled with different fluorescent dyes and mixed and loaded together on a single
gel for quantitative comparative protein expression analysis [66]. The proteins within the complex mixture are
then separated by isoelectric focusing in the first dimension and by sodium dodecyl sulfate polyacrylamide gel
electrophoresis (SDS-PAGE) that discriminates proteins based on molecular mass in the second dimension. The
relative abundance of protein in each spot is determined by scanning the gel at the wavelengths of the fluorescent
dyes, enabling differential expression analysis. The spots of interest are then further analyzed by mass spectrom-
eters (e.g., MALDI-TOF/TOF) for identification of the corresponding proteins.
Different types of chromatographic separation techniques (commonly, high-performance liquid chromatogra-
phy [HPLC]) are also used in proteomics analysis for gel-free protein separation. The basic principle behind these
techniques is to separate solute proteins or peptides depending on their distributions between mobile and station-
ary phases. A typical example of combined liquid chromatography–mass spectrometry system is multidimen-
sional protein identification technology (muDPIT) [67] as an efficient approach for protein identification lacking
the ability to provide quantitative information. Alternative gel-free labeling technologies, such as isotope-coded
affinity tag (ICAT), stable isotope labeling by amino acids (SILAC), and isobaric tags for relative and absolute
quantification (iTRAQ), are also coupled with mass spectrometry, enabling quantitative proteomics analysis.
More recently, protein microarray technology is becoming a significant tool for quantitative and functional
proteomics [68]. Protein microarrays consist of antibodies or other affinity reagents attached on a solid surface.
Each agent binds to its target protein in a complex mixture, and detected proteins are identified and quantified
subsequently. Using protein arrays, it is possible to examine the expression of hundreds of proteins simultane-
ously. In contrast, protein arrays can provide data only on preselected proteins that were included in the chip
design, whereas mass spectrometry techniques can potentially detect any protein [59].
Clinical applications of proteomics-based embryo assessment

Initial studies investigating protein expression in relation to embryo viability focused on identification of specific
molecules produced by the embryo and secreted into the surrounding culture media. Embryos leading to suc-
cessful pregnancies were shown to secrete higher levels of interleukin-1 (IL-1) alpha [69] and IL-1 beta [70], and
embryos expressing soluble human leukocyte antigen G (HLA-G) had a better chance of implantation [71]. Also,
significantly higher concentration of leptin was observed in competent blastocysts compared with the arrested
blastocytes [72].
Preliminary data revealed correlation of protein secretion profiles with embryo competence and motivated a
deeper investigation of human embryonic secretome. Using SELDI-TOF-MS, Katz-Jaffe et al. [62] successfully
analyzed secretome of individual human embryos and observed distinctive secretome profiles at each succes-
sive 24 hr intervals of preimplantation embryo development. In the same study, significant up-regulation of an
8.5 kDa protein was observed in the secretome of developing blastocysts on Day 5. A subsequent protein identi-
fication process indicated that the best candidate for this potential biomarker was ubiquitin.
In another study, Dominguez et al. [63] used protein microarrays containing 120 antibody targets to analyze
the secretome of the human blastocysts. Conditioned media were collected from implanted and nonimplanted
embryos and from blank controls in a single embryo transfer program. Expressions of proteins such as CXCL13
(BCL, B lymphocyte chemoattractant), stem cell factor (SCF), and macrophage-stimulating protein-a (MSP-a)
were significantly decreased and soluble tumor necrosis factor (TNF) receptor 1 (sTNFR1) was significantly
increased in the media where the blastocyst was present in comparison with the control media. Moreover, when
compared with nonimplanted blastocysts, granulocyte macrophage–colony-stimulating factor (GM-CSF) and
CXCL13 were significantly reduced, therefore consumed at a higher rate by the implanted blastocysts.
A subsequent study by the same group [64] investigated comparative secretome of the conditioned media from
implanted blastocysts in endometrial epithelial cell (EEC) coculture system versus sequential microdrop culture
media. Protein array analysis indicated IL-6 as the most abundant protein in the implanted embryo coculture
media, and this group suggested that the coculture system favors blastocyst development and implantation by the
contribution of the factors secreted by EEC.
A summary of the initial proteomics analysis of embryonic secretome is given in Table 13.2. Despite the
promising preliminary results, proteomics is still considered a limited source of information for assessing repro-
ductive potential of embryos, possibly due to lack of sensitivity of proteomics platforms in this domain and to
the complexity of the experimental and data analysis procedures. Nevertheless, rapid progression of proteomics
technologies is likely to overcome such limitations in the near future.
TABLE 13.2
Potential Biomarker Proteins of Embryo Viability Identified in the Embryonic Secretome
Molecule Study Technique Result
IL-1 beta Baranao et al. (1997) ELISA ↑ IL-1 beta ∝ pregnancy
HLA-G Fuzzi et al. (2002) ELISA ↑ HLA-G ∝ implantation
Ubiquitin Katz-Jaffe et al. (2006) SELDI-TOF-MS ↑ ubiquitin ∝ blastocyst development
CXCL13 Dominguez et al. (2008) Protein array ↓ CXCL13 ∝ implantation
GM-CSF Dominguez et al. (2008) Protein array ↓ GM-CSF ∝ implantation
Note: Symbols ↑, ↓, ∝ stands for higher level, lower level, and associated with, respectively. CXCL13,
C-X-C motif chemokine 13; ELISA, enzyme-linked immunosorbent assay; GM-CSF, granulocyte
macrophage colony-stimulating factor; HLA-G, human leukocyte antigen G; IL, Interleukin; SELDI-
TOF-MS, surface-enhanced laser desorption/ionization–time-of-flight–mass spectrometer.
Embryo Selection by Metabolomics

Metabolome is the entire set of small molecules (<1 kDa), including metabolic intermediates (amino acids,
lipids, and nucleotides), hormones, signaling molecules, and secondary metabolites, within a biological s ample.
Metabolites are the final downstream products of gene expression, and their inventory provides a valuable data-
base to explore genotype–phenotype relationships and genotype–environment interactions. Within the context
of IVF, the changes in the levels of metabolites associated with carbohydrate metabolism and amino acid turn-
over were investigated as indicators of normal preimplantation embryo development.
Preimplantation Embryo Metabolism as an Indicator of Viability

Preimplantation embryo development has unique metabolic characteristics, because the preferred energy source
for embryonic cellular metabolism changes during preimplantation development. Carbohydrate metabolism of
preimplantation human embryos was mainly investigated in terms of pyruvate and glucose uptake and lactate
production. Initially in 1989, Hardy et al. [73] reported higher pyruvate uptake in embryos that develop to blas-
tocyst stage, soon after to be confirmed by Gott et al. [74]. Subsequently, Conaghan et al. [75] measured pyruvate
uptake for the first time in relation to implantation and clinical pregnancy, and they demonstrated that embryo
implantation after Day 2 and Day 3 transfers was inversely correlated with pyruvate uptake by two- to eight-
cell embryos. Another study by Turner et al. [76] showed that embryos had a wide range of pyruvate uptake
values, but this variation was reduced to an intermediate range in the embryos that implanted. More recently,
Gardner et al. [77] investigated the pyruvate metabolism in relation to blastocyst development and suggested that
pyruvate uptake on Day 4 was significantly higher by embryos that formed blastocysts compared with embryos
that failed to develop before blastocyst stage. On the basis of these findings, pyruvate uptake seems to be cor-
related with blastocyst development, whereas its relationship to implantation or pregnancy outcome remains
inconclusive.
Metabolome is the analysis of metabolites that are the final downstream products of gene
expression, and in preimplantation embryo development, a distinct change in the levels of
metabolites is expected to classify a “normal” development and a viable embryo.
Glucose uptake has also been measured in correlation with the embryo’s developmental potential. According
to initial findings, glucose uptake from Day 2 to Day 4 was similar for embryos that reached the blastocyst stage
and those that were arrested during cleavage [73,74]. In contrast, Gardner et al. [77] showed a significant asso-
ciation between increased levels of glucose uptake on Day 4 and subsequent blastocyst formation and quality.
Besides the carbohydrates, amino acids are also key components of preimplantation embryo development
because in vitro–developing embryos use and produce amino acids. The improvement in extended culture to
the blastocyst stage has been achieved mainly by inclusion of amino acids into the culture medium [78,79].
Consequently, amino acid content of spent embryo culture media has been profiled to explore specific amino acid
turnover patterns during preimplantation embryo development in correlation to blastocyst formation.
Initially, Houghton et al. [80] examined 18 amino acids during in vitro embryo development and showed
that embryos that form a blastocyst displayed a different profile of amino acid metabolism than those that were
arrested. Specifically, lower uptake of glutamine, arginine, and methionine and lower release of alanine and
asparagine on Day 2 and Day 3 was found to be associated with blastocyst development. In the same study, lower
uptake of serine and lower release of alanine and glycine by eight-cell- and morula-stage embryos correlated
with blastocyst development. Moreover, consistent with the “quiet embryo hypothesis” [81], sum of depletion
and appearance of the amino acids examined, that is, amino acid turnover, was shown to be lower in developing
embryos compared with the arrested embryos.
In another study, Brison et al. [82] investigated the correlation of amino acid turnover with implantation and
pregnancy outcomes when the embryos were selected according to routine morphological criteria and transferred
TABLE 13.3
Potential Biomarker Metabolites of Embryo Viability Identified in the Culture Media Metabolome
Metabolite Study Technique Result
Pyruvate Hardy et al. (1989) Ultramicrofluorescence assay ↑ pyruvate uptake on Day 1–5 ∝ embryo development
Gott et al. (1990) Ultramicrofluorescence assay ↑ pyruvate uptake on Day 2–5 ∝ embryo development
Conaghan et al. (1993) Ultramicrofluorescence assay ↓ pyruvate uptake on Day 2–3 ∝ implantation
Turner et al. (1994) Ultramicrofluorescence assay intermediate pyruvate uptake on Day 2 ∝
implantation of morphologically good embryos
Gardner et al. (2001) Ultramicrofluorescence assay ↑ pyruvate uptake on Day 4 ∝ blastocyst development
Glucose Hardy et al. (1989) Ultramicrofluorescence assay ↑ glucose uptake on Day 5 ∝ blastocyst development
Gott et al. (1990) Ultramicrofluorescence assay ↑ glucose uptake on Day 5 ∝ blastocyst development
Gardner et al. (2001) Ultramicrofluorescence assay ↑ glucose uptake on Day 5–6 ∝ blastocyst quality
Lactate Gott et al. (1990) Ultramicrofluorescence assay ↑ lactate production on Day 3–5 ∝ blasctocyst
development
Glutamine Houghton et al. (2002) High-performance liquid ↓ glutamine uptake on Day 2–3 ∝ blastocyst
chromatography development
Arginine Houghton et al. (2002) High-performance liquid ↓ arginine uptake on Day 2–3 ∝ blastocyst
Methionine Houghton et al. (2002) High-performance liquid ↓ methionine uptake on Day 2–3 ∝ blastocyst
Alanine Houghton et al. (2002) High-performance liquid ↓ alanine production on Day 2–3 ∝ blastocyst
chromatography development ↓ alanine release at compacting 8-cell
stage ∝ blastocyst development
Asparagine Houghton et al. (2002) High-performance liquid ↓ asparagine production on Day 2–3 ∝ blastocyst
Brison et al. (2004) High-performance liquid ↑ asparagine level on Day 2 ∝ pregnancy & live birth
chromatography
Serine Houghton et al. (2002) High-performance liquid ↓ serine uptake at compacting 8-cell stage ∝ blastocyst
Glycine Houghton et al. (2002) High-performance liquid ↓ glycine release at compacting 8-cell stage ∝
chromatography blastocyst development
Brison et al. (2004) High-performance liquid ↓ glycine level on Day 2 ∝ pregnancy & live birth
chromatography
Leucine Brison et al. (2004) High-performance liquid ↓ glycine level on Day 2 ∝ pregnancy & live birth
chromatography
Glutamate Seli et al. (2008) Proton nuclear magnetic ↑ glutamate level on Day 3 ∝ pregnancy & live birth
resonance
on Day 2. This study demonstrated that decreased glycine and leucine and increased asparagine levels in the
culture media were associated with increased clinical pregnancy and live birth rates. A more recent study by Seli
et al. [83] showed an association between increased glutamate levels in the culture media and clinical pregnancy
and live birth.
Table 13.3 is a summary of the findings suggested by the studies investigating preimplantation embryo metab-
olism as an indicator of embryo viability.
Techniques for Metabolomic Analysis of Embryo Culture Media

Currently, a variety of analytical spectroscopic techniques are available for metabolomic analysis of biological
samples. Here, we provide a brief overview of common techniques and related data analysis procedures.
Analytical Spectroscopic Techniques

Common analytical approaches applied for metabolomic analysis of embryo culture media can be catego-
rized as nuclear magnetic resonance (NMR) spectroscopy, mass spectrometry, or vibrational spectroscopy.
NMR is a nondestructive analytical technique exploiting interaction of the magnetic moment of atomic nuclei
with the external magnetic field. NMR provides information on metabolites containing elements with nonzero
magnetic moments and has been efficiently used for biomarker analysis by enabling detection and quantification
of specific metabolites within a biological fluid or tissue [84,85]. Limitations of NMR were attributed to require-
ment for large amounts of sample, higher costs, and lack of sensitivity for low-abundance targets favoring the
method for analysis of high abundance metabolites.
Mass spectrometry operates by ion formation, separation of ions according to their mass-to-charge ratio (m/z),
and detection of separated ions [86] and enables simultaneous characterization of several hundred metabolites
with higher sensitivity compared with NMR approaches. Mass spectrometry is the most widely used analytical
platform in metabolomics. The sensitivity and specificity of this common technique is further enhanced when
coupled with chromatography or electrophoresis-based separation techniques.
The vibrational spectroscopy techniques include Raman and infrared (IR) approaches [87]. The main
principle behind vibrational techniques is that, when a sample is exposed to an electromagnetic radiation,
the chemical bonds within the molecules will absorb the energy and vibrate to a greater extent. The IR
spectrum is the result of absorption of electromagnetic radiation by vibrating molecules when the sample
is interrogated with an IR beam, whereas Raman spectroscopy measures the scattering of electromagnetic
radiation by the vibrating molecules under exposure to a particular wavelength light (usually in the form of
a laser).
The wavelength of the light used for radiation differs in the two IR spectroscopy techniques: near-infrared
(NIR) and Fourier transform infrared (FTIR). NIR measures the spectra in the 14,000–4000 cm –1 region,
whereas FTIR looks at the mid-IR part of the spectrum at 4000–600 cm –1. NIR has the advantage of quan-
tification with higher sensitivity in metabolomics research, whereas FTIR is a faster and higher throughput
method.
Analysis of Metabolomics Data

Analysis of the metabolomics data is strongly related to the research objective, considering three main catego-
ries, namely targeted metabolite profiling, nontargeted profiling, and metabolic fingerprinting. Targeted analysis
is based on detection and quantification of the metabolites for which chemical structures are known. In con-
trast, nontargeted profiling aims at quantification of all the peaks in the spectrum without associating the peaks
to chemical structure of certain compounds. Metabolomic fingerprinting is also a nontargeted approach that
considers the whole metabolic profile as a pattern.
Analysis of targeted metabolite profiling is relatively simple because only previously identified spectral
regions are analyzed. In contrast, nontargeted profiling and metabolic fingerprinting are complex processes
that may require bioinformatics support for efficient and accurate analysis of high dimensional metabolomics
data [88].
A common preprocessing step in the analysis of metabolomics data is normalization. The multistage
experimental setting underlying metabolomics studies is likely to introduce systematic variations in the
resulting spectra. The main source of nonbiological variations in metabolomics embryo assessment
is attributed to the culture environment. Spectral profiles of spent culture media (i.e., where individual
embryo has been cultured) need to be normalized to that of blank samples (i.e., culture media incubated
under the same conditions but without an embryo) to eliminate the possible impact of variations in cul-
ture conditions. Then, predictive models are developed using advanced statistical methods. To ensure the
robustness of the model against center-specific variations, such models should be either developed using
pooled multicenter data or validated in different c enters. Once the success and robustness of a prediction
model are established, it can be used as an embryo selection tool alone or in combination with other avail-
able criteria.
0.35
0.3
0.25
0.2
0.15
0.35
0.1
Embryo 0.05 0.3
0 0.25
500 1000 1500 2000 2500 3000 3500 4000
0.2
Embryo spectrum 0.15
0.1
0.05
0.3 0
Blank control 0.25
-0.05
0.2 0 200 400 600 800 1000 1200 1400 1600 1800
0.15 Data analysis and
0.1 Normalized spectra viability prediction
0.05
0
–0.05
500 1000 1500 2000 2500 3000 3500 4000
Culture media Control spectrum
FIGURE 13.3 Metabolomic assessment of embryo viability through spectroscopic analysis of spent culture media.
Clinical applications of metabolomics-based embryo assessment

Efficiency of metabolomics-based embryo selection strategies was first evaluated in retrospective studies and
then in subsequent randomized clinical trials (RCTs). The typical workflow of metabolomics-based embryo
assessment is shown in Figure 13.3.
Proof-of-Principal Studies
In 2007, Seli et al. [89] reported the findings of a proof-of-concept study correlating spent culture media metabo-
lome with embryo viability. In this initial study, Day 3 spent embryo culture media samples were analyzed using
Raman spectroscopy, NIR spectroscopy, or both. The mean spectrum of embryos that failed to implant was com-
pared with the mean spectrum of embryos that resulted in a live birth, and algorithms predictive of an embryo’s
reproductive potential were developed for Raman and NIR spectroscopy. Subsequently, Scott et al. [90] validated
the Raman spectroscopy algorithm using spent culture media collected at a different center, where embryos were
cultured in a different type and volume of culture medium.
In a subsequent study, Vergouw et al. [91] analyzed embryo culture media samples from single embryo trans-
fer cycles using NIR spectroscopy and developed a new algorithm predictive of embryo implantation potential.
They calculated a “viability score” for each embryo based on the NIR metabolomic profile of the corresponding
culture medium and reported that increasing viability scores were correlated with an increasing ability of the
embryo to implant. Subsequent studies with a large number of samples collected in SET cycles reported similar
findings and suggested that the metabolomic profile of embryo culture medium was a parameter independent of
morphology [92–94].
Commercialization Efforts and RCTs

The initial studies suggesting potential benefit from metabolomic profiling of spent culture media to determine
embryo viability were retrospective and were performed in a single research laboratory using frozen and trans-
ported culture media samples. Because the promise of metabolomics approach is its potential use as a rapid
on-site technology in the IVF laboratory, the algorithms developed needed to be tested using fresh samples at
different clinical sites and RCT design.
A series of prototype and commercial instruments were built and tested for this purpose by a private company,
Molecular Biometrics Inc., using an NIR system (ViaMetrics). Using the prototype instruments, two RCTs were
conducted in centers performing SET [16,36]. In these studies, in the treatment arm, embryos with good mor-
phology were identified followed by selection for transfer based on the metabolomic Viability Score™ generated
by the commercial instrument. In the control arm, embryos were selected using standard morphological embryo
assessment protocols.
In the first of these studies, Hardarson et al. [95] analyzed spent culture media from Day 2 and Day 5 SETs.
Although not statistically significant, their findings suggested a potential benefit of NIR for the assessment
of embryos transferred on Day 2. In the NIR/morphology group (n = 87) and morphology alone group (n = 83),
the pregnancy rates for Day 2 were 31% and 26.5%, respectively. In the same study, no benefit was found for
selection on Day 5 (n = 77 for NIR/morphology and n = 80 for morphology alone groups). The second random-
ized trial performed by Vergouw et al. [36] reported similar findings in women undergoing SET on Day 3, and
found no difference in live birth rates between NIR/morphology group (n = 146; live birth rate 29.5%) compared
with the morphology alone group (n = 163; live birth rate 31.3%).
Conversely, Sfontouris et al. [96] performed an RCT using the commercial platform where two or three
embryos were selected for transfer on the basis of NIR/morphology or morphology alone, and they showed an
improvement in pregnancy rates in the NIR-assisted group. Implantation rates (IRs) were significantly improved
in the NIR/morphology group (n = 39 patients; IR = 33/102 [32.4%]) compared with the morphology alone group
[n = 86 patients; 55/257 [21.4%]).
Ultimately, the commercial version of the NIR instrument was withdrawn due to the wide variability in perfor-
mance between clinics and the inconsistent results in clinical trials. The technology will hopefully be improved
and tested again in the near future.
Summary and Conclusions

Current embryo assessment strategies that rely primarily on embryo morphology and cleavage rate do not pro-
vide adequate sensitivity or specificity to achieve desired pregnancy rates in women undergoing infertility treat-
ment with IVF. Studies using emerging technologies to analyze cumulus/granulosa cell transcriptome or spent
embryo culture media protein or metabolite content report promising results. However, the transcripts or proteins
that have been identified as potential biomarkers of embryo viability or the metabolomic profiles associated
with pregnancy outcome have not yet been adequately validated. Therefore, the use of these novel noninvasive
technologies remains experimental, and their application to clinical practice awaits RCTs demonstrating benefit
from their use, alone or in combination with morphological evaluation.
REFERENCES
1. Martin JA, Hamilton BE, Ventura SJ, Osterman MJK, Wilson EC, Mathews TJ. Births: Final Data for 2010.
Hyattsville, MD: National Center for Health Statistics; 2012.
2. Practice Committee of American Society for Reproductive Medicine. Multiple gestation associated with infertil-
ity therapy: An American Society for Reproductive Medicine Practice Committee opinion. Fertility and Sterility.
2012;97(4):825–34.
3. Norwitz ER, Edusa V, Park JS. Maternal physiology and complications of multiple pregnancy. Seminars in
Perinatology. 2005;29(5):338–48.
4. Ata B, Seli E. Economics of assisted reproductive technologies. Current Opinion in Obstetrics & Gynecology.
2010;22(3):183–8.
5. Bromer JG, Ata B, Seli M, Lockwood CJ, Seli E. Preterm deliveries that result from multiple pregnancies asso-
ciated with assisted reproductive technologies in the USA: A cost analysis. Current Opinion in Obstetrics &
Gynecology. 2011;23(3):168–73.
6. Velikonja U. The costs of multiple gestation pregnancies in assisted reproduction. Harvard Journal of Law and
Gender. 2009;463:481–93.
7. Bromer JG, Seli E. Assessment of embryo viability in assisted reproductive technology: Shortcomings of
current approaches and the emerging role of metabolomics. Current Opinion in Obstetrics & Gynecology.
2008;20(3):234–41.
8. Center for Disease Control and Prevention. ART. 2010. Available from: http://www.cdc.gov/art/
9. Human Fertilisation and Embryology Authority (HFEA). 2011. Available from: http://www.hfea.gov.uk
10. Sakkas D, Gardner DK. Noninvasive methods to assess embryo quality. Current Opinion in Obstetrics &
Gynecology. 2005;17(3):283–8.
11. Wong CC, Loewke KE, Bossert NL, Behr B, De Jonge CJ, Baer TM, et al. Non-invasive imaging of human
embryos before embryonic genome activation predicts development to the blastocyst stage. Nature Biotechnology.
2010;28(10):1115–21.
12. Meseguer M, Rubio I, Cruz M, Basile N, Marcos J, Requena A. Embryo incubation and selection in a time-lapse
monitoring system improves pregnancy outcome compared with a standard incubator: A retrospective cohort
study. Fertility and Sterility. 2012;98(6):1481–9.e10.
13. Seli E, Robert C, Sirard MA. OMICS in assisted reproduction: Possibilities and pitfalls. Molecular Human
Reproduction. 2010;16(8):513–30.
14. Checa MA, Alonso-Coello P, Sola I, Robles A, Carreras R, Balasch J. IVF/ICSI with or without preimplantation
genetic screening for aneuploidy in couples without genetic disorders: A systematic review and meta-analysis.
Journal of Assisted Reproduction and Genetics. 2009;26(5):273–83.
15. Sher G, Keskintepe L, Keskintepe M, Maassarani G, Tortoriello D, Brody S. Genetic analysis of human embryos
by metaphase comparative genomic hybridization (mCGH) improves efficiency of IVF by increasing embryo
implantation rate and reducing multiple pregnancies and spontaneous miscarriages. Fertility and Sterility.
2009;92(6):1886–94.
16. Schoolcraft WB, Treff NR, Stevens JM, Ferry K, Katz-Jaffe M, Scott RT, Jr. Live birth outcome with trophec-
toderm biopsy, blastocyst vitrification, and single-nucleotide polymorphism microarray-based comprehensive
chromosome screening in infertile patients. Fertility and Sterility. 2011;96(3):638–40.
17. Uyar A, Torrealday S, Seli E. Cumulus and granulosa cell markers of oocyte and embryo quality. Fertility and
Sterility. 2013;99(4):979–97.
18. Gilchrist RB, Ritter LJ, Armstrong DT. Oocyte-somatic cell interactions during follicle development in mam-
mals. Animal Reproduction Science. 2004;82–83:431–46.
19. Roche JF. Control and regulation of folliculogenesis—a symposium in perspective. Reviews of Reproduction.
1996;1(1):19–27.
20. Heller DT, Schultz RM. Ribonucleoside metabolism by mouse oocytes: Metabolic cooperativity between the
fully grown oocyte and cumulus cells. The Journal of Experimental Zoology. 1980;214(3):355–64.
21. Brower PT, Schultz RM. Intercellular communication between granulosa cells and mouse oocytes: Existence and
possible nutritional role during oocyte growth. Developmental Biology. 1982;90(1):144–53.
22. Albertini DF, Combelles CM, Benecchi E, Carabatsos MJ. Cellular basis for paracrine regulation of ovarian
follicle development. Reproduction. 2001;121(5):647–53.
23. Tanghe S, Van Soom A, Nauwynck H, Coryn M, de Kruif A. Minireview: Functions of the cumulus ooph-
orus during oocyte maturation, ovulation, and fertilization. Molecular Reproduction and Development.
2002;61(3):414–24.
24. Veeck LL. An Atlas of Human Gametes and Conceptuses: An Illustrated Reference of Assisted Reproductive
Technology. New York: The Pathenon Publishing Group; 1999.
25. Manchester KL. Use of UV methods for measurement of protein and nucleic acid concentrations. Biotechniques.
1996;20(6):968–70.
26. Sambrook J, Russel DW. Molecular Cloning: A Laboratory Manual. 3rd ed. Cold Spring Harbor, NY: Cold
Spring Harbor Laboratory Press; 2001.
27. Schroeder A, Mueller O, Stocker S, Salowsky R, Leiber M, Gassmann M, et al. The RIN: An RNA integrity
number for assigning integrity values to RNA measurements. BMC Molecular Biology. 2006;7:3.
28. Stahlberg A, Hakansson J, Xian X, Semb H, Kubista M. Properties of the reverse transcription reaction in mRNA
quantification. Clinical Chemistry. 2004;50(3):509–15.
29. Saiki RK, Gelfand DH, Stoffel S, Scharf SJ, Higuchi R, Horn GT, et al. Primer-directed enzymatic amplification
of DNA with a thermostable DNA polymerase. Science. 1988;239(4839):487–91.
30. Higuchi R, Dollinger G, Walsh PS, Griffith R. Simultaneous amplification and detection of specific DNA
sequences. Biotechnology (N Y). 1992;10(4):413–17.
31. Higuchi R, Fockler C, Dollinger G, Watson R. Kinetic PCR analysis: Real-time monitoring of DNA amplification
reactions. Biotechnology (N Y). 1993;11(9):1026–30.
32. Andersen CL, Jensen JL, Orntoft TF. Normalization of real-time quantitative reverse transcription-PCR data:
A model-based variance estimation approach to identify genes suited for normalization, applied to bladder and
colon cancer data sets. Cancer Research. 2004;64(15):5245–50.
33. Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De Paepe A, et al. Accurate normalization of
real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biology.
2002;3(7):34–34.11
34. Mortazavi A, Williams BA, McCue K, Schaeffer L, Wold B. Mapping and quantifying mammalian transcrip-
tomes by RNA-Seq. Nature Methods. 2008;5(7):621–8.
35. Wang Z, Gerstein M, Snyder M. RNA-Seq: A revolutionary tool for transcriptomics. Nature Reviews Genetics.
2009;10(1):57–63.
36. Ozsolak F, Platt AR, Jones DR, Reifenberger JG, Sass LE, McInerney P, et al. Direct RNA sequencing. Nature.
2009;461:814–18.
37. Antoniou E, Taft R. Gene expression in mouse oocytes by RNA-Seq. Methods in Molecular Biology.
2012;825:237–51.
38. Irizarry RA, Hobbs B, Collin F, Beazer-Barclay YD, Antonellis KJ, Scherf U, et al. Exploration, normalization,
and summaries of high density oligonucleotide array probe level data. Biostatistics. 2003;4(2):249–64.
39. Bolstad BM, Irizarry RA, Astrand M, Speed TP. A comparison of normalization methods for high density oligo-
nucleotide array data based on variance and bias. Bioinformatics. 2003;19(2):185–93.
40. Irizarry RA, Bolstad BM, Collin F, Cope LM, Hobbs B, Speed TP. Summaries of Affymetrix GeneChip probe
level data. Nucleic Acids Research. 2003;31(4):e15.
41. Storey JD. A direct approach to false discovery rates. Journal of the Royal Statistical Society: Series B.
2002;64:479–98.
42. Storey JD, Tibshirani R. Statistical significance for genomewide studies. Proceedings of the National Academy of
Sciences of the United States of America. 2003;100(16):9440–5.
43. McKenzie LJ, Pangas SA, Carson SA, Kovanci E, Cisneros P, Buster JE, et al. Human cumulus granulosa cell gene
expression: A predictor of fertilization and embryo selection in women undergoing IVF. Human Reproduction.
2004;19(12):2869–74.
44. Cillo F, Brevini TA, Antonini S, Paffoni A, Ragni G, Gandolfi F. Association between human oocyte developmen-
tal competence and expression levels of some cumulus genes. Reproduction. 2007;134(5):645–50.
45. Anderson RA, Sciorio R, Kinnell H, Bayne RA, Thong KJ, de Sousa PA, et al. Cumulus gene expression as a pre-
dictor of human oocyte fertilisation, embryo development and competence to establish a pregnancy. Reproduction.
2009;138(4):629–37.
46. Gebhardt KM, Feil DK, Dunning KR, Lane M, Russell DL. Human cumulus cell gene expression as a biomarker
of pregnancy outcome after single embryo transfer. Fertility and Sterility. 2011;96(1):47–52.e2.
47. Wathlet S, Adriaenssens T, Segers I, Verheyen G, Van de Velde H, Coucke W, et al. Cumulus cell gene expres-
sion predicts better cleavage-stage embryo or blastocyst development and pregnancy for ICSI patients. Human
Reproduction. 2011;26(5):1035–51.
48. Wathlet S, Adriaenssens T, Segers I, Verheyen G, Janssens R, Coucke W, et al. New candidate genes to predict
pregnancy outcome in single embryo transfer cycles when using cumulus cell gene expression. Fertility and
Sterility. 2012;98(2):432–9.e4.
49. Zhang X, Jafari N, Barnes RB, Confino E, Milad M, Kazer RR. Studies of gene expression in human cumu-
lus cells indicate pentraxin 3 as a possible marker for oocyte quality. Fertility and Sterility. 2005;83(Suppl
1):1169–79.
50. van Montfoort AP, Geraedts JP, Dumoulin JC, Stassen AP, Evers JL, Ayoubi TA. Differential gene expres-
sion in cumulus cells as a prognostic indicator of embryo viability: A microarray analysis. Molecular Human
Reproduction. 2008;14(3):157–68.
51. Assou S, Haouzi D, Mahmoud K, Aouacheria A, Guillemin Y, Pantesco V, et al. A non-invasive test for assess-
ing embryo potential by gene expression profiles of human cumulus cells: A proof of concept study. Molecular
Human Reproduction. 2008;14(12):711–19.
52. Feuerstein P, Puard V, Chevalier C, Teusan R, Cadoret V, Guerif F, et al. Genomic assessment of human cumulus
cell marker genes as predictors of oocyte developmental competence: Impact of various experimental factors.
PLoS One. 2012;7(7):e40449.
53. Hamel M, Dufort I, Robert C, Gravel C, Leveille MC, Leader A, et al. Identification of differentially expressed
markers in human follicular cells associated with competent oocytes. Human Reproduction. 2008;23(5):1118–27.
54. Hamel M, Dufort I, Robert C, Leveille MC, Leader A, Sirard MA. Genomic assessment of follicular marker genes
as pregnancy predictors for human IVF. Molecular Human Reproduction. 2010;16(2):87–96.
55. Hamel M, Dufort I, Robert C, Leveille MC, Leader A, Sirard MA. Identification of follicular marker genes as
pregnancy predictors for human IVF: New evidence for the involvement of luteinization process. Molecular
Human Reproduction. 2010;16(8):548–56.
56. Gygi SP, Rochon Y, Franza BR, Aebersold R. Correlation between protein and mRNA abundance in yeast.
Molecular and Cellular Biology. 1999;19(3):1720–30.
57. Greenbaum D, Colangelo C, Williams K, Gerstein M. Comparing protein abundance and mRNA expression
levels on a genomic scale. Genome Biology. 2003;4(9):117.
58. Beyer A, Hollunder J, Nasheuer HP, Wilhelm T. Post-transcriptional expression regulation in the yeast
Saccharomyces cerevisiae on a genomic scale. Molecular & Cellular Proteomics. 2004;3(11):1083–92.
59. Chandramouli K, Qian PY. Proteomics: Challenges, techniques and possibilities to overcome biological sample
complexity. Human Genomics and Proteomics. 2009;2009: ii, 239204.
60. Katz-Jaffe MG, McReynolds S. Embryology in the era of proteomics. Fertility and Sterility. 2013;99(4):1073–7.
61. Garrido-Gomez T, Dominguez F, Simon C. Proteomics of embryonic implantation. Handbook of Experimental
Pharmacology. 2010;(198):67–78.
62. Katz-Jaffe MG, Schoolcraft WB, Gardner DK. Analysis of protein expression (secretome) by human and mouse
preimplantation embryos. Fertility and Sterility. 2006;86(3):678–85.
63. Dominguez F, Gadea B, Esteban FJ, Horcajadas JA, Pellicer A, Simon C. Comparative protein-profile analysis of
implanted versus non-implanted human blastocysts. Human Reproduction. 2008;23(9):1993–2000.
64. Dominguez F, Gadea B, Mercader A, Esteban FJ, Pellicer A, Simon C. Embryologic outcome and secretome pro-
file of implanted blastocysts obtained after coculture in human endometrial epithelial cells versus the sequential
system. Fertility and Sterility. 2010;93(3):774–82.e1.
65. Scherp P, Ku G, Coleman L, Kheterpal I. Gel-based and gel-free proteomic technologies. Methods in Molecular
Biology. 2011;702:163–90.
66. Unlu M, Morgan ME, Minden JS. Difference gel electrophoresis: A single gel method for detecting changes in
protein extracts. Electrophoresis. 1997;18(11):2071–7.
67. Washburn MP, Wolters D, Yates JR, 3rd. Large-scale analysis of the yeast proteome by multidimensional protein
identification technology. Nature Biotechnology. 2001;19(3):242–7.
68. MacBeath G. Protein microarrays and proteomics. Nature Genetics. 2002;32(Suppl):526–32.
69. Sheth KV, Roca GL, al-Sedairy ST, Parhar RS, Hamilton CJ, al-Abdul Jabbar F. Prediction of successful embryo
implantation by measuring interleukin-1–alpha and immunosuppressive factor(s) in preimplantation embryo cul-
ture fluid. Fertility and Sterility. 1991;55(5):952–7.
70. Baranao RI, Piazza A, Rumi LS, Polak de Fried E. Determination of IL-1 and IL-6 levels in human embryo
culture-conditioned media. American Journal of Reproductive Immunology. 1997;37(2):191–4.
71. Fuzzi B, Rizzo R, Criscuoli L, Noci I, Melchiorri L, Scarselli B, et al. HLA-G expression in early embryos is a
fundamental prerequisite for the obtainment of pregnancy. European Journal of Immunology. 2002;32(2):311–15.
72. Gonzalez RR, Simon C, Caballero-Campo P, Norman R, Chardonnens D, Devoto L, et al. Leptin and reproduc-
tion. Human Reproduction Update. 2000;6(3):290–300.
73. Hardy K, Hooper MA, Handyside AH, Rutherford AJ, Winston RM, Leese HJ. Non-invasive measurement of
glucose and pyruvate uptake by individual human oocytes and preimplantation embryos. Human Reproduction.
1989;4(2):188–91.
74. Gott AL, Hardy K, Winston RM, Leese HJ. Non-invasive measurement of pyruvate and glucose uptake and lac-
tate production by single human preimplantation embryos. Human Reproduction. 1990;5(1):104–8.
75. Conaghan J, Hardy K, Handyside AH, Winston RM, Leese HJ. Selection criteria for human embryo transfer: A com-
parison of pyruvate uptake and morphology. Journal of Assisted Reproduction and Genetics. 1993;10(1):21–30.
76. Turner K, Martin KL, Woodward BJ, Lenton EA, Leese HJ. Comparison of pyruvate uptake by embryos derived
from conception and non-conception natural cycles. Human Reproduction. 1994;9(12):2362–6.
77. Gardner DK, Lane M, Stevens J, Schoolcraft WB. Noninvasive assessment of human embryo nutrient consump-
tion as a measure of developmental potential. Fertility and Sterility. 2001;76(6):1175–80.
78. Devreker F, Hardy K, Van den Bergh M, Vannin AS, Emiliani S, Englert Y. Amino acids promote human blasto-
cyst development in vitro. Human Reproduction. 2001;16(4):749–56.
79. Lane M, Gardner DK. Nonessential amino acids and glutamine decrease the time of the first three cleavage
divisions and increase compaction of mouse zygotes in vitro. Journal of Assisted Reproduction and Genetics.
1997;14(7):398–403.
80. Houghton FD, Hawkhead JA, Humpherson PG, Hogg JE, Balen AH, Rutherford AJ, et al. Non-invasive amino
acid turnover predicts human embryo developmental capacity. Human Reproduction. 2002;17(4):999–1005.
81. Leese HJ, Baumann CG, Brison DR, McEvoy TG, Sturmey RG. Metabolism of the viable mammalian embryo:
Quietness revisited. Molecular Human Reproduction. 2008;14(12):667–72.
82. Brison DR, Houghton FD, Falconer D, Roberts SA, Hawkhead J, Humpherson PG, et al. Identification of
viable embryos in IVF by non-invasive measurement of amino acid turnover. Human Reproduction.
2004;19(10):2319–24.
83. Seli E, Botros L, Sakkas D, Burns DH. Noninvasive metabolomic profiling of embryo culture media using proton
nuclear magnetic resonance correlates with reproductive potential of embryos in women undergoing in vitro
fertilization. Fertility and Sterility. 2008;90(6):2183–9.
84. Nicholson JK, O’Flynn MP, Sadler PJ, Macleod AF, Juul SM, Sonksen PH. Proton-nuclear-magnetic-resonance
studies of serum, plasma and urine from fasting normal and diabetic subjects. The Biochemical Journal.
1984;217(2):365–75.
85. Nicholson JK, Connelly J, Lindon JC, Holmes E. Metabonomics: A platform for studying drug toxicity and gene
function. Nature Reviews Drug Discovery. 2002;1(2):153–61.
86. Dunn WB, Ellis DI. Metabolomics: Current analytical platforms and methodologies. Trends in Analytical
Chemistry. 2005;24(4):285–94.
87. Ellis DI, Goodacre R. Metabolic fingerprinting in disease diagnosis: Biomedical applications of infrared and
Raman spectroscopy. The Analyst. 2006;131(8):875–85.
88. Shulaev V. Metabolomics technology and bioinformatics. Briefings in Bioinformatics. 2006;7(2):128–39.
89. Seli E, Sakkas D, Scott R, Kwok SC, Rosendahl SM, Burns DH. Noninvasive metabolomic profiling of embryo
culture media using Raman and near-infrared spectroscopy correlates with reproductive potential of embryos in
women undergoing in vitro fertilization. Fertility and Sterility. 2007;88(5):1350–7.
90. Scott R, Seli E, Miller K, Sakkas D, Scott K, Burns DH. Noninvasive metabolomic profiling of human embryo
culture media using Raman spectroscopy predicts embryonic reproductive potential: A prospective blinded pilot
study. Fertility and Sterility. 2008;90(1):77–83.
91. Vergouw CG, Botros LL, Roos P, Lens JW, Schats R, Hompes PG, et al. Metabolomic profiling by near-infrared
spectroscopy as a tool to assess embryo viability: A novel, non-invasive method for embryo selection. Human
Reproduction. 2008;23(7):1499–504.
92. Seli E, Vergouw CG, Morita H, Botros L, Roos P, Lambalk CB, et al. Noninvasive metabolomic profiling as an
adjunct to morphology for noninvasive embryo assessment in women undergoing single embryo transfer. Fertility
and Sterility. 2010;94(2):535–42.
93. Ahlstrom A, Wikland M, Rogberg L, Barnett JS, Tucker M, Hardarson T. Cross-validation and predictive value
of near-infrared spectroscopy algorithms for day-5 blastocyst transfer. Reproductive Biomedicine Online. 2011;
22(5):477–84.
94. Seli E, Bruce C, Botros L, Henson M, Roos P, Judge K, et al. Receiver operating characteristic (ROC) analysis
of day 5 morphology grading and metabolomic Viability Score on predicting implantation outcome. Journal of
Assisted Reproduction and Genetics. 2011;28(2):137–44.
95. Hardarson T, Ahlstrom A, Rogberg L, Botros L, Hillensjo T, Westlander G, et al. Non-invasive metabolo-
mic profiling of day 2 and 5 embryo culture medium: A prospective randomized trial. Human Reproduction.
2012;27(1):89–96.
96. Sfontouris IA, Lainas GT, Sakkas D, Iliadis GS, Anagnostara K, Zorzovilis IZ, et al. Assessment of embryo selec-
tion using non-invasive metabolomic analysis as an adjunct to morphology indicates improvement in implanta-
tion and fetal cardiac activity rates. Human Reproduction. 2011;26(Suppl 1):86.
14
Invasive Techniques: Polar Body Biopsy
Introduction
Polar body (PB) biopsy dates back to the 1990s when the technique was introduced by Verlinsky et al. [1] to
diagnose chromosomal abnormalities. In contrast to preimplantation genetic diagnosis (PGD) of the embryo that
was proposed earlier by Alan Handyside et al. [2], PB biopsy enabled a genetic diagnosis before the formation of
the proper embryo, thereby establishing the term “preconception genetic diagnosis.” PBs are by-products of the
meiotic division. The first PB (PB1) is not required for successful fertilization or for normal embryonic develop-
ment. The second PB (PB2), extruded from the oocyte after initiation of the fertilization cascade by the sperma-
tozoa, is similarly not needed for subsequent embryo development. Furthermore, it is important to note that PB
diagnosis gives direct information about the first and second PBs and therefore allows only an indirect diagnosis
of the maternal genetic or chromosomal constitution of the corresponding oocyte. In contrast, analysis of blasto-
meres from the embryo gives a direct diagnosis for the embryo and allows for detection of both maternally and
paternally derived genetic or chromosomal contributions.
Polar body diagnosis gives direct information about the first and second polar body and
therefore allows only an indirect diagnosis of the maternal genetic or chromosomal constitution
of the corresponding oocyte.
These clear diagnostic limitations of the PB were a major reason for the predominance of embryo biopsy
at the six- to eight-cell stage on Day 3. PB biopsy was at that early times applied only in countries where
embryo biopsy was not in line with legal requirements (e.g., Germany) [3] and by those groups who favored
this technology for various other reasons [4]. The major diagnostic aims at that time were PGD, a technique
implemented for diagnosing genetic diseases in known carriers before embryo transfer, and screening for chro-
mosomal a neuploidies, defined by the term “preimplantation genetic screening” (PGS). PGS became a major
indication for preimplantation diagnosis and was introduced in numerous laboratories worldwide (European
Society of Human Reproduction and Endocrinology [ESHRE] PGD consortium data) with the hope to contribute
to increased success rates in assisted reproduction.
However, since 2007 numerous randomized controlled trials reported that PGS by blastomere biopsy does
not result in increased success rates [5–15], and several organizations published statements asserting that they
no longer recommended using biopsy at the blastomere stage at least for PGS [16,17]. These reflections opened
up new discussions concerning the stage of biopsy [18]. The two alternatives that were discussed in regard to
blastomere biopsy were PB and trophectoderm (TE) biopsy [19]. Meanwhile, it was widely accepted that for
PGS, both methods should be performed in combination with array-comparative genomic hybridization (CGH)
for all chromosomes. Obviously, PB biopsy has known restrictions as only the female part can be investigated,
but in view of aneuploidy screening to detect numerical chromosomal disorders that predominantly arise during
meiosis in the oocyte [20], PB biopsy is a viable alternative.
This chapter gives an overview of the relevant technical details of PB diagnosis, with special emphasis on
aneuploidy screening.
209
Zona Opening
PB biopsy requires access to the perivitelline space. For opening of the zona pellucida and subsequent removal
of PBs, chemical, mechanical, and laser-assisted opening technologies have been proposed.
Chemical Opening
Acidic Tyrode’s solution was initially used for opening of the zona pellucida by chemical means [21]. Although
acidic Tyrode’s solution can be applied at the embryo stage, there was an inhibitory effect on embryonic
development when oocytes were exposed to acid Tyrode’s solution [22]. Because both the oocyte and PB are
sensitive to the effects of acid, zona drilling by acid Tyrode’s solution is unsuitable for PB biopsy.
Mechanical Opening
The group in Chicago has established a very efficient mechanical technique [23] that is based on three-dimensional
zona dissection and subsequent biopsy. The oocyte is first affixed to the holding capillary, and then a slit is made
with a sharp needle close to the area where the PBs are located. After turning the oocytes by 90°, a second slit is
made, creating a cross-like incision in the zona that allows access to the PBs (Figure 14.1). This method is very
easy in experienced hands, but it requires multiple steps, including dissection, release, and rotation of the oocyte.
Laser-Assisted Opening
The easiest way of opening the zona pellucida is by a laser beam. The use of 1.48 μm diode laser drilling for PB
biopsy was proposed in 1997 [24]. Animal experimentation showed the potential of this method for PB biopsy and
for assisted hatching [25] and allowed investigation of its proper mode of application [2]. Such studies revealed that it
is crucial to understand the effect of laser drilling and how the size and position of laser-drilled openings can influ-
ence further embryonic development, in particular, the type of hatching at the blastocyst stage [26]. Applying laser
technology in an inappropriate way could be potentially deleterious and have an impact on overall success rate.
The easiest way of opening the zona pellucida is by a laser beam.
Laser-assisted biopsy then entered the field of PB [25,27] and embryo biopsy [28] and has helped in reducing
the rate of biopsy damage and the time required for biopsy [29]. Due to its ease, laser-assisted biopsy is now also
widely used for biopsy of blastomeres [28,30] and blastocyst cells [31], and its advantage compared with acid
Tyrode’s solution has been reported previously [29].
A first slit is made by zona drilling
A second slit is made after

turning the oocyte by 90˚
FIGURE 14.1 Schematic presentation of zona opening for subsequent mechanical biopsy.

Polar Body Biopsy 211
Laser-Assisted Biopsy Procedure

For laser-assisted PB biopsy, the size of the drilled opening is usually in the range of 15–20 μm, but it can be
easily adjusted to the diameter of the aspiration capillary (Figure 14.2). Because the capillary is introduced
through the laser-drilled opening, there is no need for a sharp aspiration needle. This procedure allows the use of
flame-polished, blunt-ended aspiration needles and greatly reduces the risk of damaging the PB, the blastomere,
or the remaining oocyte or embryo, a major reason for why the biopsy procedure becomes safer, more accurate,
and more reliable. It has even helped to significantly reduce the number of cells that cannot be reliably diagnosed
as a result of technical problems during the biopsy procedure [3]. Another benefit of laser-assisted biopsy is
that laser drilling and subsequent biopsy can be performed without changing the culture dish or the capillaries,
in contrast to zona drilling that uses acid Tyrode’s solution. This may help to prevent contamination of samples
to be diagnosed by sensitive techniques such as polymerase chain reaction (PCR).
The simultaneous removal of the first and second PB is best accomplished if the oocyte is affixed to the holding
capillary, with PB1 at the 12 o’clock position and PB2 to the right of PB1 but in the same focal plane. The reason
for this positioning is the presence of a connective strand between PB2 and the oolemma of the oocyte. Further
details regarding the timing of PB biopsy and possible consequences linked to the existence of spindle remnants
within the connective strand are discussed below.
FIGURE 14.2 Laser-assisted polar body (PB) biopsy. For biopsy, the first and second PBs were aligned with a holding capillary
so that the second PB faced the biopsy capillary (a). Using a noncontact 1.48 μm diode laser, an opening was introduced into the
zona pellucida using two or three laser shots (b) through which the biopsy capillary could be easily introduced (c). The second PB is
usually connected to the oolemma via a cytoplasmic strand. To remove the second PB without damaging the oocyte, it is not recom-
mended to suck the second PB into the capillary; instead, the capillary is pushed slowly over the second PB and toward the first PB
(d). Once the first PB enters the capillary, the strand between the second PB and the oolemma will break due to shear stress and both
PBs can be easily removed (d), leaving the oocyte without any damage. PBs should be placed in one droplet for further processing
for FISH analysis (e) or in two different droplets if a PCR-based analysis will be performed.
An opening is drilled at 2 o’clock or 3 o’clock, and the biopsy capillary is moved into the perivitelline space
toward and then over the PBs. It is crucial to move the capillary further toward the other side of the perivitelline
space to help in breaking the connective membrane strand linking PB2. This procedure is supported by using
PB1 as a counterpart that, while moving into the capillary, will shear off the connective strand of PB2. At the
proper time, both PBs can be easily removed simultaneously.
Timing of PB Biopsy
Polarization microscopy has shown that some oocytes presenting a PB1 may be still in telophase I due to the
presence of a connective spindle strand between the PB1 and the oocyte (Figure 14.3) [32]. Spindle strands
are remnants of the meiotic division, and they occur during extrusion of PB1 and PB2. Spindle fibers, also named
the spindle bridge, are present only for a limited period that is usually 1–2 hr after extrusion of the first or second
PB. During this time, chromosomal material from the oocyte is still attached to these spindle fibers. If PB biopsy
is performed within a too short of a time after the formation of PB1 and PB2, there is a substantial risk to pull out
chromosomal material during biopsy, thereby enucleating the oocyte. However, as mentioned, after dissolution
of the spindle fibers the second PB still remains connected to the surface membrane of the oocyte and developing
embryo by a connective membrane strand.
There is also an ongoing discussion as to whether biopsy of PB1 and PB2 should be done separately (sequential
approach) or at the same time (simultaneous approach). Both approaches do work and have been tested in clinical
practice. Simultaneous biopsy of the first and second PB requires only one manipulation, thereby reducing stress
to the oocyte. A good time window is 8–14 hr after fertilization. A biopsy performed too early bears the risk of
spindle remnants in the second PB, whereas a biopsy performed too late might affect the quality of PB1 caused
by its disintegration or degeneration. Interestingly, the timing of biopsy of PB2 seems to influence amplification
results in array-CGH. It was reported that biopsy of PB2 too early (4–6 hr post-intracytoplasmic sperm injection
[ICSI]) may slightly lower the amplification efficiency. This effect disappeared after adjustment to later biopsy
times (>8 hr post-ICSI) [33].
Timing of polar bodies is crucial for both sequential and simultaneous biopsies.
The timings proposed for simultaneous biopsy do equally apply to sequential biopsy. Sequential biopsy over-
comes the potential problems of distinguishing PB1 and PB2 and of accurately separating PB1 and PB2 in case
of fragmentation.
FIGURE 14.3 Presence of a meiotic spindle bridge between first polar body (PB1) and the oocytes. The presence of a connective
spindle bridge can be assessed by polarization microscopy. Spindle fibers are displayed in red and characterize the corresponding
oocyte as being in the transition phase from metaphase I to metaphase II. As long as spindle fibers are visible, chromosomes in the
oocyte are attached, and the corresponding oocyte is in anaphase/telophase I. Removal of the PB1 shown would result in the with-
drawal of chromatin material from the oocyte and could potentially lead to enucleation.
FIGURE 14.4 Degenerated PB1. A degenerated PB1 can be seen beside an intact PB2 (a). After opening the zona pellucida both
PBs can be removed without further damage or loss (b).
Independent of the time of biopsy, degeneration of PBs (Figure 14.4) seems to be of minor importance for
array-CGH, but it can be a problem for fluorescence in situ hybridization (FISH)-based analysis because it may
contribute to diagnostic failures [34].
Potential Pitfalls of PB Biopsy

In all manipulation steps and zona opening techniques, it is important to drill only one opening. Especially if
PB1 and PB2 are biopsied in a sequential approach at different times, it is tempting to introduce another open-
ing to ease the procedure. However, the presence of two openings may cause problems at the time of hatching
because the embryo could hatch through both openings simultaneously and therefore may get trapped within
the zona [35].
Another important point is to generate a sufficiently large opening that allows consecutive hatching at the
blastocyst stage. It has been shown experimentally in mouse embryos that smaller openings (<15 μm) cause trap-
ping of the blastocyst at hatching, followed by degeneration [35].
Laser-drilled openings will stay permanently in the zona; therefore, gentle handling during subsequent t ransfer
of oocytes to other media droplets and even during the embryo transfer is recommended.
In case of the presence of fragmented PBs, all fragments have to be removed. Usually fragments stick together;
therefore, there is only a minor risk of mixing fragments from PB1 and PB2 (Figures 14.5 and 14.6). However,
once fragments are transferred onto a slide for subsequent FISH analysis, there is a tendency for spreading and
hence a potential risk to loose chromosomal material.
A position paper with relevant best-practice guidelines for PB biopsy for PGD and PGS has been published
by ESHRE [36].
PB1 or PB2, or Both?

The reliability of PB biopsy is strongly dependent on the accuracy of the diagnosis of PB1 and PB2. This
accuracy is indisputable for cases with structural chromosomal rearrangements and for the diagnosis of genetic
disorders. For PGS, a recent study showed that PB1 is more prone to meiotic errors in younger women than PB2,
but the opposite was found for older women [37]. Although one could conclude that for PGS PB1 is more impor-
tant in younger women and PB2 in older women, there would be a relative risk for aneuploidy. Clinical studies
on the concordance of PB1 and PB2 analyses and the corresponding oocyte gave direct proof for the need of
analyzing both PB1 and PB2 [33,38].
FIGURE 14.5 Attached fragments of PB1. During the course of fragmentation, the fragments stay together and remain loosely
connected as long as the PB is located within the perivitelline space.
FIGURE 14.6 Biopsy of a fragmented PB1. (a) A fragmented PB1 is located beside an intact and smaller PB2. (b) The PBs remain
together while the capillary is shifted over the PBs. (c) During final aspiration, the fragments stick together and (d) can be placed in
a separate medium droplet without loosing the connectivity of the two fragments.
Transferring PBs for Subsequent Analysis (FISH and Array-CGH)

The way in which PBs are transferred for subsequent analysis by FISH or by array-CGH differs substantially;
therefore, adequate procedures are required.
Transfer for FISH

Moving PBs into capillaries for transfer does require strict visual control. The best way to transfer PBs for FISH
onto a glass slide is by using the biopsy capillary (Figure 14.7) [39]. After aspiration, both PBs are placed, under
an inverted injection microscope, into a 0.1–0.2 μL drop of water located on a clean slide. PBs must be released
at the bottom of the drop and directly on the glass surface, otherwise they will float in the droplet and rupture
during evaporation, resulting in a loss of material. To follow the evaporation process, the slide has to be moved
immediately to a stereomicroscope with good optical contrast and a reasonably high magnification (80–100×).
PBs are small in diameter, and it is nearly impossible to find them on an unmarked slide. Therefore, the area with
the PBs must be encircled after drying with a diamond or tungsten pen on top of the slide (Figure 14.8) [39].
When PBs from several oocytes are placed on one slide, it is recommended to connect all circles with a line
drawn by the tungsten scribe. It is possible to visualize the tungsten mark in bright field as well as under fluores-
cence illumination. This allows identifying the position of the encircled PB and facilitates orientation at 100× oil
immersion magnification by following the line from one circle to the next.
With experience, it is easy to distinguish PB1 and PB2 during evaluation of the FISH signals. Thus, both PBs
can be placed in the same area. It is possible to allocate PBs from up to ten oocytes in individually encircled
FIGURE 14.7 Transfer of isolated polar bodies (PBs) onto a slide. The transfer of isolated PBs from the dish (seen in the back-
ground) into the droplet on the slide must be performed on the microscope stage. The setup shown here allows sliding the dish used
for biopsy backwards. Therefore, the aspiration capillary needs only to be lowered into the droplet for release of the PB. (Reprinted
with permission from Montag M, Textbook of Assisted Reproductive Techniques Volume One: Laboratory Perspectives, 4th ed.,
Informa Healthcare, New York, 2012, pp. 336–345.)
FIGURE 14.8 Identification of the polar body (PB) on the slide. This photograph is taken with a 10× phase contrast objective,
and the diamond circle surrounding the PB can be partially seen. The PB appears gray (arrow). (Reprinted with permission from
Montag M, Textbook of Assisted Reproductive Techniques Volume One: Laboratory Perspectives, 4th ed., Informa Healthcare,
New York, 2012, pp. 336–345.)
spots within an area of 10 mm. By using a round coverslip with a diameter of 12 mm, the amount of hybridization
probe needed can be minimized [39].
Transfer for Array-CGH

For any kind of molecular-based diagnosis, such as array-CGH, both PBs must be transferred into separate
reaction tubes and in a defined volume that fits to the required amplification protocol [33]. Because array-CGH
will be the most common application, this method is primarily described here. It is important to prefill reac-
tion tubes with 2.1–2.3 μL of phosphate-buffered saline (PBS) and to transfer PBs with 0.2–0.4 μL of medium
into this solution for obtaining a final maximum volume of 2.5 μL. Pipetting PBs in whatever volume directly
into a dry test tube may result in loss of PBs and amplification failure. The transfer is best accomplished by
using a plastic capillary with a fine tip, similarly to the capillaries used for oocyte denudation. Glass capillaries
are not recommended because these can break, entrap the PB, and make the DNA inaccessible to amplifica-
tion. Releasing the transferred medium at the sidewall of the PCR tube may result in PB sticking to the wall.
Subsequent centrifugation will cause rupture of the PB and spread the DNA material along the wall, resulting in
partial amplification and inaccurate diagnosis.
Transfer of polar bodies to slides or into tubes is one of the most crucial steps of the
whole procedure.
For FISH, the use of a high-contrast stereomicroscope is recommended. PBs can be easily identified in the
medium droplets and aspirated for transfer. It is advisable to rinse the capillary after transfer under visual control
to verify that the PB has been placed in the reaction tube, because this process cannot be directly visualized due
to the plastic material of the tubes. Needless to say, transferring PBs for subsequent single-cell DNA amplifica-
tion has to be done in a manner (and using proper equipment) that avoids potential contamination.
What Is the Predictive Value of PB Diagnosis?

The discussion on the value of PGS has caused a growing interest in PB biopsy and diagnosis [18]. ESHRE has
founded a task force on PGS and started a pilot study to evaluate the potential of PB biopsy and array-CGH for
PGS [38]. The ESHRE pilot study has resulted in a series of clinical and fundamental publications [40], and the
positive perception of this technology has prompted ESHRE to initiate a multicentric, randomized controlled
trial using this technology. However, some studies based on PB biopsy and array-CGH reported a high or accept-
able correlation for predicting of aneuploidy [38,41], and others questioned the accuracy of PB diagnosis due to
the high incidence of postzygotic errors [42]. These studies were done on array-CGH, but recent data showed that
quantitative PCR has a much higher accuracy compared with array-CGH [43]. Currently, we cannot exclude
that the approach of analyzing DNA from a single PB may be more prone to methodological impact compared
with TE biopsies with multiple cells.
Another topic that is controversially discussed is reciprocal chromosome aneuploidies in PBs. A recent study
showed that this situation mostly gives rise to normal euploid embryos [44], and the birth of a healthy child
has been reported from an oocyte with reciprocal aneuploid polar bodies [45]. So far, the general understand-
ing is that reciprocal chromosome aneuploidies in PBs may give rise to mosaic embryos that should not be
transferred [46].
Conclusions
PB biopsy has been proven as sufficiently effective for the diagnosis of structural and numerical chromosome
aberrations in human oocytes by using FISH [27,47] and array-CGH [14]. Nevertheless, the use of PB biopsy and
array-CGH for PGS is still a matter of debate due to cost effectiveness, the high incidence of postmeiotic aneuploi-
dies that are undetectable by the PB approach [42], and the d iagnostic procedure [43]. There is growing advocacy
for TE biopsy and array-CGH as the new gold standard.
REFERENCES
1. Verlinsky Y, Ginsberg N, Lifchez A, et al. Analysis of the first polar body: Preconception genetic diagnosis. Hum
Reprod 1990; 5: 826–9.
2. Handyside AH, Kontogianni EH, Hardy K, et al. Pregnancies from biopsied human preimplantation embryos
sexed by Y-specific DNA amplification. Nature 1990; 244: 768–70.
3. Montag M, van der Ven K, van der Ven H. Erste klinische Erfahrungen mit der Polkörperbiopsie in Deutschland
[First clinical experience with polar body biopsy in Germany]. J Fertil Reprod 2002; 4: 7–12.
4. Verlinsky Y, Cieslak J, Freidine M, et al. Pregnancies following pre-conception diagnosis of common a neuploidies
by fluorescent in-situ hybridization. Hum Reprod 1995; 10: 1923–7.
5. Staessen C, Platteau P, Van Assche E, et al. Comparison of blastocyst transfer with or without preimplantation
genetic diagnosis for aneuploidy screening in couples with advanced maternal age: A prospective randomized
controlled trial. Hum Reprod 2004; 19: 2849–58.
6. Stevens J, Wale P, Surrey ES, et al. Is aneuploidy screening for patients aged 35 or over beneficial? A prospective
randomized trial. Fertil Steril 2004; 82 (Suppl 2): 249.
7. Mastenbroek S, Twisk M, Van Echten-Arends J, et al. In vitro fertilization with preimplantation genetic screening.
N Engl J Med 2007; 357: 9–17.
8. Blockeel C, Schutyser V, De Vos A, et al. Prospectively randomized controlled trial of PGS in IVF/ICSI patients
with poor implantation. Reprod Biomed Online 2008; 17: 848–54.
9. Hardarson T, Hanson C, Lundin K, et al. Preimplantation genetic screening in women of advance maternal age
decrease in clinical pregnancy rate: A randomized controlled trial. Hum Reprod 2008; 23: 2806–12.
10. Jansen RP, Bowman MC, de Boer KA, et al. What next for preimplantation genetic screening (PGS)? Experience
with blastocyst biopsy and testing for aneuploidy. Hum Reprod 2008; 23: 1476–8.
11. Mersereau JE, Pergament E, Zhang X, et al. Preimplantation genetic screening to improve in vitro fertilization
pregnancy rates: A prospective randomized controlled trial. Fertil Steril 2008; 90: 1287–9.
12. Staessen C, Verpoest W, Donoso P, et al. Preimplantation genetic screening does not improve delivery rate in
women under the age of 36 following single-embryo transfer. Hum Reprod 2008; 23: 2818–25.
13. Meyer L, Klipstein S, Hazlett W, et al. A prospective randomized controlled trial of preimplantation genetic
screening in the “good prognosis” patient. Fertil Steril 2009; 91: 1731–8.
14. Schoolcraft WB, Katz-Jaffe MG, Stevens J, et al. Preimplantation aneuploidy testing for infertile patients of
advanced maternal age: A randomized prospective trial. Fertil Steril 2009; 92: 157–62.
15. Debrock S, Melotte C, Spiessens C, et al. Preimplantation genetic screening for aneuploidy in embryos after
in vitro fertilization (IVF) does not improve reproductive outcome in women over 35: A prospective controlled
randomized trial. Fertil Steril 2010; 93: 364–73.
16. Anderson R, Pickering S. The current status of preimplantation genetic screening: British Fertility Society Policy
and Practice Guidelines. Hum Fertil 2008; 11: 71–5.
17. American Society for Reproductive Medicine (ASRM). Preimplantation genetic testing: A Practice Committee
opinion. Fertil Steril 2008; 90 (Suppl): S136–43.
18. Geraedts J, Collins J, Gianaroli L, et al. What next for preimplantation genetic screening? A polar body approach!
Hum Reprod 2010; 25: 575–7.
19. Harper J, Coonen E, De Rycke M, et al. What next for preimplantation genetic screening (PGS)? A position state-
ment from the ESHRE PGD Consortium Steering Committee. Hum Reprod 2010; 25: 821–3.
20. Nicolaidis P, Petersen MB. Origin and mechanisms of non-disjunction in human autosomal trisomies.
Hum Reprod 1998; 13: 313–19.
21. Gordon JW, Talansky BE. Assisted fertilization by zona drilling: A mouse model for correction of oligospermia.
J Exp Zool 1987; 239: 347–81.
22. Malter HE, Cohen J. Blastocyst formation and hatching in vitro following zona drilling of mouse and human
embryos. Gamete Res1989; 24: 67–80.
23. Cieslak J, Ivakhenko V, Wolf G, et al. Three-dimensional partial zona dissection for preimplantation genetic
diagnosis and assisted hatching. Fertil Steril 1999; 71: 308–13.
24. Montag M, van der Ven K, Delacrétaz G, et al. Efficient preimplantation genetic diagnosis using laser assisted
microdissection of the zona pellucida for polar body biopsy followed by primed in situ labelling (PRINS). J Assist
Reprod Genet 1997; 14: 455–6.
25. Montag M, van der Ven K, Delacrétaz G, et al. Laser assisted microdissection of zona pellucida facilitates polar
body biopsy. Fertil Steril 1998; 69: 539–42.
26. Montag M, Koll B, Holmes P, et al. Significance of the number of embryonic cells and the state of the zona
pellucida for hatching of mouse blastocysts in vitro versus in vivo. Biol Reprod 2000; 62: 1738–44.
27. Montag M, van der Ven K, Dorn C, et al. Outcome of laser-assisted polar body biopsy. Reprod Biomed Online
2004; 9: 425–9.
28. Boada M, Carrera M, De La Iglesia C, et al. Successful use of a laser for human embryo biopsy in preimplantation
genetic diagnosis: Report of two cases. J Assist Reprod Genet 1997; 15: 301–5.
29. Joris H, De Vos A, Janssens R, et al. Comparison of the results of human embryo biopsy and outcome of PGD
after zona drilling using acid Tyrode medium or a laser. Hum Reprod 2003; 18: 1896–902.
30. Licciardi F, Gonzalez A, Tang YX, et al. Laser ablation of the mouse zona pellucida for blastomere biopsy.
J Assist Reprod Genet 1995; 12: 462–6.
31. Veiga A, Sandalinas M, Benkhalifa M, et al. Laser blastocyst biopsy for preimplantation diagnosis in the human.
Zygote 1997; 5: 351–4.
32. Montag M, Schimming T, van der Ven H. Spindle imaging in human oocytes: The impact of the meiotic cell cycle.
33. Magli MC, Montag M, Köster M, et al. Implementing polar body biopsy and chromosome copy number analysis
by array comparative genomic hybridisation: Technical experiences from the ESHRE PGS Task Force pilot study.
Hum Reprod 2011; 26: 3181–4.
34. Munné S, Dailey T, Sultan KM, et al. The use of first polar bodies for preimplantation diagnosis of aneuploidy.
Hum Reprod 1995; 10: 1015–21.
35. Montag M, van der Ven H. Laser-assisted hatching in assisted reproduction. Croat Med J 1999; 40: 398–403.
36. Harton GL, Magli MC, Lundin K, et al. ESHRE PGD Consortium/Embryology Special Interest Group—best
practice guidelines for polar body and embryo biopsy for preimplantation genetic diagnosis/screening (PGD/PGS).
Hum Reprod 2011; 26: 41–6.
37. Fragouli E, Alfarawati S, Goodall NN, et al. The cytogenetics of polar bodies: Insights into female meiosis and
the diagnosis of aneuploidy. Mol Hum Reprod 2011; 17: 286–95.
38. Geraedts J, Montag M, Magli MC, et al. Polar body array CGH for prediction of the status of the corresponding
oocyte. Part I: Clinical results. Hum Reprod 2011; 26: 3173–80.
39. Montag M. Polar body biopsy and its clinical application. In: Textbook of Assisted Reproductive Techniques
Volume One: Laboratory Perspectives, 4th ed. (eds. Gardner D, Weissman A, Howles C, Shoham Z). New York:
Informa Healthcare, 2012, pp. 336–45.
40. Handyside AH, Montag M, Magli MC, et al. Multiple meiotic errors caused by predivision of chromatids in
women of advanced maternal age undergoing in vitro fertilization. Eur J Hum Genet 2012; 20: 742–7.
41. Christopikou D, Tsorva E, Economou K, et al. Polar body analysis by array comparative genomic hybridization
accurately predicts aneuploidies of maternal meiotic origin in cleavage stage embryos of women of advanced
maternal age. Hum Reprod 2013; 28: 1426–34.
42. Capalbo A, Bono S, Spizzichino L, et al. Sequential comprehensive chromosome analysis on polar bodies, blas-
tomeres and trophoblast: Insight into female meiotic errors and chromosomal segregation in the preimplantation
window of embryo development. Hum Reprod 2013; 28: 509–18.
43. Treff NR. qPCR based CCS. Reprod Biomed Online 2013; 26 (Suppl 1): 55.
44. Forman EJ, Treff NR, Stevens JM, et al. Embryos whose polar bodies contain isolated reciprocal chromosome
aneuploidy are almost always euploid. Hum Reprod 2013; 28: 502–8.
45. Scott RT, Jr, Treff NR, Stevens J, et al. Delivery of a chromosomally normal child from an oocyte with reciprocal
aneuploid polar bodies. J Assist Reprod Genet 2012; 29: 533–7.
46. Kuliev A, Verlinsky Y. Meiotic and mitotic non-disjunction: Lessons from preimplantation genetic diagnosis.
Hum Reprod Update 2004; 10: 401–7.
47. Verlinsky Y, Tur-Kaspa I, Cieslak J, et al. Preimplantation testing for chromosomal disorders improves reproduc-
tive outcome of poor-prognosis patients. Reprod Biomed Online 2005; 11: 219–25.
15
Invasive Techniques: Embryo Biopsy at the Cleavage Stage
Anick De Vos
Introduction
Cleavage-stage embryo biopsy of single blastomeres allows the analysis of male and female genetic contribution
to the final embryo, by means of polymerase chain reaction (PCR) or fluorescence in situ hybridization (FISH)
analysis at the single-cell level. One or two blastomeres are removed from the embryo in the morning of Day 3,
at about 68–72 hr after microinjection. The embryos are preferably in the third mitotic division, presenting at
least five cells or, more ideally, eight cells or more. Enough time for genetic or chromosomal testing is available
if blastocyst transfer is performed on Day 5.
So far, cleavage-stage biopsy is the most widely used procedure (87.9% of all cycles reported by the European
Society of Human Reproduction and Endocrinology (ESHRE) Preimplantation Genetic Diagnosis [PGD]
Consortium), far exceeding polar body biopsy (11.7%) and blastocyst biopsy (0.4%) [1]. For many years, the
method remained unchanged. Zona opening can be done mechanically; chemically by using acidic Tyrode’s solu-
tion (AT); or, more recently, by using laser energy. Blastomere removal is mainly done by aspiration; however,
extrusion and flow displacement are also used, but to a much lesser extent. Clinical application of cleavage-stage
embryo biopsy resulted in the first pregnancies [2] and birth of a normal girl [3]. These achievements were based
on the preclinical work [4] concluding that in vitro preimplantation development of biopsied human embryos is
not adversely affected by the removal of one or two cells at the eight-cell stage. Since then, the number of in vitro
fertilization (IVF) cycles, including embryo biopsy in view of PGD, has increased significantly [1,5].
This chapter presents an overview of all aspects of cleavage-stage embryo biopsy and aims to contain a very
specific practical guide on how to perform the procedure. The following aspects are addressed: developmental
stage and embryo inclusion criteria for biopsy, precautions to avoid contamination with maternal or paternal
DNA, equipment for embryo biopsy, zona opening procedures, cellular removal and maximal mass reduction,
compaction/decompaction, multinucleation and anucleates, dealing with cell lysis, postbiopsy development and
finally, advantages and disadvantages of cleavage-stage embryo biopsy.
Developmental Stage and Embryo Inclusion Criteria for Biopsy

In cases where the paternal contribution to the embryo needs to be diagnosed, biopsy at preimplantation devel-
opmental stages is needed. Obviously, the developmental stage allowing maximum retrieval of cells at biopsy
with a minimum reduction in pregnancy potential of the embryos after the procedure would be most appropriate.
Two- and four-cell human embryos do not fit into this assumption. The best moment for embryo biopsy would
then be the eight-cell stage, normally in the morning of Day 3. At this stage, all the cells are still totipotent as
confirmed by lineage tracing [6], and the embryos are not yet compacting extensively.
Unfortunately, not all embryos are present in the eight-cell stage in the morning of Day 3 (Figure 15.1). Both
delayed and advanced embryos are encountered. Embryos may also differ in quality according to the amount of
anuclear fragments present. Blastomere sizes can deviate from the stage-specific division pattern; furthermore,
vacuoles, granulation, and multinucleation may be present. It is recommended to establish biopsy criteria to
determine which embryos should be included for biopsy (Table 15.1). Mostly, embryos with <50% anuclear
219
FIGURE 15.1 Cleavage-stage embryo biopsy is usually performed in the morning of Day 3. (a) Ideally, the embryos are in the eight-
cell stage, having no fragmentation and showing stage-specific blastomere sizes. (b) An advanced-stage embryo on Day 3 with no
fragmentation. (c) An embryo with more fragmentation (between 20% and 50%); however, it would still be included for embryo biopsy.
TABLE 15.1
Inclusion Criteria for Cleavage-Stage Embryo Biopsy
• Embryos resulting from 2-pronuclear (PN) oocytes (occasionally 1-PN embryos due to a missed second pronucleus at fertilization
check, given a good morphological quality)
• ≥6 cell (occasionally 5-cell embryos, if <20% fragments and cell-stage specific)
• <50% anuclear fragments
• Preferably embryos with cell-stage-specific blastomere sizes; however, also those with cell-stage-aspecific blastomere sizes
• Preferably no granulation
• Preferably no vacuoles
• <50% multinucleated blastomeres present in the embryo
fragmentation and with at least six blastomeres are suitable for biopsy. Preferably, embryos have stage-specific
cell sizes; however, stage-aspecific cell sizes would not represent an exclusion criterion for embryo biopsy.
Occasionally, five-cell embryos could be included for one-cell removal if the embryos have <20% anuclear frag-
mentation and if the blastomere sizes are at least according to the division pattern (i.e., three larger and two small
blastomeres). The number of cells to be removed (one or two) is dictated by the diagnostic protocol, wherein
a secure result is crucial. Vacuoles and granulation are preferentially not present. Embryos with >50% of their
blastomeres multinucleated are not included for embryo biopsy.
It is common practice to include only embryos resulting from 2-pronuclear (2-PN) normal fertilization.
Occasionally, 1-pronuclear (1-PN) embryos could be included for embryo biopsy, given a good morphological
quality and given that the diagnostic protocol can distinguish between a haploid embryo and a diploid one due to
a missed second pronucleus at fertilization check.
On average, about 80% of all embryos resulting from 2-PN fertilization are suitable for embryo biopsy on
Day 3. Obviously, biopsied embryos should be cultured singly in individual droplets, or even better in individual
culture wells, thereby avoiding possible mixing of the embryos postbiopsy and diagnosis. Embryos are well
rinsed postbiopsy to remove any trace of biopsy medium, which might be Ca2+/Mg2+-free medium, or acid in
case of chemical zona opening. For this purpose, wash droplets are available in the culture dish.
Precautions to Avoid Contamination with Maternal or Paternal DNA

Intracytoplasmic sperm injection (ICSI) is recommended for all PCR cases to reduce the chance of paternal
contamination from sperm attached to the zona pellucida [7]. In contrast, both ICSI and conventional IVF are
acceptable for FISH cases [7].
Embryo Biopsy at the Cleavage Stage 221
TABLE 15.2
Precautions When Performing Cleavage-Stage Embryo Biopsy
• ICSI for all PCR cases; ICSI and IVF acceptable for FISH cases
• Careful oocyte denudation
• Careful embryo rinse after embryo biopsy (remove Ca2+/Mg2+ traces present in the culture medium)
• Always replace aspiration pipette in cases of cell lysis
• Lysed cells are not used for PCR analysis (cf. maternal DNA contamination); for FISH analysis, the nucleus can occasionally be
recovered
• Single embryo culture in individual droplets or individual culture dish wells
• Careful embryo rinse post–embryo biopsy (remove biopsy medium traces or acidic Tyrode in cases of chemical zona opening)
Cumulus cells may represent a source of maternal DNA contamination and may thus lead to misdiagnosis [8].
Therefore, extra care is taken at the moment of oocyte denudation to ensure that all these cells are removed from
the zona pellucida.
Table 15.2 summarizes all precautions to be considered when performing cleavage-stage embryo biopsy.
Equipment for Cleavage-Stage Embryo Biopsy

Embryo biopsy is performed on an inverted microscope equipped with a warmed stage, as used for
ICSI and embryo evaluations (Leica, Nikon, Olympus, and Zeiss represent well-known brands). This
m icroscope should also be equipped with micromanipulators connected to oil-filled (or air-filled) micro-
injectors: one m icroinjector is for holding the embryo and the other microinjector is for aspiration of the
blastomeres.
In case of mechanical or chemical zona opening, a double holder is needed. For mechanical zona opening, the
second holder contains the microneedle used for partial zona dissection (PZD). No additional microinjector is
needed in this case. For chemical zona opening, the second holder contains the micropipette with AT solution.
Connection to a third microinjector is needed to aspirate and expel the acidic solution.
Microtools used for embryo biopsy (e.g., holding pipette, aspiration pipette, PZD needle, assisted hatching
pipette; Table 15.3) are commercially available. Cook, Eppendorf, and Humagen microtools are well-established
brands. The holding pipette (inner diameter, 17 μm) is used to hold the embryo during the biopsy procedure.
A slightly larger inner diameter can be considered to provide greater stability and holding power for embryos
during biopsy. Blastomere biopsy micropipettes or aspiration pipettes have an inner diameter of 28–32 μm and
are used to remove the blastomeres by aspiration. PZD micropipettes or needles are pulled with a long, thin taper
to a sharp point. These micropipettes can be used for mechanical zona opening by physically piercing and rub-
bing the zona pellucida. Assisted hatching pipettes have a blunt end with a typical inner diameter of 8–10 μm.
They are used to create a hole in the zona pellucida using AT solution. For convenient manipulation in plastic
Petri dishes, all pipettes display a 30° angle.
Instead of mechanical or chemical zona opening, a laser can be used to open the zona pellucida. Of course,
a laser system represents quite expensive equipment. However, laser technology is considered less detrimental,
extremely accurate, and simple and quick to use. Three main types of lasers are currently in use (Table 15.3):
Octax Laser Shot (MTG GmbH, Germany), similar to the older version named Fertilase; Zilos-tk (Hamilton
Thorne, United States); and Saturn 5 (Research Instruments, United Kingdom). All represent 1.48 μm infrared
diode lasers, with laser power 100–150, 300, and 400 mW, respectively, and pulse lengths in the range of
milliseconds or even microseconds.
Embryo biopsy is performed in Petri dishes, containing 25–50 μL droplets of HEPES-buffered human tubal fluid
(HTF) medium supplemented with 0.5% human serum albumin or 10–20 μL droplets of Ca2+/Mg2+-free medium
(G-PGD, Vitrolife) covered with mineral oil. When using Ca2+/Mg2+-free medium, embryos to be biopsied are
rinsed twice through this medium (two separate droplets) before biopsy to remove calcium and magnesium
TABLE 15.3
Microtools for Embryo Biopsy and Laser Systems for Zona Opening
Holding pipette Inner diameter 17 μm to hold the embryo
Aspiration pipette Inner diameter 28–32 μm to aspirate blastomeres
PZD needle For mechanical zona opening
Assisted hatching pipette Inner diameter 8–10 μm for chemical zona opening
Laser systems For laser zona opening
Octax Laser Shot, MTG GmbH, Germany
Zilos-tk, Hamilton Thorne, United States
Saturn 5, Research Instruments, United Kingdom
traces present in the culture medium. Some centers manipulate with three to four biopsy droplets per dish,
whereas other centers prefer to have only one single biopsy droplet and thus one embryo per Petri dish. For
chemical zona opening, one additional droplet of AT solution (5 μL) is placed, allowing to prime the drilling
pipette. AT solution is commercially available.
Zona Opening Procedures

Cleavage-stage embryo biopsy of human preimplantation embryos always involves two steps: opening of the
zona pellucida and subsequent removal of cellular material by aspiration. Zona opening can be done in three
ways: mechanical, chemical, or using laser energy.
Human zona pellucida opening procedures date from the late 1980s and early 1990s, when they were
applied to help the hatching process of the embryo or to aid the fertilization process of the oocyte. At that
time, zona penetration or opening was performed using only mechanical or chemical means. Mechanical
zona opening for cleavage-stage embryo biopsy is applied clinically; however, in all data collections reported
by the ESHRE PGD Consortium, the amount of cycles using mechanical zona opening ranged from only
1% to 10% of all cycles with PGD (Table 15.4). For many years, AT drilling was used in the majority of clini-
cal biopsy cycles. In 2002, still two-thirds of the PGD cycles used AT to open the zona pellucida [9]. Since
then, laser zona opening became more and more popular, reaching up to 60% of the clinical PGD cycles
reported by the ESHRE PGD Consortium [1] (data collection XI, data up to 2008) at the expense of AT drill-
ing (only 30% of all PGD cycles then).
Mechanical Zona Opening

To open the zona pellucida mechanically, PZD has been described and illustrated [10]. The zona pellucida is
pierced with a sharp and closed microneedle through both sides. The size of the slit is determined by the distance
between the first and second point at which the zona is pierced. With the microneedle piercing through the zona
at both sides, the trapped area is then rubbed against the holding pipette until this area has been completely
abraded. Usually, standardized slits of ±30–40 μm in length (2 μm in width) are created as such. The procedure
in combination with embryo biopsy was described by Grifo et al. [11] for mouse embryos at the four- to eight-cell
stages. When a second slit perpendicular to the first slit is created, a larger V-shaped opening can be obtained
that is suitable for blastomere aspiration [12]. One advantage may be that the embryo remains protected until
expansion because the zona flap created in this way closes after aspiration pipette removal. This is in contradic-
tion to chemical zona opening, where an actual hole is created.
Opposite to the holding pipette, a double holder setup is used to contain the microneedle and the aspiration
pipette at the same time. Mechanical opening of the human zona pellucida by piercing and rubbing is relatively
simple and nontraumatic, and the risk of embryo damage at cleavage stages is also limited [13]. Nevertheless, the
application of PZD for the purpose of removing cleavage-stage blastomeres for genetic diagnosis has been very
limited in comparison with AT drilling and laser drilling (Table 15.4).
TABLE 15.4
Distribution of Zona Opening Procedures Used According to ESHRE PGD Consortium Data Collections
(in Percentages of PGD Cycles)
Data Collection II III IV V I VII VIII IX X XI
(up to Year) (2000) (2001) (2001) (2002) (2003) (2004) (2005) (2006) (2007) (2008)
Mechanical 1.3 1.1 5.3 5.4 6.1 8.2 9.9 9.7 9.2 8.9
AT drilling 79.4 69.4 64.5 60.7 55.0 47.1 41.7 35.8 32.5 30.5
Laser drilling 19.3 29.5 30.2 33.9 38.8 44.7 48.4 54.6 58.3 60.6
Note: AT, acidic Tyrode.
Chemical Zona Opening (AT Drilling)

Chemical zona drilling using AT (pH 2.3) represents a much cheaper option compared with the more sophis-
ticated laser systems. Chemical zona drilling creates a larger, rounder hole in the zona pellucida, and the hole
size is not always easy to control. It should be recognized that the human zona pellucida represents a stratified
bilayered structure [14]. The outer layer of the zona is indeed easily dissolved, whereas the inner layer may vary
greatly in its susceptibility to dissolution. When the inner layer is refractory to drilling, often a relatively large
quantity of AT is needed to breach the zona pellucida. As a consequence, local acidification or even deposition
of AT in the perivitelline space may occur and should be considered when evaluating the safety of this approach
of zona opening. In addition to immediate cell lysis, subtle damage to cells due to acidic exposure may interfere
with further development or implantation. To avoid acidic exposure of cells and thus reduce the blastomere lysis
rate, drilling and blastomere aspiration using a single, larger drilling/biopsy pipette [15,16] seems not a good
approach. Chemical zona drilling followed by blastomere aspiration is normally done with separate pipettes (one
drilling pipette, inner diameter 8–10 μm, and one aspiration or biopsy pipette, inner diameter 28–32 μm) using
a double holder setup [17]. An intact blastomere rate of up to 95% can be obtained [17].
Once the microtools are fixed and aligned on the inverted microscope, the embryo is visualized and fixed on
the holding pipette. The drilling pipette is filled with a small amount of AT. Drilling is then performed by releas-
ing acid onto the zona pellucida, preferentially between two blastomeres, or otherwise, in front of anuclear frag-
ments. This approach should minimize the deleterious effect of the acid on the cells. Upon rupture of the zona
pellucida, immediate aspiration is applied to remove excess acidic solution. Embryos can be washed by moving
them to another area in the medium droplet, before continuing with blastomere aspiration. In cases of cell lysis,
pipettes are always replaced in order to avoid DNA contamination to the next embryo. The present procedure is
in itself extremely simple in terms of equipment; however, some technical skill is required when exposing the
embryo to AT. The opening should be of appropriate size, while limiting the extent and duration of exposure of
the embryo. The mean diameter of the gap thus created is about 20 μm (range, 10–36 μm). The rate of dissolution
of the zona may vary between 30 s and 2 min.
In the early 2000s, representing 60%–80% of all PGD cycles reported by the ESHRE PGD Consortium [9],
the use of AT drilling started to decline at the expense of laser zona drilling (Table 15.4), with laser zona drill-
ing being certainly less detrimental [17] and extremely accurate, quick, and simple (taking only a few seconds
to perform zona openings).
Laser Zona Opening

Within the infrared region, a 1.48 μm noncontact diode laser has been described for microdissection of mouse
and human zona pellucida [18,19]. This system was used for cleavage-stage embryo biopsy [20], and its safety
and efficacy were demonstrated on the basis of one single pregnancy. Joris et al. [17] later reported similar preg-
nancy rates obtained with laser drilling compared with AT drilling. Also, more intact blastomeres were obtained
with laser drilling (up to 98% compared with 95% with AT). Additional studies compared the laser technology
with AT solution for zona opening [21–23], concluding that laser represents a suitable alternative, being quick,
easy, and safe, resulting in similar postbiopsy development. Safety evidence at the molecular level for laser
TABLE 15.5
Different Irradiation Times, According to Laser Power Specification, Have Been Used
for Cleavage-Stage Embryo Biopsy
Laser System Pulse Numbers, Pulse Length Opening Diameter References
Octax Laser Shot Two, exceptionally three, pulses of 5–8 ms 20–30 μm [17]
Zilos-tk 6–10 laser shots, 500 μs 20 μm [29]
Zilos-tk One or two laser pulses, 0.5 ms Not specified [30]
Zilos-tk 2–4 1 ms single pulses at 100% power 20–25 μm [31]
Saturn 5 One laser shot, 1 ms 16.5 μm [28]
zona pellucida perforation was obtained with mouse eight-cell embryos by measuring heats-shock protein tran-
scription [24]. These reassuring data opened the way for a steady increase in the use of laser zona drilling for
cleavage-stage biopsy (Table 15.4).
The laser uses high-energy light and consequently heat to dissolve or disintegrate the zona pellucida. The hole
formation can be explained by a local photothermolysis of the protein matrix [25]. When the light from the laser
comes in contact with the medium surrounding the embryo, heat is dissipated and transferred through the water
of the medium, creating temperature gradients in increasing concentric circles from the laser beam [26,27].
It has been estimated that the temperature in the surrounding medium can increase to 60°C–80°C [25]. Thermal
effects of the laser pulses around the embryo and on the blastomeres proper should be considered as a safety
issue [28]. Blastomeres in the immediate vicinity of the laser beam should not be damaged. Therefore, exposure
time should be limited, and a safe working distance away from blastomeres should be respected. A laser has three
properties that determine its performance and hence its effect on the embryo and surrounding medium: wave-
length, power, and pulse length. Typical wavelengths within the near-infrared light spectrum (2500–750 nm) are
considered to be nonmutagenic in contrast to ultraviolet (UV) radiation. Laser power used for embryo biopsy
ranges from 100 to 400 mW and is constant for each specific laser. Each increase or decrease in power will
influence the hole diameter and also the temperature gradient through the medium and the embryo. Pulse length
refers to the amount of time or duration of the laser beam. Pulse lengths range from 20 ms (Fertilase) to >1 μs
(Saturn Zilos). A laser with a high power and short pulse length will produce smaller temperature gradients than
a laser with low power and longer pulse lengths. A smaller temperature gradient will less likely compromise
embryo development [28].
Exposure time with laser should be limited, and a safe working distance away from blastomeres
should be respected.
The setup for laser zona pellucida drilling involves a laser beam that is tangentially guided to the zona pellu-
cida of the embryo. The embryo is positioned to place a region of the zona pellucida on the aiming spot. The hole
size can be chosen precisely by varying the irradiation time. Typically, a trench-like hole is produced. According
to laser power specification, different irradiation times have been used (Table 15.5). Given pulse numbers and
pulse lengths are indicative. According to any system used an individual approach has to be taken because pulse
length is also dependent on, for example, medium, temperature, objective coating, and dishes. It is important to
perforate the zona completely without harming the embryonic cells with the laser shot(s). The holes obtained
with laser are more precise than the ones obtained with AT.
Cellular Removal, Maximal Reduction

Once the zona pellucida of the embryo has been opened, the blastomeres inside become freely accessible.
Aspiration is most widely used to remove cells [1] (Table 15.6). The routine clinical use of flow displacement [32]
has remained very limited (0.1% of all cycles reported by the ESHRE PGD Consortium). This method requires
TABLE 15.6
Distribution of Blastomere Removal Procedures Used
According to ESHRE PGD Consortium Data Collection
ESHRE PGD Consortium No. of Cycles
Data Collection I-XI Reported %
Cleavage aspiration 26,284 93.4
Cleavage extrusion 1834 6.5
Cleavage flow displacement 38 0.1
FIGURE 15.2 (a) For the removal of human blastomeres by aspiration, an aspiration pipette is introduced into the perivitelline
space through the hole in the zona pellucida to reach a blastomere. (b, c) One blastomere is removed by gentle aspiration. Cells may
be aspirated completely and then removed; alternatively, cells are only partially aspirated and pulled out (given full decompaction
and thus no adherence to other blastomeres).
the production of two separate holes and considerable skill to displace the blastomere of choice by the injec-
tion of culture medium through the second opening. Blastomere removal by extrusion [33] is applied clinically,
however, to a much lesser extent than blastomere aspiration (6.5% vs. 93.4% of all cycles reported by the ESHRE
PGD Consortium). The blastomere is extruded through the opening by pushing against the zona pellucida at
another site using a blunt pipette.
For the removal of human blastomeres by aspiration (Figure 15.2), an aspiration pipette (inner diameter,
28–32 μm) is introduced into the perivitelline space through the hole in the zona to reach a blastomere. Close
location of the blastomeres respective to the opening allows limited penetration of the microtool. One or two
blastomeres are removed by gentle aspiration. Cells may be aspirated completely and then removed; alterna-
tively, cells are only partially aspirated and pulled out (given full decompaction and thus no adherence to other
blastomeres).
Based on preclinical work [4], up to one-quarter of the embryo can be removed without impairment of its
further in vitro development. However, there has been considerable discussion on whether the removal of two
cells from a seven- or more-cell-stage embryo reduces its capacity to implant more than if only one cell were
removed [34]. Of prime concern is that the diagnosis should be safe, that is, accurate and efficient, and misdi-
agnosis should be avoided at any price [8]. Recent evidence has accumulated that two-cell biopsy seems indeed
more detrimental than one-cell biopsy. Indirect evidence came from cryopreservation analogy [35], assuming
that cell loss from biopsy can be compared with cell loss after thaw of frozen cleaved embryos. A disproportion-
ate reduction in implantation potential according to the cell number lost has been described previously [35].
Direct evidence was obtained from a prospective randomized clinical trial [36] and from a prospective cohort
study of single blastocyst transfers [37]. Removal of two blastomeres (Figure 15.3) decreases the likelihood of
blastocyst formation compared with the removal of one blastomere. However, delivery rates with live birth per
started cycle were not significantly different (20.2% with one-cell biopsy and 17.2% with two-cell biopsy) [36].
In a prospective cohort of single blastocyst transfers, the clinical outcome from one-cell biopsy was significantly
better (37.4% live birth per transfer) than that of two-cell biopsy (22.4%) and comparable with when no interven-
tion on Day 3 was performed on the eight-cell embryos (35.0%) [37]. However, more recently, randomized single
FIGURE 15.3 Removal of two blastomeres with a clear single nucleus present in each of the two blastomeres.
blastomere biopsy of one Day 3 embryo and its transfer together with a nonbiopsied sibling embryo showed that
mostly the nonbiopsied embryo implanted (11/13) and led to the delivery [31]. In contrast, trophectoderm biopsy
at the blastocyst stage had no meaningful impact on the developmental competence of the embryo as measured
by implantation and delivery rates [31].
Removal of two blastomeres decreases the likelihood of blastocyst formation compared with the
removal of one blastomere.
Regarding cleavage-stage biopsy, the biopsy of one cell is recommended, given sufficient safeguards
for a correct diagnosis rather than the biopsy of two cells, at the expense of the implantation potential.
Technical improvements at the diagnostic level have allowed similar diagnostic accuracy to be obtained on
one blastomere compared with two blastomeres for certain indications or tests [38]. However, certain types
of analysis (e.g., translocations, aneuploidy screening) will benefit from or will even continuously necessitate
having two cells available for diagnosis. This is especially important because chromosomal mosaicism is com-
monly seen in cleavage-stage embryos. However, to alleviate chromosomal mosaicism at the cleavage stage, as
well as the invasiveness of two-cell biopsy, several laboratories are shifting toward blastocyst trophectoderm
biopsy (more cells available for diagnosis, safer than cleavage-stage biopsy [31]) in combination with array-
CGH technology.
Compaction/Decompaction
Although full compaction does not occur before the 16- to 32-cell stage, membrane adhesion mechanisms [39]
may render the biopsy procedure at the seven- or eight-cell stage rather difficult to perform because the blas-
tomeres show a strong tendency to adhere to each other. Ca2+/Mg2+-free medium has been used to loosen the
membrane adhesions between blastomeres [40,41], thereby allowing an easier removal of cells and resulting in
less blastomere lysis and a shorter procedure time. Subsequent embryo development to the blastocyst stage was
not affected by the choice of biopsy medium, even when embryos were exposed to Ca2+/Mg2+-free medium for
45 min [41]. Embryo biopsy may either be performed completely in decompaction medium, or otherwise, to limit
the exposure time, embryos can be just preincubated for 5–10 min (normally sufficient for full decompaction)
before the biopsy procedure.
Of course, the safety of embryo decompaction (using Ca2+/Mg2+-free medium) on further development, viabil-
ity, and implantation should be critically evaluated. So far, there are no indications of an adverse effect; however,
randomized controlled comparisons are lacking (Figure 15.4).
FIGURE 15.4 (a,b) Different degrees of embryo compaction can be observed in embryos to be biopsied. (c, d, e) To facilitate the
biopsy procedure, Ca2+/Mg2+-free medium is used to loosen the membrane adhesions between blastomeres. (c) represents the same
embryo as in panel (a) in a decompacted stage; (d,e) represent the same embryo as in panel (b), respectively in a less and further
decompacted stage.
Multinucleation and Anucleates

To be suitable for genetic analysis either by FISH or by PCR, the removed blastomere(s) should contain one
single, clearly visible nucleus (Figure 15.5 and 15.6a). Both multinucleation [42] and anucleate blastomeres
(Figure 15.6b) [43,44] are frequently observed in cleavage-stage embryos. The knowledge that the chromosome
constitution of multinucleated blastomeres is frequently different from that of their sibling blastomeres makes
them unsuitable for PGD [45]. The presence of multinucleated blastomeres in human embryos has been corre-
lated with chromosomal abnormalities [46]. Embryos with >50% of their blastomeres showing multinucleation
are not considered for embryo biopsy, irrespective of the developmental stage at first detection.
Efficient embryo biopsy requires a careful assessment of the nuclear status by light microscopy to exclude both
multinucleated and anucleate cells (Figure 15.6b) from genetic diagnosis. Sometimes, extreme granularity of the
blastomeres may complicate this assessment. A mistaken interpretation of the nuclear status of a blastomere may
then be anticipated if a FISH procedure follows because it would be noticed at fixation. One extra blastomere
can be removed in these cases. A PCR procedure lacks this intermediate stage because cells are immediately
transferred to PCR tubes and the amplification result is available only later.
Efficient embryo biopsy requires a careful assessment of the nuclear status by light microscopy to
exclude both multinucleated and anucleate cells from genetic diagnosis.
FIGURE 15.5 To be suitable for genetic analysis either by FISH or by PCR, the removed blastomere(s) should contain one single,
clearly visible nucleus.
FIGURE 15.6 (a) Single-nucleated blastomere. (b) Anucleated blastomere. (c) Lysed blastomere.
Cell Lysis
Blastomere lysis rate (expressed per blastomere aspirated) represents a good quality control measure for the
biopsy procedure. Cell lysis (Figure 15.6c) may occur at either step of the biopsy procedure. Cell lysis at the
moment of zona opening may be related to the method used. Laser drilling has proven to result in fewer blasto-
mere lysis (1.7%) compared with AT drilling (4.8%) [17]. Cell lysis when aspirating blastomeres may be related
to excessive mechanical stress, resulting in cell blebbing, or related to strong adherence of blastomeres to each
other [47] that may be avoided by means of Ca2+/Mg2+-free medium [41]. Lysed cells are not used for PCR
analysis because contamination with maternal DNA cannot be excluded. Only occasionally, the nucleus can
be recovered for FISH analysis. In cases of cell lysis, it is advised that the aspiration pipette be changed before
continuing biopsy of other embryos as a safety measure to avoid cross-contamination.
Postbiopsy Development
Further embryo development should not be impaired as a result of the biopsy procedure. Embryo postbiopsy devel-
opment can be evaluated on Day 4, where a doubling of cells, signs of compaction, or both represent a good pro-
gression. Otherwise, on Day 5, the quality of the inner cell mass and trophectoderm can be assessed (Figure 15.7).
Extended culture to Day 5 of biopsied embryos results in typical eight-shaped artificially hatching blastocysts that
might be in favor of implantation. Postbiopsy blastocyst formation has been well documented [36,48–51]. Embryo
developmental characteristics may however be related to the genetic condition tested for [52,53].
In addition to in vitro development, further in vivo development postbiopsy may be reflected by the implanta-
tion rates obtained post-PGD. In this respect, large data collections are very much valued to serve as a standard
reference of good laboratory and clinical practice. Live birth rates per cycle of 15.9% [54] and 15.0% [9] have
been reported.
Advantages and Disadvantages

A clear advantage of cleavage-stage embryo biopsy over polar body analysis is that the paternal contribution
to the embryo can be analyzed. In addition, enough time for diagnostic testing is available, especially when
transferring embryos on Day 5. At present, supernumerary biopsied and diagnosed embryos can be effectively
vitrified for later use, either on Day 4 [55] or in the blastocyst stage [56–58].
FIGURE 15.7 (a, b) Two examples of postbiopsy blastocyst formation on Day 5, allowing the evaluation of inner cell mass quality
and trophectoderm quality. Biopsied embryos result in typical eight-shaped artificially hatching blastocysts.
At the cleavage stage, limited amounts of cellular material are available for genetic testing. However, PCR
and FISH technologies are well established at the single-cell level. The invasive nature of the biopsy procedure
on Day 3 should be recognized, and evidence exists that two-cell biopsy is more invasive than one-cell biopsy.
Mosaicism at the cleavage stage remains a serious obstacle for chromosomal aneuploidy testing [59]. Removal of
one cell does not interrogate mosaicism, thus more normal results are encountered, albeit with a lower predictive
value. Two-cell analysis results in fewer embryos for transfer but with a higher predictive value.
REFERENCES
1. Goossens V, Traeger-Synodinos J, Coonen E, et al. ESHRE PGD Consortium data collection XI: Cycles
from January to December 2008 with pregnancy follow-up to October 2009. Hum Reprod 2012; 27: 1887–1911.
2. Handyside AH, Kontogianni EH, Hardy K, Winston RM. Pregnancies from biopsied human preimplantation
embryos sexed by Y-specific DNA amplification. Nature 1990; 344: 768–770.
3. Handyside AH, Lesko JG, Tarin JJ, Winston RM, Hughes MR. Birth of a normal girl after in vitro fertilization and
preimplantation diagnostic testing for cystic fibrosis. N Engl J Med 1992; 327(13): 905–909.
4. Hardy K, Martin KL, Leese HJ, et al. Human preimplantation development in vitro is not adversely affected by
biopsy at the 8-cell stage. Hum Reprod 1990; 5: 708–714.
5. Harper JC, Coonen E, De Rycke M, et al. ESHRE PGD Consortium data collection X: Cycles from January to
December 2007 with pregnancy follow-up to October 2008. Hum Reprod 2010; 25: 2685–2707.
6. Mottla GL, Adelman MR, Hall JL, et al. Lineage tracing demonstrates that blastomeres of early cleavage-stage
human pre-embryos contribute to both trophectoderm and inner cell mass. Hum Reprod 1995; 10: 384–391.
7. Harton GL, Magli C, Lundin K, et al. ESHRE PGD Consortium/Embryology Special Interest Group—Best prac-
tice guidelines for polar body and embryo biopsy for preimplantation genetic diagnosis/screening (PGD/PGS).
Hum Reprod 2011; 26: 41–46.
8. Wilton L, Thornhill A, Traeger-Synodinos J, et al. The causes of misdiagnosis and adverse outcomes in PGD.
Hum Reprod 2009; 24: 1221–1228.
9. Harper JC, Boelaert K, Geraedts J, et al. ESHRE PGD Consortium data collection V: Cycles from January to
December 2002 with pregnancy follow-up to October 2003. Hum Reprod 2006; 21: 1887–1911.
10. Selva J. Assisted hatching. Hum Reprod 2000; 15: 65–67.
11. Grifo JA, Boyle A, Fischer E, et al. Preembryo biopsy and analysis of blastomeres by in situ hybridization. Am J
Obstet Gynecol 1990; 163: 2013–2019.
12. Cieslak J, Ivakhnenko V, Wolf G, et al. Three-dimensional partial zona dissection for preimplantation genetic
diagnosis and assisted hatching. Fertil Steril 1999; 71: 308–313.
13. Cohen J, Elsner C, Kort H, et al. Impairment of the hatching process following IVF in the human and improve-
ment of implantation by assisting hatching using micromanipulation. Hum Reprod 1990; 5: 7–13.
14. Tucker MJ, Luecke NM, Wiker SR, Wright G. Chemical removal of the outside of the zona pellucida of day 3
human embryos has no impact on implantation rate. J Assist Reprod Genet 1993; 10: 187–191.
15. Chen SU, Chao KH, Wu MY, et al. The simplified two-pipette technique is more efficient than the conventional
three-pipette method for blastomere biopsy in human embryos. Fertil Steril 1998; 69: 569–575.
16. Inzunza J, Iwarsson E, Fridström M, et al. Application of single-needle blastomere biopsy in human preimplanta-
tion genetic diagnosis. Prenat Diagn 1998; 18: 1381–1388.
17. Joris H, De Vos A, Janssens R, et al. Comparison of the results of human embryo biopsy and outcome of PGD
after zona drilling using acid Tyrode medium or a laser. Hum Reprod 2003; 18: 1896–1902.
18. Germond M, Nocera D, Senn A, et al. Microdissection of mouse and human zona pellucida using a 1.48–microns
diode laser beam: Efficacy and safety of the procedure. Fertil Steril 1995; 64: 604–611.
19. Germond M, Nocera D, Senn A, et al. Improved fertilization and implantation rates after non-touch zona pellu-
cida microdrilling of mouse oocytes with a 1.48 microm diode laser beam. Hum Reprod 1996; 11: 1043–1048.
20. Boada M, Carrera M, De La Iglesia C, et al. Successful use of a laser for human embryo biopsy in preimplantation
genetic diagnosis: Report of two cases. J Assist Reprod Genet 1998; 15: 302–307.
21. Chatzimeletiou K, Morrison EE, Panagiotidis Y, et al. Comparison of effects of zona drilling by non-contact
infrared laser or acid Tyrode’s on the development of human biopsied embryos as revealed by blastomere
viability, cytoskeletal analysis and molecular cytogenetics. Reprod Biomed Online 2005; 11: 697–710.
22. Jones AE, Wright G, Kort HI, et al. Comparison of laser-assisted hatching and acidified Tyrode’s hatching by
evaluation of blastocyst development rates in sibling embryos: A prospective randomized trial. Fertil Steril 2006;
85: 487–491.
23. Geber S, Bossi R, Lisboa CB, et al. Laser confers less embryo exposure than acid tyrode for embryo biopsy in
preimplantation genetic diagnosis cycles: A randomized study. Reprod Biol Endocrinol 2011; 9: 58.
24. Hartshorn C, Anshelevich A, Wangh LJ. Laser zona drilling does not induce hsp70i transcription in blastomeres
of eight-cell mouse embryos. Fertil Steril 2005; 84: 1547–1550.
25. Rink K, Delacrétaz G, Salathé RP, et al. Non-contact microdrilling of mouse zona pellucida with an objective-
delivered 1.48-microns diode laser. Lasers Surg Med 1996; 18: 52–62.
26. Douglas-Hamilton DH, Conia J. Thermal effects in laser-assisted pre-embryo zona drilling. J Biomed Opt 2001;
6: 205–213.
27. Tadir Y, Douglas-Hamilton DH. Laser effects in the manipulation of human eggs and embryos for in vitro fertil-
ization. Methods Cell Biol 2007; 82: 409–431.
28. Taylor TH, Gilchrist JW, Hallowell SV, et al. The effects of different laser pulse lengths on the embryo biopsy
procedure and embryo development to the blastocyst stage. J Assist Reprod Genet 2010; 27: 663–667.
29. Lanzendorf SE, Ratts VS, Moley KH, et al. A randomized, prospective study comparing laser-assisted hatching
and assisted hatching using acidified medium. Fertil Steril 2007; 87: 1450–1457.
30. Sagoskin AW, Levy MJ, Tucker MJ, et al. Laser assisted hatching in good prognosis patients undergoing in vitro
fertilization-embryo transfer: A randomized controlled trial. Fertil Steril 2007; 87: 283–287.
31. Scott RT, Upham KM, Forman EJ, et al. Cleavage-stage biopsy significantly impairs human embryonic implantation
potential while blastocyst biopsy does not: A randomized and paired clinical trial. Fertil Steril 2013; 100: 624–30.
32. Pierce KE, Michalopoulos J, Kiessling AA, et al. Preimplantation development of mouse and human embryos
biopsied at cleavage stages using a modified displacement technique. Hum Reprod 1997; 12: 351–356.
33. Levinson G, Fields RA, Harton GL, et al. Reliable gender screening for human preimplantation embryos, using
multiple DNA target-sequences. Hum Reprod 1992; 7: 1304–1313.
34. Van de Velde H, De Vos A, Sermon K, et al. Embryo implantation after biopsy of one or two cells from cleavage-
stage embryos with a view to preimplantation genetic diagnosis. Prenat Diagn 2000; 20: 1030–1037.
35. Cohen J, Wells D, Munné S. Removal of 2 cells from cleavage stage embryos is likely to reduce the efficacy of
chromosomal tests that are used to enhance implantation rates. Fertil Steril 2007; 87: 496–503.
36. Goossens V, De Rycke M, De Vos A, et al. Diagnostic efficiency and accuracy, embryonic development and clini-
cal outcome after the biopsy of one or two blastomeres for preimplantation genetic diagnosis. Hum Reprod 2008;
23: 481–492.
37. De Vos A, Staessen C, De Rycke M, et al. Impact of cleavage-stage embryo biopsy in view of PGD on human
blastocyst implantation: A prospective cohort of single embryo transfers. Hum Reprod 2009; 24: 2988–2996.
38. Lewis CM, Pinel T, Whittaker JC, Handyside AH. Controlling misdiagnosis errors in preimplantation genetic
diagnosis: A comprehensive model encompassing extrinsic and intrinsic sources of error. Hum Reprod 2001; 16:
43–50.
39. Campbell S, Swann HR, Seif MW, et al. Cell adhesion molecules on the oocyte and preimplantation human
embryo. Hum Reprod 1995; 10: 1571–1578.
40. Santalo J, Grossmann M, Egozcue J. Does Ca2+/Mg2+-free medium have an effect on the survival of the preim-
plantation mouse embryo after biopsy? Hum Reprod Update 1992; 2: 257–261.
41. Dumoulin JC, Bras M, Coonen E, et al. Effect of Ca2+/Mg2+-free medium on the biopsy procedure for implanta-
tion genetic diagnosis and further development of human embryos. Hum Reprod 1998; 13: 2880–2883.
42. Van Royen E, Mangelschots K, Vercruyssen M, et al. Multinucleation in cleavage stage embryos. Hum Reprod
2003; 18: 1062–1069.
43. Hardy K, Winston RM, Handyside AH. Binucleate blastomeres in preimplantation human embryos in vitro:
Failure of cytokinesis during early cleavage. J Reprod Fertil 1993; 98: 549–558.
44. Johansson M, Hardarson T, Lundin K. There is a cutoff limit in diameter between a blastomere and a small
anucleate fragment. J Assist Reprod Genet 2003; 20: 309–313.
45. Munné S, Cohen J. Unsuitability of multinucleated human blastomeres for preimplantation genetic diagnosis.
Hum Reprod 1993; 8: 1120–1125.
46. Kligman I, Benadiva C, Alikani M, Munne S. The presence of multinucleated blastomeres in human embryos is
correlated with chromosomal abnormalities. Hum Reprod 1996; 11: 1492–1498.
47. De Vos A, Van Steirteghem A. Aspects of biopsy procedures prior to preimplantation genetic diagnosis. Prenat
Diagn 2001; 21: 767–780.
48. Veiga A, Gil Y, Boada M, et al. Confirmation of diagnosis in preimplantation genetic diagnosis (PGD) through
blastocyst culture: Preliminary experience. Prenat Diagn 1999; 19: 1242–1247.
49. Palmer GA, Traeger-Synodinos J, Davies S, et al. Pregnancies following blastocyst stage transfer in PGD cycles
at risk for beta-thalassaemic haemoglobinopathies. Hum Reprod 2002; 17: 25–31.
50. Magli MC, Gianaroli L, Ferraretti A, et al. The combination of polar body and embryo biopsy does not affect
embryo viability. Hum Reprod 2004; 19: 1163–1169.
51. Cieslak-Janzen J, Tur-Kapsa I, Iklevitch Y, et al. Multiple micromanipulations for preimplantation genetic diag-
nosis do not affect embryo development to the blastocyst stage. Fertil Steril 2006; 85: 1826–1829.
52. Findikli N, Kahraman S, Kumtepe Y, et al. Embryo development characteristics in Robertsonian and reciprocal
translocations: A comparison of results with non-translocation cases. Reprod Biomed Online 2003; 7: 563–571.
53. Rubio C, Simon C, Vidal F, et al. Chromosomal abnormalities and embryo development in recurrent miscarriage
couples. Hum Reprod 2003; 18: 182–188.
54. Verlinsky Y, Cohen J, Munné S, et al. Over a decade of experience with preimplantation genetic diagnosis:
A multicenter report. Fertil Steril 2004; 82: 292–294.
55. Kahraman S, Candan ZN. Outcomes of vitrified-warmed day-4 embryos after day-3 cleavage-stage biopsy.
56. Escribá MJ, Zulategui JF, Galán A, et al. Vitrification of preimplantation genetically diagnosed human blastocysts
and its contribution to the cumulative ongoing pregnancy rate per cycle by using a closed device. Fertil Steril
2008; 89: 840–846.
57. Zhang X, Trokoudes KM, Pavlides C. Vitrification of biopsied embryos at cleavage, morula and blastocyst stage.
58. Keskintepe L, Sher G, Machnicka A, et al. Vitrification of human embryos subjected to blastomere biopsy for pre-
implantation genetic screening produces higher survival and pregnancy rates than slow freezing. J Assist Reprod
Genet 2009; 26: 629–635.
59. Los FJ, Van Opstal D, van den Berg C. The development of cytogenetically normal, abnormal and mosaic
embryos: A theoretical model. Hum Reprod Update 2004; 10: 79–94.
16
Invasive Techniques: Blastocyst Biopsy
Steve McArthur
Introduction
The first report of the application of trophectoderm biopsy for the purpose of determining the genetics status of
blastocyst-stage embryos was the sexing of rabbit embryos by Edwards and Gardner in 1967 [1]. Decades later
and following in the wake of the development of human in vitro fertilization (IVF), Handyside and cowork-
ers reported the first live birth after embryo biopsy in 1990 [2]. As preimplantation genetic diagnosis (PGD)
developed, it was typical for biopsy to be performed to remove either the polar bodies or blastomeres. Polar
body biopsy required the removal of the first and second polar bodies across Day 0 and Day 1 of development,
the obvious restriction with polar body analysis being that only maternally carried genetic anomalies would be
revealed. Blastomere biopsy occurred on Day 3 postfertilization when the embryo had progressed to the six- to
eight-cell stage and typically resulted in the extraction of one or two blastomeres. Both polar bodies and blas-
tomeres were analyzed by either polymerase chain reaction (PCR) or fluorescent in situ hybridization (FISH)
techniques. PCR required the design of oligonucleotide primers to amplify allelic targets such a deletion or
insertions in a single cell or polar body. FISH targeted and labeled chromosomes in a single cell, allowing detec-
tion of chromosomal aneuploidies and translocations. Recently, comparative genome hybridization (CGH) has
displaced FISH for the purpose of chromosome aneuploidy and translocation analysis in PGD. To remove either
the polar bodies or blastomeres from the embryo, the zona pellucida was breached by either mechanical means
or by dissolving the protein with acid Tyrode’s solution, and the embryo may or may not have been incubated for
a period of time in a calcium- and magnesium-free medium to decrease cell-to-cell interactions and therefore
make the biopsy of cells easier. During the late 1990s, the development of near-infrared lasers allowed for a more
rapid and controllable opening if the zona pellucida.
Given that PGD relies upon embryo biopsy, an invasive procedure, it is critical that the impact of the biopsy
procedure is reduced to the lowest level possible. The development of a fertilized oocyte through to a blasto-
cyst (Figure 16.1) is the result of a process of selection that ultimately provides the embryo (interdependent
on the quality of the culture conditions) with an improved prospect of implantation and survival through to
live birth [3,4]. It is therefore conceivable that removal of 12%–25% of the embryos’ biomass, as occurs with
one- or two-cell biopsy on Day 3, will likely result in a decrease in the potential of the embryo to implant and
progress through to a healthy live birth. There is evidence of a capacity within human embryos to withstand
the damage of a Day 3 biopsy; there is a likelihood that embryos that may otherwise progress to implantation
and a live healthy birth will be lost as a consequence of the detrimental impact of biopsy undertaken at this
stage of embryo development. Blastocyst biopsy results in a significantly smaller loss of biomass, occurring
when an embryo has >100 cells, and importantly it leaves the inner cell mass intact. Given that the inner cell
mass will develop into the embryo proper, it is critical to minimize the effect of any invasive techniques on the
component of the embryo.
Genea (formerly Sydney IVF) introduced a comprehensive blastocyst culture, transfer, and cryostorage meth-
odology in the early 2000s that produced improved clinical pregnancy and health live baby rates, while also
reducing miscarriage and multiple pregnancy rates [3–9]. The blastocyst culture system was based upon the abil-
ity to select the healthiest embryo from a patient’s cohort for transfer and subsequent cryostorage of remaining
blastocysts. The blastocyst culture methodology led to the development and routine clinical practice of blastocyst
233
FIGURE 16.1 (a) In vitro development of a human blastocyst from 2-pronuclear stage through trophectoderm biopsy; (b) embryo
hatching occurs on Day 3; (c) the embryo develops through the expanding blastocyst, with biopsy occurring on Day 5 or Day 6.
FIGURE 16.2 Image of Day 3 embryo after hatching (using a near-infrared laser) showing a 10 μm hole at the internal margin of
the zona pellucida.
biopsy [3]. Biopsy at the blastocyst stage requires the zona pellucida to be breached on Day 3 of development
by using a near-infrared laser (Figure 16.2) and allows for the extraction of 3–10 trophectoderm cells from an
embryo generally made up of ≥100 cells. A step-by-step guide to embryo hatching is listed below.
Embryo hatching preceding blastocyst biopsy occurs either on Day 3 or on Day 5 or Day 6 of embryo
development. In Genea laboratories, all embryo hatching occurs on Day 3 of development to ensure acces-
sibility to a suitable hatching site with little risk of damaging any blastomeres. The development of this
protocol occurred as a follow-on from the early PGD practice of embryo hatching and biopsy of Day 3 of
development, when all embryos were hatched for analysis. In many cases of analysis failure, the embryos
were reviewed on Day 5 or Day 6 and found to be hatching blastocysts with trophectoderm cells protruding
from the Day 3 hatch site. It flowed into common practice once blastocyst biopsy was used for all embryo
biopsy cases after 2003. Breaching the zona pellucida on Day 5 of development is practiced in many labora-
tories worldwide and was first reported in 2006 [10]. In this report, blastocysts were hatched at 116 hr post-
fertilization and cultured for a further 4 hr to promote herniation of the trophectoderm. The 4 hr of further
culture may be restrictive in allowing for the transfer of an embryo on the day of analysis; however, with the
advent of array-CGH and vitrification of embryos for transfer in a subsequent cycle, this may no longer be
an impediment. It is often speculated that breaching the zona p ellucida on Day 3 may result in the inner cell
mass being located external to the breach site. Experience at Genea shows that this situation does eventuate;
however, given the ability to manipulate the position of the embryo on the microtools, this is no barrier to
embryo biopsy.
Blastocyst Biopsy 235
BOX 16.1 EMBRYO HATCHING

1. The hole in the zona pellucida should be made adjacent to the polar body, if identifiable.
Otherwise, hatch at a position where there is a space between the blastomeres.
2. The first pulse of the laser should be directed to the outer edge of the zona pellucida.
3. The third pulse of the laser should cut the internal wall of the zona pellucida in a direct line from
the first laser pulse.
4. A hole in the internal margin of the zona of approximately 10 μm is created.
5. Embryos should then be washed before being placed back into culture dish.
Embryos hatched on Day 3 are cultured for another 2 days to allow for blastocoel expansion and extrusion of
trophectoderm cells through the zona opening, thereby allowing for biopsy [5,9]. Embryos suitable for biopsy
are placed in 5 μL drops of blastocyst culture medium under mineral oil. The embryo is held in position by a
holding pipette, as used in intracytoplasmic sperm injection (ICSI) practice, and a 30 μm biopsy pipette is used
to collapse the blastocyst and hold the cells that will be biopsied. Three to four shots of the laser set on a low
power level reduce the cell-to-cell interactions, enabling multiple cells to be removed from the hatching trophec-
toderm. The cells remain as a single piece of tissue as the cells remain tightly coupled, thereby allowing the cells
to be transferred easily in a single pipetting movement. The embryo is then returned to culture dish containing
fresh blastocyst culture medium for further incubation while awaiting the results of the PGD analysis, at which
point the embryo is transferred, cryostored, or disposed. The embryos are cultured individually postbiopsy in
uniquely identified locations in the culture dish to ensure that embryo identity is maintained. The biopsied cells
are washed and prepared for analysis by CGH, PCR, or FISH (FISH is no longer used in Genea laboratories).
A step-by-step guide to trophectoderm biopsy procedure is listed in Box 16.2 and detailed in Figure 16.3.
The method for biopsy of a fully hatched blastocyst (Figure 16.4) is similar to that described for a stan-
dard blastocyst biopsy except there is no requirement to use the near-infrared laser for this biopsy method.
BOX 16.2 TROPHECTODERM BIOPSY

1. Ensure that the biopsy pipette tip and the leading edge of the trophectoderm are in focus.
2. Bring the biopsy pipette and touch the leading edge of the trophectoderm.
3. Draw a small number of trophectoderm cells carefully into the pipette by using suction until a
good hold is achieved.
4. Stretch out the trophectoderm (at this point, the blastocoel cavity may collapse).
5. Adjust focus until the margins of the targeted cells are in focus, and then fire the laser at the
intercellular junctions. Fire the laser three to five times across the width of the trophectoderm
cells to be biopsied.
6. Using suction on the mouthpiece and the holding pipette joystick, pull the targeted cells away
from the blastocyst.
7. Release the piece of trophectoderm from the biopsy pipette.
8. Where the stretched trophectoderm tissue does not readily separate from the remaining embryo,
it is not recommended to fire the laser excessively because this may be detrimental to embryo
recovery. In these cases, a more direct “cutting off” method may be used.
9. Gently release the embryo from the holding pipette while maintaining hold of cells in your
biopsy pipette. Do not take further cells into your pipette because this may result in excess cells
being biopsied. Ensuring that the holding and biopsy pipettes are in the same focal plane, rub
the biopsy pipette across the end of the holding pipette. Take care not to damage the embryo
protruding from the pipette as you carry out this step. If the first attempt is not successful, further
attempts may be required.
FIGURE 16.3 Photographs of blastocyst biopsy. (a) Blastocyst 2 days after laser-assisted hatching, consisting of herniating trophecto-
derm (TE) and inner cell mass (ICM). (b) Alignment of TE cells with biopsy pipette. (c) Three to ten cells aspirated from the TE while the
ICM (destined to form the embryo proper) remains intact. (d) Blastocyst and TE sample. (e) Blastocyst approximately 90 min postbiopsy.
FIGURE 16.4 Images of fully hatched blastocyst biopsy. (a) Fully hatched blastocyst 2–3 days post–laser-assisted hatching;
trophectoderm (TE) and inner cell mass (ICM) are labeled. (b) TE cells aspirated into biopsy pipette and contacted with holding
pipette. (c) TE cells cut by shearing biopsy pipette against holding pipette. (d) Blastocyst and TE sample postbiopsy. (e) Blastocyst
approximately 90 min postbiopsy.
In the method, steps 4–7 are removed, and the trophectoderm is biopsied as per step 8 by using the embryo-
holding pipette as a shearing force to remove the trophoblasts drawn into the biopsy pipette in step 3 of the
procedure.
Timing is critical to many aspects of IVF, with particular emphasis placed on the embryo transfer procedure.
Embryo biopsy on Day 5 or Day 6 of development places time pressures on delivering analysis outcomes in time
for an embryo transfer. To deliver the optimum number of embryos for biopsy, it is important to ensure coordi-
nation of all aspects of the IVF cycle, especially the timing of oocyte collection and insemination. It is ideal to
undertake blastocyst biopsy on the morning of Day 5 or Day 6; therefore, it is suggested that oocyte collections
are scheduled early in the theatre list to ensure early insemination time and maximum culture length through to
the morning of Day 5 or Day 6. Blastocysts are routinely assessed for suitability for biopsy on the morning of
Day 5 or Day 6 and between 12 pm and 1 pm on both days. Embryos not suitable for biopsy at the final check on
Day 6 are excluded from the analysis.
The most advanced PGD technologies are reliant upon molecular biology techniques to provide material for
analysis. Therefore, in a practical sense, having multiple cells and therefore multiple copies of DNA as targets
for the amplification provides benefit to the quality of the amplification product generated compared with that
provided by the single-copy DNA available from polar body or Day 3 embryo biopsy [8]. DNA amplification has
inherent technical risks, such as amplification failure of the target DNA. The amplification risks in single cells
(blastomeres) are likely to be higher than that seen in other sources of genetic material, such as polar bodies and
lymphocytes [11]. Consequently, blastocyst biopsy allows increased confidence in diagnostic potential of PGD
testing methods that require DNA amplification.
Use of PGD has increased in many clinics across the globe in the last decade. The 2010 report from the European
Society of Human Reproduction and Endocrinology (ESHRE) PGD Consortium indicated that Day 3 biopsy
accounted for 90% of embryo biopsies, whereas blastocyst biopsy accounted for just 0.4% [12]. At Genea clinics,
PGD cycles now account for >15% of all cycles undertaken and the rate of uptake continues to increase. The increased
use of PGD worldwide is not unexpected given reports of improved outcomes [13,14]. The technologies applied to
cells biopsied from preimplantation embryos have grown in complexity as the uptake of PGD has expanded.
Chromosome enumeration, historically undertaken using FISH techniques, has in recent times been replaced
by CGH techniques, allowing for the complete karyotyping of the preimplantation embryo. In the decade or so
since the first reports of CGH and a live birth from the analysis of an embryo via the technique [15,16], there have
been many advances in molecular analysis technology. CGH on metaphase chromosomes spread on glass slides
has been replaced by molecular or microarray-CGH based on whole-genome amplification [17]. The complete
karyotyping of an embryo has ensured that embryos are not transferred with aneuploidies previously undetected
due to the limitations of FISH technologies. Figure 16.5 compares aneuploidies detected by microarray-CGH
25.0
20.0
Aneuploidy rate (%)
15.0
IVF miscarrage samples

10.0
PGD embryos
5.0
0.0
1
2
3
4
5
6
7
8
9
X/Y
10
11
12
13
14
15
16
17
18
19
20
21
22
Chromosome
FIGURE 16.5 Comparison of chromosome aneuploidy rates between embryos biopsied on Day 5 or Day 6 with those detected in
miscarriage samples post-IVF treatment. (Data courtesy of Genea Fertility, Sydney, Australia.)
with aneuploidies identified in miscarriage samples after an IVF cycle. Many of the aneuploidies detected by
microarray-CGH would not have been revealed by FISH analysis, and subsequently such embryos would have
been transferred.
The improvements in pregnancy and live birth outcomes associated with PGD have been attributed to the
ability to produce a comprehensive chromosome karyotype of an individual embryo. Pregnancy and live birth
outcomes have also improved with the advent of vitrification technologies postvitrification being introduced dur-
ing the mid-2000s, and groups now report outcomes postembryo warming equivalent and in many clinics better
than those achieved with a fresh embryo transfer [18]. Using blastocyst biopsy with microarray-CGH poses a
challenge in achieving a fresh embryo transfer that is not otherwise an issue when either polar body biopsy or
Day 3 embryo is applied. It is therefore important to be able to ensure that pregnancy and birth outcomes are
maintained and improved with the application of more advanced PGD technologies combined with blastocyst
biopsy. Wells et al. [19] reported on a strategy combining PGD and microarray-CGH analysis with vitrification
of biopsied embryos before obtaining an analysis outcome. After analysis and assignment of the chromosomal
status of the embryo, a normal embryo is identified and transferred in a subsequent cycle. The combination of
blastocyst culture–biopsy techniques, vitrification, and advanced molecular technologies offers patients excel-
lent clinical outcomes.
The last decade has shown that trophectoderm biopsy on Day 5 or Day 6 of embryo development delivers
high implantation and live birth rates compared with both polar body and Day 3 blastomere biopsy. Improved
implantation rates combined with a reduced impact of known embryological phenomena, such as mosaicism,
have resulted in a wider move away from polar body and blastomere biopsy. The advent and use of whole-
genome amplification technologies ensure that more detailed information is delivered from the PGD process,
further adding to the implantation potential of embryos and consequently increasing the likelihood of success
for patients accessing PGD technologies combined with blastocyst biopsy.
REFERENCES
1. Edwards RG, Gardner RL. Sexing of live rabbit blastocysts. Nature 1967;214(5088):576–7.
2. Handyside AH, Lesko JG, Tarín JJ, Winston RM, et al. Birth of a normal girl after in vitro fertilization and pre-
implantation diagnostic testing for cystic fibrosis. N Engl J Med 1992;327(13):905–9.
3. Henman M, Catt JW, Wood T, Bowman MC, et al. Elective transfer of single fresh blastocysts and later transfer
of cryostored blastocysts reduces the twin pregnancy rate and can improve the in vitro fertilization live birth rate
in younger women. Fertil Steril 2005;84(6):1620–7.
4. Jansen RPS. Benefits and challenges brought by improved results from in vitro fertilization. Intern Med J
2005;35(2):108–17.
5. de Boer KA, Catt JW, Jansen RPS, Leigh D, et al. Moving to blastocyst biopsy for preimplantation genetic diag-
nosis and single embryo transfer at Sydney IVF. Fertil Steril 2004;82(2):295–8.
6. Jansen RPS. Female age and the chance of a baby from one in-vitro fertilization treatment. Med J Aust
2003;178:258–61.
7. Jansen RPS, Bowman MC, de Boer KA, Leigh DA, et al. What next for preimplantation genetic screening (PGS)?
Experience with blastocyst biopsy and testing for aneuploidy. Hum Reprod 2008;23(7):1476–8.
8. McArthur SJ, Leigh D, Marshall JT, de Boer KA, et al. Pregnancies and live births after trophectoderm biopsy
and preimplantation genetic testing of human blastocysts. Fertil Steril 2005;84(6):1628–36.
9. McArthur SJ, Leigh D, Marshall JT, Gee AT, et al. Blastocyst trophectoderm biopsy and preimplanta-
tion genetic diagnosis for familial monogenic disorders and chromosomal translocations. Prenat Diagn
2008;28(5):434–42.
10. Kokkali G, Traeger-Synodinos J, Vrettou C, Stavrou D, et al. Blastocyst biopsy versus cleavage stage biopsy
and blastocyst transfer for preimplantation genetic diagnosis of β-thalassaemia: A pilot study. Hum Reprod
2007;22(5):1443–9.
11. Rechitsky S, Strom C, Verlinsky O, Amet T, et al. Allele dropout in polar bodies and blastomeres. J Assist Reprod
Genet 1998;15:253–7.
12. Harper JC, Coonen E, De Rycke M, Harton G, et al. ESHRE PGD Consortium data collection X: Cycles from
January to December 2007 with pregnancy follow-up to October 2008. Hum Reprod 2010;25:2685–707.
13. Forman EJ, Hong KH, Ferry KM, Tao X, et al. In vitro fertilization with single euploid blastocyst transfer: A
randomized controlled trial. Fertil Steril 2013;100(1):100–7.
14. Treff NR, Ferry KM, Zhao T, Su J, et al. Cleavage stage embryo biopsy significantly impairs embryonic repro-
ductive potential while blastocyst biopsy does not: A novel paired analysis of cotransferred biopsied and non-
biopsied sibling embryos. Fertil Steril 2011;96(3):S2.
15. Wells D, Fragouli E, Stevens J, Munne S, et al. High pregnancy rate after comprehensive chromosomal screening
of blastocysts. Fertil Steril 2008; 90:S80.
16. Wilton L, Williamson R, McBain J, Edgar D, et al. Birth of a healthy infant after preimplantation confirmation of
euploidy by comparative genomic hybridization. N Engl J Med 2001;345(21):1537–41.
17. Hu DG, Webb G, Hussey N. Aneuploidy detection in single cells using DNA array-based comparative genomic
hybridization. Mol Hum Reprod 2004;10(4):283–9.
18. Roy T, Peura T, McArthur S. Effect of embryo development and grade on pregnancy outcomes following fresh
and vitrified single blastocyst embryo transfer. Fertil Steril 2011;96(3):S44–5.
19. Wells D, Fragouli E, Stevens J, Munne S, et al. High pregnancy rate after comprehensive chromosomal screening
of blastocysts. Fertil Steril 2008;90:S80.
17
Invasive Techniques: Aneuploidy Testing by FISH
Introduction
Preimplantation Genetic Diagnosis for Aneuploidy Testing
Preimplantation genetic diagnosis (PGD) is a preventive method to identify genetic disorders and chromosomal
abnormalities in preimplantation embryos, thus giving couples the opportunity to avoid a pregnancy termination
after the prenatal diagnosis of such disorders and abnormalities. With PGD, only embryos having a normal geno-
type and chromosomal constitution are selected for transfer to the uterus, providing the birth of a healthy baby.
There are three main indication groups in which PGD could be applied: (1) aneuploidy screening for numerical
chromosomal abnormalities, (2) diagnosis of structural chromosomal abnormalities in translocation and inver-
sion carriers, and (3) diagnosis of genetic disorders or human leukocyte antigen (HLA) compatibility for a child
in need of stem cell transplantation (Figure 17.1) [1–8].
Although first applications of the PGD technique were for selecting disease-free embryos for an X-linked
single gene disorder [1], soon after the introduction of the fluorescence in situ hybridization (FISH) technique
for chromosomal disorders [2], the number of PGD cycles rose rapidly to detect chromosomal abnormalities in
embryos generated from patients undergoing in vitro fertilization (IVF) treatments for the purpose of improving
success [9,10]. The most frequent reasons for aneuploidy testing are advanced maternal age, recurrent miscar-
riages, repeated implantation failures, previous fetal chromosomal abnormality, and severe male factor infertility.
Chromosomal Abnormalities in Preimplantation Embryos

Chromosomally normal human somatic cells contain 46 chromosomes (22 pairs of autosomes and 1 pair of
gonosomes, thus 2n, n = 23) (Figure 17.2). Any deviation from this chromosome set is called as abnormality.
Aneuploidy is known as abnormalities in chromosome number that can originate from an excess number of
chromosomes (e.g., trisomy) or from missing chromosomes (e.g., monosomy), whereas haploidy (n) and triploidy
(3n) are associated with the abnormalities of the whole chromosome set.
A significant proportion of human preimplantation embryos contain aneuploidy as one of the major causes of
implantation failures, pregnancy losses, and abnormal live births [11]. Although the rate of aneuploidy increases
with the maternal age, PGD reveals a high incidence of aneuploidy and mosaicism in both young and advanced
maternal age patients [12,13].
The mechanisms of chromosomal abnormalities in embryos could be divided into two major categories
according to the onset of formation as meiotic and postmeiotic. Although both mechanisms are common in
preimplantation embryos, most of the aneuploidies originate from meiotic divisions in oogenesis [14–17],
increasing with advancing maternal age. In normal meiosis, homologous chromosomes are separated in

metaphase I (MI), and sister chromatids are separated in metaphase II (MII). Meiotic segregation errors in
oocytes and polar bodies are formed either by premature separation of sister chromatids (PSSC) or nondisjunc-
tion of homologous chromosomes (Figure 17.3). Recent studies using single-nucleotide polymorphism arrays
(SNP-arrays) in which the origin of individual alleles could be labeled and tracked demonstrate the former
mechanism is responsible for almost all errors in the first meiotic division [16,17].
241
Numerical Structural Single gene disorders

Chromosomal Chromosomal w/o HLA genotyping
Abnormalities Abnormalities
Advanced maternal Translocation

age Disease carrier
Reciprocal,
couples
Robertsonian,
Insertional, and
Recurrent Complex
pregnancy losses
Couples with an
Inversion affected child
Chromosomally Paracentric and
abnormal fetus Pericentric
history
Child requiring stem
cell transplantation
Repeated
implantation failures
FIGURE 17.1 Main indication groups for PGD.
1 2 3 4 5
6 7 8 9 10 11 12
13 14 15 16 17 18
19 20 21 22 X Y
(a)
FIGURE 17.2 Normal set of chromosomes in human somatic cells. GTG banding image of chromosome sets in (a) normal
female, 46,XX.
Aneuploidy Testing by FISH 243
1 2 3 4 5
6 7 8 9 10 11 12
13 14 15 16 17 18
19 20 21 22 X Y
(b)
FIGURE 17.2 (Continued) Normal set of chromosomes in human somatic cells. GTG banding image of chromosome sets in
(b) male, 46,XY, peripheral lymphocytes induced by conventional culturing and karyotyping techniques.
Normal PB1
N N Normal PB2
N
FIGURE 17.3 Normal meiosis. Chromosomal segregations in oocytes in two successive meiotic divisions.
Aneuploidy Testing by FISH

There are several methods for the detection of aneuploidy in embryos: FISH, array comparative genomic hybrid-
ization (a-CGH), SNP-arrays, and real-time polymerase chain reaction (RT-PCR).
In the FISH technique, DNA probes that are labeled with different colored fluorescent tags that are specific for
chromosomal regions are hybridized to interphase nuclei or metaphase chromosomes. The advantage over con-
ventional cytogenetic methods is that it can be applied to the interphase cell; thus, it is not necessary to culture
cells to achieve metaphases. Because it is a rapid test and eliminates the need of culture, this technique is suitable
for use in prenatal and preimplantation genetic diagnosis.
To date, the majority of PGD analyses for aneuploidy screening and rearrangements have been performed
using FISH, which has still wider application among other techniques. Aneuploidy testing by FISH consists of
the following steps: fixation of biopsied cells, pretreatment and probe application, denaturation, hybridization,
Embryo Blastomere Denaturation
Renaturation and
hybridization
Analysis Probe DNA
FIGURE 17.4 Steps of FISH.
stringent washing, and analysis after counterstain application (Figure 17.4) [2,18]. The most critical step for FISH
is the cell fixation; each cell should be informative, and the loss of genetic material should be avoided [19].
Preparing for FISH

Fixation Methods
Mainly, there are two different methods of fixation of cells that are biopsied from embryos. Both involve the
use of Carnoy’s fixative that contains methanol:acetic acid in a 3:1 proportion. The methods differ by solutions
used to swell and breakdown the cell membrane: a detergent-based (Tween 20) solution or a hypotonic buffer
containing either KCl or bovine serum albumin (BSA) with sodium citrate. Although the former method requires
less skill and is easier to handle, the resulting nuclear diameter is small and signal overlaps and FISH errors are
more frequent. In contrast, the hypotonic method gives a larger diameter nucleus that decreases the probability
of signal overlaps and nuclear loss during fixation.
BOX 17.1 FIXATION OF CELLS

Materials
Microscope slides (Superfrost)
Falcon tubes (TreffLab)
Glass Pasteur pipettes
Flame source
Mouth pipette (Drummond Scientific)
Carbide marker
Methanol (Sigma)
Glacial acetic acid (Sigma)
Sterile distilled H2O
Sodium citrate (tribasic:dihydrate) (Sigma)
Albumin, bovine (Sigma)
Stereomicroscope (Olympus SZX12), inverted microscope (Olympus CK40)
Method
1. Fixative solution (3:1) should be prepared freshly just before fixation.
2. Hand-drawn glass pipettes with a 60 μL inner diameter are prepared by using a flame source.
3. Glass slides are cleaned with fixative solution.
4. The proper function of the mouth pipette is checked by aspirating of a little amount of hypotonic
solution (6 mg/mL BSA, 1% sodium citrate). Another mouth pipette should be ready to avoid oil.
5. A small amount of hypotonic solution is aspirated into the pipette. The blastomere/trophecto-
derm tissue is aspirated and transferred into hypotonic solution, and then with another mouth
pipette, the blastomere/trophectoderm tissue is taken onto the slide.
6. Just before the nucleus dries, one drop of fixative solution is added to digest the cytoplasm. The
drop containing fixative can be added with a Pasteur pipette pulled in a flame, and the tip of the
pipette has to be very narrow so the drop of fixative added is tiny.
7. At the time the fixative is dropped, the blastomere/trophectoderm tissue may move while the
fixative and hypotonic are mixing on the slide. After a short distance, it will stop and attach to
the slide. At this moment if needed, another drop of fixative can be added to lyse the cytoplasm
completely.
8. The place of fixed chromatin is carefully marked using a carbide marker.
9. The sample number and case name are written on the slide, and slides are maintained at room
temperature until hybridization.
In polar body fixation, distilled water is used instead of hypotonic solution. The rest of the method is as
same as the fixation method for blastomeres and trophectoderm samples [26–29].
Pretreatment and Dehydration

Dehydration steps are common for polar body, blastomere and trophectoderm tissues, and they are done before
hybridization to clean the fixed chromatin from the cytoplasm and fixative artifacts that remain after fixation.
This step provides effective hybridization with a clean nucleus.
BOX 17.2 DEHYDRATION AND HYBRIDIZATION

Materials
Coplin jars
Ethyl alcohol (70%, 85%, 100%)
Method
1. The slides are immersed in solutions as follows: 1 min each in 70% ethanol, 85% ethanol, and
100% ethanol.
2. Slides are air-dried.
Probe mix and hybridization (first round)
Materials
Slide warmer (Hybrite-Vysis), microcentrifuge (Eppendorf), vortex (Velp)
2–10 μL micropipette (Eppendorf)
18 × 18 mm coverslipsa (Isolab)
Parafilm
Direct labeled probes (Vysis): MultiVysion PB [26–29].
The coverslips can be resized via cutting by a carbide marker according to the area size for hybridization.
a
FISH Method
Single-cell interphase FISH is a rapid, reliable, and efficient technique capable of detecting up 9–12 chromo-
somes in two or three rounds of hybridization on a single nucleus. Currently, fluorescent DNA probes for chro-
mosomes X, Y, 13, 14, 15, 16, 17, 18, 21, and 22 are being used in PGD for aneuploidy screening because they
are involved in >50% of all chromosomally abnormal miscarriages.
There are commercially available probe panels for aneuploidy screening. With the five-chromosome probe
set, chromosomes 13, 16, 18, 21, and 22 can be screened simultaneously. The number of chromosomes screened
could be increased to nine (Figure 17.5) with the addition of a panel with four chromosomes, 15, 17, X, and Y, and
further to 12 with the addition of locus-specific probes for three more chromosomes, 9, 19, and 20. Evaluation
of these chromosomes offers the selection of normal embryos for the most common aneuploidies found in chro-
mosomally abnormal miscarriages. Alternatively, for translocations and other chromosomal rearrangements,
different probe combinations are used that are specific for the type of abnormality (Figure 17.6).
FIGURE 17.5 First and second round results of the same blastomere. (a) Normal for the first round study, including chromosomes 13, 16,
18, 21, and 22. Chromosome 13 (red), chromosome 16 (light blue), chromosome 18 (blue), chromosome 21 (green), and chromosome 22
(yellow). (b) Abnormal for the second round study, including chromosomes 15, 17, X, and Y. Chromosome 15 (yellow), chromosome 17
(aqua), chromosome X (green), and chromosome Y (blue). This blastomere was diagnosed as “monosomy 17.” Notice that there is a signal
overlap between aqua and one of the green signals. In addition, for the yellow signal, the signal at the bottom-middle is an artifact, evident
from the color and the intensity. (Courtesy of Reproductive Genetics Laboratory, Istanbul Memorial Hospital, Istanbul, Turkey, 2009.)
FIGURE 17.6 Preimplantation genetic diagnosis (PGD) for translocation carriers by FISH. (a) In this example, the patient was a
carrier of a reciprocal translocation: 46,XY,t(17;22)(q23;q11.2). Probes were tested on patient’s blood samples to ensure translocation
and test the efficiency of probes. CEP 17 (spectrum aqua), Telomeric 17q (spectrum orange), and Telomeric 22q (spectrum green)
probes were used on metaphase chromosomes obtained after peripheral blood culture. (b) Detection of unbalanced products by
using same probe mixture on trophectoderm sample in the PGD study. This embryo was diagnosed as partial monosomy for 17q and
partial monosomy for 22q and hence as unbalanced for translocation.
1. MultiVysion PB probe panel is ready to use.

2. Before hybridization, humidified paper towels are placed on the slide warmer and the slide warmer set
up to 37°C.
3. The working probes are warmed up to 37°C, mixed by vortex, and centrifuged for a few seconds.
4. Then, 3 μL of probe mix is applied onto the marked area where the fixed chromatin is located.
5. A coverslip is carefully placed on the probe. It is important to avoid air bubbles.
6. The coverslip is sealed with parafilm or rubber cement.
7. Slides are placed onto the slide warmer (Hybrite), and the hybridization program starts with the dena-
turation step at 75°C for 5 min and the hybridization step at 37°C for 3 hr.
Washing and Counterstaining

N O T E: Because fluorophores that label the probes may fade under light, all steps after the hybridization should
be performed under dim light.
BOX 17.3 WASHING AND COUNTERSTAINING

Materials
Waterbath (Memmert)
Coplin jars
Coverslips (24 × 60 mm) (IsoLab)
2–10 μL micropipette (Rainin)
0.45 μm Minisart filters (Sartorius)
Thermometer
20× standard saline citrate (SSC) (Vysis)
Nonidet P-40 (NP-40) (Vysis)
Antifade solution (Vysis)
4,6-Diamidino-2-phenylindole (DAPI) (Vysis)
Washing solution (0.4× SSC, 0.3% NP-40)
For wash solution, combine 10 mL of 20× SSC (pH 5.3) with 1.5 mL of NP-40 and add distilled water to
the final volume of 500 mL. Adjust the pH to 7–7.5 with NaOH, and then filter with 0.45 μm filter before use.
Method
1. A Coplin jar is filled with wash solution and placed in the waterbath; another jar filled with wash
solution stays at room temperature. The waterbath is warmed to 71°C, and the temperature of the
solution is checked by a thermometer.
2. The parafilm and coverslip are carefully removed from the slide and immersed in 71°C wash
solution for 3 min incubation.
3. After 3 min of incubation, slides are removed from the wash solution and placed in the Coplin
jar containing wash solution at room temperature and incubated for a minute.
4. Slides are removed from wash solution, rinsed with distilled water, and allowed to dry vertically
on a paper towel in a dark area.
5. After the slides have dried, 10 μL of DAPI or Antifade solution is added onto the hybridization
area and the slide is covered with a 24 × 60 mm coverslip. For MultiVysion PB and MultiVysion
4CC probe panels, Antifade is used instead of DAPI because some probes have blue fluorophores.
6. Slides are covered with aluminum foil and evaluated as soon as possible by using a fluorescent
microscope that contains appropriate filters for the probe fluorophores [26–29].
Evaluation of Signals
Evaluation of signals is performed in a dark area using a fluorescent microscope. This step is the most important
step in the FISH-based PGD technique. Visualization of signals is performed using the appropriate single band-
pass filters (red, fluorescein isothiocyanate [FITC], aqua, blue, gold, and DAPI).
There are some points that are very important in the signal evaluation:
1. Dual or triple bandpass filters are used to distinguish artifacts from specific signals.
2. Cross-hybridizations may occur between centromeric regions of repetitive sequences. The unspecific
cross-hybridizations can be detected easily because their signal intensity is lower than that of the spe-
cific cross-hybridizations.
3. High-intensity signals may appear in other filters; aqua signals may leak in the blue filter and yellow
signals may leak in green filter because they have excitation wavelengths close to each other depending
on the choice of bandpass filters.
4. For a signal to be considered as “split,” the centers of the two signals should not be far from two signal-
size and the shapes should be similar to each other.
5. Because the evaluation is subjective, at least one witness is needed in the interpretation of the
signals.
6. When there is no consensus for a signal, the chromosome that was doubted should be reevaluated after
rehybridization with a new probe set (Figure 17.7).
BOX 17.4 APPLICATION OF PROBE MIX AND HYBRIDIZATION (SECOND ROUND)

Materials
Slide warmer (Hybrite-Vysis)
Incubator (Heraeus)
Microcentrifuge (Eppendorf)
Vortex (Velp)
2–10 μL micropipette (Rainin)
18 × 18 mm coverslips (Isolab)
Parafilm
MultiVysion 4 Color Custom probe set for chromosomes X, Y, 15, and 17 (Vysis)
Method
After the evaluation of the first round hybridization, the fixed blastomere chromatin is prepared for second
round hybridization.
1. The coverslip is removed by placing slide under water faucet until it falls.
2. The slides are immersed in solutions as follows:
a. 1 min in 70% ethanol
b. 1 min in 85% ethanol
c. 1 min in 100% ethanol
3. After the slides are dried, 3 μL of working probe (MultiVysion 4cc) is applied onto the
chromatin.
4. A coverslip is carefully placed on the probe. It is important to avoid air bubbles.
5. The coverslip is sealed with parafilm or rubber cement.
6. Slides are placed onto the slide warmer (Hybrite-Vysis), and the hybridization program starts
with the denaturation step at 75°C for 5 min and the hybridization step at 42°C for 1.5 hr.
Washing, Couterstaining, and Evaluation (Second Round)

1. Washing step is the same as in the previous section.
2. After the slides have dried, 10 μL of Antifade solution is added onto the hybridization area, and
the slide is covered with a 24 × 60 mm coverslip. For 4cc probe panels, Antifade is used instead
of DAPI because some probes have blue fluorophores.
3. Slides are covered with aluminum foil and evaluated as soon as possible with a fluorescent
microscope that contains appropriate filters for the probe fluorophores. For the 4cc probe, chro-
mosome 15 is seen in orange, chromosome 17 is seen in aqua, chromosome X is seen in green,
and chromosome Y is seen in blue (Figure 17.5b) [26–29].
FIGURE 17.7 Distinguishing an artifact. (a) In aqua and blue filters, the cell seems to have three aqua (as shown by the arrows) and
two blue signals. (b) In the triple filter, the signal in the 11 o’clock position remains but has a whitish color that is very different from
the rest of the specific aqua signals. Thus, the cell was diagnosed as “normal.” Chromosome 13 (red), chromosome 16 (light blue),
chromosome 18 (blue), chromosome 21 (green), chromosome 22 (yellow). (Courtesy of Reproductive Genetics Laboratory, Istanbul
Memorial Hospital, Istanbul, Turkey, 2009.)
Third Round Hybridization

1. After the evaluation of the second round hybridization, if there is a doubt in first and second analyses,
the fixed blastomere chromatin can be reanalyzed with telomeric- and locus-specific probes. This situ-
ation is often present in embryos of 16qh polymorphism carriers (Figure 17.8).
2. If a probe mix is to be used for the third round, 1 μL of each probe and 7 μL of buffer are added to the
mixture, and it can be stored at –20°C.
3. Probe application, hybridization conditions, and washing procedures are the same as described for the
first round, except for the time for hybridization; it should be 6–14 hr.
There are some difficulties regarding the interpretation of signals in polar bodies, blastomeres, and troph-
ectoderm samples. For polar bodies, the frequent problem would be the fragmentation of the first polar body
(Figure 17.9) and the signal overlap and pulverization (Figure 17.10) due to the compact nature of the polar bod-
ies. Because the first polar body contains “2n,” it should have two signals for each chromosome, and second polar
body should contain only one signal. Another difficulty is the discrimination of signal splitting in the first polar
bodies. An interpretation of a split signal as two distinct signals may lead to the diagnosis of a trisomic embryo
as normal (Figure 17.11).
In cleavage-stage embryo analysis, the most frequent difficulty that may be encountered is the low representa-
tive ability of a blastomere for the embryo due to mosaicism. A significant proportion of cleavage-stage embryos
have mosaicism, representing the most frequent reason of misdiagnosis. In addition, because the diagnosis
FIGURE 17.8 (a) For monosomic embryos such as monosomy 16, only one aqua signal specific for chromosome 16 is observed
in the first round. (b) When additional third round hybridization is performed, specific for telomeres of chromosome 16 (Tel 16q,
spectrum orange), it is identified that this blastomere is indeed normal for chromosome 16, thus “rescued.”
FIGURE 17.9 (a) An intact polar body that is easily interpretable. (b) Although this polar body is fragmented, it is interpretable,
but caution should be taken to ensure that no other parts belonging to the polar body have been missed during fixation.
FIGURE 17.10 (a) This polar body has normal signals that were concentrated on the bottom-left side of the nucleus, resulting in
partial overlaps. (b) This second polar body has a blue signal that was pulverized due to being located on the edge of the nucleus.
FIGURE 17.11 In this first polar body, the chromosome 22 (spectrum gold) signals are split and should be counted as one chromo-
some. This means one extra copy of chromosome 22 will remain in the oocyte unless a reciprocal aneuploidy occurs in the second
division in this gamete.
FIGURE 17.12 (a, b) Good spreading of cells after fixation of a trophectoderm tissue. Each cell has clear borders, and the nuclear
quality is high, with no residual cytoplasm. (c) Cells overlapped, and the nuclear material was lost during fixation, probably due to
excessive use of fixative.
depends only on one blastomere, the efficiency is limited by the efficiency of hybridization of that blastomere.
In contrast, blastocyst-stage biopsy provides approximately five cells (Figure 17.12a, b), reducing the error rates
and increasing the diagnosis rate by providing more nuclei for double-checking the evaluations. However, there
are also difficulties in the interpretation of signals in trophectoderm tissues that may result in overlap of cells and
residual cytoplasm due to improper fixation (Figure 17.12c).
Advantages and Disadvantages of FISH

Although it is a relatively low cost and less complex, FISH suffers from being subjective and error prone; thus,
quality control guidelines should be followed for every step from fixation to signal interpretation.
The most important disadvantage of FISH is that it is not able to detect all abnormalities because the num-
ber of chromosomes that can be analyzed is limited. Comprehensive chromosomal screening methods such as
a-CGH is able to detect aneuploidies related to all 23 pairs of chromosomes. In the a-CGH technique, DNA
from a test sample (green) and a normal reference sample (red) is labeled differentially and hybridized to several
25
20
15
% rate
10
0
2 3 4 6 7 8 9 12 13 15 16 17 18 20 21 22
Trisomy
FIGURE 17.13 Summary of cytogenetic analysis of 410 abortus materials. The chart shows the rate of trisomic conceptions by the
specific chromosome involved. (Courtesy of Istanbul Memorial Hospital.)
thousand probes that are imprinted on a glass slide. The fluorescence intensity of the test and the reference DNA
is measured to calculate the copy number changes for each chromosome. The analysis lasts 12–24 hr.
Studies using a comprehensive chromosomal screening (e.g., CGH) technique on embryos demonstrated
that abnormalities of the chromosomes that are not present in the FISH panel are also quite frequent [20].
Furthermore, FISH is not efficient in detecting all trisomies that end up with spontaneous abortions. Although
the most frequent trisomy found is 16, trisomies of chromosomes 2, 3, 4, 6, 7, 8, 9, 12, and 20 could also be
found in abortus materials that are not in the regular FISH panel. Approximately 30% of all trisomies would be
undiagnosed with the nine-probe panel and 45% would be undiagnosed with the five-probe panel (Figure 17.13).
On the other hand, FISH technique has some advantages over other novel techniques. Comprehensive chromo-
somal screening methods are yet to be improved on some aspects on which FISH is still superior. For example,
a-CGH is not able to detect all abnormalities, such mosaicism, polyploidy and haploidy, and cryptic transloca-
tions. SNP-array analysis [21–23] suffers from high noise and unspecific results. RT-PCR [24] is quite fast but
it is limited to analysis of several chromosomes. Although next generation sequencing is the most promising
technique among them, it is time-consuming and expensive, and the prices have to be reduced before it can enjoy
wider application in clinical use [25]. Furthermore, all these recently developed techniques suffer from interpre-
tational difficulties due to the large volume of data that is often beyond what is needed.
The future direction of PGD should be to obtain all required genetic information of the embryo with maximal
accuracy at the earliest time possible. The technical limitations are gradually being reduced, and simultane-
ous screening of all chromosome pairs and identification of polyploidy and uniparental disomy, diagnosis of
polygenic disorders and imprinting disorders, HLA genotyping, and copy number repeats are not too far. In the
future, although FISH is going to be used less frequently, the routine use of comprehensive chromosomal screen-
ing and next generation sequencing methods will launch a new era of PGD.
REFERENCES
1. Handyside AH, Kontogianni EH, Hardy K, Winston RML. Pregnancies from biopsied human preimplantation
embryos sexed by Y-specific DNA amplification. Nature. 1990;344:768–70.
2. Munné S, Weier HU, Stein J, Grifo J, et al. A fast and efficient method for simultaneous X and Y in situ hybridiza-
tion of human blastomeres. J Assist Reprod Genet. 1993;10(1):82–90.
3. Scriven PN, Handyside AH, Ogilvie CM. Chromosome translocations: Segregation modes and strategies for
preimplantation genetic diagnosis. Prenat Diagn. 1998;18(13):1437–49.
4. Verlinsky Y, Rechitsky S, Schoolcraft W, Strom C, et al. Preimplantation diagnosis for Fanconi anemia combined
with HLA matching. JAMA. 2001;285(24):3130–3.
5. Kuliev A, Pakhalchuk T, Verlinsky O, Rechitsky S. Preimplantation genetic diagnosis for hemoglobinopathies.
Hemoglobin. 2011;35(5–6):547–55.
6. Kahraman S, Benkhalifa M, Donmez E, Biricik A, et al. The results of aneuploidy screening in 276 couples
undergoing assisted reproductive techniques. Prenat Diagn. 2004;24(4):307–11.
7. McArthur SJ, Leigh D, Marshall JT, Gee AJ, et al. Blastocyst trophectoderm biopsy and preimplantation genetic
diagnosis for familial monogenic disorders and chromosomal translocations. Prenat Diagn. 2008;28(5):434–42.
8. Kahraman S, Beyazyurek C, Ekmekci CG. Seven years of experience of preimplantation HLA typing: A clinical
overview of 327 cycles. Reprod Biomed Online. 2011;23(3):363–71.
9. Munné S, Sandalinas M, Escudero T, Velilla E, et al. Improved implantation after preimplantation genetic
diagnosis of aneuploidy. Reprod Biomed Online. 2003;7(1):91–7.
10. Voullaire L, Collins V, Callaghan T, McBain J, et al. High incidence of complex chromosome abnormality in
cleavage embryos from patients with repeated implantation failure. Fertil Steril. 2007;87(5):1053–8.
11. Hassold T, Hunt P. To err (meiotically) is human: The genesis of human aneuploidy. Nat Rev Genet.
2001;2(4):280–91.
12. Baart EB, Martini E, van den Berg I, Macklon NS, et al. Preimplantation genetic screening reveals a high incidence
of aneuploidy and mosaicism in embryos from young women undergoing IVF. Hum Reprod. 2006;21(1):223–33.
13. Garrisi JG, Colls P, Ferry KM, Zheng X, et al. Effect of infertility, maternal age, and number of previous
miscarriages on the outcome of preimplantation genetic diagnosis for idiopathic recurrent pregnancy loss. Fertil
Steril. 2009;92(1):288–95.
14. Verlinsky Y, Ginsberg N, Lifchez A, Valle J, et al. Analysis of the first polar body: Preconception genetic diagnosis.
Hum Reprod. 1990;5(7):826–9.
15. Verlinsky Y, Cieslak J, Ivakhnenko V, Evsikov S, et al. Chromosomal abnormalities in the first and second polar
body. Mol Cell Endocrinol. 2001;183(Suppl 1):S47–9.
16. Christopikou D, Handyside AH. Questions about the accuracy of polar body analysis for preimplantation genetic
screening. Hum Reprod. 2013;28(6):1732–3.
17. Handyside AH, Montag M, Magli MC, Repping S, et al. Multiple meiotic errors caused by predivision of chromatids
in women of advanced maternal age undergoing in vitro fertilisation. Eur J Hum Genet. 2012;20(7):742–7.
18. Griffin DK, Handyside AH, Harper JC, Wilton LJ, et al. Clinical experience with preimplantation diagnosis of sex
by dual fluorescent in situ hybridization. J Assist Reprod Genet. 1994;11(3):132–43.
19. Velilla E, Escudero T, Munné S. Blastomere fixation techniques and risk of misdiagnosis for preimplantation
genetic diagnosis of aneuploidy. Reprod Biomed Online. 2002;4(3):210–17.
20. Wilton L. Preimplantation genetic diagnosis and chromosome analysis of blastomeres using comparative genomic
hybridization. Hum Reprod Update. 2005;11(1):33–41.
21. Handyside AH. PGD and aneuploidy screening for 24 chromosomes by genome-wide SNP analysis: Seeing the
wood and the trees. Reprod Biomed Online. 2011;23(6):686–91.
22. Brezina PR, Benner A, Rechitsky S, Kuliev A, et al. Single-gene testing combined with single nucleotide
polymorphism microarray preimplantation genetic diagnosis for aneuploidy: A novel approach in optimizing
pregnancy outcome. Fertil Steril. 2011;95(5):1786.
23. van Uum CM, Stevens SJ, Dreesen JC, Drüsedau M, et al. SNP array-based copy number and genotype analyses
for preimplantation genetic diagnosis of human unbalanced translocations. Eur J Hum Genet. 2012;20(9):938–44.
24. Treff NR, Scott RT, Jr. Four-hour quantitative real-time polymerase chain reaction-based comprehensive chro-
mosome screening and accumulating evidence of accuracy, safety, predictive value, and clinical efficacy. Fertil
Steril. 2013;99:1049–53.
25. Desai AN, Jere A. Next-generation sequencing: Ready for the clinics? Clin Genet. 2012;81(6):503–10.
26. Verlinsky Y, Kuliev A. 2000. An Atlas of Preimplantation Genetic Diagnosis. Section 4, 31–39. Parthenon
Publishing Group. New York, USA.
27. Abdelhadi I, Colls P, Sandalinas M, Escudero T, et al. Preimplantation genetic diagnosis of numerical abnormali-
ties for 13 chromosomes. Reprod Biomed Online. 2003;6(2):226–31.
28. Munné S, Gianaroli L, Tur-Kaspa I, Magli C, et al. Substandard application of preimplantation genetic screening
may interfere with its clinical success. Fertil Steril. 2007;88(4):781–4.
29. Colls P, Escudero T, Cekleniak N, Sadowy S, et al. Increased efficiency of preimplantation genetic diagnosis for
infertility using “no result rescue”. Fertil Steril. 2007;88(1):53–61.
18
Invasive Techniques: Aneuploidy Testing by Array-CGH
Introduction
Aneuploidy is the term used to describe gross chromosomal imbalance in an organism or embryo, presenting
as either additional (e.g., trisomy) or missing (e.g., monosomy) chromosomes. Aneuploidy arises during cell
division when chromosomes fail to separate equally between the two new daughter cells [1]. Aneuploidy may
be present in all the cells of the body and extra embryonic membranes, or it may be represented in a concep-
tus having both normal and aneuploid cells (so-called mosaicism). The consequences of aneuploidy constitute
a wide phenotypic spectrum from early embryonic arrest to mild infertility, with the best-known being Down
syndrome (trisomy 21) and first trimester spontaneous abortion (mostly trisomies and monosomy X). The origins
of aneuploidy lie in the meiotic divisions (principally in the ovary) and the early cleavage divisions of the pre-
implantation embryo. Arising by either nondisjunction, precocious separation of sister chromatids, or anaphase
lag [1], the impact of aneuploidy on families can be devastating, for example, when faced with pregnancy loss,
stillbirth, or a severely affected child. In all cases, aneuploidy results in an unfavorable outcome for the family
in question and is undoubtedly a major contributing factor to the relatively low fecundity of humans compared
with other species.
The concept of preimplantation genetic screening (PGS) for aneuploidy in oocytes or preimplantation embryos
to both lower the risk of the above-mentioned phenotypic consequences and improve in vitro fertilization (IVF)
success rates is not new. Indeed it was proposed alongside the earliest developments of preimplantation genetic
diagnosis [2]. The ability to do this effectively has required rapid evolution of diagnostic technologies to com-
bine speed, accuracy, and reliability. To date, only direct analysis of chromosome copy number has been applied
clinically because indirect approaches (e.g., metabolomic analysis of embryonic products or detailed morpho-
kinetic analysis using time-lapse imaging technology) have yet to be convincingly associated with aneuploidy
incidence.
Explanations for the failure of a fluorescence in situ hybridization (FISH) approach on blastomeres from
cleavage-stage embryos to demonstrate a clinical benefit in a large randomized controlled trial [3] and in
subsequent trials (see meta-analysis [4]) are well documented [5]. Potential reasons proposed for the reported
failure include the safety of biopsy, the importance of low error rates on the diagnostic efficiency of PGS [6],
and the need to detect all chromosomes simultaneously for aneuploidy. Notwithstanding the ability now to
detect all 24 chromosomes by FISH using successive rounds of hybridization [7], the issues of mosaicism, sig-
nal interpretation, clinical trial data, and the development of microarray-based methods for detecting 24 chro-
mosome copy number are now signaling the demise of FISH-based cleavage-stage embryo biopsy approaches
to PGS. Microarray-based tests are now the standard, and these tests have been made possible through the
advancement of whole-genome amplification (WGA) technology. Despite the undoubtedly superior technical
capabilities of array comparative genomic hybridization (aCGH) compared with FISH, and several prospec-
tive clinical trials showing benefit and widespread clinical use of aCGH, the policy position regarding PGS
among both professional and regulatory bodies is regrettably out of date and refers only to the historic and
flawed FISH approach [8,9].
The classical approaches to embryo biopsy for preimplantation genetic diagnosis (polar body, cleavage
stage, trophectoderm [10–12] and reviewed in [13]) are all in current clinical application for PGS, with first
255
polar body (PB1) [14], combined PB1 and second polar body (PB2) [15], and trophectoderm biopsy [16]
increasingly finding favor. The more well-established cleavage-stage approach, although once thought to
be harmless [17], is now considered to reduce implantation potential, especially when two cells are biop-
sied [3,18], and should ideally be used only after conducting a cost–benefit analysis. The invasive nature of
embryo or oocyte biopsy means that PGS has historically been targeted to specific high-risk patient groups
(e.g., advanced maternal age, repeated implantation failure, recurrent pregnancy loss, and elevated sperm
aneuploidy levels [common in patients presenting with severe male factor infertility]). More recently, PGS
has been used to improve the effectiveness of elective single embryo transfer in good prognosis patients to
reduce multiple birth rates while maintaining high success rates [19]. The complex spectrum of chromosome
abnormalities in the human preimplantation embryo has yet to be fully described, and diagnostic procedures
can be expensive to implement. Moreover, embryo biopsy is invasive; thus, a robust cost–benefit analysis
is essential to achieve widespread patient benefit through the use of PGS [20]. This chapter explores the
current methodologies used to perform PGS using aCGH and provides a description of alternative and future
approaches.
Biopsy Strategies
Several different strategies have been proposed and used to detect aneuploidy using aCGH in oocytes and
embryos. The relative merits of each are listed in Table 18.1. PB1 biopsy alone and combined PB1 and PB2
strategies have both been used clinically for PGS. However, it is increasingly evident that PB1 alone has limited
applicability for PGS because up to 30% of aneuploidy of maternal origin will not be diagnosed if only PB1
is analyzed [26]. Because precocious separation of sister chromatids appears to be the predominant cause of
maternal meiotic aneuploidies [26,27], PB2 must be biopsied to accurately identify all maternal aneuploidies and
to ensure even abnormal segregations in PB1 are not corrected in the second meiotic division. The timings of
both PB1 and PB2 biopsy are critical to the efficiency of diagnosis. This was relevant when aneuploidy screening
using the FISH approach was used [28] and is equally critical when using aCGH [29].
Theoretically, blastocyst-stage biopsy is the optimal stage for aneuploidy screening because it partially
negates the problem of mosaicism and gives maximum aneuploidy information from maternal, paternal, and
postzygotic events at the latest possible stage of embryo development possible in current in vitro culture sys-
tems. In addition, the biopsy of three or more cells virtually eliminates the problem of allelic dropout (ADO)
after WGA [30]. Historically, it has been viewed as a downside that embryos may need cryopreservation
after trophectoderm biopsy while awaiting genetics results. However, it is becoming increasingly apparent
that vitrification is a viable strategy to maintain or even potentially increase live birth rates after biopsy [31].
Furthermore, embryo vitrification may be considered for all PGS cases to overcome logistic issues with sample
transportation and diagnostic timings. It should be noted that TE biopsy does not eliminate the problem of
mosaicism. Furthermore, it is possible that some embryos failing to reach blastocyst stage in vitro may be viable
in vivo [32].
Principles of aCGH
Originally designed for molecular karyotyping of tumor cells [33], CGH is a method whereby the chromosomal
genotype of an unknown DNA sample can be inferred according to its relative ability to competitively hybrid-
ize with reference DNA of known genotype to either (1) metaphase chromosomes from a karyotypically normal
reference male (metaphase CGH) or (2) a series of specified DNA sequences at specified spots or positions on
a glass slide (aCGH). A schematic representation of the principles of CGH is shown in Figure 18.1. Like its
more time-consuming predecessor, metaphase CGH, which has been used for clinical PGS [34,35], aCGH is a
relatively DNA-hungry technique and commonly requires nanogram to microgram amounts of DNA for opti-
mal performance. It is estimated that a single blastomere contains approximately 6 pg of DNA; therefore, it is
essential to perform WGA before the aCGH procedure itself. Although aCGH can be performed using either
Aneuploidy Testing by Array-CGH 257
TABLE 18.1
Advantages and Disadvantages of Different Biopsy Strategies for PGS of Aneuploidy
Biopsy Stage Advantages Disadvantages
TE biopsy Fewer embryos are ultimately tested reducing diagnostic costs. Blastocyst culture potentially not suitable
Only the highest quality embryos (suitable for embryo transfer for all patients. Some viable embryos may
[ET]) are tested. not reach blastocyst in vitro.
Fresh ET on Day 6 (or Day 7) is possible in many cases. Higher “no biopsy” rate compared with PB
Maternal, paternal, and postzygotic errors can be detected. biopsy and cleavage-stage biopsy.
Multicellular samples result in better amplification rates than Depending on availability of local
single cell and enable detection of mosaicism. diagnostic services, vitrification of all
Mosaicism may not be major issue for TE biopsy due to low biopsied blastocysts may be required.
prevalence of mosaic diploid/aneuploid blastocysts and the high Significance of mosaicism is unclear.
detection rate of clinically relevant mosaicism. Aneuploidy
detected at the earlier stages may not affect the blastocyst [21,22].
Cleavage-stage Fresh ET from Day 4 onward is possible in many cases. High rates of mosaicism are present in even
(blastomere) Maternal, paternal, and post zygotic errors detected. good-quality cleavage-stage embryos [23].
biopsy Inability to distinguish between aneuploidy
mosaic and uniformly aneuploid embryos
leads to false-positive results and
subsequent discard of potentially
chromosomally normal viable embryos.
Biopsy of one or two cells affects
implantation rate [18].
PB biopsy Removal of PBs from a human oocyte has no deleterious effect on Biopsy of first and second polar bodies is
subsequent embryo, fetal, and infant development as neither is relatively labor-intensive (many oocytes
required for successful fertilization or embryogenesis [24]. may not even develop into therapeutic
Detection of constitutive embryonic aneuploidies of maternal quality embryos).
origin without confounding mosaicism. Extensive testing may be expensive.
Majority of patients will have a biopsy. Does not allow the detection of
Screening of whole cohort of oocytes provides comprehensive aneuploidies of paternal origin or those
diagnostic information. arising after fertilization in the embryo.
May be suitable to patients for whom blastocyst culture is
typically unsuccessful.
Flexibility of fresh ET timing, in many cases from Day 2 onward.
High concordance with zygote aneuploidy [14] and cleavage-
stage aneuploidy [25].
bacterial artificial chromosome (BAC) DNA clones or specific oligonucleotides across the genome, this chapter
focuses on the BAC clone approach because this approach has been validated and used in >300,000 clinical pre-
implantation genetic samples to date. The current 24Sure™ microarray contains 2900 unique BAC clones spaced
approximately 1 Mb apart that have been extensively validated in >2000 postnatal clinical array experiments to
exclude copy number polymorphisms. In addition, the exact genomic location of each probe has been confirmed
by reverse painting of labeled single chromosome preparations onto arrays, FISH mapping, and sequencing
verification. aCGH for preimplantation testing has been pioneered by BlueGnome (Fulbourn, Cambridge, United
Kingdom), and all reagents for WGA, labeling, hybridization, washing, reference DNA, microarray slides, and
analytical software described below are available from BlueGnome except where noted.
aCGH for Chromosome Enumeration

After egg (PB1, PB2, or both), embryo (cleavage-stage), or blastocyst biopsy, the biopsied material is
washed in a buffer, typically phosphate-buffered saline (PBS), with an additive such as polyvinyl alco-
hol (PVA) to reduce cell stickiness [36]. After washing, the biopsy material is picked up in a very small
Biopsied cell(s)
Biopsied cell(s)
DNA
Whole genome amplification
Combined labeled
& control and biopsied
cell(s) DNA
Control DNA Biopsied cell(s)
DNA
A B
Metaphase Array
CGH CGH
1:1 2:1 2:3

Normal Monosomy Trisomy Normal Monosomy Trisomy
FIGURE 18.1 Schematic of comparative genomic hybridization.
volume (<2 μL) and placed into a sterile 0.2 mL Eppendorf tube for transport to the testing laboratory.
A more detailed description of this process is given in Chapter 14. Most laboratories perform a quick cen-
trifugation step to ensure that the cellular material and all of the fluid are collected at the bottom of the
tube. Most labs then freeze each sample before transport to the testing lab; however, this is not essential,
depending on the length of time the sample will be in transit before further processing. It is very important
to note that the m ineral oil that is typically used as an overlay on top of biopsy material destined for poly-
merase chain reaction (PCR) testing should never be used before WGA and aCGH because it inhibits the
amplification process.
WGA and Labeling

Several different WGA methods have been used historically for aCGH, with the current, most often used
method being SurePlex™ (Rubicon Genomics Inc., Ann Arbor, MI; and BlueGnome). After fragmentation of
the sample DNA, self-inert degenerative primers are annealed at multiple sites along the genome. Extension
then displaces downstream strands to generate multiple fragments spanning each region. The reaction pro-
duces large fragment sizes that are reproducible between samples and that are optimized for aCGH. Many
of the other WGA techniques have been adapted for use in aCGH, but they were originally used for other
purposes (e.g., single-locus PCR and mutation detection). SurePlex is suitable because of its simple, short
protocol; highly representative amplification; and low allele dropout rates. In brief, after sample receipt in the
lab, each tube is opened in a dedicated DNA amplification cleanroom, under laminar flow conditions, and the
WGA reagents are added (SurePlex kit). Amplification is performed according to the manufacturer’s instruc-
tions because these kits have been validated using single cells. For SurePlex, there is a 15 min cell lysis (DNA
extraction) step, followed by the preamplification steps (90 min), and finally amplification (30 min). All of these
steps are performed in a single tube, thereby reducing the likelihood of sample switches and contamination.
In addition, all of these steps require the use of a PCR thermal cycler machine because they are time and tem-
perature dependent. Because the arrays are the most expensive consumable in the process, it is best to ensure
amplification before taking the sample further through the process. After amplification, most laboratories run
an agarose gel electrophoresis step to confirm amplification. A smear of high-molecular-weight DNA, observed
on the gel after electrophoresis, is indicative of positive amplification, and all such samples may be taken
forward to the fluorescent labeling steps. Low-molecular-weight DNA, or the absence of any smear, indicates
poor or failed amplification. In such cases, it may be prudent to avoid running such samples on the microarrays.
Successfully amplified WGA product is labeled through nick translation with either Cy3 (green) or Cy5 (red)
fluorescence and purified.
Hybridization
Samples with unknown genotype (i.e., embryo biopsy) labeled in one fluorescent color and control reference
DNA (typically a karyotypically normal male) labeled in the opposite color are separately denatured (to render
them single stranded) at 74°C before being mixed together in equal proportions in hybridization buffer contain-
ing formamide and cot-1 human DNA before adding to each 24Sure microarray. Microarrays are hybridized at
47°C for at least 4 hr or overnight in a humidified chamber. The length of hybridization time varies depending
on the number of samples in the lab on any given day, the time samples are received during the day, and the
local staffing levels and shift patterns. During validation of the array in the lab, hybridization times as short
as 3 hr and as long as 16 hr (overnight) were tested with no differences noted (BlueGnome, unpublished data).
On the basis of these results, hybridization for at least 4 hr and no longer than 16 hr is deemed to be interchange-
able (Reprogenetics Ltd., unpublished data).
Posthybridization Washing
After hybridization, each microarray is washed as follows: 10 min in 2× standard saline citrate (SSC)/0.05%
Tween 20 at room temperature, 10 min in 1× SSC at room temperature, 5 min in 0.1× SSC at 60°C, and 2 min in
0.1× SSC at room temperature to remove unbound DNA.
Scanning
Each microarray slide is scanned using a dual-channel fluorescent laser scanner that creates TIFF images
(e.g., ClearScan™, BlueGnome) for green fluorescence at 632 nm and for red fluorescence at 587 nm. Raw
images are loaded automatically into BlueFuse™ software allowing for automated evaluation of fluorescent
signals (ratio analysis).
Scoring
Each sample is scored by a trained technologist who assesses traces for all 24 chromosomes, noting all
gains and losses as well as determining the sex of each sample. A second technologist then scores the
sample blindly, with no knowledge of the initial score by the first technologist. A final score for each
sample is assigned by comparing the two scores. Any discrepancies are noted and are adjudicated by a third
technologist, the laboratory supervisor or director, or both. Single chromatid errors can be distinguished
from whole chromosome errors through examination of the mean per-chromosome hybridization ratios
(Figure 18.2) [27].
1.20
0.88
0.60 Chromatid gain Chromatid gain
0.24
Log2 ratio Ch1/Ch2
0.00
–0.40
–0.72
Chromosome loss
–1.04
–1.36
–1.68
21
10
11
12
13
14
15
16
17
18
19
20
22
1
Y
X
Chromosomal position
FIGURE 18.2 Determining chromatid versus chromosome loss in first polar body samples by aCGH. For most chromosomes
(i.e., not the sex chromosomes or the aneuploid chromosomes), a clear and consistent 1:1 ratio is observed along the chromosome
length. Because the polar body sample was cohybridized with male genomic DNA, a hybridization pattern representing a 2:1 ratio
for the X chromosome and a “0:2” ratio for the Y chromosome is observed. This polar body clearly shows multiple aneuploidies
with chromatid gains on chromosomes 1 and 10 (single chromatid gains are consistent with a 3:2 [or 1.5:1] ratio, i.e., approximately
half that of the X chromosome shift) and a loss of whole chromosome 15 (similar to the shift seen for the absent Y chromosome).
Reporting
Once results for all samples from each patient are finalized, a diagnostic report is prepared, signed off by an
appropriately qualified person (on site or remotely), and shared with the referring laboratory and physician before
embryo transfer [36].
Validation
In extensive validation using single cells from known cell lines against the gold standard of karyotyping, 24Sure
demonstrated 98% accuracy. In contrast to the use of cell lines, validation for embryo aneuploidy is difficult.
To obtain reliable and robust results, one needs truth data (i.e., samples of known genotype). Although the same
is true of human oocytes, in that they are of unknown genotype, the ability to biopsy both the PB1 and PB2
from the oocyte provides the opportunity to obtain relatively robust validation data comparing results from
so-called “trios” of both polar bodies and their corresponding oocyte. The expectation is to see reciprocal results
(i.e., chromosomal gains and losses) from aneuploid polar body and oocyte (Figure 18.3). The presence of chromo-
somal mosaicism in human embryos makes it difficult to categorically identify embryos as having a single s pecific
genotype. Thus, from a mosaic embryo, individual single-cell results may appear to be unrepresentative and
multicellular results (e.g., from trophectoderm biopsy) potentially difficult to interpret (Figure 18.4) and therefore
challenging to make decisions regarding embryo selection.
To date, 24Sure and 24Sure-plus™ arrays have been validated using several different cell types to evaluate both
technical and biological performance, some examples of which are listed in Table 18.2. After clinical implemen-
tation, it is essential to maintain accuracy and overall quality assurance of the test(s) offered in your laboratory.
2.00
1.60
1.20
Log2 ratio Ch1/Ch2
0.80
0.40
0.00
–0.40
–0.80
–1.20
–1.60
10
11
12
13
14
15
16
17
18
19
20
22
21
Y
1
9
(a)
2.00
1.60
1.20
Log2 ratio Ch1/Ch2
0.80
0.40
0.00
–0.40
–0.80
–1.20
–1.60
1
10
11
12
13
14
15
16
17
18
19
20
21
22
Y
(b)
FIGURE 18.3 aCGH trace from second polar body and corresponding oocyte sample. Multiple aneuploidies (gains or losses)
detected in (a) the PB2 (–2, –5, –9, –15, and –21) are seen in the reciprocal form in (b) the corresponding oocyte (+2, +5, +9, +15, +21,
and also –19). Note the increased amplitude of the signal-to-noise ratio for the oocyte versus the polar body sample. In part, this is
due to the quantity and quality of DNA available within the respective cells.
This task is often performed by means of internal quality control measures and external quality assessment (EQA)
schemes. The sex chromosomes provide a good internal control by observing the X and Y chromosome devia-
tion from the autosomes within a euploid DNA complement. This level of deviation can be used as a guide to
assess aneuploid and euploid positions on the aCGH plot. For EQA, a scheme for single-cell aCGH is currently in
development based on earlier single-cell PGD schemes [38], and in many countries such schemes are required for
laboratory accreditation and to meet regulatory requirements.
High-Resolution aCGH for Detection of Chromosome Imbalance

Where aCGH is used to detect chromosomal imbalance in embryos derived from couples where at least one
partner carries a balanced translocation, an assessment of (1) the likely unbalanced outcomes should be made
by a qualified genetics professional and (2) the likely size of all unbalanced products should be performed.
A prediction tool (available from BlueGnome) can be used to ensure the microarray has sufficient resolu-
tion to detect all potential products. For example, the current version of 24Sure-plus contains 4800 BAC
clones and claims to be able to accurately detect products as small as 10 Mb (with possible detection to the
2.5 Mb r esolution level in regions with good clone coverage). The protocol for use of 24Sure-plus is essen-
tially the same as for 24Sure, with the primary difference being the higher resolution microarray slide used in
2.00
1.60
1.20
0.80
Log2 ratio Ch1/Ch2
0.40
0.00
–0.40
–0.80
–1.20
–1.60
1
6
7
8
9
10
11
12
13
14
165
19
17
18
20
21
22
X
Y
1
(a)
2.00
1.60
1.20
0.80
Log2 ratio Ch1/Ch2
0.40
0.00
–0.40
–0.80
–1.20
–1.60
165
10
11
12
13
14
1
6
7
8
9
17
18
19
20
21
X
22
Y
1
(b)
FIGURE 18.4 Detection of mosaicism in trophectoderm samples using aCGH. Trophectoderm samples from two different blas-
tocysts show likely chromosomal mosaicism as follows: (a) euploid female except for mosaic monosomy 8 and (b) euploid male
except for mosaic trisomy 7. Mosaicism is suspected because the signals for the remaining chromosomes are relatively uniform
and the log2 ratio shift is uniform along the length of the chromosome but does not exceed the threshold (bottom red and top green
line, respectively) required to call as a uniform aneuploidy. It is unclear whether the presence of these cells identifies this embryo as
chromosomally abnormal and potentially nonviable for this reason. Note the generally high signal-to-noise ratio for this multicel-
lular sample compared with that seen for polar body samples in Figure 18.2. (Data courtesy of Dr. Francesco Fiorentino, Genoma
Laboratories, Rome, Italy.)
the 24Sure-plus test. Aside from improved accuracy and effectively eliminating the need to provide couple-
specific test development, an additional benefit of using aCGH for translocation carriers is the simultaneous
detection of aneuploidy for all the other chromosomes not involved in the translocation [39]. An example of
the array trace showing reciprocal gains and losses in the predicted chromosomes (involved in the transloca-
tion) is shown in Figure 18.5.
Limitations of aCGH
aCGH is highly accurate (98%), and in competent hands it delivers a 98% result rate. Despite its proven accu-
racy, aCGH has clinically relevant limitations in the field of preimplantation genetics. For example, it cannot
TABLE 18.2
Formal Preclinical Validation of 24Sure BAC Microarrays
Cell/Sample Type Known Genotype Accuracy of aCGH Diagnosis (%)
Blinded euploid/aneuploid ovarian carcinoma cell lines Various aneuploid lines 51/51 (100)
(single cells) (Dr. Joyce Harper, UCL Centre for PGD,
Personal Communication)
Blinded aneuploidic cell lines (single cells) (BlueGnome, 45, X; 47, XY +13; 47, XY 118/121 (98)
unpublished data) +21; 47, XYY; 47, XXY
Human embryonic blastomeres (reanalyzed embryos) [37] Various genotypes established 53/54 (98)
by 12-color FISH
Human oocytes and polar bodies [15] Various genotypes deduced by 90% concordance in 226 trios (PB1/2
reciprocity between PB1/2 and oocyte)
and corresponding oocyte
2:0kb 5:0kb
–2.00 –1.60 –1.20 –0.80–0.40 0.00 0.40 0.80 1.20 1.60 –2.00 –1.60 –1.20 –0.80 –0.40 –0.00 0.40 0.80 1.20 1.60 2.0
2:242,951 kb 5:180,858 kb Log ratio Ch1/Ch2
Log2 ratio Ch1/Ch2 2
(a) (b)
FIGURE 18.5 Detection of partial aneuploidies in embryonic samples from a reciprocal translocation carrier, t(2;5)(q21;q31), using
array CGH. Embryos from reciprocal translocation carriers can be used to validate 24Sure detection of segmental aneuploidies
because each affected embryo will have segmental aneuploidies based at two specific breakpoints. Detecting gains (chromosome 2q)
and losses at these two specific breakpoints internally validates the accuracy of the test. In this case, one parent carried the translo-
cation t(2;5)(q21;q31), resulting in gains and losses (chromosome 5q) of chromosomal material in the single blastomeres of affected
embryos. Even though the deletion/duplications can be relatively small, the combination of a higher resolution microarray, known
breakpoints, and their predicted meiotic products and a multicellular sample makes it relatively straightforward to detect reliably.
discriminate between maternal and paternal errors. Nor can it distinguish between meiotic and mitotic errors of
chromosome segregation. For the purposes of the cycle in which the testing is being done, this is not so relevant.
However, such additional information may provide clues as to how to treat the patient in the future. Finally,
because aCGH is a method predicated on high-quality DNA and successful hybridization, the possibility exists
that borderline results could be miscalled (as with FISH).
Future Opportunities
Although aCGH has become the gold standard for direct chromosome enumeration in embryos and oocytes,
there are alternatives available. As with all competing technologies, there are advantages and disadvantages to
each [40,41]. Comprehensive chromosomal screening using multiplex quantitative fluorescent PCR [42] may pro-
vide a cheaper alternative, but it is not currently commercially available, thereby restricting its use. Aneuploidy
can also be detected with single-nucleotide polymorphism (SNP) arrays using a combination of loss of hetero-
zygosity (monosomy), quantitation of specific SNP loci, and incongruous SNP calls compared with predicted
Mendelian results with parental information that are incompatible with normal disomy [43–47]. Next generation
sequencing (NGS) promises to supersede all other methodologies [41], but currently it is cheaper than array-
based methods only if large numbers of samples are processed simultaneously. At this time, NGS solutions are
not commercially available.
Noninvasive Indirect Methods of Determining Aneuploidy

Weak correlations exist between the presence of embryonic aneuploidy and morphological aspects of embryo
development after retrospective analysis [48,49]. Such findings have stimulated the field of morphokinetic analy-
sis of embryos in an attempt to identify aneuploidy in a real-time clinical setting. Time-lapse imaging and
analysis appear to demonstrate that morphological features and developmental timings of the embryo have some
relationship to aneuploidy, with an algorithm based on the onset and duration of blastulation correlating well
with implantation rates [50,51]. This finding, if confirmed in larger data sets and with appropriate subgroup
analysis stratified by maternal age, provides some useful prioritization criteria but cannot, at present, replace
the specificity and accuracy of aneuploidy testing using aCGH. Another promising morphokinetic approach is
to assess dynamic fragmentation patterns within early embryos; but again, it currently does not identify specific
aneuploidies and cannot be regarded as a threat to direct genetic analysis [52]. Indeed, to date, no morphokinetic
parameter has been observed to discriminate between euploid and simple aneuploid (e.g., trisomy 21) embryos.
Moreover, although there have been some useful predictors of viability related to specific metabolites, none so
far has been linked specifically to aneuploidy rather than viability or been able to differentiate between general
chromosomal aneuploidy and specific aneuploidy [53,54].
Summary and Essentials

At present, aCGH is considered the gold standard for detecting aneuploidy in single cells or multicellular
samples from oocytes and embryos because of its reliability, reproducibility, and accuracy and the large
worldwide experience with it. It is possible to obtain results within 12 hr after biopsy, making it acces-
sible to most laboratories regardless of the biopsy stage selected. Although there remains some contro-
versy over the benefit of PGS for specific patient indications, there appears to be a recent shift toward
trophectoderm (multicellular) biopsy of the highest quality embryos from good prognosis patients in
contrast to the historic focus on cleavage-stage biopsy of few, poor-quality embryos in poor prognosis
patients. Irrespective of the approach taken, the following tips should be considered critical for effective use
of this technology.
Quality of the IVF Laboratory

Overall quality of the IVF program is critical. Whenever an oocyte or embryo is subjected to a procedure
outside of the incubator, there is a risk involved. Thus, a fundamental principle underpinning the use of aneu-
ploidy screening is that the benefit gained should outweigh the harm, if any, caused. Thus, if the success rates
of the p rogram are already suboptimal, it is difficult to see any procedure providing sufficient benefit to rescue
the cycle.
Embryo biopsy
Embryo biopsy has become simpler with the availability of the noncontact infrared laser that is common in
many IVF laboratories (see Chapters 14 and 15). With this laser, it is relatively simple to perform oocyte or
embryo biopsy, in many cases without apparent damage to the subject. However, after indiscriminate use,
invisible thermal damage might be caused that, although not immediately lethal, could have later conse-
quences, compromising development and subsequent implantation. For this reason, it is vital to perform the
embryo biopsy as quickly as possible with minimal exposure to laser energy to complete the task safely and
effectively. The issue of embryo selection for biopsy is not straightforward because there is inevitably a balance
between the cost and efficacy of biopsying all embryos in the cohort against the need for all information and
the small chance that apparently delayed embryos (either at cleavage or blastocyst stages) may still have some
implantation potential.
Sample Preparation and Transportation

Proper preparation of the biopsied sample, although mundane, is of critical importance. Focus on the steril-
ity of the working area and solutions; precise volume of buffer in the microcentrifuge tube, with minimal
carryover of embryo culture medium, which can reduce amplification efficiency; and subsequent storage of
the sample preanalysis is paramount to ensure high diagnostic success rates. Particular care must be taken
not to contaminate the sample with foreign DNA by means of good laboratory practice and appropriate
apparel and dedicated cleanroom, equipment, and consumables for amplification steps. A negative control
of the embryo media and collection buffer should always be taken at this point to check for the absence of
contamination because aCGH is not able to identify the origin of any contamination. Although published
protocols exist and diagnostic service laboratories generally provide their own standard operating proce-
dures, a series of laboratory-specific validation experiments are extremely useful before offering the service
clinically.
Whole-Genome Amplification
Many different methods are available and have been reported for use in preimplantation genetic testing. However,
the specifications of the amplified DNA optimal for aCGH are DNA fragments of a specific size correspond-
ing to the specific array type. For 24Sure BAC arrays, SurePlex (a PCR-based method) is used in preference to
multiple displacement amplification (MDA).
aCGH Procedure
Hybridization is fairly robust but care should be taken to ensure microarray slides do not dry out before wash-
ing. The high-temperature, high-stringency wash posthybridization must be temperature controlled. Lower
temperatures prevent the removal of nonspecific-labeled DNA from the array and generate noisy results. If the
temperature is too high, too much of the labeled DNA will be stripped from the array and could result in too few
probes per chromosome to accurately call the result, particularly for smaller chromosomes. Drying of the micro-
array slides is also critical; the most effective way is to mechanically remove wash buffer by centrifugation.
If wash buffer is left to dry on the slides, it fluoresces and will reduce data quality. After drying, slides should
be scanned immediately because in some circumstances the fluorescent dyes can be degraded by atmospheric
ozone. Although the software is highly accurate, a combination of both automated and manual calling of results
is recommended.
Target Population for Testing

Regardless of the testing method used, it is important to properly identify the appropriate patients who would
benefit from the test. Indications for testing vary widely, and it is crucial for both providers and patients to
understand the difference between using the test to provide diagnostic information (e.g., in the case where
there are very few embryos present but they are highly likely to be grossly aneuploidy) or information to
enable selection of euploid embryos (e.g., to improve the likelihood of success in that embryo transfer cycle).
Essentially, in the absence of comprehensive prospective randomized controlled clinical trials for each
putative indication for PGS, clinics must conduct a cost–benefit exercise weighing the potential prognostic
(selection) and diagnostic (closure or alternative therapy) benefits against the financial cost to the patient and
biological cost to the biopsied embryo [20].
REFERENCES
1. Hassold T, and Hunt P. (2001). To err (meiotically) is human: The genesis of human aneuploidy. Nat Rev Genet
2, 280–91.
2. Penketh R, and McLaren A. (1987). Prospects for prenatal diagnosis during preimplantation human development.
Baillieres Clin Obstet Gynaecol 1(3), 747–64.
3. Mastenbroek S, Twisk M, van Echten-Arends J, Sikkema-Raddatz B, Korevaar JC, Verhoeve HR, et al. (2007).
In vitro fertilization with preimplantation genetic screening. N Engl J Med 357, 9–17.
4. Mastenbroek S, Twisk M, van der Veen F, and Repping S. (2011). Preimplantation genetic screening: A system-
atic review and meta-analysis of RCTs. Hum Reprod Update 17(4), 454–66.
5. Cohen J, and Grifo JA. (2007). Multicentre trial of preimplantation genetic screening reported in the New England
Journal of Medicine: An in-depth look at the findings. Reprod Biomed Online 15, 365–6.
6. Summers MC, and Foland AD. (2009). Quantitative decision-making in preimplantation genetic (aneuploidy)
screening (PGS). J Assist Reprod Genet 26, 487–502.
7. Ioannou D, Fonseka KGL, Meershoek EJ, Thornhill AR, Abogrein A, Ellis M, et al. (2012). Twenty four chromo-
some FISH in human IVF embryos reveals patterns of post-zygotic chromosome segregation and nuclear organi-
zation. Chromosome Res 20, 447–60.
8. Anderson RA, and Pickering S. (2008). The current status of preimplantation genetic screening: British Fertility
Society Policy and Practice Guidelines. Hum Fertil (Camb) 11, 71–5.
9. American Society of Reproductive Medicine (ASRM) Practice Committee. (2008). Preimplantation genetic
testing: A Practice Committee opinion. Fertil Steril 90, S136–43.
10. Magli MC, Gianaroli L, Ferraretti AP, Toschi M, Esposito F, and Fasolino MC. (2004). The combination of polar
body and embryo biopsy does not affect embryo viability. Hum Reprod 19, 1163–9.
11. Cieslak-Janzen J, Tur-Kaspa I, Ilkevitch Y, Bernal A, Morris R, and Verlinsky Y. (2006). Multiple micromanipula-
tions for preimplantation genetic diagnosis do not affect embryo development to the blastocyst stage. Fertil Steril
85, 1826–9.
12. Kokkali G, Traeger-Synodinos J, Vrettou C, Stavrou D, Jones GM, Cram DS, et al. (2007). Blastocyst biopsy
versus cleavage stage biopsy and blastocyst transfer for preimplantation genetic diagnosis of beta-thalassaemia:
A pilot study. Hum Reprod 22, 1443–9.
13. Thornhill AR, Ottolini C, and Handyside AH. (2012). Human embryo biopsy procedures. In Textbook of Assisted
Reproductive Technologies, 4th edn. (eds. Gardner DK, Howles CM, Weissman A, and Shoham Z). London, UK:
Informa Healthcare, pp. 197–211.
14. Wells D, Escudero T, Levy B, Hirschhorn K, Delhanty JD, and Munne S. (2002). First clinical application of
comparative genomic hybridization and polar body testing for preimplantation genetic diagnosis of aneuploidy.
Fertil Steril 78, 543–9.
15. Geraedts J, Montag M, Magli MC, Repping S, Handyside A, Staessen C, et al. (2011). Polar body array CGH for
prediction of the status of the corresponding oocyte. Part I: Clinical results. Hum Reprod 26(11), 3173–80.
16. Schoolcraft WB, Fragouli E, Stevens J, Munne S, Katz-Jaffe MG, and Wells D. (2010). Clinical application of
comprehensive chromosomal screening at the blastocyst stage. Fertil Steril 94, 1700–6.
17. Hardy K, Martin KL, Leese HJ, Winston RM, and Handyside AH. (1990). Human preimplantation development
in vitro is not adversely affected by biopsy at the 8-cell stage. Hum Reprod 5, 708–14.
18. Cohen J, Wells D, and Munne S. (2007). Removal of 2 cells from cleavage stage embryos is likely to reduce the
efficacy of chromosomal tests that are used to enhance implantation rates. Fertil Steril 87, 496–503.
19. Yang Z, Liu J, Collins GS, Salem SA, Liu X, Lyle SS, et al. (2012). Selection of single blastocysts for fresh
transfer via standard morphology assessment alone and with array CGH for good prognosis IVF patients: Results
from a randomised pilot study. Mol Cytogenet 5, 24.
20. Ottolini C, Griffin DK, and Thornhill AR. (2012). The role of aneuploidy screening in human preimplantation
embryos. In Aneuploidy in Health and Disease (ed. Storchova Z). InTech. ISBN 978-953-51-0608-1, pp. 217–44.
21. Capalbo A, Bono S, Spizzichino L, Biricik A, Baldi M, Colamaria S, et al. (2013). Sequential comprehensive
chromosome analysis on polar bodies, blastomeres and trophoblast: Insights into female meiotic errors and
chromosomal segregation in the preimplantation window of embryo development. Hum Reprod 28(2), 509–18.
22. Capalbo A, Wright G, Elliott T, Ubaldi FM, Rienzi L, and Nagy ZP. (2013). FISH reanalysis of inner cell mass
and trophectoderm samples of previously array-CGH screened blastocysts shows high accuracy of diagnosis and
no major diagnostic impact of mosaicism at the blastocyst stage. Hum Reprod 28(8), 2298–307.
23. Mertzanidou A, Wilton L, Cheng J, Spits C, Vanneste E, Moreau Y, et al. (2013). Microarray analysis reveals
abnormal chromosomal complements in over 70% of 14 normally developing human embryos. Hum Reprod
28(1), 256–64.
24. Gianaroli L. (2000). Preimplantation genetic diagnosis: Polar body and embryo biopsy. Hum Reprod 15(Suppl 4),
69–75.
25. Christopikou D, Tsorva E, Economou K, Shelley P, Davies S, Mastrominas M, et al. (2013). Polar body analysis
by array comparative genomic hybridization accurately predicts aneuploidies of maternal meiotic origin in cleav-
age stage embryos of women of advanced maternal age. Hum Reprod 28(5), 1426–34.
26. Handyside AH, Montag M, Magli MC, Repping S, Harper J, Schmutzler A, et al. (2012). Multiple meiotic
errors caused by predivision of chromatids in women of advanced maternal age undergoing in vitro fertilisation.
Eur J Hum Genet 20(7), 742–7.
27. Gabriel AS, Thornhill AR, Ottolini CS, Gordon A, Brown AP, Taylor J, et al. (2011). Array comparative genomic
hybridisation on first polar bodies suggests that non-disjunction is not the predominant mechanism leading to
aneuploidy in humans. J Med Genet 48, 433–7.
28. Verlinsky Y, Cieslak J, Freidine M, Ivakhnenko V, Wolf G, Kovalinskaya L, et al. (1996). Polar body diagnosis of
common aneuploidies by FISH. J Assist Reprod Genet 13, 157–62.
29. Magli MC, Montag M, Koster M, Muzi L, Geraedts J, Collins J, et al. (2011). Polar body array CGH for predic-
tion of the status of the corresponding oocyte. Part II: Technical aspects. Hum Reprod 26(11), 3181–5.
30. Ling J, Zhuang G, Tazon-Vega B, Zhang C, Cao B, Rosenwaks Z, et al. (2009). Evaluation of genome
coverage and fidelity of multiple displacement amplification from single cells by SNP array. Mol Hum Reprod
15, 739–47.
31. Schoolcraft WB, and Katz-Jaffe MG. (2013). Comprehensive chromosome screening of trophectoderm with vit-
rification facilitates elective single-embryo transfer for infertile women with advanced maternal age. Fertil Steril
100(3), 615–19.
32. Glujovsky D, Blake D, Farquhar C, and Bardach A. (2012). Cleavage stage versus blastocyst stage embryo trans-
fer in assisted reproductive technology. Cochrane Database Syst Rev 7, CD002118.
33. Kallioniemi A, Kallioniemi OP, Sudar D, Rutovitz D, Gray JW, Waldman F, et al. (1992). Comparative genomic
hybridization for molecular cytogenetic analysis of solid tumors. Science 258, 818–21.
34. Voullaire L, Slater H, Williamson R, and Wilton L. (2000). Chromosome analysis of blastomeres from human
embryos by using comparative genomic hybridization. Hum Genet 106, 210–17.
35. Wells D, and Delhanty JD. (2000). Comprehensive chromosomal analysis of human preimplantation embryos
using whole genome amplification and single cell comparative genomic hybridization. Mol Hum Reprod 6,
1055–62.
36. Harton GL, Magli MC, Lundin K, Montag M, Lemmen J, Harper JC, et al. (2011). ESHRE PGD Consortium/
Embryology Special Interest Group—best practice guidelines for polar body and embryo biopsy for preimplanta-
tion genetic diagnosis/screening (PGD/PGS). Hum Reprod 26(1), 41–6.
37. Gutierrez-Mateo C, Colls P, Sanchez-Garcia J, Escudero T, Prates R, Ketterson K, et al. (2011). Validation of
microarray comparative genomic hybridization for comprehensive chromosome analysis of embryos. Fertil Steril
95, 953–8.
38. Deans Z, Fiorentino F, Biricik A, Traeger-Synodinos J, Moutou C, De Rycke M, et al. (2013). The experience of
3 years of external quality assessment of preimplantation genetic diagnosis for cystic fibrosis. Eur J Hum Genet
21(8), 800–6.
39. Alfarawati S, Fragouli E, Colls P, and Wells D. (2011). First births after preimplantation genetic diagnosis of
structural chromosome abnormalities using comparative genomic hybridization and microarray analysis. Hum
Reprod 26, 1560–74.
40. Bisignano A, Wells D, Harton G, and Munné S. (2011). PGD and aneuploidy screening for 24 chromosomes:
Advantages and disadvantages of competing platforms. Reprod Biomed Online 23(6), 677–85.
41. Handyside AH. (2013). 24–chromosome copy number analysis: A comparison of available technologies.
Fertil Steril 100(3), 595–602.
42. Treff NR, Tao X, Ferry KM, Su J, Taylor D, and Scott RT, Jr. (2012). Development and validation of an accurate
quantitative real-time polymerase chain reaction-based assay for human blastocyst comprehensive chromosomal
aneuploidy screening. Fertil Steril 97(4), 819–24.
43. Handyside AH, Harton GL, Mariani B, Thornhill AR, Affara N, Shaw MA, et al. (2010). Karyomapping:
A universal method for genome wide analysis of genetic disease based on mapping crossovers between parental
haplotypes. J Med Genet 47, 651–8.
44. Johnson DS, Gemelos G, Baner J, Ryan A, Cinnioglu C, Banjevic M, et al. (2010). Preclinical validation
of a microarray method for full molecular karyotyping of blastomeres in a 24-h protocol. Hum Reprod 25,
1066–75.
45. Treff NR, Su J, Tao X, Levy B, and Scott RT, Jr. (2010). Accurate single cell 24 chromosome aneuploidy
screening using whole genome amplification and single nucleotide polymorphism microarrays. Fertil Steril 94,
2017–21.
46. Gabriel AS, Hassold TJ, Thornhill AR, Affara NA, Handyside AH, and Griffin DK. (2011). An algorithm for
determining the origin of trisomy and the positions of chiasmata from SNP genotype data. Chromosome Res
19, 155–63.
47. Brezina PR, Benner A, Rechitsky S, Kuliev A, Pomerantseva E, Pauling D, et al. (2011). Single-gene testing
combined with single nucleotide polymorphism microarray preimplantation genetic diagnosis for aneuploidy:
A novel approach in optimizing pregnancy outcome. Fertil Steril 95, 1786.e5–8.
48. Munne S. (2006). Chromosome abnormalities and their relationship to morphology and development of human
embryos. Reprod Biomed Online 12, 234–53.
49. Alfarawati S, Fragouli E, Colls P, Stevens J, Gutierrez-Mateo C, Schoolcraft WB, et al. (2011). The relationship
between blastocyst morphology, chromosomal abnormality, and embryo gender. Fertil Steril 95, 520–4.
50. Campbell A, Fishel S, Bowman N, Duffy S, Sedler M, and Hickman CF. (2013). Modelling a risk classification
of aneuploidy in human embryos using non-invasive morphokinetics. Reprod Biomed Online 26(5), 477–85.
51. Campbell A, Fishel S, Bowman N, Duffy S, Sedler M, and Thornton S. (2013). Retrospective analysis of out-
comes after IVF using an aneuploidy risk model derived from time-lapse imaging without PGS. Reprod Biomed
Online 27(2), 140–6.
52. Chavez S, Loewke K, Han J, Moussavi F, Colls P, Munne S, et al. (2012). Dynamic blastomere behaviour reflects
human embryo ploidy by the four-cell stage. Nature Comm 3, 1251.
53. Picton HM, Elder K, Houghton FD, Hawkhead JA, Rutherford AJ, Hogg JE, et al. (2010). Association between
amino acid turnover and chromosome aneuploidy during human preimplantation embryo development in vitro.
Mol Hum Reprod 16, 557–69.
54. McReynolds S, Vanderlinden L, Stevens J, Hansen K, Schoolcraft WB, and Katz-Jaffe MG. (2011). Lipocalin-1:
A potential marker for noninvasive aneuploidy screening. Fertil Steril 95, 2631–3.
19
Summary: Comprehensive Summary of Main Points by Topic
Markus Montag
This chapter contains the key points and images from the preceding chapters for ease of quick reference.
269
Chapter 1: Handling Gametes and Embryos: Sperm Collection

and Preparation Techniques
The analysis of a semen sample is crucial for the treatment decision in an assisted reproduction technique (ART)
and should be performed in the following order:
• Liquefaction
• Record of volume and appearance
• Measurement of the pH
• Motility assessment
• Identification of aggregation or agglutination
• Determination of vitality
• Evaluation of concentration and morphology
The right technique for sperm preparation can be determined only after appropriate analysis of the native semen
sample. For ART, sperm preparation should select the most motile and morphologically normal spermatozoa.
The most common preparation techniques are as follows:
• Simple washing
• Swim-up
• Density-gradient centrifugation
Semen samples that possess sperm with parameters within the normal range are processed primarily by swim-
up; samples with lower concentrations benefit from density-gradient centrifugation preparation.
Swim-up does not provide as high recovery rates as gradient centrifugation, but it is quick and easy to p erform,
and yields a population enriched with the best motile spermatozoa. Gradient centrifugation results in the retrieval
of high numbers of motile spermatozoa; however, standardization of the method is difficult due to the inherent
variation of semen samples.
Testicular or epididymal spermatozoa can be retrieved using standard surgical techniques. Sperm retrieval
rates from testicular tissue depend on the surgical technique applied and the status of the testis and its underly-
ing spermatogenic state. In cases of obstructive azoospermia, high retrieval rates are achieved. The method of
choice for surgical sperm retrieval in nonobstructive azoospermia is a microsurgical method wherein focal areas
with spermatogenic activity are selected. In nonobstructive cases where spermatogenic abnormalities are the
expected cause, spermatozoa retrieval might be laborious and time-consuming and should always be handled
with care.
Comprehensive Summary of Main Points by Topic 271
TABLE 1.1
Comparison of Different Sperm Preparation Techniques
Type of Ejaculate Quality Pros Cons
Washing (W) Normozoospermia Quick-and-easy technique Only for sperm counts with high
Yields high numbers of sperm numbers
spermatozoa Not useful for samples with
contamination with other cells,
debris, or blood cells
Swim-up (SU) Normozoospermia Quick and easy to perform Lower recovery rates compared
Moderate OAT High rates of highly progressive with DG
motile spermatozoa
Density gradient (DG) Moderate-to-severe OAT High numbers of motile Difficult to standardize
spermatozoa High-molecular-weight
Best removal of debris or other cell compounds of unknown influence
types occurring in the semen Costs more because DG solutions
sample have to be purchased
Chapter 2: Handling Gametes and Embryos: Oocyte Collection and Embryo Culture
Lars Johansson
Oocyte retrieval should be performed in a way that avoids damaging the oocyte via too strong aspiration and
exposure to suboptimal temperature.
Prepare only one dish at a time if microdrop cultures are to be used and avoid repeated retrieval of culture
media from the vial or flask.
Use embryo-tested material for all steps.
The optimal temperature must be checked at the location of the oocyte (e.g., in a dish, tube).
Work fast, concentrate, and avoid too long exposure of cumulus–oocyte complex (COC)/oocytes to the
environment.
Control and maintenance of proper pH and temperature are key to success.
TABLE 2.1
Classification of Cumulus Oophorous Complexes
Quality Cumulus Corona Oocyte
Immature (I) Small, compact, grey, Dark ring around the zona pellucida Not visible, if visible GV
(Figure 2.8) non-expanded stage
Mature (M) Small, bright, expanded Not fully radiated Partially visible
(Figure 2.9–2.11)
Excellent (E) Large, bright, expanded Radiated Clear and visible
(Figure 2.12–2.13)
Overmature (O) Small, thin, patches of dark cells. Light or dark Visible, granular and dark
(Figure 2.14–2.15)
Atretic (A) Patches of dark cells or absent None or clumps Dark
TABLE 2.2
Classification of Oocyte Maturational Stages
Stage Polar Body Nucleus
Germinal vesicle (GV) None Large halo with nucleoli
Meiosis I (MI) None None
Meiosis II (MII) Present None
TABLE 2.3
Timing of Procedures in Relationship to Injection of HCG (hpHCG)
Procedure Egg Collection Oocyte Vitrification Insemination (IVF) Denudation ICSI
hpHCG 36–38 38–40 40–41 40–41 41–42
Chapter 3: Handling Gametes and Embryos: Quality Control for Culture Conditions
Jason E. Swain
• When optimizing in vitro fertilization (IVF) results in the laboratory, the devil is in the details.
• Poor air quality in the lab can compromise embryo development and outcome, but air quality in the
incubator should also be addressed.
• CO2 measurement is a poor substitute for actually measuring pH.
Proper quality control (QC) is essential to optimize function of the IVF lab and to maximize reproductive
outcomes. Although purchasing of high-quality materials and equipment can aid improving culture conditions,
proper oversight and implementation are crucial. Despite having the same equipment and media in place, two
laboratories can have dramatically different outcomes because several variables within the culture system are
controlled directly within the individual lab. Examples include pH, osmolality, air quality, and incubator work-
flow. However, another variable that needs to be examined includes variation between technicians to ensure
consistency in lab practices and approaches. This applies not only to procedures, such as embryo grading, intra-
cytoplasmic sperm injection (ICSI), or catheter loading but also to how daily QC readings are taken, how QC
assays are performed, and how media and dishes are prepared. Subtle deviations may introduce an environmen-
tal stressor into the culture system or miss a slight variation that could affect outcome.
TABLE 3.1
Recommended Monitoring and Filter Exchange Schedules to Maintain or Improve IVF Lab
Air Quality
Equipment Recommended Maintenance
Heating, ventilating, and air-conditioning (HVAC)/air handler Confirm daily
(positive pressure, if present)
HVAC/air handler (HEPA filters, charcoal/permanganate filters) Replace 3–6 months or as needed
Incubator HEPA filters Replace 6–12 months
Incubator/gas line VOC filters Replace 3–6 months
Laminar flow hood prefilters Replace or clean annually or as needed
Laminar flow hood (flow rate inspection) Inspect annually
Lab air quality assessment (particle counts, VOC levels) Inspect annually
Note: Lab air exchange rates may be modified to improve certain variables, such as particle counts.
Additional maintenance may be required to maintain air quality if other systems are in place
(i.e., UV photocatalytic oxidation).
Chapter 4: Morphological Selection of Gametes and Embryos: Sperm

Pierre Vanderzwalmen, Magnus Bach, Olivier Gaspard, Bernard Lejeune,
Anton Neyer, Françoise Puissant, Maximilian Schuff, Astrid Stecher,
Sabine Vanderzwalmen, Barbara Wirleitner, and Nicolas H. Zech
Optimized sperm selection techniques are important in the era of ICSI because ICSI does bypass the natural
barriers of conception. The introduction of intracytoplasmic morphologically selected sperm injection (IMSI)
made embryologists aware that in times of ICSI, the selection of sperm has to be given proper attention.
The type of spermatozoa selected for injection influences the outcome in terms of embryo development,
pregnancy, miscarriage, and malformation.
Large vacuoles reflect a pathological situation, probably correlated with sperm chromatin immaturity.
Practical and technical aspects of sperm selection need to be considered to facilitate the workflow while per-
forming IMSI. In particular, a proper setup of the dish for selecting and capturing sperm facilitates the routine
application of IMSI.
The application of IMSI leads to more and better quality blastocysts, and as a consequence, it increases the
chance to select the proper embryo for transfer with the highest implantation potential.
A lot of questions are still pending and unclear:
(1) The terminology of vacuoles, their classification, and their location on the sperm head and their origin
and formation
(2) The application of motile-sperm organelle-morphology examination (MSOME)–IMSI for specific
indications such as teratozoospermia or to a large population
(3) The application of IMSI instead of IVF in cases of unexplained infertility
(4) The oocyte repairing factors
D
D
E E
A A A E E A A A E
C E
C E E
C C E
C
B B B C E
C
B B B
FIGURE 4.9 Position of the drops in an IMSI dish (A, D) 7.5%–10% PVP. (B, C, E) Culture medium. (A) Sperm selection
PVP drops. (B) Sperm aliquots drops. (C) Host-selected spermatozoa microdrops. (D) Sperm immobilization drops. (E) Oocytes
injection drops.
IMSI IMSI
(a) (b)
IMSI
Hoffman
microscope
microscope
(c)
(d)
FIGURE 4.10 IMSI. (a–c) Description of the different steps for sperm selection on the IMSI microscope followed by (d) oocyte
injection on the Hoffman ICSI microscope.
Chapter 5: Morphological Selection of Gametes and Embryos: Oocyte

• Check the maturity stage of the oocyte after cumulus-corona removal.

• If the oocyte is at germinal vesicle (GV) stage, and if the patient does not have any other available
mature oocytes, GV stage oocytes can be in vitro cultured for up to 24 hr. If the oocyte is in vitro
matured, it could be used for microinjection procedure. However, the patient should be informed that
the viability and the implantation potential of embryos derived from such oocytes are significantly
lower than those of embryos derived from oocytes that are at metaphase II (MII) stage after oocyte
collection. It is recommended to use only morphologically normal MII oocytes that are obtained after
in vitro maturation of GV stage oocytes.
• If the oocyte is at metaphase I (MI) stage, it could be in vitro cultured and if MII stage is reached it
could be used for microinjection. The patient should be informed that MI oocytes at the time of oocyte
retrieval can have similar viability as MII oocytes if they can be in vitro matured for up to 6 hr post-
collection. Same morphological recommendations for MII oocytes can be used for in vitro matured
MI oocytes.
• If the oocyte is at MII stage, morphological deviations that should be examined with high priority are
in the following order:
• Oocytes that are large in size (giant oocytes), and oocytes that have a large first polar body should
not be used for insemination because of the high risk of chromosomal abnormalities. If the
patient has only such oocytes, preimplantation genetic screening for the derived embryo can be
recommended.
• Oocytes should be observed for the presence of smooth endoplasmic reticulum cluster(s) within
the cytoplasm. The patient should be informed that embryos derived from such oocytes may have
significantly reduced rates of healthy offspring.
• Oocytes should be observed for the presence of vacuole(s) within the cytoplasm. Patients should
be informed that MII oocytes with vacuole(s) ≥14 μm have a significantly lower chance of getting
fertilized compared with oocytes with normal morphological appearance.
• Oocytes should be observed for the presence of organelle clustering/centrally located condensed
granulation within the cytoplasm. The patient should be informed that embryos derived from such
oocytes may have a higher risk of chromosomal abnormalities.
• Oocytes defined with other cytoplasmic deviations such as refractile bodies/cytoplasmic inclu-
sions or with dark cytoplasm/dark cytoplasm–granular cytoplasm/dark cytoplasm with slight
granulation/dark granular appearance of the cytoplasm/diffused cytoplasmic granularity should
be documented.
• Ovoid oocytes with ovoid zona and normally shaped oolemma, or ovoid zona and ovoid oolemma,
should be observed because the blastocyst formation rate of embryos derived from such oocytes
may be detrimentally affected and delayed.
• Oocytes with extremely large perivitelline space (PVS) may result with reduced fertilization rates
and higher degeneration rates after ICSI.
• Dysmorphic zona pellucida, discoloration of the oocyte, first polar body morphology, and debris
in PVS should be documented.
FIGURE 5.4 Oocyte with central refractile body and FIGURE 5.5 Vacuolized MII oocyte.
granule in the perivitelline space.
FIGURE 5.6 Oocyte with an aggregation of the smooth FIGURE 5.14 Metaphase II oocyte with large perivitelline
endoplasmic reticulum. space.
FIGURE 5.16 Oocyte with large first polar body. FIGURE 5.17 Diploid giant MII oocyte showing two
first polar bodies.
Chapter 6: Morphological Selection of Gametes and Embryos: 2PN/Zygote

• A regular fertilized oocyte shows in general two pronuclei.

• Pronuclear stage oocytes with three pronuclei or more after IVF or ICSI should be discarded and not
be used for transfer or cryopreservation.
• Pronuclear stage with two pronuclei that are not of the same size or are not aligned in the center of the
oocyte show a reduced development potential.
• Pronuclear stages with one pronuclei after ICSI should be discarded.
• Pronuclear stages with one pronuclei after IVF: if there are no other cells and the patient are informed
about the situation it is possible to transfer them; however, there should be no cryopreservation.
• Criteria for the selection of the pronuclear stages as a function of the distribution of the nucleoli in the
pronuclei.
• Always remember the development is time dependent and the correct timing is important.
It is not recommended to transfer or cryopreserve pronuclear stage oocytes with score Z4 or Rang 3.
(a) (b) (c) (d)
Rang 1/Z1-Score Rang 1/Z2-Score
Rang 2/Z3-Score Z4-Score
Rang 3 (ghost) Rang 3 (bull eye)

Chapter 7: Morphological Selection of Gametes and Embryos: Embryo

Day 1 At 26 ± 1 hr postinjection or 28 ± 1 hr postinsemination, the embryo should have completed the first
mitotic division and resulted in a two-cell embryo with blastomeres of even size and have a single
nucleus evident in each of the blastomeres. Embryos showing more than one nucleus per blastomere
should be preferentially excluded from selection for transfer on subsequent days of development. At
this stage, very little fragmentation is usually evident, and fragments observed at this time may be
reabsorbed into the embryo, resulting in a different appearance on Day 2 of development.
Day 2 At 44 ± 1 hr postinsemination, the embryo should have completed the second round of cleavage,
resulting in a four-cell embryo with blastomeres of even size and have a single nucleus evident in each
of the blastomeres. Again, any embryos showing more than one nucleus per blastomere should be pref-
erentially excluded from selection for transfer on this or subsequent days of development. Four-cell
embryos showing ≥25% fragmentation should also be excluded from selection for transfer. Four-cell
embryos showing more than one-third difference in cell size should be preferentially excluded from
transfer on this and subsequent days of development. Embryos showing very rapid cleavage (more than
five cells) or very slow cleavage (fewer than three cells) should be preferentially excluded from selec-
tion for transfer on this or subsequent days of development because rapid cleavage and slow cleavage
have been linked to an increase in chromosomal aneuploidy. Three-cell or five-cell embryos at 44 ±
1 hr postinsemination showing stage-specific cell sizes may be suitable alternatives when no four-cell
embryos are available for transfer, provided multinucleation has never been observed during develop-
ment and fragmentation is minimal.
Day 3 At 68 ± 1 hr postinsemination, the embryo should have completed the third round of cleavage, result-
ing in an eight-cell embryo with blastomeres of even size. Eight-cell embryos showing ≥25% fragmen-
tation or having a difference in size of more than one-third in any blastomere should be preferentially
excluded from selection for transfer. Seven-cell embryos and nine-cell embryos at 68 ± 1 hr postin-
semination, showing stage-specific cell sizes, may be suitable alternatives when no eight-cell embryos
are available for transfer, provided multinucleation has never been observed during development and
fragmentation is minimal. Very rapidly cleaving embryos and very slow cleaving embryos should be
excluded from selection for transfer because these embryos are more likely to be chromosomally aneu-
ploid. If an embryo is showing early signs of compaction, this is not detrimental to outcome.
Day 4 At 92 ± 1 hr postinsemination, the embryo should have entered the fourth round of cleavage, that is, be
more than eight cells, and should be partially or preferably completely compacted. Embryos that have
more than eight cells but that are showing no signs of compaction at 92 ± 1 hr postinsemination should
be preferentially excluded from selection for transfer if a more ideal embryo is available. Embryos that
have not entered the fourth round of cleavage should be excluded from transfer unless a more suitable
embryo is unavailable for transfer. Embryos that are fully compacted and showing early signs of cavi-
tation should be preferentially selected for transfer.
The criteria for each of the major morphological selection paramters are indicated in the heirarchical order for
selection. The ideal embryo at each time of assessment is indicated in redand exclusion criteria are indicated in blue.
Hierarchy for Selection

Time of
Observation Cell Number Cell Size Fragmentation Multinucleation Compaction
26±1 hr • 2 cells • Stage-specific • <10% • Single nucleus/ • Cavitating
postinjection or • Syngamy • Not • 10%–25% blastomere
28±1 hr • 2PN or >2 cells stage-specific • >25% • No nucleus
postinsemination observed
• >1 nucleus/
blastomere
44±1 hr • 4 cells • Stage-specific • <10% • Single nucleus/ • Completely
postinsemination • 3 cells or 5 cells • Not • 10%–25% blastomere compacted
• <3 cells or >5 cells stage-specific • >25% • No nucleus
observed
• >1 nucleus/
blastomere
68±1 hr • 8 cells • Stage-specific • <10% • Multinucleation • Partially
postinsemination • 7 cells or 9 cells • Not • 10%–25% observed in compacted
• 6 cells or 10 cells stage-specific • >25% earlier
• <6 cells or >10 cells development
92±1 hr • >8 cells • Multinucleation • No
postinsemination • < 8 cells observed in compaction
earlier
development
FIGURE 7.1 Examples of embryos showing normal cleavage kinetics and normal cytokinesis on Day 1 (a), Day 2 (b), Day 3 (c),
and Day 4 (d). Original magnification, 300×.
Chapter 8: Morphological Selection of Gametes and Embryos: Blastocyst

Thomas Ebner
• If a clinical embryologist plans to change in vitro culture from a cleavage stage to a blastocyst transfer
program, it is recommended to start working with supernumerary embryos to check whether culture
conditions actually work properly. Thus, good prognosis patients with sufficient embryos in abundance
should preferably be considered for training purposes.
• At least 40%–50% blastocyst formation rate should be used as KPI.
• In case blastocysts are available, an adequate scoring system is an absolute prerequisite.
• It is of utmost importance to distinguish between trophoblastic vesicles (“pseudoblastocysts”) and
real blastocysts. Preceding cleavage behavior might give a clue.
• The applied scoring scheme should at least deal with expansion as well as the quality of the inner
cell mass and the trophectoderm.
• There is growing evidence that in terms of live birth rate the quality of the trophectoderm out-
weighs the quality of the inner cell mass.
• Additional morphological parameters should be noted, for example, presence of apoptotic areas,
vacuoles, cytoplasmic strings, and exclusion of fragments and/or blastomeres.
• Spontaneous hatching is a positive predictor of implantation. Usually no artificial hatching is applied
at blastocyst stage.
• Prolonged culture to Day 5 might be associated with the phenomenon of monozygotic twinning.
• An increase in birth weight has been observed in blastocyst transfers.
FIGURE 8.2 Full IVF blastocyst with optimal cell

lineages (3AA). FIGURE 8.5 Suboptimal full blastocyst (3BC) devel-
oped after conventional IVF. Trophectoderm is not
cohesive between 6 o’clock and 2 o’clock position.
Inner cell mass consists of few cells only.
FIGURE 8.12 Expanded IVF blastocyst (4AA) with

FIGURE 8.10 Expanded blastocyst of poor quality necrotic area in trophectoderm (8 o’clock position).
(4CB). Blastomere extruded into blastocoel should not Two cytoplasmic processes bridge the blastocoel.
be mixed up with inner cell mass.
FIGURE 8.14 Expanded blastocyst of ovoid shape

(4AB). FIGURE 8.15 Pseudoblastocyst presumably consist-
ing of <12 cells.
Chapter 9: Noninvasive Techniques: Gamete Selection—Sperm

The ultimate aim of any sperm selection method is to provide the best quality sperm possible so as to maximize
the outcome of whatever ART procedures are to be undertaken. For artificial insemination and IVF, the main
requirement is the provision of a sample enriched with progressive motile sperm because these techniques are
dependent upon the ability of sperm to find and penetrate the oocyte by themselves.
Numerous techniques have been developed to select the best possible sperm for ART. Although many of the
described methods may be considered novelties and lack the robust verification and validation essential for rou-
tine clinical use, they nonetheless constitute a great improvement on the existing selection process that is solely
dependent upon the subjective choice of the embryologists. The main disadvantage, however, is that although
most of the techniques provide information upon which sperm quality can be better classified, the practical
aspects of the procedures either destroy or alter sperm in such a way that renders them unusable for ART. Be that
as it may, until the full potential of upcoming techniques based on advanced technology has been realized, the
procedures described in this chapter, regardless of their limitations, represent the best options presently available
to the ART clinic.
TABLE 9.1
Overview of Different Technologies Used for Sperm Selection
Technique Emerging Used Diagnostic Therapeutic Studies
Polarization microscopy X X X 2–4
MACS X X X 9, 30
FACS X X 13
PICSI X X X 31
HA medium X X X 32
Zona pellucida binding X X X 15, 33, 34
Zeta potential X X X 35
Electrophoresis X X X 21, 21, 23
Raman microspectroscopy X X 26–29
Note: HA, hyaluronic acid.
FIGURE 9.2 Raman spectroscopy for sperm. (a) Averaged Raman spectra from sperm (blue) and after ultraviolet (UV) i rradiation
(red). Arrow indicates main spectral feature in damaged sperm (peak at 1047 cm–1). Spectra have been normalized, baseline-
corrected, and smoothed. (b) Single-point Raman spectra are acquired from the post–acrosomal region of the sperm head. The red
dot represents the position on which the laser is focused.
Chapter 10: Noninvasive Techniques: Gamete Selection—Oocyte

Early selection of oocytes is considered one major goal of contemporary IVF worldwide, allowing the identifica-
tion of the most competent gametes to inseminate. In turn, this would help reduce the number of embryos pro-
duced in vitro and progress to single embryo transfer. Unfortunately, standard morphological evaluation is not
precise, and a consensus is lacking. However, new noninvasive tools for oocyte selection are gaining increasing
interest from the scientific community, from the more classic polarized light microscopy analysis to the evalu-
ation of PFV and the ground-breaking OMICS technology. The results obtained so far are really intriguing and
encouraging, but it would be wise to rise some concerns. For polarized light microscopy analysis, the contradic-
tory results underline the need of more intense study to reach a consensus. For OMICS and, in part, PFV evalu-
ation, the high costs, the difficulty of the techniques, and the time required for testing are limiting the routine
applicability. Moreover, even if these approaches are all considered “noninvasive,” we still need more evidence
about the safety of the techniques (i.e., the possible effect of additional time required for each oocyte outside
the incubator or unindicated removal of CCs and, as a consequence, ICSI performance). Finally, prospective
randomized studies are required to determine their predictive power, alone or in combination with other factors,
so that further efforts enrich our current knowledge.
FIGURE 10.5 MII oocyte observed at polarized light microscopy (400× magnification). The birefringent MS is visible just below
the extruded IPB at about the 6 o’clock position.
FIGURE 10.6 Telophase I oocyte observed at polarized light microscopy (400× magnification). The MS is visible at the 6 o’clock
position. It is interposed between the extruded IPB and the ooplasm, indicating that the first meiotic division is not yet concluded.
FIGURE 10.7 MII oocyte observed at polarized light microscopy (400× magnification). The MS is visible at the 9 o’clock position,
and it is clearly dislocated about 90° from the IPB (placed at 6 o’clock position).
Chapter 11: Noninvasive Techniques: Embryo Selection by Oxygen Respiration

• Because the embryo development is directly related to a gamete’s health, oxygen uptake could be a
noninvasive marker of oocyte and embryo quality.
• Oocyte OC is affected by different ovarian stimulation regimens, and it increases 10% when we obtain
a successful fertilization.
• OC levels vary depending on the embryonic stage.
• Best quality embryos show higher OC levels during embryo development, displaying a correlation
between quality and OC.
• Higher levels of OC correlate with embryos that achieve implantation and give rise to ongoing
pregnancy.
• There is a peak of OC during the first division, corresponding to a high energy demand period that is
significantly higher in implanted embryos and defines cytokinesis as the best time period to consider
OC as a marker of embryo quality.
Oxygen Oxygen
microsensor concentration
Depth
∆X
∆C
Glass well ∆C
Respiration = –D*A* ∆X
FIGURE 11.1 Schematic overview of incubator with a microsensor that can be used to measure the oxygen consumption through
the gradient created by oocyte uptake, as explained in the text.
Embryo O2 uptake (transferred embryos)

Not implanted
6.5 *
6.27 * 100 implanted
6.0 5.84 *
5.75 * 5.75
5.5 5.55
5.38
5.0 4.98
4.86
4.5
4.0
≤17.2 17.2–35.0 35.0–52.1 >52.2
p < .001
FIGURE 11.7 OC averages from transferred embryos in each of the four time ranges depending on implantation success.
(*) indicates a significant difference (p < .05) between implanted embryos and nonimplanted embryos, with bigger differences from
52.1 hr post-ICSI.
Chapter 12: Noninvasive Techniques: Embryo Selection by Time-Lapse Imaging

Alison Campbell
Implementation and Routine Use of Time-Lapse Technology in IVF

Selection of a Time-Lapse System or Device
Before introducing a time-lapse system, the following should be considered to justify the decision: evidence base
for positive impact on clinical outcome; opportunities for development and continuous improvement; device
specification, for example, focal planes, image quality, and capacity; user friendliness; degree of validation
conducted by the supplier and required in-house; certification and licensing (where required), limitations; space
requirement; cost; customer support; servicing; and access to training.
Installation
Before installation, a detailed plan must be devised and approved that ensures that all requirements for installation of
the equipment have been considered and that the laboratory and staff are prepared. Depending on the device selected,
requirements may include gas supply, uninterrupted power supply (UPS), incubator capacity and accessibility, and
space for additional hardware. A validation plan should be prepared to outline the process of qualifying the instal-
lation, operation, and performance of the time-lapse device before the introduction of time lapse for clinical use.
Practical Aspects
Standard operating procedure should be defined before the introduction of the time-lapse system for clinical use
to which all users should acknowledge and adhere. As with any new technology, robust and rigorous training and
quality assurance processes are required.
Dish or slide preparation methodologies must be established and practiced. The operating procedure should
detail these methodologies, ensuring that the media and consumables to be used are appropriate and standardized.
Annotation guidelines, where required for nonautomated systems, must be defined strictly because it will be
the accuracy of these guidelines downstream that could limit or support the development of embryo selection
algorithms, based on the clinic’s own data. This is crucial and buy-in of staff is essential. Users are encouraged
to consider with care which morphokinetic variables, already defined or novel, should be routinely recorded and
whether nonrecording assumes that a phenomenon was not seen. Table 12.1, within the body of the chapter, may
assist with this task, and users are encouraged to follow consensus guidelines where they are available.
Time lapse allows great flexibility within the IVF lab. Monitoring of embryo development, traditionally per-
formed at fixed times daily, can now be performed to suit the laboratory workflow. When manual annotation is
required, it is advised that this annotation is performed regularly by a review of the images accumulated since
the previous session of annotation. For flexible and efficient practice, annotation should be performed daily
rather than at the end of the extended culture period, but users may find their own way of incorporating this
annotation according to the workflow through their laboratories and find that it is dependent on the amount of
morphokinetic variables to be recorded.
Quality Assurance and Review

Although the direct use of published algorithms for embryo selection may be desirable to new users, this practice
may not be effective in varied settings, and as such, it is not recommended. The use of “known implantation
data” from within the clinical setting is arguably the most robust method for developing embryo selection algo-
rithms for in-house application. Regular review of time-lapse scientific literature is recommended, and commu-
nication with other time-lapse users may be beneficial, particularly during the early stages of implementation.
Close monitoring of clinical results is necessary to ensure that the new technology is working well, and regular
quality assurance activity will ensure that users are working in the same way.
TABLE 12.1
Summary of Morphokinetic Variables and Proposed Definitions
Description
Morphokinetic Variables
Time (t)
t0 IVF or midtime of ICSI/IMSI
tPB2 The second polar body is completely detached from the ooplasm
tnPN Fertilization status is confirmed
(tPN1a) The first pronucleus is first visible
(tPN2a) The second pronucleus is first visible
tPNf All pronuclei have faded (see Figure 12.1)
t2-t9 Two (see Figure 12.2) to nine sequential, distinguished cells are present
tSC The first two cells merge; initiation of compaction observed (see Figure 12.3)
tMx/w Morula is formed or compaction goes no further; “x” corresponds to fully compacted, and “w” corresponds
to partially compacted or cells excluded
tSB The first sign of a cavity is observed as blastulation begins (see Figure 12.4)
tByz Full blastocyst stage is reached; the last frame before the zona pellucida starts to thin; “y” corresponds to
morphology of inner cell mass cells, and “z” corresponds to trophectoderm cells (see Figure 12.4)
tEyz Initiation of expansion is confirmed; the zona pellucida starts to thin
tHNyz Extrusion of cells from the zona pellucida is present
tHDyz Blastocyst is fully hatched from the zona pellucida
Calculated Variables
VP tPNf-tPN1a (period of visible pronuclei)
Cell Cycle
CC1 t2-tPB2
The end of the second meiosis to the formation of two discrete cells
CC2 The time for a two-cell embryo to form a four-cell embryo
The two blastomeres (a and b) can be considered individually
CC2a = t3-t2
CC2b = t4-t2
CC3 The time for a four-cell embryo to form an eight-cell embryo
The four blastomeres can be considered individually.
CC3a = t5-t4
CC3b = t6-t4
CC3c = t7-t4
CC3d = t8-t4
Synchronization
S2 The duration of the transition from two sister cells, each dividing to reach the four-cell stage
t4-t3
S3 As above, but from four to eight cells
t8-t5
Duration of Compaction (Morula Stage)

tMx-tSC Full compaction
tMy-tSC Partial compaction
Blastocyst Stage
tHN-tSB Duration of blastulation
Note: Each time point defines the time-lapse frame in which the phenomena described are first observed or recorded.
Chapter 13: Noninvasive Techniques: Embryo Selection by Transcriptomics,

Proteomics, and Metabolomics
Current embryo assessment strategies that rely primarily on embryo morphology and cleavage rate do not
provide adequate sensitivity or specificity to achieve desired pregnancy rates in women undergoing infertility
treatment with IVF. Studies using emerging technologies to analyze cumulus/granulosa cell transcriptome or
spent embryo culture media protein or metabolite content report promising results. However, the transcripts or
proteins that have been identified as potential biomarkers of embryo viability or the metabolomic profiles associ-
ated with pregnancy outcome have not yet been adequately validated. Therefore, the use of these novel nonin-
vasive technologies remains experimental, and their application to clinical practice awaits randomized clinical
trials demonstrating benefit from their use, alone or in combination with morphologic evaluation.
Key Messages
Omics research basically looks for answers to two questions: what distinct molecules and how many copies of
each molecule exist in a biological sample?
Gene expression research evolved from single-gene analysis to genome-wide transcription profiling with the
advance of microarray technology; however, next generation sequencing technologies provide an alternative
transcriptomics approach called RNA Sequencing that appears to be a more precise detection and quantification
of RNA transcripts.
Proteomics aims at (1) profiling proteome content of a biological sample, (2) performing comparative protein
expression analysis, (3) localizing and identifying of posttranslational modifications, and (4) exploring protein–
protein interactions.
Metabolome is the analysis of metabolites that are the final downstream products of gene expression, and
in preimplantation embryo development, a distinct change in the levels of metabolites is expected to classify a
“normal” development and a viable embryo.
DNA RNA Protein Metabolites
Transcription Translation Metabolism
~3×109 bp nucleotides mRNA, tRNA, rRNA, miRNA Small molecules (<1 kDa)
~1,000,000 proteins
~30,000 genes ~200,000 transcipts ~3000 metabolites
Genomics Transcriptomics Proteomics Metabolomics
FIGURE 13.1 Omics approaches investigate the molecular constitution of biological samples at genomics, transcriptomics,
proteomics, and metabolomics levels.
Cumulus/granulosa Isolated Reverse transcribed, labeled, Hybridization to Image scanning and Differential
cells RNA and amplified cDNA microarray chip quality control expression analysis
FIGURE 13.2 Major steps of a typical microarray experiment. Total RNA is extracted from the follicular cell samples, and mRNA
is reverse transcribed into cDNA. Amplified and labeled samples are hybridized to a microarray slide, allowing the labeled targets
to bind to their complementary oligonucleotides attached to the microscopic probes. The array is then washed and scanned to obtain
the fluorescent image that is further processed to get the intensity values for differential expression analysis.
Chapter 14: Invasive Techniques: Polar Body Biopsy

• Polar body diagnosis gives direct information about the first and second PB and therefore allows only
an indirect diagnosis of the maternal genetic or chromosomal constitution of the corresponding oocyte.
• Polar body biopsy can be done by mechanical means using micropipettes for zona drilling or by using
a laser for contact-free opening of the zona pellucida.
• The easiest and safest way of opening the zona pellucida is by laser.
• Although laser application seems easy, certain dead ends need to be avoided:
• Openings in the zona pellucida should be large enough to enable undisturbed hatching at later
stages.
• Too small openings may trap the blastocyst during hatching.
• Zona opening must be done at one site only: two openings will lead to blastocyst splitting or degen-
eration at the time of hatching.
• Timing of polar bodies is crucial for sequential as well as simultaneous biopsy.
• Too early biopsy in the presence of spindle remnants may lead to enucleation.
• Too late biopsy bears the risk of degeneration of PB1 and compromised quality of amplification (array-
CGH) or signals (FISH).
• Transfer of polar bodies for subsequent analysis onto slides or into tubes is one of the most crucial steps
of the whole procedure.
• For FISH, always release polar bodies in the droplet on the slide at or close to the bottom to avoid
floating and rupture.
• For array-CGH, always release polar bodies in the fluid droplet on the bottom of the tube.
• Openings in the zona pellucida require precautions during all subsequent steps that involve pipetting
or handling of the embryos.
• After zona opening, embryos do hatch earlier.
• Embryo transfer should be done smoothly due to the presence of the zona opening.
Summary
PB biopsy has been proven as sufficiently effective for the diagnosis of structural and numerical chromosome
aberrations in human oocytes using FISH and array-CGH. Nevertheless, the use of PB biopsy and array-CGH for
PGS is still a matter of debate due to cost effectiveness, the high incidence of postmeiotic aneuploidies that are
undetectable by the polar body approach, and the debate on the accurate diagnostic procedure. There is growing
advocacy for trophectoderm biopsy and array-CGH as the new gold standard.
FIGURE 14.2 Laser-assisted polar body (PB) biopsy. For

biopsy, the first and second PBs were aligned with a holding FIGURE 14.3 Presence of a meiotic spindle bridge between
capillary so that the second PB faced to the biopsy capillary (a). first polar body (PB1) and the oocytes. The presence of a con-
Using a noncontact 1.48 μm diode laser, an opening was intro- nective spindle bridge can be assessed by polarization micros-
duced into the zona pellucida using two or three laser shots (b) copy. Spindle fibers are displayed in red and characterize the
through which the biopsy capillary could be easily introduced corresponding oocyte as being in the transition phase from
(c). The second PB is usually connected to the oolemma via a metaphase I to metaphase II. As long as spindle fibers are vis-
cytoplasmic strand. To remove the second PB without damaging ible, chromosomes in the oocyte are attached, and the corre-
the oocyte, it is not recommended to suck the second PB into the sponding oocyte is in anaphase/telophase I. Removal of the PB1
capillary; instead, the capillary is pushed slowly over the second shown would result in the withdrawal of chromatin material
PB and toward the first PB (d). Once the first PB enters the cap- from the oocyte and could potentially lead to enucleation.
illary, the strand between the second PB and the oolemma will
break due to shear stress and both PBs can be easily removed (d),
leaving the oocyte without any damage. PBs should be placed in
one droplet for further processing for FISH analysis (e) or in two
different droplets if a PCR-based analysis will be performed.
FIGURE 14.7 Transfer of isolated polar bodies (PBs) onto a FIGURE 14.8 Identification of the polar body (PB) on the
slide. The transfer of isolated PBs from the dish (seen in the back- slide. This photograph is taken with a 10× phase contrast objec-
ground) into the droplet on the slide must be performed on the tive, and the diamond circle surrounding the PB can be partially
microscope stage. The setup shown here allows sliding the dish seen. The PB appears gray (arrow). (Reprinted with permission
used for biopsy backwards. Therefore, the aspiration capillary from Montag M, Textbook of Assisted Reproductive Techniques
needs only to be lowered into the droplet for release of the PB. Volume One: Laboratory Perspectives, 4th ed., Informa
(Reprinted with permission from Montag M, Textbook of Assisted Healthcare, New York, 2012, pp. 336–345.)
Reproductive Techniques Volume One: Laboratory Perspectives,
4th ed., Informa Healthcare, New York, 2012, pp. 336–345.)
Chapter 15: Invasive Techniques: Embryo Biopsy at the Cleavage Stage

Anick De Vos
• Cleavage-stage embryo biopsy still remains the most widely practiced method for embryo biopsy.
• Cleavage-stage embryo biopsy offers the advantage over polar body biopsy that the paternal contribution
to the genetic content of the embryo can be analyzed.
• The use of ICSI avoids paternal contamination from sperm attached to the zona pellucida, whereas a
careful oocyte denudation should avoid maternal contamination resulting from cumulus cells.
• One or two blastomeres are removed from the embryo in the morning of Day 3, at about 68–72 hr after
microinjection, when the embryos are preferably in the eight-cell stage.
• It is common practice to include only embryos resulting from 2-pronuclear (2PN) normal fertilization.
Embryos with <50% of anuclear fragmentation and with at least six blastomeres are suitable for biopsy.
Preferably, the embryos have stage-specific cell sizes.
• Laser zona opening is most often used, although chemical zona opening using acidic Tyrode’s solution
or mechanical zona opening are still in use too.
• Aspiration is most widely used to remove the blastomeres.
• Ca2+/Mg2+-free biopsy medium can be used to facilitate the biopsy procedure by reducing the existing
junctions between blastomeres.
• Care is taken to remove mononucleate intact blastomeres.
• Lysed cells are not used for PCR analysis because contamination with maternal DNA cannot be
excluded. Occasionally, the nucleus can be recovered for FISH analysis. In cases of cell lysis, it is
advised that the aspiration pipette is changed before continuing biopsy of other embryos as a safety
measure to avoid cross-contamination.
• Postbiopsy, single embryo culture in individual droplets or individual culture dish wells is adopted to
ensure tracking of blastomeres removed and easy identification of embryos postdiagnosis.
• Further embryo development should not be impaired as a result of the biopsy procedure.
• Supernumerary biopsied and diagnosed embryos can be effectively vitrified for later use.
• The invasive nature of the biopsy procedure on Day 3 should be recognized, and evidence exists that
one-cell biopsy is less invasive than two-cell biopsy. Based on the observation and direct proof that
blastocyst trophectoderm biopsy safeguards the developmental competence of the embryo even better,
a shift toward blastocyst trophectoderm biopsy might be anticipated in the near future. As such, chro-
mosomal mosaicism at the cleavage stage could be alleviated as well.
FIGURE 15.1 Cleavage-stage embryo biopsy is usually performed in the morning of Day 3. (a) Ideally, the embryos are in the
eight-cell stage, having no fragmentation and showing stage-specific blastomere sizes. (b) An advanced-stage embryo on Day 3 with
no fragmentation. (c) An embryo with more fragmentation (between 20% and 50%); however, it would still be included for embryo
biopsy.
FIGURE 15.2 (a) For the removal of human blastomeres by aspiration, an aspiration pipette is introduced into the perivitelline
space through the hole in the zona pellucida to reach a blastomere. (b, c) One blastomere is removed by gentle aspiration. Cells may
be aspirated completely and then removed; alternatively, cells are only partially aspirated and pulled out (given full decompaction
and thus no adherence to other blastomeres).
FIGURE 15.5 To be suitable for genetic analysis either by FISH or by PCR, the removed blastomere(s) should contain one single,
clearly visible nucleus.
Chapter 16: Invasive Techniques: Blastocyst Biopsy

Steve McArthur
Blastocyst biopsy is preceded by embryo hatching on either Day 3 or on Day 5 or Day 6 of embryo development.
Hatching on Day 3 of development ensures accessibility to a suitable hatching site with little risk of damaging
any blastomeres.
Embryo Hatching
1. The hole in the zona pellucida should be made adjacent to the polar body, if identifiable; otherwise,
hatch at a position where there is a space between the blastomeres.
2. The first pulse of the laser should be directed to the outer edge of the zona pellucida.
3. The third pulse of the laser should cut the internal wall of the zona pellucida in a direct line from the
first laser pulse.
4. A hole in the internal margin of the zona of approximately 10 μm is created.
5. Embryos should then be washed before being placed back into culture dish.
Trophectoderm Biopsy
1. Ensure that the biopsy pipette tip and the leading edge of the trophectoderm are in focus.
2. Bring the biopsy pipette and touch the leading edge of the trophectoderm.
3. Draw a small number of trophectoderm cells carefully into the pipette using suction until a good hold
is achieved.
4. Stretch out the trophectoderm (at this point, the blastocoel cavity may collapse).
5. Adjust focus until the margins of the targeted cells are in focus and fire the laser at the intercellular
junctions. Fire the laser three to five times across the width of the trophectoderm cells to be biopsied.
6. Using suction on the mouthpiece and the holding pipette joystick, pull the targeted cells away from the
blastocyst.
7. Release the piece of trophectoderm from the biopsy pipette.
8. Where the stretched trophectoderm tissue does not readily separate from the remaining embryo, it is
not recommended to fire the laser excessively because this may be detrimental to embryo recovery. In
these cases, a more direct “cutting off” method may be used.
9. Gently release the embryo from the holding pipette while maintaining hold of cells in your biopsy
pipette. Do not take further cells into your pipette because this may result in excess cells being biop-
sied. Ensure the holding and biopsy pipettes are in the same focal plane, and rub the biopsy pipette
across the end of the holding pipette. Take care not to damage the embryo protruding from the pipette
as you carry out this step. If the first attempt is not successful, further attempts may be required.
FIGURE 16.3 Photographs of blastocyst biopsy. (a) Blastocyst 2 days after laser-assisted hatching, consisting of herniating troph-
ectoderm (TE) and inner cell mass (ICM). (b) Alignment of TE cells with biopsy pipette. (c) Three to ten cells aspirated from the
TE while the ICM (destined to form the embryo proper) remains intact. (d) Blastocyst and TE sample. (e) Blastocyst approximately
90 min postbiopsy.
FIGURE 16.4 Images of fully hatched blastocyst biopsy. (a) Fully hatched blastocyst 2–3 days post–laser-assisted hatching;
trophectoderm (TE) and inner cell mass (ICM) are labeled. (b) TE cells aspirated into biopsy pipette and contacted with holding
pipette. (c) TE cells cut by shearing biopsy pipette against holding pipette. (d) Blastocyst and TE sample postbiopsy. (e) Blastocyst
approximately 90 min postbiopsy.
Chapter 17: Invasive Techniques: Aneuploidy Testing by FISH

In the FISH technique, DNA probes that are labeled with different-colored fluorescent tags that are specific for
chromosomal regions are hybridized to interphase nuclei or metaphase chromosomes.
Aneuploidy testing by FISH consists of the following steps: fixation of biopsied cells, pretreatment and probe
application, denaturation, hybridization, stringent washing, and analysis after counterstain application.
The most critical step for FISH is the cell fixation; each cell should be informative and the loss of genetic
material should be avoided.
The preferred method for fixation is by the hypotonic method that gives a larger diameter nucleus and decreased
probability of signal overlap and nuclear loss.
Dehydration steps are common for blastomere and trophectoderm tissues and are done before hybridization to
clean the fixed chromatin from the cytoplasm and any fixative artifacts that remain after fixation.
Denaturation prepares for effective hybridization and is mostly done with prelabeled, ready-to-use DNA
probes.
Washing off unspecific bound probes is crucial to enable subsequent signal evaluation. The detection probes
are labeled by fluorophores that may fade under light.
Evaluation of signals is performed in the dark area using a fluorescence microscope equipped with appropriate
filters for the fluorophores used.
FISH can be performed at a relatively low cost, and the procedure is less complex. However, QC guidelines
should be followed for every step from fixation to signal interpretation.
Embryo Blastomere Denaturation
Renaturation and
hybridization
Analysis Probe DNA
FIGURE 17.4 Steps of FISH.
FIGURE 17.5 First and second round results of the same blastomere. (a) Normal for the first round study, including chromosomes
13, 16, 18, 21, and 22. Chromosome 13 (red), chromosome 16 (light blue), chromosome 18 (blue), chromosome 21 (green), and
chromosome 22 (yellow). (b) Abnormal for the second round study, including chromosomes 15, 17, X, and Y. Chromosome 15 (yel-
low), chromosome 17 (aqua), chromosome X (green), and chromosome Y (blue). This blastomere was diagnosed as “monosomy 17.”
Notice that there is a signal overlap between aqua and one of the green signals. In addition, for the yellow signal, the signal at the
bottom-middle is an artifact, evident from the color and the intensity. (Courtesy of Reproductive Genetics Laboratory, Istanbul
Memorial Hospital, Istanbul, Turkey, 2009.)
Chapter 18: Invasive Techniques: Aneuploidy Testing by Array-CGH

Currently, array-CGH is considered the gold standard for detecting aneuploidy in single cells or multicellular
samples from oocytes and embryos because of its reliability, reproducibility, and accuracy. In addition, it enjoys
large worldwide experience (>250,000 clinical samples tested to date) and results can be obtained within 12 hr
after biopsy.
A fundamental principle in applying aneuploidy screening is that the benefit gained should outweigh the harm,
if any, caused. Thus, if the success rates of the program are already suboptimal, it is difficult to see any procedure
providing sufficient benefit to rescue the cycle.
Noncontact laser technology is the primary choice for assisting oocyte and embryo biopsy. Proper preparation
of the biopsied sample is of critical importance to ensure high diagnostic success rates, with a focus on the steril-
ity of the working area and solutions, precise volume of buffer in the microcentrifuge tube, minimal carryover
of embryo culture medium, and subsequent storage of the sample preanalysis.
Good laboratory practice, appropriate apparel, and a dedicated cleanroom, equipment, and consumables for
amplification steps are mandatory.
A negative control of the embryo media and collection buffer should always be taken to check for the absence
of contamination.
Laboratory-specific validation experiments are extremely useful before offering the service clinically.
For whole-genome amplification, a PCR-based method should be preferred to multiple displacement amplifi-
cation (if working with 24Sure).
Care should be taken to ensure microarray slides do not dry out during hybridization and before washing. The
high-temperature, high-stringency wash posthybridization must be temperature controlled.
Drying of the microarray slides is critical after washing, and the most effective way is to mechanically remove
wash buffer by centrifugation.
Biopsied cell(s)
Biopsied cell(s)
DNA
Whole genome amplification
Combined labeled
& control and biopsied
cell(s) DNA
Control DNA Biopsied cell(s)
DNA
A B
Metaphase Array
CGH CGH
1:1 2:1 2:3

Normal Monosomy Trisomy Normal Monosomy Trisomy
FIGURE 18.1 Schematic of comparative genomic hybridization.
1.20
0.88
0.60 Chromatid gain Chromatid gain
0.24
Log2 ratio Ch1/Ch2
0.00
–0.40
–0.72
Chromosome loss
–1.04
–1.36
–1.68
21
10
11
12
13
14
15
16
17
18
19
20
22
1
X
Y
FIGURE 18.2 Determining chromatid versus chromosome loss in first polar body samples by aCGH. For most chromosomes (i.e.,
not the sex chromosomes or the aneuploid chromosomes), a clear and consistent 1:1 ratio is observed along the chromosome length.
Because the polar body sample was cohybridized with male genomic DNA, a hybridization pattern representing a 2:1 ratio for the X
chromosome and a “0:2” ratio for the Y chromosome is observed. This polar body clearly shows multiple aneuploidies with chroma-
tid gains on chromosomes 1 and 10 (single chromatid gains are consistent with a 3:2 [or 1.5:1] ratio, i.e., approximately half that of
the X chromosome shift) and a loss of whole chromosome 15 (similar to the shift seen for the absent Y chromosome).
RepRoductive Medicine
Selecting gameteS
and embryoS
Edited by
Markus Montag, PhD
iLabCoMM GmbH, International Reprolab Consulting
St. Augustin, Germany
Among the many recent advances in assisted reproduction therapies (ART), improved
technologies for identifying viable oocytes, sperm, and embryos are of primary importance.
Paradoxically, the latest advances presented at conferences and symposia are often slow
to become part of the daily routine in IVF laboratories. Detailing established and developing
techniques, A Practical Guide to Selecting Gametes and Embryos provides a user-friendly
text of ready-to-use ARTs that can be utilized effectively in the lab.
In this volume, renowned embryologist and educator Markus Montag and his expert panel
highlight sophisticated and proven selection strategies and emphasize the importance of
proper lab practice in handling gametes and embryos.
topics include
• Steps undertaken for the analysis of a semen sample
• Quality control and prevention of exposure to toxins in oocyte collection
and embryo culture
• Morphological selection of gametes and embryos
• Both commonly used and innovative techniques for gamete and embryo
selection, such as oxygen respiration and time-lapse imaging
• Invasive techniques, including polar body, embryo, and blastocyst biopsies
as well as aneuploidy testing by FISH and array-CGH
Accompanied by numerous figures and descriptions, this guide to selecting gametes and
embryos brings the insight of international authors with knowledge and expertise, highlighting
practical tips and key points. The book offers a starting point for applying successful selection
strategies for reducing the rate of high-risk multiple gestations while maintaining or increasing
viable pregnancy rates.
H100479
ISBN-13: 978-1-84214-547-0
90000
9 781842 145470

Markus Montag - A Practical Guide To Selecting Gametes and Embryos (2014, Taylor and Francis, CRC Press)

Uploaded by

Copyright:

Available Formats

Markus Montag - A Practical Guide To Selecting Gametes and Embryos (2014, Taylor and Francis, CRC Press)

Uploaded by

Document Information

Original Title

Copyright

Available Formats

Share this document

Share or Embed Document

Sharing Options

Did you find this document useful?

Is this content inappropriate?

Copyright:

Available Formats

Markus Montag - A Practical Guide To Selecting Gametes and Embryos (2014, Taylor and Francis, CRC Press)

Uploaded by

Copyright:

Available Formats

a Practical guide to

EditEd by Markus Montag

Boca Raton London New York

CRC Press is an imprint of the

© 2014 by Taylor & Francis Group, LLC

No claim to original U.S. Government works

International Standard Book Number-13: 978-1-84214-548-7 (eBook - PDF)

and the CRC Press Web site at

1. Handling Gametes and Embryos: Sperm Collection and Preparation Techniques�������������������������������1

2. Handling Gametes and Embryos: Oocyte Collection and Embryo Culture����������������������������������������� 17

3. Handling Gametes and Embryos: Quality Control for Culture Conditions����������������������������������������� 39

4. Morphological Selection of Gametes and Embryos: Sperm������������������������������������������������������������������� 59

5. Morphological Selection of Gametes and Embryos: Oocyte������������������������������������������������������������������ 81

6. Morphological Selection of Gametes and Embryos: 2PN/Zygote����������������������������������������������������������97

7. Morphological Selection of Gametes and Embryos: Embryo��������������������������������������������������������������� 115

8. Morphological Selection of Gametes and Embryos: Blastocyst����������������������������������������������������������� 131

9. Noninvasive Techniques: Gamete Selection—Sperm���������������������������������������������������������������������������� 143

10. Noninvasive Techniques: Gamete Selection—Oocyte��������������������������������������������������������������������������� 155

11. Noninvasive Techniques: Embryo Selection by Oxygen Respiration��������������������������������������������������� 165

12. Noninvasive Techniques: Embryo Selection by Time-Lapse Imaging������������������������������������������������� 177

13. Noninvasive Techniques: Embryo Selection by Transcriptomics, Proteomics,

15. Invasive Techniques: Embryo Biopsy at the Cleavage Stage���������������������������������������������������������������� 219

17. Invasive Techniques: Aneuploidy Testing by FISH������������������������������������������������������������������������������� 241

18. Invasive Techniques: Aneuploidy Testing by Array-CGH�������������������������������������������������������������������� 255

19. Summary: Comprehensive Summary of Main Points by Topic�����������������������������������������������������������269

Carmela Albert Martin Greuner

Olivier Gaspard Maria Köster

Bernard Lejeune Françoise Puissant

Jana Liebenthron Laura Rienzi

Marcos Meseguer Astrid Stecher

Asli Uyar Barbara Wirleitner

• Volume (or weight)

Analysis of the Native Ejaculate

Low Sperm Numbers

Washing Centrifuge Add medium,

FIGURE 1.2 Sketch of protocol for simple washing.

BOX 1.1 SIMPLE SPERM WASHING

Overlay/ Swim-up Enriched

FIGURE 1.3 Sketch of protocol for direct swim-up.

BOX 1.2 DIRECT SWIM-UP

Swim-Up with an Additional Washing Step

Sperm medium Sperm medium

Dilution/ Centrifuge Swim-up Enriched

FIGURE 1.4 Sketch of protocol for swim-up with dilution/washing.

BOX 1.3 SWIM-UP WITH AN ADDITIONAL WASHING STEP

Discontinuous Density Gradient (DG) Centrifugation

Ejaculate Sperm medium

FIGURE 1.5 Sketch of protocol for density gradient centrifugation.

BOX 1.4 DISCONTINUOUS DG CENTRIFUGATION

FIGURE 1.7 Mechanical maceration by fine needles.

Mechanical Preparation of Testicular Sperm

BOX 1.5 MECHANICAL PREPARATION OF TESTICULAR SPERM

Enzymatic Preparation of Testicular Sperm

FIGURE 1.8 Digested testicular tissue after enzymatic treatment.

BOX 1.6 ENZYMATIC PREPARATION OF TESTICULAR SPERM

Collecting Testicular Spermatozoa for ICSI

BOX 1.7 COLLECTING TESTICULAR SPERMATOZOA FOR ICSI (FIGURE 1.9)

Vitality Tests for Immotile Spermatozoa

Oocytes can be fertilized by immotile sperms if these sperms are viable.

1. Handling Gametes and Embryos: Sperm Collection and Preparation Techniques��1

2. Handling Gametes and Embryos: Oocyte Collection and Embryo Culture�� 17

3. Handling Gametes and Embryos: Quality Control for Culture Conditions�� 39

4. Morphological Selection of Gametes and Embryos: Sperm�� 59

5. Morphological Selection of Gametes and Embryos: Oocyte�� 81

6. Morphological Selection of Gametes and Embryos: 2PN/Zygote��97

7. Morphological Selection of Gametes and Embryos: Embryo�� 115

8. Morphological Selection of Gametes and Embryos: Blastocyst�� 131

9. Noninvasive Techniques: Gamete Selection—Sperm�� 143

10. Noninvasive Techniques: Gamete Selection—Oocyte�� 155

11. Noninvasive Techniques: Embryo Selection by Oxygen Respiration�� 165

12. Noninvasive Techniques: Embryo Selection by Time-Lapse Imaging�� 177

15. Invasive Techniques: Embryo Biopsy at the Cleavage Stage�� 219

17. Invasive Techniques: Aneuploidy Testing by FISH�� 241

18. Invasive Techniques: Aneuploidy Testing by Array-CGH�� 255

19. Summary: Comprehensive Summary of Main Points by Topic��269