Prenatal Diagnosis (Brynn Levy)

Methods in
Molecular Biology 1885
Brynn Levy Editor
Prenatal
Diagnosis
Second Edition
METHODS IN MOLECULAR BIOLOGY
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John M. Walker
School of Life and Medical Sciences
University of Hertfordshire
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Prenatal Diagnosis
Second Edition
Edited by
Brynn Levy
Department of Pathology and Cell Biology, Vagelos College of Physicians and Surgeons, Columbia
University Irving Medical Center, New York, NY, USA
Editor
Brynn Levy
Department of Pathology and Cell Biology
Vagelos College of Physicians and Surgeons
Columbia University Irving Medical Center
New York, NY, USA
ISSN 1064-3745 ISSN 1940-6029 (electronic)

Methods in Molecular Biology
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Preface
It has been ten years since the first edition of this book was published. Over this period, the
concept of “genetic testing,” which historically conjured up the notion of single assays and
pinpoint precision in clinical diagnostics, has evolved to include a broader whole genome
testing approach that often leverages big data and complex algorithms. As such, we can now
consider ourselves to be deeply entrenched in the genomics era where bioinformatics and
terabytes of data are an integral part of analysis and diagnostics. Indeed, the American
College of Medical Genetics officially changed their name to the American College of
Medical Genetics and Genomics to recognize this paradigm shift.
The biggest change in the field of prenatal diagnosis has been observed in the area of
cytogenomics where microarray and next-generation sequencing technologies have become
the preferable genomic tool for the assessment of preimplantation embryos and first and
second trimester fetuses. The introduction of these newer genomic technologies has had a
major effect on the success rates of IVF as demonstrated in recent prospective randomized
clinical trials. In traditional prenatal diagnosis, chromosomal microarray analysis (CMA) has
been recommended for all pregnancies with a fetal structural anomaly and currently offers
the most comprehensive assessment of fetal aneuploidy, aneusomy, and microdeletions and
microduplications possible.
While microarray diagnostics currently provides the most complete cytogenomic assess-
ment of the fetus, the appeal of a highly sensitive noninvasive blood test that screens for the
common aneuploidies has made noninvasive prenatal testing (NIPT) one of the fastest-
adopted genetic tests. In fact, in the United States, the mass adoption of NIPT is believed to
be the primary reason for the decline in the number of invasive prenatal diagnosis proce-
dures performed. It is important to emphasize that NIPT, in its current form, is a screening
test while CMA is a diagnostic test. As a screening test, NIPT offers superior detection of
Down syndrome and the other common aneuploidies compared to traditional biochemical
markers and nuchal translucency (NT) measurements. However, as a screening test, it does
not cover the vast scope of genomic abnormalities that are detectable by CMA. NIPT has
also been developed to predict the fetal RhD gene status in order to guide targeted prenatal
anti-D prophylaxis and prevent hemolytic disease of the fetus and the newborn. This
approach has been particularly well received in Europe, especially in Scandinavia.
Next-generation sequencing (NGS) entered the prenatal world predominantly for
aneuploidy detection in preimplantation embryos and for noninvasive copy number assess-
ment of cell-free DNA. It is now increasingly utilized to evaluate single nucleotide variations
(SNVs) in fetuses with structural anomalies as well as assess cryptic complex rearrangements
and imbalances in fetuses carrying apparently balanced translocations. It is likely that in the
future NGS will serve to assess both SNVs and copy number changes in a single assay. In
order to replace CMA, NGS as a single test will need to match the current resolution and
accuracy of CMA testing. This will certainly require greater coverage of the genome which
will only become a reality for routine testing when whole genome sequencing costs decrease.
Over the past decade, novel technologies have been utilized for the development of new
diagnostic tests. However, the newer prenatal tests have not necessarily replaced the older
ones. A prime reason pertains to payment/reimbursement of the new and often more
expensive tests. Payment/reimbursement in countries with national health systems and
v
vi Preface
even by private health insurance companies is often not approved as the new tests are usually
deemed “experimental” with “insufficient evidence” to support clinical utility. In addition,
there is often a large time gap before these tests are universally adopted worldwide and even
within individual countries. In some instances, the time gap is directly related to the amount
of time it takes for the appropriate clinical trials to be performed to support clinical utility. In
other cases, there may be a lack of the necessary expertise to interpret the more complex
genomic assays. In many cases, the delay in adoption has to do with the economic resources
available to purchase the genomic equipment as well as validate and implement the new
tests. As such, there remains tremendous utility for many of the old style and less expensive
targeted tests like FISH, QF-PCR and MLPA.
This second edition of Prenatal Diagnosis is divided into three major sections; preim-
plantation genetic testing, traditional prenatal testing and finally non-invasive prenatal
testing. The first part of the book begins with a historical introduction to each of the
three major sections. Traditional prenatal testing methodologies that have served as the
gold standard for decades remain an important aspect of this book as they continue to serve
as the primary testing assays in many regions of the world. New to this book are methodol-
ogies that employ next generation sequencing techniques and these can be found in each of
the three primary sections.
My appreciation and thanks goes to the authors for their individual contributions. Their
willingness to share their protocols and experience provides a valuable resource to clinical
laboratories around the globe.
New York, NY, USA Brynn Levy

Contents
Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v
Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix
PART I HISTORICAL INTRODUCTION

1 Traditional Prenatal Diagnosis: Past to Present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3
Brynn Levy and Melissa Stosic
2 Overview of Preimplantation Genetic Diagnosis (PGD):
Historical Perspective and Future Direction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23
Joe Leigh Simpson, Anver Kuliev, and Svetlana Rechitsky
3 Noninvasive Approaches to Prenatal Diagnosis: Historical
Perspective and Future Directions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45
Lisa Hui
PART II PREIMPLANTATION GENETIC TESTING
4 Molecular Testing for Preimplantation Genetic Diagnosis

of Single Gene Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61
Rebekah S. Zimmerman, Jennifer Eccles, Chaim Jalas,
Nathan R. Treff, and Richard T. Scott Jr.
5 Detection of Aneuploidy and Unbalanced Rearrangements
Using Comparative Genomic Hybridization Microarrays . . . . . . . . . . . . . . . . . . . . 73
Lorena Rodrigo Vivo and Carmen Rubio Lluesa
6 Aneuploidy Screening using Next Generation Sequencing . . . . . . . . . . . . . . . . . . . 85
Cengiz Cinnioglu, Refik Kayali, Tristan Darvin, Adedoyin Akinwole,
Milena Jakubowska, and Gary Harton
PART III TRADITIONAL PRENATAL DIAGNOSIS

7 DNA Extraction from Various Types of Prenatal Specimens . . . . . . . . . . . . . . . . . . 105
Odelia Nahum, Amanda Thomas, and Brynn Levy
8 Assessment of Maternal Cell Contamination in Prenatal Samples
by Quantitative Fluorescent PCR (QF-PCR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117
Christie M. Buchovecky, Odelia Nahum, and Brynn Levy
9 Rapid Prenatal Aneuploidy Screening by Fluorescence In Situ
Hybridization (FISH) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129
Anja Weise and Thomas Liehr
10 Prenatal Detection of Chromosome Aneuploidy by Quantitative
Fluorescence PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139
Kathy Mann, Erwin Petek, and Barbara Pertl
vii
viii Contents
11 Multiplex Ligation-Dependent Probe Amplification (MLPA)

for Prenatal Diagnosis of Common Aneuploidies . . . . . . . . . . . . . . . . . . . . . . . . . . . 161
Jan Schouten, Paul van Vught, and Robert-Jan Galjaard
12 Chromosomal Microarray Analysis Using Array Comparative
Genomic Hybridization on DNA from Amniotic Fluid
and Chorionic Villus Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171
Ankita Patel
13 Prenatal Diagnosis Using Chromosomal SNP Microarrays . . . . . . . . . . . . . . . . . . . 187
Mythily Ganapathi, Odelia Nahum, and Brynn Levy
14 Rapid Detection of Fetal Mendelian Disorders: Thalassemia
and Sickle Cell Syndromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207
Joanne Traeger-Synodinos, Christina Vrettou,
and Emmanuel Kanavakis
15 Prenatal Diagnosis of Cystic Fibrosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221
Anastasia M. Fedick, Jinglan Zhang, Lisa Edelmann,
and Ruth Kornreich
16 Prenatal Diagnosis of Tay-Sachs Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233
Jinglan Zhang, Hongjie Chen, Ruth Kornreich, and Chunli Yu
17 Next Generation Sequencing of Prenatal Structural
Chromosomal Rearrangements Using Large-Insert Libraries . . . . . . . . . . . . . . . . . 251
Benjamin B. Currall, Caroline W. Antolik,
Ryan L. Collins, and Michael E. Talkowski
18 Prenatal Diagnosis by Whole Exome Sequencing in Fetuses
with Ultrasound Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
Vanessa Felice, Avinash Abhyankar, and Vaidehi Jobanputra
19 Isolation and Characterization of Amniotic Fluid-Derived
Extracellular Vesicles for Biomarker Discovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287
Blake Ebert and Alex J. Rai
PART IV NON-INVASIVE PRENATAL TESTING
20 Quad Screen Test, A Multiplexed Biomarker Assay for Prenatal Screening

to Assess Birth Defects: The Columbia University Experience
Using the Beckman Access2 Immunoassay Analyzer and Benetech PRA . . . . . . . 297
Awet Tecleab, Alex K. Lyashchenko, and Alex J. Rai
21 Isolation of Cell-Free DNA from Maternal Plasma . . . . . . . . . . . . . . . . . . . . . . . . . . 309
James Stray and Bernhard Zimmermann
22 Noninvasive Detection of Fetal Aneuploidy Using Next
Generation Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325
Kirsten J. Curnow, Rebecca K. Sanderson, and Sue Beruti
23 Noninvasive Antenatal Screening for Fetal RHD in RhD Negative
Women to Guide Targeted Anti-D Prophylaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 347
Frederik Banch Clausen, Klaus Rieneck, Grethe Risum Krog,
Birgitte Suhr Bundgaard, and Morten Hanefeld Dziegiel
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361
Contributors
AVINASH ABHYANKAR Molecular Diagnostics, New York Genome Center, New York, NY,
USA
ADEDOYIN AKINWOLE IGENOMIX USA, Torrance, CA, USA
CAROLINE W. ANTOLIK Massachusetts General Hospital, Boston, MA, USA; Broad Institute,
Harvard Medical School, Cambridge, MA, USA
SUE BERUTI Illumina, Inc., San Diego, CA, USA
CHRISTIE M. BUCHOVECKY Department of Pathology and Cell Biology, Vagelos College of
Physicians and Surgeons, Columbia University Irving Medical Center, New York, NY,
USA
BIRGITTE SUHR BUNDGAARD Department of Clinical Immunology, Section 2034,
Copenhagen University Hospital, Copenhagen, Denmark
HONGJIE CHEN Mount Sinai Genomics, Inc., DBA Sema 4, New York, NY, USA;
Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai,
New York, NY, USA
CENGIZ CINNIOGLU IGENOMIX USA, Torrance, CA, USA
FREDERIK BANCH CLAUSEN Department of Clinical Immunology, Section 2034, Copenhagen
University Hospital, Copenhagen, Denmark
RYAN L. COLLINS Massachusetts General Hospital, Boston, MA, USA; Broad Institute,
KIRSTEN J. CURNOW Illumina, Inc., Foster City, CA, USA
BENJAMIN B. CURRALL Massachusetts General Hospital, Boston, MA, USA; Broad Institute,
TRISTAN DARVIN IGENOMIX USA, Torrance, CA, USA
MORTEN HANEFELD DZIEGIEL Department of Clinical Immunology, Section 2034,
Copenhagen University Hospital, Copenhagen, Denmark; Institute of Clinical Medicine
(IKM), Copenhagen University, Copenhagen, Denmark
BLAKE EBERT Department of Pathology and Cell Biology, Vagelos College of Physicians and
Surgeons, Columbia University Irving Medical Center, New York, NY, USA
JENNIFER ECCLES Baylor Genetics, Houston, TX, USA
LISA EDELMANN Department of Genetics and Genomic Sciences, Icahn School of Medicine at
Mount Sinai, New York, NY, USA; Mount Sinai Genomics, Inc., DBA Sema4 Genomics,
New York, NY, USA
ANASTASIA M. FEDICK Department of Genetics and Genomic Sciences, Icahn School of
Medicine at Mount Sinai, New York, NY, USA; Mount Sinai Genomics, Inc., DBA Sema4
Genomics, New York, NY, USA
VANESSA FELICE Molecular Diagnostics, New York Genome Center, New York, NY, USA
ROBERT-JAN GALJAARD Department of Clinical Genetics, Erasmus University Medical
Center, Rotterdam, The Netherlands
MYTHILY GANAPATHI Department of Pathology and Cell Biology, Vagelos College of
USA
GARY HARTON IGENOMIX USA, Torrance, CA, USA
ix
x Contributors
LISA HUI Department of Perinatal Medicine, Mercy Hospital for Women, Heidelberg, VIC,
Australia; Department of Obstetrics and Gynaecology, University of Melbourne, Parkville,
VIC, Australia; Reproductive Epidemiology, Murdoch Children’s Research Institute,
Parkville, VIC, Australia
MILENA JAKUBOWSKA IGENOMIX USA, Torrance, CA, USA
CHAIM JALAS Foundation for Embryonic Competence, Baskign Ridge, NJ, USA
VAIDEHI JOBANPUTRA Molecular Diagnostics, New York Genome Center, New York, NY,
USA; Department of Pathology and Cell Biology, Vagelos College of Physicians and
EMMANUEL KANAVAKIS Department of Medical Genetics, St. Sophia’s Children’s Hospital,
National and Kapodistrian University of Athens, Athens, Greece; Genesis Genoma Lab,
Athens, Greece
REFIK KAYALI IGENOMIX USA, Torrance, CA, USA
RUTH KORNREICH Department of Genetics and Genomic Sciences, Icahn School of Medicine
at Mount Sinai, New York, NY, USA; Mount Sinai Genomics, Inc., DBA Sema4
Genomics, New York, NY, USA
GRETHE RISUM KROG Department of Clinical Immunology, Section 2034, Copenhagen
University Hospital, Copenhagen, Denmark
ANVER KULIEV Florida International University, Miami, FL, USA; Reproductive Genetics
Institute, Inc. (RGI), Northbrook, IL, USA
BRYNN LEVY Department of Pathology and Cell Biology, Vagelos College of Physicians and
THOMAS LIEHR Jena University Hospital, Institute of Human Genetics, Friedrich Schiller
University, Jena, Germany
CARMEN RUBIO LLUESA Igenomix, Valencia, Spain
ALEX K. LYASHCHENKO Department of Pathology and Cell Biology, Vagelos College of
USA
KATHY MANN Viapath Analytics, Guy’s Hospital, London, UK
ODELIA NAHUM Department of Pathology and Cell Biology, Vagelos College of Physicians
and Surgeons, Columbia University Irving Medical Center, New York, NY, USA
ANKITA PATEL Lineagen, Salt Lake City, UT, USA
BARBARA PERTL Prenatal Centre, Ragnitz Hospital, Graz, Austria
ERWIN PETEK Institute of Human Genetics, Medical University of Graz, Graz, Austria
ALEX J. RAI Department of Pathology and Cell Biology, Vagelos College of Physicians and
SVETLANA RECHITSKY Florida International University, Miami, FL, USA; Reproductive
Genetics Institute, Inc. (RGI), Northbrook, IL, USA
KLAUS RIENECK Department of Clinical Immunology, Section 2034, Copenhagen University
Hospital, Copenhagen, Denmark
REBECCA K. SANDERSON Illumina, Inc., Foster City, CA, USA
JAN SCHOUTEN Department of Clinical Genetics, Erasmus University Medical Center,
Rotterdam, The Netherlands
RICHARD T. SCOTT JR. Thomas Jefferson University, Basking Ridge, NJ, USA; Rutgers-
Robert Wood Johnson Medical School, Piscataway Township, NJ, USA
JOE LEIGH SIMPSON March of Dimes Foundation, White Plains, NY, USA; Florida
International University, Miami, FL, USA; Reproductive Genetics Institute, Inc. (RGI),
Northbrook, IL, USA
Contributors xi
MELISSA STOSIC Department of Obstetrics and Gynecology, Vagelos College of Physicians and
JAMES STRAY Natera Inc., San Carlos, CA, USA
MICHAEL E. TALKOWSKI Massachusetts General Hospital, Boston, MA, USA; Broad
Institute, Harvard Medical School, Cambridge, MA, USA
AWET TECLEAB Department of Pathology and Laboratory Medicine, Staten Island
University Hospital, Staten Island, NY, USA
AMANDA THOMAS Department of Pathology and Cell Biology, Vagelos College of Physicians
and Surgeons, Columbia University Irving Medical Center, New York, NY, USA
JOANNE TRAEGER-SYNODINOS Department of Medical Genetics, St. Sophia’s Children’s
Hospital, National and Kapodistrian University of Athens, Athens, Greece
NATHAN R. TREFF Genomic Prediction, North Brunswick, NJ, USA
PAUL VAN VUGHT Department of Clinical Genetics, Erasmus University Medical Center,
Rotterdam, The Netherlands
LORENA RODRIGO VIVÓ Igenomix, Paterna, Valencia, Spain
CHRISTINA VRETTOU Department of Medical Genetics, St. Sophia’s Children’s Hospital,
National and Kapodistrian University of Athens, Athens, Greece
ANJA WEISE Jena University Hospital, Institute of Human Genetics, Friedrich Schiller
University, Jena, Germany
CHUNLI YU Mount Sinai Genomics, Inc., DBA Sema 4, New York, NY, USA; Department
of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY,
USA
JINGLAN ZHANG Department of Molecular and Human Genetics, Baylor College of
Medicine, Houston, TX, USA
BERNHARD ZIMMERMANN Natera Inc., San Carlos, CA, USA
REBEKAH S. ZIMMERMAN Sema4, New York, NY, USA; Department of Genetics and
Genomic Sciences, The Icahn School of Medicine at Mount Sinai, New York, NY, USA
Part I
Historical Introduction
Chapter 1
Traditional Prenatal Diagnosis: Past to Present

Brynn Levy and Melissa Stosic
Abstract
In the nearly 60 years since prenatal diagnosis for genetic disease was first offered, the field of prenatal
diagnosis has progressed far past rudimentary uterine puncture to provide fetal material to assess gender and
interpret risk. Concurrent with the improvements in invasive fetal sampling came technological advances in
cytogenetics and molecular biology that widened both the scope of genetic disorders that could be
diagnosed and also the resolution at which the human genome could be interrogated. Nowadays, routine
blood work available to all pregnant women can determine the risk for common chromosome abnormal-
ities; chorionic villus sampling (CVS) and amniocentesis can be used to diagnose nearly all conditions with a
known genetic cause; and the genome and/or exome of a fetus with multiple anomalies can be sequenced in
an attempt to determine the underlying etiology. This chapter will discuss some of the major advances in
prenatal sampling and prenatal diagnostic laboratory techniques that have occurred over the past six
decades.
Key words History of prenatal diagnosis, Chorionic villus sampling, Amniocentesis, Ultrasound
1 Introduction
It was less than a lifetime ago that, in 1956, Tijo and Levan first
correctly determined that humans have 46 chromosomes, rather
than 48 as was thought for over 30 years [1]. The establishment of
the proper number of human chromosomes laid the groundwork
for defining the various common chromosomal aneuploidies that
were just waiting to be discovered. The association of chromosome
abnormalities and specific clinical phenotypes led to a new era in
pediatric diagnosis and it was not long before chromosome analysis
was applied to prenatal testing. One of the early scientific reports on
transabdominal sampling of amniotic fluid dates back to 1897
when Prochownick reported on the chemical components of amni-
otic fluid [2]. However, it was not until 1955 and 1956 that the
first analyses of amniotic fluid for genetic information were per-
formed utilizing Barr bodies to determine fetal sex [3, 4]. Three
years later, in 1959, the underlying causes of common aneuploidies
Brynn Levy (ed.), Prenatal Diagnosis, Methods in Molecular Biology, vol. 1885, https://doi.org/10.1007/978-1-4939-8889-1_1,
3
4 Brynn Levy and Melissa Stosic
including Down syndrome, Turner syndrome, Klinefelter syn-

drome, and XXX were identified and shortly thereafter, XYY [5–9].
In 1960, Riis and Fuchs described performing the amniocente-
sis procedure on two pregnant women who were Hemophilia A
carriers [10]. This is the first report of performing prenatal testing
on the basis of an inherited risk for genetic disease. These women
would have terminated their pregnancies due to the risk of Hemo-
philia in their sons, but instead, underwent amniocentesis to deter-
mine fetal sex with the plan of continuing the pregnancy if a female
fetus was identified.
While the idea of culturing amniotic fluid for chromosome
analysis was advanced by Fuchs and Riis in 1960 [11], Steele and
Breg published the first proof of concept study in 1966 showing
that culturing and karyotyping of amniotic fluid cells was indeed
possible [12]. The following year, Jacobsen and Barter initiated
cultures on amniotic fluid obtained from 85 human cases. They
showed a 67% success rate (67/85) for obtaining viable cell cultures
and a 38% success rate (33/85) for yielding a final genetic evalua-
tion [13]. Their cohort included 6 cases which presented with
“genetic high risk pregnancies” prompting a “diagnostic” amnio-
centesis. A successful karyotype was obtained on all 6 cases. From
that point onward, the vast field of prenatal diagnosis opened up as
major efforts were directed at the development of and improve-
ment upon both invasive testing and the laboratory analysis of
prenatal samples. Here, we take a brief look at the progress of
routine prenatal screening as well as invasive testing techniques
including amniocentesis, fetoscopy, chorionic villus sampling
(CVS), and percutaneous umbilical blood sampling. In addition,
we will review the major milestones in the technologies developed
to diagnose genetic disease in prenatal specimens, taking you on a
journey from cytogenetics to sequencing.
2 Noninvasive Screening
2.1 Ultrasound Case reports identifying fetal and pregnancy abnormalities via ultra-
sonography began to appear in the 1960s [14–17]. Advances in
ultrasonography over the next 20 years provided the ability to
better date pregnancies, identify twins, locate the placenta, visualize
intrauterine growth restriction, and diagnose anomalies such as
spina bifida and limb reduction defects. When real time scanners
with improved resolution were introduced alongside color Doppler
in the mid-1980s, the potential for imaging all pregnancies became
a reality and hospitals began to perform anatomy scans at 20 weeks
gestation. The detection rate for fetal anomalies at 20 weeks varied
by study but was potentially as low as 17% [18]. This rate has
steadily increased over time as has the capacity to detect major
anomalies in the first trimester. Dane et al. in 2007, performed a
History of Traditional Prenatal Diagnosis 5
large study of 1290 cases and demonstrated a 70% detection rate for
major anomalies after the first trimester scan [19]. When a second
trimester ultrasound was added, the detection rate increased to 95%
[19]. Furthermore, the ability to perform invasive testing with
placental localization, and later, under continuous ultrasound mon-
itoring, lead to reduced complication rates [18, 20–23].
2.2 Biochemical In 1977, Wald et al. published a large multi-center collaborative

Screening study to determine the accuracy of maternal serum alpha-
fetoprotein (AFP) in screening for open neural tube defects
(ONTDs) [24]. The study included nearly 19,000 unaffected preg-
nancies and 301 pregnancies with ONTDs. They determined that
16–18 weeks is the best window for screening, allowing for 88% of
cases of anencephaly and 79% of cases of open spina bifida to be
detected at a 3% false positive rate. This has improved with mod-
ifications to the AFP measurement and algorithm [25]. In addition,
the sensitivity of ultrasound to identify cases of spina bifida and all
cases of anencephaly is now reported to be as high as 95%, making
AFP less widely utilized [26].
Benacerraf et al. reported on the association of an increased
nuchal fold measurement in the second trimester with Down syn-
drome in 1985 and shortly after, additional markers such as short-
ened femurs were noted [27–29]. By 1992, the earlier association
of increased nuchal translucency in the first trimester and Down
syndrome had also been identified by Nicolaides et al. [30].
In the mid to late 1980s, the association between aneuploidy
and abnormal amounts of AFP, unconjugated estriol, and human
chorionic gonadotropin (HCG) in maternal serum was uncovered
[31–35]. This lead to the creation of the triple screen in 1988
which was able to identify 60% of Down syndrome pregnancies at
a 5% false positive rate [36]. Several years later, in 1995, 79% of
laboratories were using multiple markers to screen for Trisomy 21;
the remainder were using AFP alone [37]. In the early 1990s,
abnormal levels of free ß-HCG and pregnancy-associated plasma
protein A (PAPP-A) in the first trimester were correlated with
aneuploidy [38, 39]. By 1997 it was shown that these two markers,
when combined with nuchal translucency measurement, provide an
80% detection rate for Down syndrome for a 5% false positive rate
[40]. Around the same time, Inhibin A was found to be a fourth
marker in the second trimester and was added to the others to
create the quad test, replacing the triple screen [41–43]. With
both first and second trimester screening possible, the integrated
screen was developed in 1999, combining first and second trimester
screens for an 85% detection rate and 0.9% false positive rate
[44]. Integrated screening requires waiting until the second trimes-
ter for results and therefore, the sequential screen was proposed,
which provides results for very high risk women in the first trimes-
ter, and those with low risk results proceeded to subsequent
additional screening in the second trimester [45]. With the avail-

ability of multiple options for maternal serum screening, the First
and Second Trimester Evaluation of Risk (FASTER) consortium
published findings on the reliability of first and second trimester
screening as well as combined screening in 2005 [46].
2.3 Noninvasive In 2011, after decades of reports of fetal cells and cell-free fetal
Prenatal Testing (NIPT) DNA in maternal circulation, the first noninvasive prenatal tests
Using Cell-Free using cell-free fetal DNA for the detection of Trisomy 21 became
Fetal DNA commercially available [47–49]. Now, NIPT is able to screen for
the common aneuploidies (13, 18, 21, X & Y), large deletions and
duplications, and the common microdeletion syndromes. The sen-
sitivity of NIPT for Down syndrome is >99% for a very low false
positive rate (<1% even when screening for multiple conditions).
With NIPT clearly showing superiority over traditional biochemi-
cal/sonographic markers for aneuploidy screening [50], the Amer-
ican College of Obstetricians and Gynecologists (ACOG) released
guidelines in 2016 stating that NIPT is an appropriate screening
test for pregnant women. The same year, the American College of
Medical Genetics and Genomics (ACMGG) issued a statement
strongly suggesting NIPT replaces serum screening for aneuploidy
in women of any age [51, 52].
3 Invasive Procedures
3.1 Amniocentesis Amniocentesis is the technique for withdrawing amniotic fluid

transabdominally from the amniotic sac using a needle guided by
ultrasound. Amniotic fluid contains cells that have been sloughed
off from the amnion and from the fetus. In 1968, Nadler showed
that enzyme levels could be measured in amniotic fluid, thus open-
ing the door for diagnosing metabolic conditions in utero
[53]. Together, he and his colleague Walsh, would go on to prena-
tally diagnose the first cases of cystic fibrosis (one of the most
common autosomal recessive genetic diseases in Caucasians) using
quantitative and qualitative measurements of methylumbelliferyl-
guanidinobenzoate (MUGB) reactive proteases in amniotic fluid
[54]. The earliest description of an increase in alpha-fetoprotein
(AFP) levels in the amniotic fluid of fetuses affected with open
neural tube defects was by Brock and Sutcliffe [55]. In 1970 Nadler
and Gerbie published an amniocentesis series of 162 high risk
women, diagnosing 10 cases of Down syndrome and 3 additional
conditions for a total of 13 affected pregnancies with no reported
fetal or maternal complications [56]. Several years later, in 1976,
the National Institute of Child Health and Human Development
(NICHD) published on the safety and accuracy of amniocentesis,
with a quoted diagnostic accuracy of 99.4% and a fetal loss rate of
well below 1% [57]. Once the results were published, it became
standard of care to recommend an amniocentesis during the second

trimester to women over the age of 35 in order to detect fetal
chromosome abnormalities. Reports of early amniocenteses per-
formed prior to 15 weeks began to appear in 1987 but the fetal
loss rate was found to be 2.3% and a discernable increase in con-
genital abnormalities, particularly talipes, was observed [58]. As
such, early amniocentesis was discouraged and is no longer rou-
tinely performed [58–61]. It was not until 2007 that ACOG
changed the recommendations for invasive testing, no longer lim-
iting it to pregnant women over the age of 35, but rather offering
diagnostic testing and screening to all women, regardless of age
[62, 63].
3.2 Fetoscopy Fetoscopy is an endoscopic procedure that provides access (for

biopsy and/or surgery) to the fetus, the amniotic sac, the umbilical
cord, and the fetal side of the placenta. As early as 1954, Westin was
able to visualize three fetuses in utero via hysteroscope [64]. It
wasn’t until 1973 that Valenti collected fetal blood and skin biop-
sies with the goal of eventually using the technique to diagnose
hemoglobinopathies and ultimately perform intrauterine therapy
[65]. The same year, Scrimgeour used an endoamnioscope to
identify central nervous system anomalies in three fetuses [66]
and a year later, Hobbins and Mahoney reported on fetal blood
collection from a small cohort of eight women [67]. Fetoscopy was
readily introduced for the diagnosis of inherited blood disorders,
beginning with hemoglobinopathies in 1975 and then hemophilia
in 1979 [68, 69]. This method was also utilized to identify genetic
conditions by visualizing the fetal anatomy, as was shown in 1977
for the case of a fetus with skeletal dysplasia [70]. Incipient exam-
ples of obtaining fetal blood and/or skin biopsy for the diagnosis of
skin diseases such as ichthyosis and epidermis bullosa as well as
metabolic diseases including galactosemia and Tay-sachs were first
reported between 1977 and 1980 [70–74]. Given the fetal loss rate
of 3–5% and the relative safety of amniocentesis, fetoscopy is no
longer utilized for prenatal diagnosis except in rare instances, such
as mosaicism [75]. However, it has become a mainstay in the
treatment of prenatal anomalies and complications such as twin to
twin transfusion syndrome, diaphragmatic hernia, and amniotic
band syndrome.
3.3 Chorionic Villus Chorionic villi are part of the placenta and can be obtained by two
Sampling (CVS) different ultrasound guided methods depending upon the uterine
position and the location of the placenta. The transcervical method
involves inserting a catheter through the vagina and cervix while
the transabdominal method involves inserting a thin needle
through a sterile area of the abdomen (similar to amniocentesis).
The concept of first trimester transcervical sampling of the
placenta was first proposed by Mohr in 1968 [76]. Five years
later, in 1973, Sandahl and Kullander biopsied the placenta in

39 patients undergoing termination of pregnancy. Karyotype was
successful in half of them and was shown to truly represent the fetal
genetic constitution [77]. The following year, Hahnemann devel-
oped new equipment and had a 38% biopsy success rate [78]. In
1975, a group in China reported 99 successful procedures for fetal
sex determination [79]. In their case series, only six incorrect fetal
sex determinations were made and four fetal losses occurred fol-
lowing the procedure [79]. 1982–3 saw groundbreaking work on
transcervical CVS resulting in the procedure being offered in many
centers around the globe with each center having a slightly varied
technique and instrument [80–85]. When tissue culture of chori-
onic villi was perfected, it became possible to offer routine cyto-
genetic testing of the fetus from a CVS. In 1983, Pergament and
colleagues performed simultaneous enzyme and chromosome test-
ing on a CVS sample from a woman at risk for Tay-Sachs disease
[86]. The same year, Brambati and Simoni reported the prenatal
diagnosis of trisomy 21 following a first trimester CVS performed
on a woman at 50% risk for a male child with Duchenne muscular
dystrophy [87]. In the Brambati case report, they utilized a newly
developed technique [88] for direct analysis of the villi which
resulted in a karyotype within hours of the procedure.
Transabdominal techniques were unsuccessful initially, as they
could only be performed in the second trimester, and amniocente-
sis was the better option at that gestational age [89, 90]. In 1984,
the transabdominal route became possible in the first trimester
[91]. Today, both transcervical and transabdominal CVS are com-
monly performed.
The safety and efficacy of CVS was assessed in large studies
toward the end of the 1980s and early 1990s with the finding that
the rates of miscarriage after CVS were only slightly higher than
those associated with amniocentesis [92, 93]. These studies indi-
cated the rates of miscarriage to be around 0.5%–1.0%
[92, 93]. Despite the somewhat higher risk for miscarriage, CVS
has various emotional and medical advantages over amniocentesis
as it provides an opportunity for earlier decision making in the wake
of a diagnosis of a chromosome abnormality (almost 6 weeks earlier
than amniocentesis). In the early 1990s, there was considerable
worry regarding the causal link between CVS and limb-reduction
defects. After much debate and several cohort studies, it was deter-
mined that CVS performed before 9 weeks was indeed a risk factor
for limb-reduction defects [94]. In 1992, a WHO-sponsored com-
mittee recommended that CVS be performed at 9–12 weeks after
the last menstrual period [94]. CVS is now safely performed in
most institutions between 10 and 12 weeks for the diagnosis of a
wide range of genetic conditions.
3.4 Percutaneous Percutaneous Umbilical Blood Sampling (PUBS) involves obtain-

Umbilical Blood ing a blood sample from the fetus. It was originally developed in
Sampling (PUBS) conjunction with fetoscopy in mid 1960s and early 1970s as a
means to do prenatal diagnosis of hemoglobinopathies
[95, 96]. PUBS is also referred to as Fetal Blood Sampling, Cor-
docentesis, and Umbilical Vein Sampling. PUBS involves the inser-
tion of a thin needle through the abdomen and uterine walls into
the umbilical cord at the site where the cord inserts into the
placenta.
In 1983 Daffos et al. extracted fetal blood from the umbilical
vein near the cord insertion using ultrasound to guide the proce-
dure in 63 pregnancies [97]. Nicolaides perfected the technique he
named cordocentesis, and used it in cases of Rhesus disease to
determine the severity of anemia in the fetus [98, 99]. PUBS was
also performed to assess for amino acid status in fetuses with
intrauterine growth restriction [100].
PUBS carries a risk of fetal death or spontaneous abortion
which is higher than that of CVS or amniocentesis and is estimated
to be around 1–2% when performed by an experienced operator
[101–103]. Today, PUBS is mainly reserved for cases in which
diagnostic information cannot be obtained through amniocentesis,
CVS, or ultrasound, or the results of these tests are inconclusive
and further information is required for appropriately interpreting
the clinical significance of the prior results. Examples include chro-
mosomal mosaicism discovered after CVS or amniocentesis, the
necessity for a rapid fetal karyotype, and the identification of fetal
infection or hematologic condition. However, the procedure is also
often performed for therapeutic indications such as transfusions in
the case of Rhesus isoimmunization or anemia. The procedure is
best performed at or after 18–20 weeks.
4 Techniques for Assessing Fetal Genetic Information
4.1 Karyotype The ability to diagnose chromosome abnormalities from prenatal

specimens dates back to end of the 1960s [12, 13]. In these early
studies, chromosomes were grouped based on their length and
centromere position since not all of them could be individually
identified [104]. Starting in 1970, multiple types of fluorescent
dyes such as quinocrine and stains such as Giemsa were utilized to
stain the chromosomes and display a banding pattern that allowed
unique identification of individual chromosomes and structural
changes [105–109]. When the safety and accuracy of amniocentesis
was established in the mid-1970s [57], the improved karyotype
techniques developed during that same decade became a routine
tool for the prenatal diagnosis of fetal chromosome abnormalities.
This was fueled primarily by the recognition that advanced maternal
age is directly related to an increased risk of chromosome
abnormalities in pregnancy. As such, advanced maternal age, gen-

erally defined in the United States as 35 or older at delivery, became
the most common indication for prenatal cytogenetic diagnosis
[110, 111]. To this day, karyotyping remains one of the most
commonly performed techniques utilized on prenatal specimens.
4.2 Molecular DNA The first DNA-based genetic tests were developed in the late 1970s
Testing and used the principles of Restriction Fragment Length Poly-
morphisms (RFLP). RFLP analysis for genetic disease relies on a
mutation (DNA base changes/small insertion/deletion) that
removes, inserts, or rearranges a restriction enzyme site, thus gen-
erating a different sized DNA fragment compared to the wild-type
fragment. The classic illustration is that of sickle cell disease where
the wild-type β-globin gene fragment is 7.7 Kb and the mutant
β-globin gene fragment is 13 Kb when cut with the restriction
enzyme Hpa I [112]. In 1978, Kan and Dozy demonstrated that
these RFLP differences could be utilized for prenatal diagnosis of
sickle cell disease from DNA isolated from amniotic fluid
[112]. This proved to be the first example of a DNA-based prenatal
diagnostic test [112]. From 1978 to 1980, the principles of RFLP
analysis were applied for prenatal diagnosis of α- and
β-Thalassemias [113–115]. Linkage analysis using RFLPs began
to emerge around 1983 when investigators began to track the
inheritance of multiple restriction fragments that cosegregated
with the disease of interest. This required large family studies to
identify informative RFLPs that were in linkage disequilibrium with
the disease under consideration.
RFLP analysis at that time was a labor intensive and time-
consuming technique that required multiple technical processes
such as probe labeling, DNA fragmentation, electrophoresis,
Southern blotting, hybridization, washing, and autoradiography.
In some instances, it could take up to a month to get results back
and in a prenatal setting, this was a major disadvantage. When Kary
Mullis conceived of the polymerase chain reaction (PCR) technique
in 1983 [116], it set the stage for one of the most significant
scientific advances of the twentieth century. For his efforts, Mullis
became one of the recipients of the 1993 Nobel Prize for Chemis-
try. PCR allowed for the rapid generation of millions of copies of a
particular DNA sequence [117] making it a highly attractive tech-
nique for prenatal diagnosis. Indeed, soon thereafter in 1985, Saiki,
together with Mullis and colleagues, reported on two new molecu-
lar methods that allowed for a rapid and highly sensitive prenatal
diagnostic test for sickle cell anemia [118]. The first method uti-
lized PCR to amplify specific β-globin target sequences, resulting in
the exponential increase of target DNA copies. The second tech-
nique used end-labeled oligonucleotide probes to hybridize to the
amplified β-globin DNA fragments that had been digested with a
restriction endonuclease [118]. The two methods, coupled
together, dramatically increased the speed and sensitivity of the

assay and allowed for a result to be generated overnight instead of
after several days [118]. In 1987, Kogan et al. improved the PCR
method by substituting the Klenow fragment of Escherichia coli
DNA polymerase I (Klenow) with a heat-stable DNA polymerase
from Thermus aquaticus (Taq polymerase) [119]. By utilizing the
heat-stable Taq polymerase, Kogan and colleagues showed that
they could amplify DNA sequences at an elevated temperature
resulting in a marked improvement of the specificity of amplifica-
tion [119]. This specificity generated an enormous surplus of the
desired DNA molecule of the appropriate size, which they could
then detect by visual inspection after ethidium bromide staining
and without the use of radioactive probes [119]. Kogan et al. used
their modified PCR technique to perform prenatal diagnosis of
hemophilia and fetal sex determination [119]. In 1989, Gasparini
and coworkers used Kogan’s updated PCR method to perform
prenatal diagnosis for cystic fibrosis [120]. In their report, they
described their experience using PCR on eight cases for first-
trimester prenatal diagnosis of cystic fibrosis, following fetal sam-
pling by transabdominal CVS [120]. Interestingly, at that time, the
gene causing cystic fibrosis was unknown and prenatal studies could
only be done by linkage analysis. Utilization of the newly developed
PCR technique was not limited to the diagnosis of genetic disorders
as it was also embraced for the detection of fetal infectious diseases.
In 1990, Grover et al. used PCR for rapid prenatal diagnosis in
amniotic fluid of congenital toxoplasma infection [121] while
Ho-Terry and colleagues used it for prenatal detection of fetal
rubella infection [122]. There are now over 30 different variations
of the basic PCR technique, two of which play a major role in rapid
prenatal diagnosis of the common aneuploidies. The first is quanti-
tative fluorescent PCR (QF-PCR) which calculates chromosome
dosage in real time using polymorphic small tandem repeats (STR)
markers. It was developed in 1993 by Mansfield who demonstrated
that allele dosage for the affected chromosome in patients with
trisomy 18 and trisomy 21 could be distinguished from normal
individuals [123]. In 1994, Pertile and coworkers used a similar
approach to detect aneuploidies in fetal amniotic fluid samples
[124]. The second technique, called multiplex ligation-dependent
probe amplification (MLPA), was developed by Schouten and col-
leagues in 2002, and allows relative quantification of chromosome-
specific DNA sequences in a single reaction [125]. Both techniques
are widely utilized, primarily outside the USA, for quick assessment
of aneuploidy of chromosomes 13, 18, 21, X, and Y. In the USA,
fluorescence in situ hybridization (FISH) is the primary method for
rapid aneuploidy detection (see below).
4.3 Fluorescence In The combination of traditional cytogenetic and molecular techni-

Situ Hybridization ques yielded a new area in the field of human genetics which is
(FISH) referred to as molecular cytogenetics. Fluorescence in situ hybridi-
zation (FISH) can be considered the hallmark of molecular cytoge-
netics and its development added a powerful new dimension to the
clinical diagnostic testing arena. Molecular cytogenetics really
began in the early 1970s when various groups described a tech-
nique for detecting highly repetitive nucleic acid sequences in cell
preparations [126–128]. The in situ hybridization (ISH) proce-
dures utilized in the 1970s and early 1980s used radioactive probes
that were readily substituted for by fluorescence in situ hybridiza-
tion (FISH) techniques that instead applied non-radioactive probes
labeled with haptens or fluorochromes [129–132].
From a clinical perspective, FISH could be performed using
both interphase nuclei and metaphase spreads [133–137] and
provided a way to detect cryptic and submicroscopic rearrange-
ments that remained undetected or undecipherable by conven-
tional cytogenetic analysis [134, 138–143]. In the prenatal world,
it was readily utilized on uncultured cells (interphase nuclei)
derived from chorionic villi or amniotic fluid to rapidly screen for
the common aneuploidies [144, 145]. The primary advantage of
prenatal FISH was the ability to get preliminary results in as little as
1–2 days. In 2001, a 2-year multi-center retrospective study by
Tepperberg and colleagues showed an extremely high concordance
rate between FISH and standard cytogenetics (99.8%) as well as
sensitivity, specificity, and positive and negative predictive values all
being greater than 99.5% [146]. In the Tepperberg study, abnor-
mal ultrasound and advanced maternal age (though not mutually
exclusive) followed by an abnormal maternal serum analyte screen
were the most common indications for performing interphase pre-
natal FISH [146]. The clinical use of FISH has not changed much
over the years and today, it is still utilized for rapid preliminary
results after invasive testing and to test for deletions or duplications
in specific regions associated with disease. FISH has also proven
useful for follow-up studies assessing the suspicion of chromosomal
mosaicism.
4.4 Comparative The development of chromosomal comparative genomic hybridi-

Genomic Hybridization zation (cCGH) by Kallioniemi and coworkers in 1992 can be
and Microarray viewed as the next major leap in molecular cytogenetic diagnostic
Analysis technologies [147]. cCGH was initially utilized as a cancer research
tool to uncover novel oncogenes and tumor suppressor genes
[147–153] but was quickly adapted, a few years later, for clinical
purposes in constitutional genetics [154–156]. The power of
cCGH lays in its ability to reveal any DNA sequence copy number
change (i.e., gains, amplifications, or losses) in a particular speci-
men, and map these changes on normal chromosomes [147, 153,
157, 158]. For the first time, clinical cytogeneticists had a tool that
could scan the entire genome without having to preselect specific

targeted regions. This proved particularly useful in the prenatal
setting for identifying chromosomal material of unknown origin
such as marker chromosomes and derivative chromosomes
[154, 159–162]. While the clinical utility of cCGH was readily
apparent, the technique itself was particularly challenging and
tedious and did not lend itself to routine clinical testing. In addi-
tion, the resolution of the technique was limited to that of the
metaphase chromosome which is theoretically around 2–4 Mb
when utilizing high resolution prometaphase chromosomes
[163]. This meant that cCGH could not readily detect submicro-
scopic imbalances like those observed in the microdeletion syn-
dromes such as Prader-Willi. In 1997/8, Solinas-Toldo et al. and
Pinkel et al. leveraged sequence data from the Human Genome
project and built DNA microarrays by spotting multiple cloned
DNA fragments onto glass microscope slides [164, 165]. The use
of spatially separated DNA clones on glass slides effectively replaced
metaphase chromosomes as the hybridization targets and made it
possible to detect imbalances ~100 kilobases in size
[164, 165]. As with cCGH, the clinical utility of array comparative
genomic hybridization (aCGH) soon became apparent and in the
early to mid-2000s, investigators began to use the new technique
for clinical purposes [166–172]. One of the first reports of the use
of aCGH in a prenatal setting was in 2004 by Tachdjian and cow-
orkers who demonstrated the origin of the extra chromosomal
material observed in a fetus with a reported 46,X,Xq + karyotype
[168]. In 2005, Le Caignec et al. reported the first large prenatal
case series using aCGH [171]. Their cohort comprised 49 fetuses
with three or more significant anomalies and normal karyotype.
Their report demonstrated that aCGH on fetuses with anomalies
could increase the diagnostic yield over karyotype, which in their
study was around 8% [171]. In 2007, a Columbia University lead
multi-center trial was initiated in the USA to assess the clinical
utility of chromosomal microarray analysis (CMA) in prenatal diag-
nosis [173]. The trial lasted 5 years and was funded by the National
Institute of Child Health and Human Development (NICHD)
[173]. The NICHD trial demonstrated that CMA could detect all
of the abnormalities that karyotype could, plus additional copy
number variants, leading to incremental yields of 1.7% in low risk
pregnancies and 6% in anomalous fetuses [173].
The mid-2000s also saw evolutionary advances in array tech-
nology with DNA clones (typically bacterial artificial chromo-
somes) being replaced by short oligonucleotide probes, 25–80 bp
in size [174, 175]. Around the same time, investigators also began
to use oligonucleotide probes that included single nucleotide poly-
morphisms (SNPs) [176, 177]. The use of oligonucleotide probes
improved the resolution of imbalances that could be detected by
CMA to far below 100 Kb, essentially only being limited by the
number of probes used on a given array. Today, most clinical

laboratories prefer to use SNP-based oligonucleotide arrays as it
offers the ability to detect uniparental disomy (UPD), consanguin-
ity, maternal cell contamination, and triploidy, in addition to DNA
copy number changes [178–180].
4.5 Next Generation Traditional Sanger sequencing, developed in 1977, is a reliable way
Sequencing (NGS) to sequence individual genes or small regions of DNA [181]. Sanger
sequencing, while extremely useful, does not lend itself to high
throughput of multiple genes. Next Generation Sequencing
(NGS) was first developed in 1990 and provided a way to increase
the number of genes being sequenced in parallel [182]. The first
type of NGS was done by capillary sequencing which parallelized
Sanger sequencing into machines that could run 96 or 384 reac-
tions at once in microplate format [182]. NGS platforms are now
capable of sequencing millions or even billions of small fragments
of DNA at the same time [183]. Whole genome sequencing (WGS)
by NGS allows for the discovery of novel mutations and disease
causing genes without requiring prior knowledge of the gene or
locus being investigated [183]. Since WGS is expensive, strategies
for exclusively assessing coding (exon/gene) regions became an
attractive proposition, as this requires sequencing of only 1.5% of
the genome. This NGS method is referred to as whole exome
sequencing (WES) and its role in gene discovery was demonstrated
in 2010 when Ng and colleagues uncovered the genes associated
with Miller-Dieker [184] and Kabuki syndromes [185]. Other
studies quickly followed along with 100’s of additional gene dis-
coveries [186]. WGS and WES have recently made their way onto
the prenatal scene and are especially useful in identifying the under-
lying cause of genetic disease when karyotype, microarray, and
FISH or Sanger sequencing of known genes is not fruitful
[187]. However, WES and WGS are only recommended prenatally
in the case of ultrasound anomalies where other diagnostic tests
have failed to reveal an answer [188].
5 Concluding Comments
Amidst the progression in both procedural sampling and analysis

techniques in prenatal diagnosis, there has been a strong need to
support physicians in educating them and their patients about the
benefits and limitations of the new technologies. The first genetic
counseling program opened in New York in 1969 and began to
provide resources to physicians involved in clinical genetics and
prenatal diagnosis. In North America, there are currently 45 accre-
dited programs with more than 4000 genetic counselors across the
United States and Canada. As the field continues to expand and the
available tests become more technically advanced and nuanced, the
need for additional geneticists and genetic counselors to interpret

data and correlate genotypic and phenotypic information will defin-
itively increase. The prenatal tests available to women 50 years ago
were drastically different to what they are today and it is likely that
the future holds even more discoveries, options, and answers.
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Chapter 2
Overview of Preimplantation Genetic Diagnosis (PGD):

Historical Perspective and Future Direction
Joe Leigh Simpson, Anver Kuliev, and Svetlana Rechitsky
Abstract
Preimplantation genetic diagnosis (PGD) can be considered the earliest form of prenatal testing. It was first
used in humans over 26 years ago. At its inception, PGD could only be performed for a limited number of
genetic disorders. Technological advances in molecular biology and cytogenomics have been utilized in the
field of PGD to greatly expand the spectrum of genetic disorders that can now be detected in early human
embryos.
Key words Preimplantation genetic diagnosis, PGD, PGT, PGT-A, PGT-M, PGT-SR, PGT-HLA,
Single gene disorder, Biopsy, Blastomere, Trophectoderm, Mosaicism
1 Introduction
Preimplantation genetic diagnosis (PGD)1 was first accomplished

in humans over 26 years ago. The first 1000 PGD births were not
reached until 2004 [2], but since then cases have progressively
increased. PGD is now accepted as not simply an earlier extension
of traditional prenatal genetic diagnosis, but also offering novel
indications. This chapter will review the history of PGD in humans,
describing some of the key events and major breakthroughs that
have occurred over the past three decades.
1
Preimplantation genetic diagnosis (PGD) is the term traditionally applied. For aneuploidy, preimplantation
genetic screening (PGS) is often used. However, preimplantation genetic testing (PGT) is now considered by
WHO to be more appropriate [1]. The reason is that rarely does a confirmatory “test” follow PGD, and especially
after interrogation for aneuploidy testing to increase pregnancy rates. That “screening” is a misnomer is
increasingly recognized. Here we use the terms interchangeably given historical literature but accept that PGT
is the term to be used going forward.
23
24 Joe Leigh Simpson et al.
2 Pioneers in Detection of Single Gene Disorders and Chromosomal Abnormalities
PGD can be said to have begun in 1968 when Gardner and

Edwards [3] biopsied a rabbit blastocyst and performed
X-chromatin analysis. The application to X-linked recessive traits
was noted. This was then followed by Modlinsky and McLaren’s
[4] work on the visualization of the mouse second polar body
chromosomes, which, however, did not result in a sufficiently reli-
able method for practical application [4]. Over the next decade,
mouse geneticists demonstrated occasional success in generating
metaphase chromosomes from blastomeres [5]. Human in vitro
fertilization (IVF) was finally achieved in 1978 [6], and thereafter
animal studies in the preparation of human PGD accelerated.
Biopsy and analysis of mouse blastomeres soon showed feasibility
for detecting single gene disorders [7].
Pioneering animal work was performed in the United Kingdom
by Marilyn Monk and, Alan Handyside; in Australia by Leanda
Wilton and Alan Trounson; in Brussels by Andre Van Steirteghem
and colleagues; and in the United States by Yury Verlinsky and
colleagues.
Development of polymerase chain reaction (PCR) in the 1980s
had made human PGD feasible. This allowed molecular analysis of a
single cell (6 pg of DNA). In Europe, emphasis in the late 1980s
focused on blastomere biopsy. Handyside, Braude, and Winston
pursued blastomere biopsy and analysis, determining in 1990 sex in
two couples known to be at risk of transmitting adrenoleukody-
strophy and X-linked mental retardation. Two female embryos
were transferred after in vitro fertilization (IVF), biopsy of a single
cell at the six- to eight-cell stage, and sexing by DNA [8]. Soon
cystic fibrosis was detected, using PCR [9].
In the United States, Verlinsky and colleagues preferred polar
body biopsy, reporting their first clinical case at the 1987 Interna-
tional Congress on IVF in Kyoto [10].The first peer publication of
that case (α 1-antitrypsin deficiency) was delayed until 1990
[11]. In 1990, PGD for cystic fibrosis was also reported by this
same group [12].
PGD for chromosomal abnormalities had to await develop-
ment of chromosome-specific fluorescence in situ hybridization
(FISH) probes. In the United Kingdom, Darrel Griffin successfully
performed FISH on blastomeres [13], whereas in the United States
Grifo [14, 15] working with Cohen reported a pregnancy following
embryo biopsy subjected to both X and Y FISH [16]. Munné and
this same group later applied multicolor FISH on blastomeres [17],
setting the stage for modern PGD aneuploidy testing. In 1998,
Munné used FISH to identify unbalanced chromosomal transloca-
tions [18, 19], using breakpoint-specific probes. In the United
Kingdom, Delhanty and Harper performed rapid FISH [20] and
Historical Perspective and Future Directions in PGD 25
observed mosaicism in cleavage stage embryo; Munné also recog-

nized this problem, but considered it manageable with optimal
diagnostics. Verlinsky et al. [21] likewise applied FISH but to
polar bodies.
3 Professional and Scientific Organizations
Scientific presentations and clinical application of preimplantation

genetic diagnosis (PGD)—now properly called preimplantation
genetic testing (PGT)—has progressively become more organized.
Early cooperation was demonstrated by yearly international meet-
ings arranged by Verlinsky [22]. Beginning in 1990, an annual
“International Working Group on PGD” held in sites around the
world, often in conjunction with other major international meet-
ings. Materials of First International Conference on Preimplanta-
tion Genetics in Chicago in 1990 yielded a 1991 volume by
Verlinsky and Kuliev that exercised traction for the broader scien-
tific community [23].These meetings focused early PGT investiga-
tors on technology development until clinical application caught up
with their vision. In a few years, PGD had evolved from a boutique
component of prenatal genetic diagnosis to mainstream, having its
own distinct indications [24]. This set the stage for rapid progress
that followed [25–27].
In 2003, the Preimplantation Genetic Diagnosis International
Society (PGDIS) was incorporated in the United States (Illinois),
de facto the direct continuation of the yearly International Working
Group on PGD. The first meeting after incorporation was in
London in 2005. Annual meetings have been also held in Chicago,
Bologna, Cyprus, Antalya, Miami, Montpellier, Bergenz, Istanbul,
and Canterbury. The membership elected Yury Verlinsky as a
founding President of PGDIS. Terms of offices were set for
2 years. Subsequent Presidents have been Luca Gianaroli, Joe
Leigh Simpson, Robert G. Edwards, Renée Martin, Semra Kahra-
man, and Svetlana Rechitsky. Anver Kuliev serves as the Executive
Director (www.pgdis.org).PGDIS meetings are robust, with
pre-congress workshops. PGDIS now serves as the largest aggrega-
tion of dedicated PGD investigators globally.
In Europe, PGD workshops and formal communication are
organized through the European Society for Human Reproduction
and Embryology (ESHRE). PGD activities exist under aegis of the
ESHRE Special Interest Group (SIG) in Genetics, which in 1997
formed the ESHRE PGD Consortium. This Consortium collates
clinical results annually, and places data on the ESHRE web site and
in peer review publications [1, 28, 29]. Founding leaders were
Karen Sermon, Joep Geraedts, and Joyce Harper.
The American Society for Reproductive Medicine (ASRM)
formed its own SIG on PGD in 2005. Participation is open to all
ASRM members. Presidents (currently Chairmen) serve 2 year

terms and to date have been Yury Verlinsky, Jacques Cohen, Marcus
Hughes, Santiago Munné, Luca Gianaroli, and Anver Kuliev, with
Semra Kahraman Chairman Elect, and Svetlana Rechitsky Vice
Chairman. ASRM PGD SIG members provide recommendations
for PGD activities at the annual ASRM meeting. This includes
postgraduate courses, abstract review, and other educational and
scientific events.
The Reproductive Health and Research Program of the WHO
recently codified terms used to designate diagnosis of preimplanta-
tion embryos. This was undertaken through WHO NGOs
ICMART and as well as ASRM, IFFS, FIGO. In particular, the
term preimplantation genetic screening (PGS) to connote aneu-
ploidy testing was considered inappropriate because not screening
but “testing” was performed (see the footnote 1 to this Chapter).
The concept of a “screen” has long been accepted (National Acad-
emy of Science, 1975) as implying follow-up with a definite test
(T). That is, screening merely identifies individuals in the general
population or in at-risk population who could benefit from a defin-
itive test not offered otherwise. In prenatal genetic diagnosis, of
which preimplantation genetics is a component, the “test” is an
invasive test, e.g., amniocentesis. The term preimplantation genetic
screening thus implies that it will be followed by an invasive “test.”
The 2016 ICMART and WHO Glossary for reproductive has codi-
fied these terms [30]:
Preimplantation Genetic Testing (PGT)
PGT-A: Aneuploidy.
PGT-M: Monogenic (Single Gene).
PGT-SR: Structural (Chromosomal) Rearrangement.
PGT-HLA: Human Leukocyte Antigen.
4 Selecting the Stage for Analysis of Gamete or Embryonic Stage for Analysis
PGD requires access to DNA from gametes or embryos within

6 days of conception, when implantation occurs. DNA can be
obtained at three stages: (1) polar body I or II or both; (2) cleavage
stage, a single blastomere from the 3 day six- to eight-cell embryos;
and (3) blastocyst, trophectoderm from the 5- to 6-day embryo. All
three are still used, but preferences have changed with respect to
stages preferentially utilized.
4.1 Polar Body Polar body biopsy received the most focus in early human PGD,
Biopsy being espoused by the pioneer Yury Verlinsky. Genotype of the
oocyte can be deduced by the analysis of the first or second polar
biopsy. The underlying principle is fully explained elsewhere
[31]. Briefly, let us suppose the first polar body from a heterozy-
gous individual shows a mutant maternal allele; the complementary
primary oocyte should have the normal allele. Oocytes deduced to

be genetically normal can then be fertilized in vitro to be trans-
ferred for potential implantation. Conversely, a normal polar body
indicates an abnormal oocyte having the mutant allele; thus, fertili-
zation would not proceed. Similarly, if the first polar body showed
no chromosome 21, the oocyte would be presumed to have two
21 chromosomes and, hence, lead to a trisomic zygote. The above
assumes lack of recombination, which if present requires delaying
definitive diagnosis until assessment of the second polar body.
As additional laboratories embarked upon PGD in the late
1990s and early 2000s, relatively few chose polar body biopsy as
their preferred biopsy method. Cleavage stage was more often
utilized. One reason was the obvious disadvantage of being unable
to assess paternal genotype, precluding in part application if the
father had an autosomal dominant disorder, thus making analysis
less efficient in managing couples at risk for autosomal recessive
traits. Another reason seems to be technical difficulty.
Still, polar biopsy retains an indispensable role. Information can
be obtained on genotype of potential offspring from the first polar
body, which is present before fertilization. Only biopsy of the first
polar body allows normal oocytes to be deduced to be fertilized as
normal for single gene diagnosis. Polar body is the only viable
option if regulatory statutes require a provider to inseminate
every oocyte that can be fertilized or restrict the number and
preclude cryopreservation. Identifying and fertilizing only euploid
oocytes allows pregnancy rates to be maintained despite onerous
legislation [32].
Centers experienced in polar body biopsy appear to have preg-
nancy rates comparable to those achieved using blastomere biopsy.
However, no attempts have been made to compare relative safety of
blastomere or trophectoderm biopsy versus polar body biopsy.
Given relatively few centers performing polar body biopsy, any
RTC conducted at present might be misleading if technical exper-
tise were not comparable for both techniques.
4.2 Cleavage Stage Until approximately 2010, most PGT-A cases were performed by
Embryo blastomere biopsy of the cleavage state embryo. The zona pellu-
cida—the glycoprotein layer that surrounds a cleavage stage embry-
o—is breached by mechanical, laser, or chemical means to extract a
cell containing DNA (blastomere). Typically, only a single cell is
removed because even one fewer cell at this stage is believed to
reduce embryo survival. Actually, the frequently cited 10% reduc-
tion in survival after blastomere biopsy was initially deduced by
correlating pregnancy rates as a function of numbers of blastomeres
remaining after thawed cryopreserved embryos [33] were trans-
formed. Loss of one cell after thawing was correlated with10%
reduced pregnancy rates, and loss of two cells even more [34]. In
a later direct study, one experienced center reported livebirth rates
of 22.4% and 37.4% after removal of one versus two cells,

respectively [35].
A notable decrease in number of cleavage stage biopsies per-
formed in the 2007–2010 interval occurred following concerns
related to safety and accuracy. During this interval, fluorescence in
situ hybridization (FISH) using 5–9 chromosome-specific probes
was used for diagnosis. This was at that time the only diagnostic
approach possible for detecting of chromosomal abnormalities;
however, so few chromosomes were obviously not optimal. At
present, 24 chromosome array CGH and NGS-based technologies
are being performed with considerable increase in pregnancy rates.
However, the embryonic stage selected for biopsy is now more
often trophectoderm biopsy.
4.3 Blastocyst Biopsy of trophectoderm in the 5- to 6-day approximately 120-cell

Biopsy blastocyst as stated above is increasingly the approach used at
present. More cells (DNA) can be removed at this stage, potentially
facilitating diagnosis. Trophectoderm forms the placenta; thus,
embryonic cells per se are not removed. In fact, blastocyst was
initially envisioned as the approach for PGD. Gardner and Edwards
recovered blastocysts for PGD in rabbits [3], and in humans Buster
et al. [36] recovered human blastocysts by uterine lavage. Lavage to
recover blastocysts was envisioned to be an approach by which cells
removed by trophectoderm biopsy could allow genetic diagnosis
[37]. Prior to PCR, no other approach seemed feasible. Ultimately,
lavage for PGT was not pursued because of fear of retained, undi-
agnosed embryos.
By 2005 McArthur, Jansen, and their Australian team demon-
strated feasibility of blastocyst biopsy for PGT-A [38, 39], as did
Schoolcraft (2010)** and others in the USA [40]. Usually 5–10
cells are obtained, facilitating diagnosis by next generation
sequencing. The additional 2 to 3 days in vitro beyond that
required for the 8-cell cleavage stage embryo allows self-selection
against non-thriving embryos. Approximately one third of embryos
with chromosomal abnormalities do not survive the day 3 to 5 tran-
sition. Still, PGT-A remains necessary to exclude remaining aneu-
ploidies, the frequency of which depends upon maternal age.
4.4 Embryo Biopsy Cryopreservation of biopsied embryos was once not considered
and Cryopreservation possible. Biopsied embryos thus had to be transferred by day 6;
all diagnostic results had to be completed by that time.
With development of cryopreservation by vitrification, biopsied
embryos could be thawed and transferred with considerable success
[41–43]. In fact, a viable approach is blastocyst biopsy and vitrifica-
tion of all embryos (“freeze all”). In the intervening time, 24 chro-
mosome and other diagnostic applications can be performed. One
may then thaw and transfer normal embryos one or more months
later. “Freeze-all” approaches with later transfer may even result in
higher pregnancy rates than “same-cycle” transfer. The presump-

tive explanation is a more hospitable endometrium, no longer
perturbed by ovarian hyperstimulation.
5 Single Gene Disorders (PGT-M)
PGT-M was the original intent of PGD. Perhaps 20% or more of all
PGT cases currently still involve couples at risk for one or more
single gene disorders. PGT-M can be performed whenever the
chromosomal location of the gene causing a given disorder is
known. The causative mutation need not be known so long as
affected and unaffected family members are available to determine
phase (cis/trans) of markers linked to a mutant allele. This allows
one to deduce whether a given embryo has or has not inherited the
gamete containing the mutation. Close to 500 different conditions
have now been tested worldwide.
By 2009, Reproductive Genetic Innovations (Chicago) had
tested over 202 different conditions [44], the most frequent
being hemoglobinopathies, cystic fibrosis and fragile X syndrome;
in 2010 Verlinsky and Kuliev [45] enumerated disorders tested by
that time. The latest tabulation from this group came from
Rechitsky et al. [46], 2982 cycles involving 1685 patients. These
yielded 1095 pregnancies and 1118 live births; 47 pregnancies were
still ongoing at the time of publication [47].
In Europe, the ESHRE PGD Consortium [29] reported that in
2005 cycles the most common single gene indications to be myo-
tonic dystrophy (N ¼ 76 of 500), Huntington disease (N ¼ 56),
cystic fibrosis (N ¼ 55), fragile X syndrome (N ¼ 51), spinal
muscular atrophy (N ¼ 27), tuberous sclerosis (N ¼ 15), and
Marfan syndrome (N ¼ 13); β-thalassemia/sickle cell anemia
(N ¼ 61). A total of 110 other conditions had by that time been
interrogated. Several disorders seem more common in this cohort
than in the ESHRE PGD consortium than RGI, examples being
neurofibromatosis and familial dysautonomia; however, the spec-
trum of disorders tested was overall similar.
Gutierrez-Mateo et al. reported the Reprogenetics experience
over a 51-month period ending March 2008 [48]. A total of
162 couples in 224 cycles were at risk for 46 different single gene
disorders. Biopsy was performed in 59 different ART centers, blas-
tomeres being sent from IVF center to Reprogenetics labs for
laboratory analysis. By far the most common indication was cystic
fibrosis (N ¼ 73 cycles).
5.1 Unique As already discussed, only PGT allows information to be obtained

Indications for Single prior to implantation, thus obviating necessity for pregnancy ter-
Gene Disorder mination. In the 2001 ESHRE PGD Consortium, 36% of 1561
couples stated that their reason for undergoing PGD was avoiding
5.1.1 Avoiding Clinical clinical pregnancy termination [49]. Some were in their initial
Termination (PGT) pregnancy, but others probably reflected the disquiet expected of
any couple having previously undergone pregnancy termination. In
the 2001 report, 21% of the absolute number of women under-
going PGD had a previous termination and wished to avoid
another. Since 2001, PGT-A has increased considerably in fre-
quency and, hence, the proportion (but probably not absolute
number) of women undergoing PGT-M to avoid clinical pregnancy
termination has decreased.
Increasing number of PGT-M is presently performed for
conditioned determined by de novo mutations (DNM). Develop-
ment of special PGD designs is always required for DNM, detected
in a parent or affected children, as neither origin, nor relevant
haplotypes are available for tracing the inheritance of DNM in
single cells biopsied from embryos or oocytes. With the improved
awareness of PGT, increasing number of couples request PGT,
without any family history of the genetic disease that has been
first diagnosed in one of the parents or in their affected children.
The first systematic PGD for DNM was reported by RGI, describ-
ing a large series of 38 different genetic disorders, determined by
dominant, recessive, and X-linked DNM, of both maternal and
paternal origin [50]. PGD strategies for these families depend on
the origin of DNM, and included an extensive DNA analysis of the
parents and affected children prior to PGT, with the mutation
verification, polymorphic marker evaluation, whole and single
sperm testing and PB analysis in order to establish the normal and
mutant haplotypes, without which PGT-M cannot be performed.
In cases of DNM of paternal origin, the DNM is first confirmed on
the paternal DNA from blood and total sperm, followed by single
sperm typing to determine the proportion of sperm with DNM and
relevant normal and mutant haplotypes. For a higher reliability of
testing, the relevant maternal linked markers are also detected, to
be able to trace for possible shared maternal and paternal markers.
In cases of DNM of maternal origin, DNM is first confirmed in
maternal blood, and PGD was performed, when possible, by PB
analysis, to identify the normal and mutant maternal haplotypes.
Also, in order to trace the relevant paternal haplotypes, single
sperm typing is performed, whenever possible, for avoiding misdi-
agnosis caused by possible shared paternal and maternal markers. In
cases of DNM detected first in children, the mutation is verified in
their whole blood DNA, followed by testing for the mutation in
paternal DNA from blood, total and single sperm, if the DNM
appears of paternal origin. In DNM of maternal origin, PGD is
performed by PB approach, with confirmation of the diagnosis by
embryo biopsy, if necessary. So despite the complexity of PGT-M

for DNM, the applied strategies are highly accurate, representing
an important addition to the practice of PGT-M for Mendelian
diseases, as it makes now possible to offer PGD to any couple at risk
for producing the offspring with genetic disease, despite the tradi-
tional requirement of family data.
5.1.2 Non-Disclosure of PGT-M is the only practical approach if a person at risk for an adult-
Parental Genotype (PGT-M) onset disorder wishes to avoid transmitting to his/her offspring a
mutant gene he/she may (or may not) [51]. In addition, they wish
to remain unaware of his/her own genotype. Such an individual can
still avoid transmitting a mutant allele to offspring without learning
their own genotype. The prototypic example is the son or daughter
of a parent known to be affected with an autosomal dominant
disorder, for example Huntington disease or autosomal dominant
Alzheimer disease.
All this PGT-M and transfer of a genetically normal embryo
without knowledge of parental genotype can thus be accomplished.
The caveat is that the couple needs to remain clueless as to details
on their own ART cycle—number of embryos, number available for
transfer, certainly diagnosis per embryo. In subsequent cycles, the
protocol must then be repeated even if the person in question has
proved unaffected. Otherwise, the at-risk patient would readily
deduce his or her genotype.
The number of PGT-M cases performed for non-disclosure is
about 5 to 10% of single gene disorders in the ESHRE PGD
Consortium and in large U.S. centers. Not significantly different
proportion of cases has been at RGI, mainly for HD (61 HD of a
total 1685 patients) [47].
5.1.3 Cancer and Adult- Performing prenatal genetic diagnosis for any adult-onset Mende-
Onset Disorders (PGT-M) lian conditions was once considered highly arguable [52]. In the
United States, little controversy now exists, but there remains
reticence in much of Europe. The first PGT case for inherited
adult onset cancer was performed by Verlinsky et al. [53] to avoid
transmitting an adult-onset autosomal dominant cancer involved
Li-Fraumeni syndrome, caused by a p53 perturbation. Diagnosis of
other disorders soon followed and by 2002 the RGI group [54]
had performed 337 cycles for 169 patients, resulting in 113 births
free of predisposition to cancer. PGT-M for BRCA1 and BRCA
2, multiple endocrine neoplasia, familial adenomatous polyposis
(FAP), Li-Fraumeni syndrome, Fanconi anemia, retinoblastoma
and Von Hippel-Lindau (VHL) syndrome, neurofibromatosis
1 (NF1) and neurofibromatosis 2 (NF2), and Gorlin syndrome
are now well accepted indications. Given diagnosis of cancer is
usually known in a person to be at risk or under active clinical
surveillance, non-disclosure PGT-M protocol is usually unneces-
sary; however, this could be applicable if only a less closely related
relative were affected. PGT-M to exclude transmission of any adult

onset autosomal trait potentially affecting offspring a prospective
parent now seems well accepted worldwide.
PGD-M also provides a relief for the at-risk couples for inher-
ited cardiac disease, another group of late onset disorders which
may manifest despite pre-symptomatic diagnosis and follow-up.
The majority of inherited cardiac disorders are dominant, for
which no cure may be administered, because their first and only
clinical occurrence may be a premature or sudden death. One of
such conditions is the familiar hypertrophic cardiomyopathy
(HCM), which clinically manifests at different ages, with no symp-
toms observed for years until provoked by different factors, such as
excessive exercise. PGD-M was applied to different conditions
leading to HCM, including HCM4 and HCM7, which may cause
an asymmetric ventricular hypertrophy and defect in interventricu-
lar septum, with high risk of cardiac failure and sudden death.The
other condition for which PGD is strongly indicated is dilated
cardiomyopathy (CMD), which is an autosomal dominant disease
caused by different mutations in LMNA gene located on chromo-
somes 1. This cardiac disease is characterized by ventricular dilation
and impaired systolic function, resulting in a heart failure and
arrhythmia, which causes premature or sudden death. While the
large phenotypic variability of patients may be determined by dif-
ferent mutations in LMNA gene, differences from one family to
another may be observed within the same mutation, with possible
involvement of skeletal muscles that leads to the muscles weakness.
At RGI, 51 PGD cycles were performed for 14 cardiac diseases,
determined by 23 different gene mutations. This resulted in the
embryo transfer in 44 of 51 PGD cycles, yielding 29 (66%) unaf-
fected pregnancies and birth of 27 healthy, disease predisposition-
free children [55, 56]. Results show that PGD may be a realistic
option for couples at risk for producing offspring with cardiac
disease, determined by inherited predisposition. Inheritance of
such susceptibility factors places the individual at risk of serious
cardiac disease, clinically manifested from as early as the first year
of life such as in cardioencephalopathy, to as late as later in life, with
the only clinical realization of premature or sudden death, as in
CMD and CMH.
Among the conditions in the family history of the couples at
risk that may indicate to a possible need of PGD may be a heart
attack and sudden death at young ages, family members with pace-
makers or internal cardiac defibrillators, arrhythmia, and heart
surgery. The chances that the offspring of these patients will
develop the same heart disease will differ depending on the mode
of inheritance, but their penetrance is difficult to predict, because
many inherited cardiac conditions are difficult to diagnose and will
develop with age and may be induced by certain medications or
activities, such as excessive exercise, which may lead to cardiac arrest
or sudden death, justifying the parents’ requests for PGD-M.
5.1.4 HLA-Compatible Among sibs, one in four is HLA compatible (identical).

Embryos (PGT-HLA) HLA-compatibility among siblings becomes attractive if there
exists an older, moribund sibling with a lethal disease who could
benefit from stem cell transplantation to repopulate his or her bone
marrow. PGT-HLA is the only practical way to identify the few
embryos that are HLA compatible with an affected individual
[57, 58]. Sometimes, the affected individual is a child of a couple
of reproductive age. Given this, one attractive strategy is using
umbilical cord blood from a neonate to repopulate the bone mar-
row of an older, affected sibling. Odds that a couple can both avoid
another genetically abnormal child for an autosomal recessive dis-
order (3/4) and also utilize HLA compatible umbilical cord stem
cells to repopulate a bone marrow is 3 in 16 (1 in 4
HLA-compatible embryos multiplied by the 3 in 4 likelihood of
also being affected ¼ 3/16). Actually, the likelihood is less because
recombination can occur within the HLA locus, established by RGI
to be 4.3% [57]; thus a 100% HLA match is not possible for the
affected child because the exact recombination event cannot be
expected to recur. A similar proportion of embryos recovered in
any cycle will not be HLA identical.
PGT-H for the transferring HLA-compatible embryos was first
performed by Verlinsky and colleagues in a couple of at risk for
Fanconi anemia [58]. Like inherited PGT-HLA for autosomal dis-
orders like cancer, this generated considerable controversy at the
time (2001). By 2004, however, the RGI group reported 45 cycles
for HLA typing had been performed [2, 59]; 17.5% embryos were
genetically suitable for transfer, very near the expected (3/16)
(16%). The most common genetic indication is β-thalassemia; how-
ever, the strategy is applicable for any bone marrow infiltrative
disorder [60].
Testing for HLA-compatible embryos without risk of genetic
disease is now also widely accepted in United States and Turkey,
with the prime indication being an older sib with sporadic Black-
fan–Diamond anemia or leukemia, first performed in 2004 [61]. In
the United States, this now accounts for approximately one third of
PGD-HLA cases. A 2004 compilation by RGI showed 11% of their
single cell PGD cases involved HLA testing. By 2009, 15% of 1666
single gene PGD cases were performed for HLA typing [44]. In
Istanbul, 261 HLA cases had been performed by 2009, all but
61 for β-thalassemia [61]. In Europe, however, PGT-HLA for
this indication has remained uncommon [28]. Because the majority
of patient requesting PGT-HLA are of an advanced reproductive
age, PGT-HLA is now increasingly performed concurrently with
PGT-A, and it was demonstrated that PGT-M or PGT-H concur-
rent with PGT-A increases pregnancy rates by 20% or more com-
pared to age-matched cases (<35 years) not concurrently
undergoing PGT-A [46].
By the present time, the list of disorders for which PGT-HLA

has been applied with successful stem cell transplantation treatment
includes thalassaemia, Wiscott–Aldrich syndrome, X-linked hyper
IgM syndrome, X-linked hypohidrotic ectodermal dysplasia with
immune deficiency and incontinentia pigmenti, and Blackfan–Dia-
mond anemia, with the list continuously expanding [2, 57–60,
62–65]. The accumulated experience of approximately one thou-
sand cases of PGT-HLA shows that this represents an important
prospect for radical treatment of the affected members of the
families with congenital and acquired bone marrow failures and
immunodeficiencies. Still the majority of PGT-HLA was performed
for thalassemia, with over 400 cases performed in Istanbul and
Chicago, resulting in birth of dozens unaffected HLA matched
children, who served as donors for stem cell transplantation to
their affected sibling, resulting in a complete recovery in almost
all the cases.
6 Chromosomal Rearrangements (PGT-SR)
Chromosomal rearrangements (translocation or inversion) are well

recognized as leading to unbalanced gametes and, hence, an unbal-
anced zygote. Couples with rearrangements are often detected after
repeated spontaneous abortions, reflecting lethality conferred by
unbalanced gametes. Not long after FISH became available and
PGT possible, PGT-SR was predictably performed, initially using
chromosome-specific probes [19, 66]. This was recognized to be
an attractive approach for assuring transfer of only cytogenetically
normal embryos. Euploid embryos not only precluded another
miscarriage and avoided abnormal liveborn, but increased repro-
ductive efficiency. Relatively few gametes/embryos of any translo-
cation heterozygote are normal or balanced. PGT-SR can identify
and transfer only the relative few embryos that are normal.
Breakpoint-specific probes were initially devised unique to each
case [18]. PGD-SR translocation analysis using this approach was
accurate, but costs prohibitive [18]. Thus, commercially available
probes are now used. A limitation is that usually one cannot distin-
guish an embryo with a balanced translocation from an embryo
without a translocation.
No RCTs have been conducted to determine whether PGT-SR
increases pregnancy rates in couples having a balanced translocation
who present with repeated pregnancies losses. However, surrogate
evidence is ample. Otani et al. [67] observed only 5.3% miscarriages
after PGT-SR for translocations, much less than expected even in
the general population. Second, pregnancy can be achieved more
rapidly. Otani et al. reported the lifetime cumulative pregnancy rate
using PGT-SR to be 57.6% [67]. This is per se not remarkable, but
that this was achieved with an average of 1.24 cycles is unexpected

for translocation heterozygotes. The short time-frame before
which translocation couples undergoing PGD can achieve preg-
nancy contrasts with the mean 4–6 years necessary for translocation
couples who do not use PGD [68–70]. In 2008, the Society for
Assisted Reproductive Technology (SART) guidelines thus recom-
mended PGD for this indication [71].
7 PGT-Aneuploidy (PGT-A) to Improve Pregnancy Rates
PGT aneuploidy testing is increasingly performed solely to improve

pregnancy rates in women who require ART and have no other
indication. The logic is irrefutable. Pregnancy rates in ART decline
precipitously beginning late in the fourth decade; the primary
reason is high embryonic loss due to aneuploidy; thus, transferring
only euploid embryos is attractive, and capable of being assured by
PGT-A. That endometrial factors were not paramount had, more-
over, been shown by successful pregnancies following transfer of
donor embryos from younger women to recipients in their late fifth
and sixth decades. From the late 1990s onward, favorable results
for PGT-A were reported frequently from experienced centers
worldwide [31, 72–77]. Most reports compared outcome to his-
torical expectations for age-matched women not undergoing PGD.
Two small RCTs showing favorable results were conducted in the
United States [78, 79], both showing increased pregnancy rates
although neither sufficiently powered.
During 2000–2007, larger PGD and ART centers in the
United States and Europe increasingly offered PGD to improve
pregnancy rates in older women. However, these large centers were
not able to complete an RCT. Patients were not willing, and in the
U.S. embryo research could not be funded federally (National
Institutes of Health). Other centers, with less volume and experi-
ence, did conduct RCTs. Results showed no significant improve-
ment in pregnancy rates [80–83], and at least one [81] showed
harmful effects. Questionable technical prowess, diagnostic diffi-
culties, and arguable indications were widely criticized by this
author and others [84–87]. Not surprisingly however, the increas-
ing trajectory of increasing PGT-aneuploidy cycles to improve ART
pregnancy rates was blunted. Perhaps a 20% drop in PGT-A cycles
occurred during the years 2007–2010.
Two explanations were commonly accepted as explanations for
the disappointing RTC results. One is the technical difficulty in
performing cleavage stage embryo. Some embryologists appear
more skilled than others, not surprisingly for any surgical proce-
dure. (If so, this explanation would not refute proof of principle for
the value of PGT-A but would impede generalizability). Similarly,
diagnostic methods available at the time were less than optimal.
Using FISH with chromosome-specific probes, only 5–7 chromo-

somes or at best 9–12 could be interrogated.
The single RCT generating the most publicity concerning
harmful effects of aneuploidy testing was conducted by Masten-
broek et al. [81]. This well-cited report warrants a few comments.
One major problem was applying an intent to treat statistical
design. When PGT-A was successful (defined as seven chromo-
somes tested and at least one euploid embryo transferred), the
pregnancy rate was 16.8% per embryo. When no biopsy was per-
formed (true control), the pregnancy rate was 14.7% per embryo,
13% lower than with PGT-A. In some 20% of embryos, however,
there were no diagnostic results, a rate higher than in other centers
at the time [85, 86, 88]. Given lack of results in these 20%, a third
(unintended) group evolved: biopsy performed, no diagnostic
result, transfer nonetheless. In this group, the pregnancy rate was
only 6%. Intent-to-treat statistical analysis dictated that all cases
remain assigned to their original group even if the assigned proto-
col (i.e., PGD) was not completed or even reversed. Insisting on
this, the true PGD group (16.8% pregnancy per embryo) was
merged with the de facto sham group (6%). This yielded a merged
“PGD” live birth rate of 24% per cycle, compared to the statistically
higher 35% in the non-biopsied, non-PGD group.
Given this situation, efforts were directed in the cytogenetic
community to increase diagnostic robustness and accuracy. Technol-
ogies introduced included multiple hybridization cycles to test all
24 chromosomes, testing 11–12 chromosomes tailored expressly
per age group, orre-testing to minimize “no result” cases. Still, the
obvious need was reliable 24 chromosome testing. Actually, genome-
wide molecular approaches for 24 chromosome analysis had begun
earlier with metaphase CGH [88–90]. Later, Wells et al. [91, 92]
used metaphase (CGH) to interrogate all chromosomes in a small
series of blastocysts: 90% gave informative results, and 36 to 42 cycles
(86%) resulted in a clinical pregnancy. By contrast, non-PGD blasto-
cysts in the same series showed a 60% pregnancy rate. Single cell
genome-wide SNPs (in 2000–2010) were also used reliably on a
single blastomere [93–95]. Results were available quickly enough
for normal embryos to be transferred in the same cycle: using SNPs.
Treff et al. [96] accomplished this in 4 h.
The technique that ultimately became used most commonly for
24 chromosome interrogation proved to be array CGH, using a
BAC-platform (BlueGnome). BlueGnome utilized selected
sequences from all 24 chromosomes, placed on the proverbial
“chip.” Reference sequences were derived from bacterial artificial
chromosomes (BACs), which cover very large chromosomal
regions. This method provides less sensitivity than arrays that utilize
smaller sequences and thus broader “coverage.” That PGT-A used
BAC arrays paradoxically proved advantageous clinically. Results on
a single blastomere could be stratified simply into euploid,
aneuploid, or no result bins. Of course, it was realized that some

aneuploidy embryos were actually mosaic and containing an unrec-
ognized euploid cell line, yet undetectable given analysis restricted
to a single blastomere. The single cell tested could have been
trisomic whereas the remaining 7–8 cells unknowingly euploid,
(i.e., mosaicism). Still, clinical utility was pragmatically achieved
with BlueGnome. Based on data from disaggregated cleavage
stage embryos, the frequency of mosaicism was concluded to
1–2% [97].
By the 2010s there had been further transformative movement
in expanding use of PGT-A [40, 98]. One advance is moving from
cleavage stage biopsy to trophectoderm biopsy. Improved embryo
culture, facile laser trophectoderm biopsy, and vitrification were
combined to make this feasible. By 2012 several different groups
had conducted RCTs with salutary results. These RCTs observed
improved ART pregnancy rates by transferring euploid
embryos vs. simply morphologically normal embryos. Harton
et al. definitely showed that with euploid embryos the “take home
baby” rates did not begin to decrease until age 42 years [99]. Multi-
ple gestations could be avoided because transfer of a single euploid
embryo protected livebirth rates (without twins) equal to transfer of
two morphologically normal non-PGD embryos [100, 101].
BAC Array CGH by next generation sequencing (NGS) tech-
nologies had encompassed SNPs, copy number variants (CNVs),
and quantitative PCR. With NGS, greater laboratory productivity is
possible; however, mosaicism is detected more often. Increased
detection of mosaicism thus occurs, presumably reflecting both
greater sensitivity of NGS and greater amounts of DNA. In the
six-day embryo the number of cells biopsied is 5–15; in cleavage
stage blastomere biopsy only one cell was available. Not surpris-
ingly, mosaicism was detected more often in trophectoderm biopsy.
Mosaicism can be detected at level of 20% [97]. The frequency of
mosaicism in trophectoderm is 20–25% using validated NGS tech-
nologies; the frequency on reanalyzed cleavage stage embryos was
<2% [97]. Although some mosaic embryos may also be mosaic as
well in this inner cell mass (to be developing embryo), other
embryos could have normal cells only in their ICM and thus
would be suitable for transfer.
It follows that trophectoderm biopsy results characterized by a
normal and a monosomic line could be candidates for transfer,
given that the monosomic line is probably lethal and would not
persist [102]. Greco et al. reported livebirths in four mosaic mono-
somy cases but none in mosaic trisomies [102]. At present, 7%
mosaic aneuploid embryos by Munné et al. [97] are estimated as
leading to a livebirth [102]. If a couple lacks non-mosaic euploid
embryos, one might thus perform transfer of a mosaic aneuploid
embryo with patient consent. The need for clarity is recognized by
Commentaries [99, 103–105].
2016 Guidelines by PGDIS (http://www.pgdis.org/docs/

newsletter_071816.html) provide recommendations on signifi-
cance of a given level of mosaicism, on transferring mosaic mono-
somy embryos, and on priority of transferring mosaic trisomy
embryos for a specific chromosome.
8 Repeated Pregnancy Loss (Recurrent Aneuploidy)
Observational studies utilizing PGT-A have long been accepted as

decreasing miscarriage rates [31, 72–77]. No RCTs exist, but only
surrogate data exist. One can compare results to objective criteria,
such as the Brigham formula [106]. This formula takes into
account maternal age and the number of prior abortions to derive
the priori likelihood of a pregnancy loss. Using this comparison,
Munné et al. [107] observed losses in 13% of couples who used
PGT-A, compared to the Brigham-expected rate of 33%. Benefit
was, not surprisingly, greatest for women older than 35 (39 vs.
expected 13%; P > 0.001). Thus, using array CGH, and transferring
euploid embryos, the miscarriage rate is now 10% [99].
Recurrent miscarriage has multiple etiologies—genetic and
nongenetic. How might the above data apply to couples having
experienced recurrent miscarriage, but not actually infertile?
Non-random distribution appears to exist with respect to successive
miscarriages. That is, abortuses tend to be either successively aneu-
ploid or successively euploid [108]. Non-random distribution also
occurs in preimplantation embryos in successive cycles [109].
Given all the above, the rationale for performing PGT aneu-
ploidy testing in women having repeated miscarriages is strong if
the goal is preventing recurrent miscarriage. Less clear is whether
the livebirth rate is improved and if so whether this justifies ART
with PGT-A. Successive aneuploidy carries implications for diag-
nostic testing and management. However, there is not consensus
[110]. If recurrent aneuploidy is an indication for PGT aneuploidy,
at least one loss should have documented aneuploidy if one is to
perform PGT-A solely for this purpose.
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mosaic aneuploid blastocysts. N Engl J Med
Chapter 3
Noninvasive Approaches to Prenatal Diagnosis: Historical

Perspective and Future Directions
Lisa Hui
Abstract
The field of prenatal screening and diagnosis has undergone enormous progress over the past four decades.
Most of this period has been characterized by gradual improvements in the technical and public health
aspects of prenatal screening for Down syndrome. Compared to the direct analysis of fetal cells from
amniocentesis or chorionic villus sampling, noninvasive approaches using maternal blood or ultrasound
have the great advantage of posing no risk of miscarriage to the pregnancy. Recent advances in molecular
genetics and DNA sequencing have revolutionized both the accuracy and the range of noninvasive testing
for genetic abnormalities using cell-free DNA in maternal plasma. Many of these advances have already been
incorporated into clinical care, including diagnosis of fetal blood group and aneuploidy screening. The
accelerated pace of these recent developments is creating not just technical and logistical challenges, but is
also magnifying the ethical and public policy issues traditionally associated with this field.
Key words Noninvasive prenatal testing, Noninvasive prenatal diagnosis, Cell-free DNA, Fetal cells,
Nuchal translucency, Serum screening, Next-generation sequencing
1 Historical Perspective of Noninvasive Prenatal Diagnosis
The field of noninvasive prenatal screening and diagnosis has

undergone enormous progress over the past four decades (Fig. 1).
Compared to the direct analysis of fetal cells from amniocentesis or
chorionic villus sampling, noninvasive approaches using maternal
blood or ultrasound have the great advantage of posing no risk of
miscarriage to the pregnancy.
1.1 Screening for Prenatal screening for fetal anomalies has traditionally focused on
Chromosome chromosome abnormalities because they are major causes of perina-
Abnormalities by tal morbidity and mortality. Aneuploidies are also amenable to
Maternal Age invasive prenatal diagnosis from fetal cells using standard cyto-
genetic techniques, allowing a definitive diagnosis before 20 weeks
gestation. The first fetal diagnosis of Down syndrome was per-
formed using cultured amniotic fluid cells in 1968 [1].
45
46 Lisa Hui
Fig. 1 Timeline of prenatal testing for fetal aneuploidy AFP alphafetoprotein, US ultrasound, NIPT noninvasive
DNA-based prenatal testing
Down syndrome, or trisomy 21, is the most common autosomal

aneuploidy leading to live birth and consequently is the most impor-
tant genetic cause of intellectual disability. As invasive testing with
amniocentesis or chorionic villus sampling (CVS) carries an appre-
ciable risk of miscarriage, estimated at 0.5 to 1.0% [2], it has been a
long-standing goal of prenatal diagnosis to develop a noninvasive
test for trisomy 21.
The most important risk factor for trisomy 21 is maternal age.
The risk of an affected newborn at term is approximately 1 in
300 for a woman aged 35 years, increasing to 1 in 100 by the
maternal age of 40 years [3]. Consequently, the prevalence of
trisomy 21 has increased due to trends to later childbearing in
most developed counties. The estimated birth prevalence of tri-
somy 21 is now around 1.4 to 1.7 per 1000 births [4, 5].
For several decades, screening for Down syndrome was based
on maternal age alone, with women at increased risk (typically using
an age threshold of 35 years or more) being offered invasive prena-
tal testing. This method only detects 30–40% of all fetuses with
trisomy 21 as the majority of affected pregnancies occur in women
less than 35 years old. This aged-based approach also exposes older
women to high rates of invasive testing (about 50% uptake) for a
relatively low diagnostic yield.
1.2 Second In the late 1970s, maternal serum screening for neural tube defects
Trimester Screening was introduced into clinical practice [6]. From this screening pro-
for Aneuploidy Using gram arose the observation that maternal serum alpha fetoprotein
Maternal Serum (AFP) was reduced in pregnancies affected by chromosome
Biochemical Markers abnormalities [7]. This serendipitous discovery represented a land-
mark in prenatal screening for aneuploidy [8]. Subsequently, other
Historical Perspective of NIPT 47
serum markers of aneuploidy were discovered and combined with

AFP to improve the performance of serum screening for both
trisomy 21 and 18. The addition of human chorionic gonadotropin
(hCG) [9] and unconjugated estriol [10] to AFP measurement
created the “triple test” [11]. This could identify approximately
70% of pregnancies affected by trisomy 21 for a 5% screen-positive
rate [12, 13]. The later addition of inhibin-A created the “quadru-
ple test,” increasing detection rates to approximately 80% [12]. Sec-
ond trimester serum screening thus allowed women of all ages to
have a risk assessment for aneuploidy, not just those women of
advanced maternal age. Screening was confined to a specific gesta-
tional age window (ideally 15–18 weeks), and those who received a
high risk result (arbitrarily set at around 1/300 risk or higher) were
offered a definitive diagnosis with an invasive test.
1.3 Second The field of prenatal ultrasound underwent major developments in

Trimester Ultrasound parallel to those of serum screening in the 1980s. A second trimes-
Screening ter morphology ultrasound for fetal structural abnormalities
for Aneuploidy became a routine part of prenatal care, enabling the detection of
aneuploid fetuses with structural malformations. Major structural
malformations that are strongly associated with Down syndrome
include atrioventricular septal defects, tetralogy of Fallot, duodenal
atresia, cystic hygroma, and hydrops fetalis [14]. The presence of a
major anomaly is an indication for invasive testing, however the
utility of second trimester ultrasound as a stand-alone screening test
for trisomy 21 is limited by its relatively low sensitivity [15].
The second trimester morphology scan also can detect anato-
mical variants known as “soft markers” that are present in increased
frequency in aneuploid fetuses. The sonographic “soft markers” of
Down syndrome include increased nuchal fold, hypoplastic nasal
bone, echogenic bowel, and short femur. Individually, these mar-
kers have poor sensitivity and specificity for Down syndrome, but
their predictive value increases if multiple markers are present
[16]. These markers have been used to calculate likelihood ratios
for an affected pregnancy [17], but the use of this approach is now
being questioned in the era of widespread first trimester screening
for aneuploidy [18].
1.4 First Trimester In the early 1990s an important observation that first trimester
Screening with fetuses with excessive fluid accumulation in the neck were more
Biochemical and likely to be aneuploid was reported [19]. The measurement of the
Ultrasound Markers fetal nuchal translucency thickness at 11–13 weeks gestation was
subsequently established as the single most powerful sonographic
marker of trisomy 21 [20]. For a fixed screen positive rate of 5%, and
a risk cutoff of 1 in 300, up to 80% of affected pregnancies could be
detected by the combination of nuchal translucency measurement
and maternal age. The added attraction of this test was the earlier
stage of diagnosis compared with second trimester serum screening.
48 Lisa Hui
The performance of the first trimester ultrasound was

improved by the addition of first trimester serum markers. In
combination with maternal serum measurement of free β-hCG
and pregnancy-associated protein-A (PAPP-A), detection rates of
85–90% could be achieved for a false positive rate of 5%
[12, 13]. Risk results for trisomy 13 and 18 were also incorporated
into the first trimester combined screening algorithm. This first
trimester combined test has now largely replaced second trimester
maternal serum screening as the standard of care in most developed
countries, due to its dual advantages of improved accuracy and
earlier diagnosis.
Other first trimester sonographic markers of aneuploidy have
been developed, including assessment of the nasal bone [21], duc-
tus venosus waveform [22], and tricuspid valve flow [23]. The
addition of these markers to the first trimester combined test
increases detection rates to 96% and lowers the false positive rate
to 2.5% [24]. However, the practical barriers to universal imple-
mentation of these advanced ultrasound markers are even greater
than those faced by nuchal translucency measurement alone and as
such they have not been uniformly adopted outside of specialist
centers [25].
1.5 Combined First Many models have been investigated to determine the utility of
and Second Trimester combining first and second trimester screening strategies. These
Screening Strategies include the integrated, stepwise, and contingent screening
approaches [12, 13, 26]. However, serial testing with independent
risk results in first and second trimesters is not generally recom-
mended due to the higher cumulative false positive rate.
2 DNA-Based Tests Using Maternal Blood
Fetal cells and cell-free nucleic acids derived from the placenta are
consistently detectable in maternal blood. This phenomenon pro-
vides the opportunity to directly sample fetal genetic material in
maternal blood for noninvasive DNA-based testing. It is in this area
that the most exciting recent advances in noninvasive testing have
taken place.
2.1 Fetal Cells in The presence of fetal cells in maternal tissues is a long-recognized
Maternal Blood phenomenon, first described in 1893 when syncytiotrophoblastic
cells were documented in the lungs of women who had died from
eclampsia [27]. Fetal lymphocytes were detected in maternal blood
many years later in 1969 and their potential for prenatal diagnosis
was immediately recognized [28]. However, it was several decades
until technology could capitalize on this fetomaternal cell traffick-
ing for the purposes of prenatal diagnosis. In 1990 DNA was
successfully isolated from circulating fetal cells in maternal blood
to diagnose fetal sex [29]. It was also discovered that some fetal
cells, namely lymphocytes, could remain in the maternal body for
decades postpartum [30], potentially confounding prenatal testing
results from subsequent pregnancies. Fetal erythroblasts, or nucle-
ated red cells, were therefore considered the most suitable for
noninvasive prenatal diagnosis (NIPD) because they are detectable
from early pregnancy, have short life spans, and can be distin-
guished from maternal cells using embryonic hemoglobin markers.
However, fetal cells are only present in very small numbers in
maternal blood despite enrichment protocols [31] and they are
difficult to culture without maternal cell contamination [32]. The
first major attempt to ascertain the clinical feasibility of using fetal
cells for prenatal diagnosis was the NIFTY study [33]. This study
reported a sensitivity rate for aneuploidy of 74.4% with a false
positive rate 0.6–4.1% and a fetal gender detection rate of 41.4%.
The investigators concluded that significant technical challenges
needed to be overcome before it was a feasible method for clinical
practice. A variety of enrichment methods have continued to be
explored, including fluorescence-activated cell sorting, magnetic-
activated cell sorting, density gradient centrifugation, and more
recently, nuclear reprogramming of fetal erythrocytes [34].
2.2 Cell-Free DNA in The detection of cell-free fetal DNA in maternal plasma in 1997
Maternal Plasma was a watershed moment in noninvasive prenatal testing [35]. The
major source of circulating cell-free fetal DNA is the trophoblast,
which releases DNA into the circulation as a result of apoptosis
[36–38]. Cell-free DNA of placental origin is detectable as early as
5 weeks of gestation, prior to the establishment of the fetal circula-
tion [39], and it is clearly rapidly after delivery [40, 41]. These
circulating nucleic acids have been the subject of remarkable
achievements in the past decade.
2.2.1 Obstetric Obstetric complications due to abnormal placentation may be

Outcomes Using accompanied by altered levels of circulating nucleic acids of placen-
Quantitative Analysis of tal origin due to increased trophoblast microparticle release
Cell-Free Fetal DNA [42]. Quantitative changes in total cell-free fetal DNA in maternal
plasma are potential predictors of adverse obstetric complications
such as preeclampsia and intrauterine growth restriction
[43–45]. Early studies quantifying circulating fetal DNA were
based on the detection of Y chromosome-specific sequences, but
this approach was limited to pregnancies in which the fetus was
male. The lack of a gender-independent, fetus-specific DNA marker
was a major barrier to clinical translation. Sequences that are differ-
entially methylated in the fetus compared with the mother, such as
MASPIN [46] or the promoter region of RASSF1A [47], have
been successfully used as universal fetal DNA markers; however,
the technical challenges of studying differential methylation are
considerable. An alternative approach to study obstetric disorders
50 Lisa Hui
using cell-free nucleic acids is to use RNA markers of placental

origin [48–50], but this appears some way from application in a
clinical setting [45]. More recently, mechanisms involving the
TLR9 receptor that link alterations in cell-free DNA levels and
parturition have been postulated [51]. This suggests future avenues
for non-genetic obstetric disorders such as preterm birth to be
investigated using cell-free fetal DNA.
2.2.2 Noninvasive The term noninvasive prenatal diagnosis (NIPD) is generally used
Prenatal Diagnosis Using to refer to DNA-based tests of fetal health that do not require direct
Cell-Free Fetal DNA access into the uterus and that are sufficiently accurate not to
require follow-up confirmation with amniocentesis or CVS. The
earliest successful clinical applications of cell-free fetal DNA were
for the noninvasive diagnosis of fetal Rhesus D antigen status and
fetal sex.
The detection of a paternally inherited allele in the fetus is a
relatively simple problem of detecting the unique sequence in
maternal blood. The detection of Y-chromosome-specific
sequences in maternal plasma using real-time qPCR was used to
first demonstrate the presence of cell-free fetal DNA in maternal
blood [35]. Medical indications for early prenatal diagnosis of fetal
sex include pregnancies at risk of X-linked disease, or in utero
virilization from congenital adrenal hyperplasia. NIPD using
qPCR for the male-specific SRY or DYS 14 loci is reliable for fetal
sex from 7 weeks gestation, much earlier than is possible for either
invasive testing or ultrasound. The accuracy of NIPD for fetal sex is
very high [52] and can achieve sensitivity >99.5% with stringent
quality control [53]. NIPD for diagnosis of fetal sex has been
clinically available in the UK for over 8 years and has substantially
reduced the number of women undergoing invasive
procedures [53].
NIPD for fetal Rhesus blood group is the other major success-
ful application of cell-free fetal DNA. The basis of this test is the
amplification of the RHD gene in Rhesus D negative pregnant
women to allow early identification of fetuses at risk of hemolytic
disease. NIPD for fetal D antigen status became widely available
more than 10 years ago in the United Kingdom through the
International Blood Group Reference Laboratory [54]. Other
countries in Europe quickly adopted this technology, aided by the
Special Non-invasive Advances in Fetal and Neonatal Evaluation
Network [55]. In Denmark, NIPD testing is now implemented on
a national basis to restrict anti-D immunoglobulin prophylaxis to
those carrying a Rhesus positive fetus. NIPD at 25 weeks of gesta-
tion can detect 99.9% of Rhesus positive fetuses and reduce unnec-
essary prenatal RhD prophylaxis in 97.3% of women carrying an
RhD-negative fetus [56].
NIPD for other single gene disorders has received relatively less
attention than Rhesus typing due to the need for disease-specific or
patient-specific assay development. However, a number of disor-
ders have been detected using cell-free fetal DNA in proof-of-
principle studies, including hemoglobinopathies [57] and fetal/
neonatal alloimmune thrombocytopenia [58]. The use of digital
PCR may improve detection of paternally inherited mutations and
facilitate further clinical applications [59, 60].
However, it is the enormous capabilities of next generation
sequencing that may rapidly advance NIPD for single gene disor-
ders. A recent proof of principle study of the noninvasive diagnosis
of congenital adrenal hyperplasia before 9 weeks gestation using
next generation DNA sequencing suggests that NIPD for a wider
range of autosomal recessive disorders may become a clinical reality
in the near future [61].
2.2.3 Aneuploidy The diagnosis of fetal aneuploidy using cell-free DNA in maternal
Detection Using Cell-Free plasma poses a much greater technical challenge than the detection
DNA of paternally inherited fetal sequences. Detection of fetal aneu-
ploidy from maternal plasma requires the determination of relative
chromosome dosage in the fetus from a mixed population of
maternal and placental cell-free DNA fragments. Furthermore,
DNA fragments of fetal origin are not readily distinguishable
from those of maternal origin. The recent application of next
generation sequencing techniques to this task has led to revolution-
ary advances in noninvasive prenatal testing for aneuploidy.
Sequencing is the determination of the individual base order of
nucleotide bases in a sample of DNA or RNA. “First generation,”
or Sanger sequencing, was the primary method of sequencing
dating from the 1980s. This method is very accurate and can
sequence very long stretches of DNA (500–600 bases per read),
but is relatively slow and not conducive to high throughput. Next
generation sequencing is based on similar approach of synthesizing
DNA from single stranded genomic DNA template but it performs
multiple short reads simultaneously, in a “massively parallel” fash-
ion. In massively parallel sequencing (MPS), only a fraction of each
DNA fragment is actually sequenced. A 36-base pair read is typi-
cally sufficient to map a DNA fragment to a unique location in the
human genome and thus identify its chromosome of origin. Hence,
sequencing for noninvasive prenatal testing for fetal aneuploidy
(NIPT) does not aim to reconstruct the actual fetal or maternal
genome, but simply determines the relative contribution of chro-
mosomes to the cell-free population in maternal plasma. This
approach is based on the assumption that the entire genome is
represented in maternal plasma at a constant relative proportion.
If there is more than expected cell-free DNA originating chromo-
some 21, this is evidence of a pregnancy affected by trisomy 21.
However, the mean proportion of cell-free DNA of placental origin
52 Lisa Hui
in maternal plasma is only 10% at 13 weeks gestation [62]. The high

background level of maternal cell-free DNA means that high preci-
sion counting is required to detect an affected pregnancy.
Since 2011, multiple groups have demonstrated the clinical
validity of a DNA-based sequencing approach for fetal aneuploidy
in high risk women. The various terms noninvasive prenatal testing
(NIPT), noninvasive prenatal screening (NIPS), and noninvasive
DNA-based testing (NIDT) have been coined to distinguish this
test from NIPD, as it is not considered a diagnostic test.
Various methods exist, including MPS of the whole genome
(random sequencing) [63–68], chromosome selective approaches
using targeted assays [69–72], or single nucleotide polymorphism-
based assays [73, 74]. The MPS method has the potential to
provide information across the entire genome, including detection
of whole chromosome and subchromosomal abnormalities [64, 75,
76]. However, targeted approaches have much lower sequencing
requirements as only DNA fragments unique to the chromosomes
of interest are sequenced, and therefore have advantages in cost and
throughput.
The extremely high sensitivity (98.58–100%) and specificity
(97.95–100%) of NIPT outperform any screening method for
aneuploidy to date [77]. It is also effective from 10 weeks gestation
on, with no upper gestational age limit, greatly improving the
flexibility of timing of testing. Professional societies have now
endorsed its use in high risk women, but emphasize that it remains
a screening, not a diagnostic test [78, 79]. Given the relatively low a
priori risk of an affected fetus even in high risk groups, and the
potentially serious consequences of a false positive result, confirma-
tory invasive testing is still required after an abnormal NIPT result
[80]. As test performance is not affected by maternal age, NIPT in
low risk populations appears to perform just as well as in high risk
populations [81–83].
In a direct clinical comparison between standard prenatal
screening and DNA sequencing-based methods, the anticipated
advantages of NIPT were confirmed [84]. Specifically, there was a
much lower false positive rate with cell-free DNA-based testing vs
routine screening for trisomy 21 (0.3% vs. 3.6%), with a
corresponding higher positive predictive value of 45.5% vs. 4.2%.
The major limitation of NIPT is the high cost compared with
current screening techniques. In most countries, this cost has been
directly born by the patient or their private insurer, creating sub-
stantial issues with equity of access. Reductions in invasive testing
rates have been widely reported since the introduction of NIPT
into clinical practice but the many ethical and public health chal-
lenges associated with its implementation remain to be solved
[85, 86].
3 Future Directions
The past 40 years have seen prenatal screening and diagnosis for
aneuploidy become an integral part of pregnancy care. All women,
regardless of background risk, are now offered some form of non-
invasive testing for trisomy 21 [87]. The field has grown to encom-
pass the various disciplines of epidemiology, laboratory medicine,
obstetric ultrasound, clinical genetics, bioinformatics, genomics,
genetic counseling, bioethics, and health economics. The final
result of these developments has been the steady reduction in the
number of women undergoing invasive testing, and an increase in
the prenatal detection rate of Down syndrome [5, 88, 89].
The knowledge accumulated over several decades of screening
with maternal serum biochemistry and ultrasound is now being
rapidly eclipsed by advances in molecular genetics and direct analy-
sis of fetal DNA using noninvasive methods. Noninvasive prenatal
testing is no longer just about trisomy 21, 18, and 13. Noninvasive
DNA-based testing for fetal aneuploidy has already expanded
beyond these three common autosomal trisomies to include fetal
sex, sex chromosome aneuploidy [90], selected microdeletion syn-
dromes [91, 92], and genome-wide abnormalities [75]. It is now
possible to interrogate the fetal genome at the same resolution as a
chromosomal array [93] and to even reconstruct the entire fetal
genome from maternal blood [94–96].
This represents the final convergence of the major themes in
noninvasive prenatal testing from the past decades (Fig. 2). This
paradigm shift has enormous ethical, medical, and resource impli-
cations for the field. We stand at the crossroads where noninvasive
Fig. 2 Converging trends in screening for chromosomal abnormalities

54 Lisa Hui
prenatal testing is posed to move beyond the detection of common

aneuploidies to encompass the whole range of human genetic
variation. The major future challenges in the noninvasive detection
of genetic abnormalities are therefore not simply technical, but
ethical and social.
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Part II
Preimplantation Genetic Testing

Chapter 4
Molecular Testing for Preimplantation Genetic Diagnosis

of Single Gene Disorders
Rebekah S. Zimmerman, Jennifer Eccles, Chaim Jalas, Nathan R. Treff,
and Richard T. Scott Jr.
Abstract
Preimplantation genetic testing has evolved tremendously from the early days of FISH detection for a select
few chromosome aneuploidies to now combining the detection of all whole chromosome imbalances in
conjunction with single gene disorder testing for inherited diseases. As universal carrier screening and
exome or genome studies become more commonplace, more and more families are becoming interested in
reducing the risk of having a child with a severe disease using preimplantation genetic testing. We describe
here the use of quantitative PCR (qPCR) for the custom construction of single gene disorder testing plans
for families, the validation of the probes designed, and the protocol for diagnosing an embryo biopsy. qPCR
has been shown to have the lowest risk of failed amplification and allele dropout and thus the lowest risk of a
misdiagnosis, while also currently providing the fastest protocol to allow for rapid turnaround of results.
Key words Single gene disorder, PGD, IVF, Preimplantation genetic diagnosis, qPCR
1 Introduction
During the earliest stage of preimplantation genetic diagnosis

(PGD), the first patients to benefit were those at risk to have
offspring with single gene disorders (SGD) and first attempts
were aimed at ruling out X-linked disorders by targeting
Y-specific probes [1]. Following those efforts, a successful outcome
using PGD to address an increased risk of cystic fibrosis was
reported [2]. Over 20 years later, the scope of PGD has expanded
to include over 200 single disorders including adult-onset condi-
tions, cancer predisposition syndromes, and HLA genotyping to
identify sibling matches for bone marrow transplants [3, 4]. The
advent of preimplantation genetic screening (PGS) for aneuploidy
further broadened the applications of these technologies [5].
Unlike PGS for aneuploidy, which employs the same testing
paradigm for use in multiple populations, PGD for SGD requires a
case-specific workup that is designed to address the testing variables
61
62 Rebekah S. Zimmerman et al.
associated with each unique gene, mutation(s), inheritance pattern,

and family history. Of particular importance, is whether or not the
laboratory is able to obtain additional samples from informative
family members that are needed to build supporting linkage
probes. This extensive and complex workup is required to address
the diagnostic challenges that are inherent in testing small numbers
of cells. Such challenges include amplification failure, allele drop
out (ADO), and contamination [6, 7]. As a result, the testing
approach for two couples seeking PGD for the same single gene
disorder may be quite different.
Currently, the most rapidly advancing standard for PGD for
SGD involves DNA amplification via quantitative polymerase chain
reaction technology (qPCR) to detect mutations directly (when
applicable) along with amplification of one or more closely linked
polymorphic markers [8]. This strategy is designed to provide both
a direct and indirect means of assessing genotype with the goal of
increasing accuracy by reducing the risk of a false result related to
ADO and increasing the tests ability to detect contamination.
Validation studies have demonstrated the validity and diagnostic
value of this approach and accuracy using this methodology is
greater than 98% [8, 9].
The first step of the process involves creating a custom valida-
tion for a family (Fig. 1) which includes: (1) extraction of DNA
from blood collected from both members of a couple who are
going to be the biological parents of the embryo, (2) possible
extraction of DNA from buccal swab samples collected from perti-
nent family members, (3) SNP microarray to screen multiple SNP
probe candidates simultaneously for SNPs that are linked to the
mutation/gene of interest, (4) selection of SNPs from the micro-
array and ordering of TaqMan PCR primers, (5) testing of the
TaqMan PCR probes on the couple and possible family members
to both determine the accuracy of the probe as well as determining
the configuration of alleles, or phase, for the linkage probes to
ensure correct interpretation of results in the future embryo biop-
sies. When possible the specific mutation(s) are targeted and then at
least one linkage probe is selected upstream and downstream from
the mutation site, usually within 1 Mb. For repeat expansion dis-
orders, or when the mutation probe is not performing up to
standards, a “linkage only” approach is used where at least four
linkage markers are selected (two upstream and two downstream).
In the case of microdeletions, two markers are selected within the
deletion and at least one marker is selected upstream and down-
stream of the deletion of interest.
Once the laboratory has completed the validation process,
genetic material from the embryo can then be analyzed. While it
is possible to perform embryo biopsies during different stages of
embryonic development, there is currently a shift away from cleav-
age stage (blastomere, day 3) biopsy, toward trophectoderm
Molecular Testing for Preimplantation Genetic Diagnosis of Single Gene Disorders 63
1. Obtain DNA from patient and partner
2. Receive buccal swabs from family members

(if needed)
3. Run SNP microarray on patient and partner

(occasionally other family members, too)
4. Select informative SNPs and order Taqman

probes
5. Validate the Taqman probes on parental and

familial samples
Fig. 1 Single gene disorders validation workflow
(blastocyst, days 5–6) biopsy, which has been shown to provide

improved performance with regards to ADO and implantation rates
[9, 10]. Current technology also allows for analysis of single gene
disorders to be performed concurrently with PGS for aneuploidy,
which provides further information regarding whether an embryo
is suitable for transfer [8, 11].
The following report presents the protocol for the workup and
embryo PGD for single gene disorders in a clinical diagnostic
laboratory using real-time quantitative PCR.
2 Materials
2.1 Genomic DNA 1. QIAamp DNA blood maxi kit (Qiagen, Germantown, MD,
Extraction from USA).
Peripheral Blood 2. QIAGEN Protease (Qiagen, Germantown, MD, USA).
3. Prepare the protease by pipetting 5.5 mL distilled water into a
vial of lyophilized QIAGEN Protease. Label the cap with date it
was constituted (see Notes 1 and 2).
4. 100% ethyl alcohol (ethanol).
2.2 Genomic DNA 1. Gentra Puregene Buccal Cell Kit for buffers (Qiagen, German-
Extraction from Buccal town, MD, USA).
Swabs 2. DNeasy Blood and Tissue Kit for collection tubes and spin
columns (Qiagen, Germantown, MD, USA).
3. 100% ethyl alcohol (ethanol).
2.3 TaqMan 1. TaqMan® SNP Genotyping Assays, custom ordered for each
Genotyping PCR for family (Life Technologies, Carlsbad, CA, USA).
Probe Validation 2. Molecular Biology Grade water.
3. TaqMan® Gene Expression Master Mix (Life Technologies,
Carlsbad, CA, USA).
2.4 TaqMan 1. TaqMan® SNP Genotyping Assays, custom ordered for each
Genotyping PCR for family (Life Technologies, Carlsbad, CA, USA).
Embryo Testing 2. Molecular Biology Grade water.
3. TaqMan® Gene Expression Master Mix (Life Technologies,
Carlsbad, CA, USA).
3 Methods
3.1 Genomic DNA Peripheral blood is strongly recommended for patients for which
Extraction from SNP arrays will be performed.
Peripheral Blood
1. Mix Buffer AL from the QIAamp DNA blood maxi kit thor-
oughly by shaking before use.
2. Make sure ethanol was added to buffers AW1 and AW2.
3. Preheat the water bath to 70 C.
4. Pipet 500 μL QIAGEN Protease into the bottom of 50 mL
centrifuge tubes.
5. Add 10 mL blood and mix briefly.
6. Bring the volume of the samples up to 10 mL with PBS
(if necessary) before adding to the centrifuge tube.
7. Add 12 mL Buffer AL, and mix by inverting the tubes 15 times,
followed by additional vigorous shaking for at least 1 min.
8. Incubate at 70 C for 10 min.
9. Add 10 mL ethanol (96–100%) to the samples, and mix by
inverting the tubes 10 times, followed by additional vigorous
shaking.
10. Insert a VacConnector into each VacValve on the QIAvac
manifold that is to be used. Insert the QIAamp Maxi column
into the VacConnector. Open the corresponding VacValve.
Close the main vacuum valve and switch on the vacuum pump.
11. Carefully transfer one half of the solution from the previous
step onto the QIAamp Maxi column, taking care not to
moisten the rim. Open the vacuum valve. If sample flow rates
differ, close the VacValves where the lysate has already passed
through. After all lysates have been drawn through the col-
umns, close the main vacuum valve. Repeat this step.
12. Open the VacValve, and add 15 mL Buffer AW1 to the

QIAamp Maxi columns, taking care not to moisten the rim.
Open the main vacuum valve. After all of the Buffer AW1 has
been drawn through the columns, close the main vacuum
valve.
13. Open the VacValve, and add 15 mL Buffer AW2 to the
QIAamp Maxi columns, take care not to moisten the rim.
Open the main vacuum valve. After all of the Buffer AW1 has
been drawn through the columns, close the main vacuum
valve.
14. To dry the membranes, place the QIAamp Maxi column into
the 50 mL centrifuge tube. Close the cap and centrifuge at
4500 g for 15 min.
15. Place the QIAamp Maxi column in a new 50 mL centrifuge
tube. Pipet 1.3 mL Buffer AE directly onto the membrane of
the QIAamp Maxi column and close the cap. Incubate at room
temperature for 5 min, and centrifuge at 4500 g for 2 min.
16. Pipet 1.3 mL Buffer AE directly onto the membrane of the
QIAamp Maxi column and close the cap. Incubate at room
temperature for 5 min, and centrifuge at 4500 g for 2 min.
17. Measure the concentration using NanoDrop ND-1000 and
calculate the yield.
18. Aliquot 200 μL into 1.5 mL microtubes labeled with the
sample ID. Quantify the DNA by Nanodrop and store at
20 C.
3.2 Genomic DNA Extraction of DNA from buccal swabs is a modified protocol that
Extraction from Buccal combines the use of the Gentra Puregene Buccal Cell Kit with the
Swabs Qiagen DNeasy Blood and Tissue kit.
1. Preheat water bath to 55 .
2. For each buccal swab, dispense 300 μL Cell Lysis Solution into
a 1.5 mL microcentrifuge tube. Remove the collection brush
from its handle using sterile scissors or a razor blade, and place
the detached head in the tube containing the lysis solution.
3. Add 1.5 μL Puregene Proteinase K, mix by inverting 25 times,
and incubate at 55 C for at least 1 h (up to overnight for
maximum yield).
4. Remove the collection brush head from the Cell Lysis Solu-
tion, scraping it on the sides of the tube to recover as much
liquid as possible.
5. Add 300 μL 100% ethanol to the sample. Vortex mix for 15 s.
6. Carefully apply the entire sample (including any precipitate) to

the DNeasy Mini Spin column placed in a 2 mL collection tube.
Centrifuge at 6000 g for 1 min. Discard the filtrate and the
collection tube.
7. Place the DNeasy Mini Spin Column into a clean 2 mL collec-
tion tube. Add 500 μL Buffer AW1 without wetting the rim.
Close the cap and centrifuge at 6000 g for 1 min. Discard
the filtrate and collection tube.
8. Place the DNeasy Mini Spin Column into another clean 2 mL
collection tube. Carefully open the DNeasy Mini Spin Column
and add 500 μL of Buffer AW2 without wetting the rim. Close
the cap and centrifuge at full speed 20,000 g for 1 min.
Discard the filtrate, put the column back to the collection
tube, close the cap, and centrifuge at full speed 20,000 g
for 2 min.
9. Discard the filtrate and collection tube. Place the DNeasy Mini
Spin Column into a clean, sterile, 1.5 mL microfuge tube
labeled with the sample ID. Carefully open the column and
add 50 μL AE buffer. Incubate at room temperature
(15–25 C) for 2 min. Centrifuge at 6000 g for 1 min.
10. Discard the filter column, close the 1.5 mL microcentrifuge
tube, and store the DNA at 4 C until needed for testing.
3.3 Informative Any SNP array is suitable for this step. The greater the number of
Marker Selection genome-wide SNPs, the better the likelihood of capturing infor-
mative SNPs.
1. Obtain the genotyping calls from the SNP array. Confirm that
the coordinates of the SNPs on the array are from reference
set hg19.
2. Find the closest informative marker that is upstream of the
mutation/gene of interest (see Fig. 2 for examples of an infor-
mative marker). Be sure to select markers with the most signifi-
cant p values (or confidence scores), as the DNA obtained from
buccal swabs may have lower quality calls. Try to select markers
within 1 Mb of the mutation/gene of interest to comply with
ESHRE guidelines [9] and reduce the risk of recombination
between markers.
3. Similarly, find the closest informative marker that is down-
stream of the mutation/gene of interest.
4. If the mutation will not be detected directly then select addi-
tional markers upstream and downstream.
5. Order all probes for the selected markers from the vendor,
including the mutation specific assays.
Fig. 2 Examples of informative markers. (a) Autosomal dominant condition with child with known genotype. (b)
Autosomal dominant condition with no child, but affected parent one generation up. (c) X-linked condition
(e.g., Fragile X) with child with known genotype (e.g., affected male). (d) X-linked condition with no children,
but positive parent one generation up. (e) Autosomal recessive condition where both parents are carriers of the
same mutation, with a child with a normal (non-carrier) or affected genotype
3.4 Taqman For each probe being used the following steps are required: Table 1
Genotyping PCR for shows the volumes required and includes a 20% volume excess for
Probe Validation each reagent to allow for pipetting variation.
3.4.1 Preparation of 20 1. Add TaqMan Gene Expression Mastermix to a labeled
Probes/Assays 1.5 mL tube.
2. Add Molecular Biology Grade water to the 1.5 mL tube.
3. Add the specific TaqMan assay to the 1.5 mL tube.
4. Vortex the tube for 10 s and spin down on short spin setting for
5 s.
Table 1
Example of volumes required for preparation of Taqman 20 probes/assays
Amount for one Amount for 38 samples/ 20% Total

reaction (μL) reactions excess volume
TaqMan gene expression 2.5 95 19 114
master mix
Probe assay (20) 0.125 4.75 0.95 5.7
MGB water 0.375 14.25 2.85 17.1
Total volume 3 114 22.8 136.8
3.4.2 Normalize gDNA 1. Thaw genomic DNA.

2. Calculate the amount of DNA required to normalize the sam-
ple to 5 ng/μL (if extracted from blood) or 10 ng/μL (if ex-
tracted from a buccal swab—see Note 3) in a total volume of
50 μL.
3. Calculate the amount of water required to normalize the sam-
ple by subtracting the amount in step 2 from the total volume
of 50 μL.
4. After the DNA master tube has thawed vortex the tube and
spin down for 5 s.
5. Label one 1.5 mL tube for each sample with the sample ID.
6. Add water to each labeled tube according to the amount
calculated in step 3.
7. Add DNA to each labeled tube according to the amount calcu-
lated in step 2.
8. Close each tube, vortex for 10 s and spin down.
3.4.3 PCR Setup for 1. Label a 384-well plate with the date the PCR is being
Probe Validation performed.
2. Using a multi-pipettor, add 3 μL of the assay mixture into each
designated well.
3. Using a multi-pipettor, add 2 μL of normalized DNA to each
assigned well.
4. Include two “wild-type” controls and at least one “no template
control” (NTC) for each assay being run.
5. Seal the plate with optical adhesive film.
6. Vortex the plate for 10 s and spin down in a microfuge for
1 min at 2000 rpm.
7. Place the plate inside a thermal cycler (see Note 4) previously
programmed with the following program:
l Hold: 50 C for 2 min, 95 C for 10 min.

l Forty cycles: 95 C for 15 s, 60 C for 1 min.
l 4 C Hold.
8. After the PCR program is complete remove the plate from the
thermal cycler and transfer it to the 7900HT Fast Real-Time
PCR System.
9. Perform allelic discrimination post-read per the manufacturer’s
instructions.
10. Analyze the genotyping calls using either SDS or TaqMan
Genotyper.
3.4.4 TaqMan This section of the protocol is intended to start on samples that
Genotyping PCR for Embryo have already been biopsied from an embryo on day 5 or 6 of
Testing development and lysed using standard lysing protocols.
1. Prepare Primer Pool: Mix 2.5 μL of each mutation assays and
linkage assays for a specific SGD patient so that the final con-
centration for each assay will be 0.2.
2. Prepare precoated plates containing the probes designed and
validated previously.
3. Dilute all of the probe assays from 40 to 20 first with MBG
water (50 μL of assay and 50 μL of water).
4. Then dilute the assays to 1 in a 96-well plate (7 μL of 20
assay and 133 μL of MBG water).
5. The 1 assay plate is aliquot into 6384-well plates by EP
motion and dried by Genevac machine for 40 min.
6. Prepare the preamplification (PreAmp) reaction by combining
the following for each reaction: 25 μL TaqMan PreAmp Master
Mix, 12.5 μL PreAmp Primer Pool, 2.5 μL MBG Water.
7. Vortex and spin the combined reagents before use.
8. Add 40 μL of the mix to the side of the tube containing 10 μL
of lysed embryo biopsy sample.
9. Cap the tube, spin down, vortex the tube, and spin down again.
10. Add the tubes to the thermal cycler (see Note 4) and run the
following program:
l Hold: 95 C for 10 min.
l Eighteen cycles: 95 C for 15 s, 60 C for 4 min.
l 4 C Hold.
11. Remove the tubes from the thermal cycler and spin down.
12. Using a 12-channel repeat pipette, aliquot 5 μL of each sample
into precoated 384-well SGD plate.
13. Tap plate after adding samples to bring complete volume to
bottom of well.
14. Seal the plate with optical film and spin down. After the spin
seal edges tightly.
15. Run amplification in the thermal cycler (see Note 4) using the
following program:
l Hold: 50 C for 2 min, 95 C for 10 min.
l Forty cycles: 95 C for 15 s, 60 C for 1 min.
l 4 C Hold.
16. After the PCR program is complete remove the plate from the
thermal cycler and transfer it to the 7900HT Fast Real-Time
PCR System.
17. Perform allelic discrimination post-read per the manufacturer’s
instructions.
18. Analyze the genotyping calls using either SDS or TaqMan
Genotyper.
4 Notes
1. Dissolved QIAGEN Protease is stable for 2 months when

stored at 2–8 C.
2. For storage at 20 C, prepare and freeze QIAGEN Protease
stock solution in aliquots.
3. DNA from buccal swabs can often be contaminated with DNA
from bacterial sources, so to account for those contaminants a
higher concentration of DNA is used if the DNA was isolated
from buccal swabs.
4. The GeneAmp PCR System 9700 is the preferred PCR thermal
cycler.
Acknowledgments
Thank you to David Gabrielle, Anastasia Fedick, Xin Tao, Heather

Garnsey, and Andrew Behrens for the assistance in the development
and implementation of these protocols.
References
1. Handyside AH, Kontogianni EH, Hardy K and preimplantation diagnostic testing for cys-
et al (1990) Pregnancies from biopsied tic fibrosis. N Engl J Med 327(13):905–909
human preimplantation embryos sexed by 3. Findlay I, Ray P, Quirke P et al (1995) Allelic
Y-specific DNA amplification. Nature 344 drop-out and preferential amplification in sin-
(6268):768–770. https://doi.org/10.1038/ gle cells and human blastomeres: implications
344768a0 for preimplantation diagnosis of sex and cystic
2. Handyside AH, Lesko JG, Tarin JJ et al (1992) fibrosis. Hum Reprod 10(6):1609–1618
Birth of a normal girl after in vitro fertilization
4. Gianaroli L, Magli MC, Ferraretti AP et al 8. Scott RT Jr, Upham KM, Forman EJ et al

(1999) Preimplantation diagnosis for aneuploi- (2013) Cleavage-stage biopsy significantly
dies in patients undergoing in vitro fertilization impairs human embryonic implantation poten-
with a poor prognosis: identification of the tial while blastocyst biopsy does not: a rando-
categories for which it should be proposed. mized and paired clinical trial. Fertil Steril 100
Fertil Steril 72(5):837–844 (3):624–630. https://doi.org/10.1016/j.fer
5. Zimmerman RS, Jalas C, Tao X et al (2016) tnstert.2013.04.039
Development and validation of concurrent pre- 9. Harton GL, De Rycke M, Fiorentino F et al
implantation genetic diagnosis for single gene (2011) ESHRE PGD consortium best practice
disorders and comprehensive chromosomal guidelines for amplification-based PGD. Hum
aneuploidy screening without whole genome Reprod 26(1):33–40. https://doi.org/10.
amplification. Fertil Steril 105(2):286–294. 1093/humrep/deq231
https://doi.org/10.1016/j.fertnstert.2015. 10. Simpson JL (2010) Preimplantation genetic
10.003 diagnosis at 20 years. Prenat Diagn 30
6. Verlinsky Y, Rechitsky S, Schoolcraft W et al (7):682–695. https://doi.org/10.1002/pd.
(2001) Preimplantation diagnosis for Fanconi 2552
anemia combined with HLA matching. JAMA 11. Tao X, Su J, Pepe R, Northrop LE, Ferry KM,
285(24):3130–3133 Treff NR (2011) PGD for monogenic disease
7. Thornhill AR, Snow K (2002) Molecular diag- by direct mutation analysis alone in 2 or more
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Mol Diagn 4(1):11–29. https://doi.org/10. ysis in single cells. Fertil Steril 96(3):S21
1016/S1525-1578(10)60676-9
Chapter 5
Detection of Aneuploidy and Unbalanced Rearrangements

Using Comparative Genomic Hybridization Microarrays
Lorena Rodrigo Vivó and Carmen Rubio Lluesa
Abstract
Comparative genomic hybridization arrays (aCGH) allow the analysis of all 24 chromosome aneuploidies
and chromosome rearrangements in the same single (or few) biopsied cells in a short period (less than 24 h).
When applied to preimplantation genetic diagnosis (PGD) and screening (PGS) this technique can improve
the selection of embryos for transfer and therefore also the reproductive outcomes. In this chapter, we
describe the CGH microarray protocol for PGS and PGD used in our laboratory.
Key words Preimplantation genetic screening (PGS), Preimplantation genetic diagnosis (PGD),
Aneuploidy, Chromosomal rearrangement, Array CGH, Whole genome amplification
1 Introduction
Aneuploidy may be a contributing factor to the reproductive pro-

blems in normal-karyotype infertile couples with advanced mater-
nal age, recurrent pregnancy loss, repetitive implantation failure, or
even male infertility [1]. For this reason preimplantation genetic
screening (PGS) is often used to select euploid embryos before
their placement into the uterus in in vitro fertilization (IVF) treat-
ments. Over the last 20 years the technology used for chromosome
screening has dramatically changed; whereas in 1995 the fluores-
cence in situ hybridization (FISH) technique could only be applied
in single cells for the analysis of a limited number of chromosomes,
from 2008 comparative genomic hybridization arrays (aCGH) have
allowed all 24 chromosomes to be analyzed, not only in single cells
but also in trophectoderm biopsies. Additional microarray plat-
forms using different techniques such as single nucleotide poly-
morphisms (SNPs, in 2010) or quantitative polymerase chain
reaction (qPCR, in 2013) have also been successfully applied in
PGS [2]. Consequently, embryologists can now use PGS in several
groups of infertile couples as a tool to be used in addition to
73
74 Lorena Rodrigo Vivó and Carmen Rubio Lluesa
morphological criteria to help select embryos with the best chance

of giving rise to a successful pregnancy.
Patients who carry chromosomal rearrangements have a high
risk of having unbalanced pregnancies resulting in miscarriages or
abnormal offspring. Preimplantation genetic diagnosis (PGD) is a
potent tool which helps to avoid the transfer of embryos carrying
unbalanced rearrangements into the uterus. Selecting a normal or
balanced embryo for transfer increases the chances of having a
healthy, live, newborn child, and decreases the potential for stress
and difficult decisions that these couples usually have to make in the
case of an unbalanced pregnancy. For many years the FISH tech-
nique has been used to analyze embryos in chromosomal rearrange-
ment carriers. Specific strategies using different fluorescent DNA
probes must be tested and applied in each particular couple, and no
chromosomes other than those implicated in the rearrangement
can usually be analyzed. In contrast, the aCGH approach offers the
advantage of detecting rearrangement and aneuploidy unbalances
for every chromosome at the same time and in the same single cell
or trophectoderm biopsy [3]. However, unfortunately, only chro-
mosome rearrangements with translocated fragment sizes larger
than the resolution of the microarray platform can be addressed
with this technique, meaning that the remaining cases must still be
analyzed using the FISH technique.
The aim of this chapter is to describe, in detail, the Illumina®
aCGH microarray 24sure and 24sure + protocols (for PGS and
PGD respectively) used in our laboratory. These microarray plat-
forms are based on BACs (bacterial artificial chromosomes), and
contain spots with normal DNA fragments representing the whole
genome. The technology uses a microarray-based genomic hybri-
dization approach to investigate whole chromosome copy number
and sub-chromosomal structural imbalances in a 12-h protocol.
Regardless of whether a single cell or trophectoderm biopsy is
used, the genome must be amplified to obtain sufficient DNA for
the analysis. This amplified DNA is labeled with Cyanine 3 (Cy3) or
Cyanine 5 (Cy5) fluorophores and is hybridized to the array plat-
form. Female and male euploid DNA are also labeled and hybri-
dized and used as references. After hybridization, the arrays are
washed and scanned using a dual-channel scanner: the green chan-
nel (532 nm) for excitation and reading the Cy3 signal, and the red
channel (635 nm) for excitation and reading of the Cy5 signal. The
images generated are analyzed by the software provided by the array
manufacturer, which generates a plot representing the sample DNA
quantity compared to the reference DNA. Interpretation of the
plots allows DNA gains or losses in the sample to be determined
by comparing them to the hybridized references.
PGS and PGD by aCGH 75
2 Materials
1. 100–130 μm diameter micro glass capillaries.

2. Mouth aspiration system/striper.
3. Stereomicroscope for cell manipulation.
4. Hybridization chamber.
2.1 Cell Loading 1. 20 PBS.

Consumables 2. Double-distilled autoclaved water.
3. Mineral oil.
2.2 aCGH 1. SurePlex Single Cell Whole Genome Amplification Kit (Illu-
Consumables mina, San Diego, CA, USA): Contains cell extraction buffer;
extraction enzyme dilution buffer; extraction enzyme;
pre-amplification buffer; pre-amplification enzyme; amplifica-
tion buffer; amplification enzyme; nuclease-free water.
2. Fluorescent dCTP Labelling System (Illumina, San Diego, CA,
USA): Contains reaction buffer; dCTP-labeling mix; Cy3
dCTP; Cy5 dCTP; Klenow enzyme; dextran sulfate Hybridiza-
tion Buffer.
3. SureRef DNA (Illumina, San Diego, CA, USA): Contains male
and female amplified genomic DNA ready for use as hybridiza-
tion references.
4. COT Human DNA (Illumina, San Diego, CA, USA).
5. 20 SSC Buffer, pH 7.0.
6. 15 pg genomic DNA for positive amplification control (pro-
duced in house).
7. 24sure and 24sure + BAC array slides (Illumina, San Diego,
CA, USA).
2.3 Solutions (See 1. 1 PBS: Autoclave the 20 PBS. Prepare 1 mL of 1 PBS in a
Notes 1 and 2) sterile, autoclaved 1.5 mL tube by mixing 50 μL of the auto-
claved 20 PBS in 950 μL of the double-distilled autoclaved
2.3.1 Solutions for Cell
water; vortex and briefly centrifuge it. Dispense 2.5 μL of 1
Loading
PBS into autoclaved 0.2 mL PCR tubes. Prepare as many tubes
as the number of embryo biopsies. Prepare two additional
tubes to be used as positive and negative PCR controls.
2. 1% Polyvinylpyrrolidone (PVP) Solution: In an autoclaved,
sterile 1.5 mL tube, dilute 0.01 g of polyvinylpyrrolidone in
1 mL of 1 PBS. Vortex and briefly centrifuge.
2.3.2 Hybridization 1. 50% Formamide/2 SSC Solution: 3 mL formamide +3 mL

2 SSC solution.
2.3.3 Slide Washing 1. 20 SSC, adjusted to pH 7.0. Store all of the buffers at room
Buffers (See Notes 3 temperature.
and 4) 2. 2 SSC/0.05%Tween 20: 100 mL 20 SSC + 899.5 mL
double-distilled water +0.5 mL Tween 20.
3. 1 SSC: 25 mL 20 SSC + 475 mL double-distilled water.
4. 0.1 SSC: 5 mL 20 SSC + 995 mL double-distilled water.
3 Methods
The protocols described in this chapter are based on the protocols

recommended by the array manufacturer, Illumina [4, 5] (see Note 5).
3.1 Cell Loading (See The single cell, or the few biopsied trophectoderm cells, should be
Notes 6–8) washed before amplification to minimize non-cellular DNA con-
tamination. After this, the samples are loaded into 0.2 mL PCR
tubes in order to start the amplification.
1. Use one capillary for each individual cell or trophectoderm
biopsy.
2. Briefly centrifuge the 0.2 mL PCR sterile tubes containing
2.5 μL 1 PBS to avoid the formation of bubbles inside them.
3. Prepare a petri dish (for each biopsied embryo) containing a
row with three droplets of 5 μL 1% PVP solution and cover
with mineral oil. Capture the cell or trophectoderm biopsy
with the capillary and rinse it by passing it through each of
these droplets (release it in the first droplet, recapture it and
transfer it into the second droplet; repeat for the third droplet).
4. Deposit each biopsied cell, or trophectoderm cell sample, into
a 0.2 mL PCR sterile tube under the stereoscope. The cell
(s) release should be observed by the embryologist and done
with a minimal volume of washing media (maximum 0.5 μL).
Cap the PCR tube and place inside a cold rack.
5. Store at 20 C until amplification, or maintain at 4 C if the
amplification procedure is immediate.
3.2 Sample 1. Defrost the positive control genomic DNA; vortex and briefly
Amplification centrifuge it. Store it in a rack.
3.2.1 Cell Lysis and DNA 2. Organize the batch of samples to be amplified and calculate the
Extraction in the Pre-PCR buffer quantities for the different master mixes (see quantities
Lab (See Notes 7–10) indicated in the protocol).
3. Defrost a vial of Cell Extraction Buffer and Extraction Enzyme
Dilution Buffer, vortex and briefly centrifuge it.
4. Transfer the 0.2 PCR tubes containing the samples from the
batch into a cold rack. Prepare two additional tubes containing
2.5 μL 1 PBS for positive and negative PCR controls.
5. Briefly centrifuge all the tubes before starting the amplification.
6. Prepare a master mix in a 1.5 mL Eppendorf tube as follows:
For each sample, use 3 μL cell extraction buffer, 4.8 μL extrac-
tion enzyme dilution buffer, and 0.2 μL extraction enzyme.
Scale the amounts according to the number of samples being
processed. Add the Extraction Enzyme at the last minute and
mix by inversion (or a very short vortex burst). Briefly centri-
fuge and keep the master mix in a cold rack.
7. Add 8 μL of freshly prepared master mix to each sample and the
controls. Briefly centrifuge.
8. Take the samples in the cold rack to the post-PCR lab and add
1 μL of control DNA to the positive control tube. Incubate the
samples in a PCR thermo-cycler with the lid preheated to 95 C
(for approximately 15 min) prior to starting the reaction, and
use the following program settings: 75 C for 10 min, 95 C for
4 min and hold at room temperature.
3.2.2 Pre-Amplification 1. In pre-PCR lab, defrost a vial of Pre-amplification Buffer,

in the Pre-PCR Lab (See vortex and briefly centrifuge it.
Notes 7–10) 2. Prepare a master mix in a 1.5 mL Eppendorf tube as follows:
For each sample, use 4.8 μL pre-amplification buffer and
0.2 μL pre-amplification enzyme. Scale the amounts according
to the number of samples being processed. Add the
pre-amplification Enzyme at the last minute and mix by inver-
sion (or a very short vortex burst). Briefly centrifuge and keep
the master mix in a cold rack.
3. Collect the PCR tubes from thermo-cycler in the post-PCR lab
into a cold rack.
4. In pre-PCR lab, add 5 μL of freshly prepared master mix to
each sample and the controls. Briefly centrifuge.
5. Take the samples to the post-PCR lab in cold rack and incubate
them in a PCR thermal cycler with the lid preheated to 99 C
(for 1 h 10 min) and use the following program settings:
l One cycle: 95 C for 2 min.
l Twelve cycles: 95 C for 15 s, 15 C for 50 s, 25 C for 40 s,
35 C for 30 s, 65 C for 40 s, 75 C for 40 s.
l 4 C Hold.
3.2.3 Amplification (See 1. In pre-PCR lab, defrost a vial of Amplification Buffer and a vial
Notes 7–10) of Nuclease-free water, vortex and briefly centrifuge it.

For each sample, use 34.2 μL nuclease-free water, 25 μL ampli-
fication buffer, and 0.8 μL amplification enzyme. Scale the
amounts according to the number of samples being processed.
Add the amplification enzyme at the last minute and mix by
inversion (or a very short vortex burst). Briefly centrifuge and
keep the master mix in a cold rack.
3. Collect the PCR tubes from the thermo-cycler in the post-PCR
lab into a cold rack.
4. In pre-PCR lab, add 60 μL of freshly prepared master mix to
each sample and the controls. Briefly centrifuge.
5. Take the samples in a cold rack to the post-PCR lab and
incubate the samples in a PCR thermal cycler with the lid
preheated to 99 C (for approximately 40 min) and then
apply the following program settings:
l Fifteen cycles: 95 C for 15 s, 65 C for 1 min, 75 C for
1 min.
l 4 C Hold.
6. Place the PCR tubes into a cold rack (in the fridge at 4 C) and
check for sample amplification by agarose gel electrophoresis
(see Note 11).
3.3 Sample Labeling After amplification, samples and reference DNAs are labeled with
Cy3 and Cy5 fluorophores using random primers. The 24sure®
microarray used for PGS is a single channel platform in which the
test samples are labeled and hybridized against each other and are
compared to male and female DNA references which are hybridized
in a different sub-array. In contrast, the 24sure + ® microarray used
for PGD is a dual channel platform in which the test and reference
samples are hybridized onto the same sub-array. The manufacturer
provides an excel workbook template for planning the sample
labeling and distribution in the 96-well plate, and automatically
calculates the proportion of labeling reagents to prepare for the
mix. Moreover, by introducing the microarray slide barcodes into
the planning software it specifies the slide position for hybridization
of each sample.
1. Defrost the Reaction Buffer, Primer Solution, dCTP labeling
mix, Cy3, Cy5, and SureRef Male® and SureRef Female®,
vortex and briefly centrifuge them, and keep them in a
cold rack.
2. Introduce the sample batch and microarray barcodes into the
planning software, and distribute the 0.2 mL tubes with the
amplified samples into the rack according to the planning
software specifications.
3. Add 5 μL of Primer solution to the 96-well plate.

4. Add 8 μL of male and female DNA reference samples according
to the order specified by the planner.
5. Using a multichannel pipette add 8 μL of amplified DNA
product following the order specified by the planner.
6. Seal the plate with adhesive film and briefly centrifuge (20 s).
7. Denature the DNA at 94 C for 5 min in the thermo-cycler
and then keep it at 4 C (in a cooled rack) for 5 min.
8. Prepare the Cy3 master mix in a 1.5 mL Eppendorf tube as
follows: For each sample use 5 μL reaction buffer, 5 μL dCTP
mix, 1 μL Cy3, 1 μL Klenow. Scale the amounts according to
the number of samples being processed. Remove the Klenow
enzyme from the freezer and add it to the master mix at the last
minute. Mix by inversion (or a very short vortex burst). Briefly
centrifuge and keep the master mix in a cold rack.
9. Prepare the Cy5 master mix in a 1.5 mL Eppendorf tube as
follows: For each sample use 5 μL reaction buffer, 5 μL dCTP
mix, 1 μL Cy5, 1 μL Klenow. Scale the amounts according to
the number of samples being processed. Remove the Klenow
enzyme from the freezer and add it to the master mix at the last
minute. Mix by inversion (or a very short vortex burst). Briefly
centrifuge and keep the master mix in a cold rack.
10. Dispense 12 μL of the Cy3 or Cy5 mix into each well according
to the planning software specifications, and always work with
the plate in a cooled rack.
11. Seal the plate with adhesive film and briefly centrifuge (20 s).
12. Incubate at 37 C for 2 h in the thermal cycler to allow labeling
to take place, and then store the plate in a cold rack.
3.4 Combination, 1. Defrost the COT Human DNA; vortex and briefly centrifuge it
Volume Reduction, and keep it in a cold rack.
and Hybridization 2. Warm the dextran sulfate Hybridization Buffer to 75 C.
3. In a fume cabinet, prepare the hybridization chambers by lining
the bottom of the chamber with a paper towel soaked in the
50% Formamide/2 SSC solution.
4. Pulse spin the 96-well plate from the labeling step to collect the
content at the bottom of the wells, and transfer the plate to a
cold rack.
5. Add 25 μL COT Human DNA to each well of the 96-well plate
containing Cy5 labeling products.
6. Using a multichannel pipette, combine Cy3 with the
corresponding Cy5-labeled DNA as set out in the schema.
7. Transfer the plate to a pre-warmed centrifugal evaporator with

the wells uncovered. Evaporate under centrifuge at 75 C until
around 3 μL remains in each tube/well (approximately 1 h).
8. Transfer the plate to a pre-warmed thermo-block (75 C) and
add 21 μL of DS Hybridization Buffer to each well. Resuspend
the pellet, ensuring that it is completely dissolved. Pulse centri-
fuge to collect the contents into the bottom of the wells.
9. Seal the plate and denature it at 75 C for a further 10 min,
centrifuge it for 20 s, and allow it to cool to room temperature.
10. Apply 18 μL of labeled DNA solution to each cover slip
(22 22) and lower the slide (barcode-down) onto the cover
slip for each hybridization area in a fume cabinet.
11. Use the hybridization template to position the cover slips and
to confirm which labeled DNAs are loaded onto each
hybridization area.
12. Place the arrays into the hybridization chamber. Seal them and
incubate in a water bath set at 48 C for 3–16 h.
3.5 Slide Washing To remove the unhybridized DNA from the array, the slides must
be washed under stringent salt concentration and temperature
conditions.
1. Warm the 0.1 SSC solution to 60 C in a Clear Hybex®
Hybridization System.
2. Remove the cover slips from each slide by manually agitating
them in 2 SSC/0.05%Tween20 at room temperature in a
fume cabinet. Immediately transfer them to a stainless steel
rack sitting in a staining jar containing 2 SSC/0.05%
Tween20 solution. Repeat this process for all the slides.
3. Once the rack is fully loaded replace lid, turn on the stirrer, and
clean the slides for 10 min at room temperature. Transfer the
stainless steel rack to a staining jar containing 1 SSC solution,
turn on the stirrer, and clean them for 10 min at room
temperature.
4. Transfer the stainless steel rack to the Clear Hybex® Hybridiza-
tion System containing 0.1 SSC solution at 60 C and wash
them for 5 min without agitation.
5. Transfer the stainless steel rack to a staining jar containing 0.1
SSC solution, turn on the stirrer, and clean them for 1 min at
room temperature.
6. Immediately dry the slides by centrifugation at 180 g for
3 min and store them in an opaque box. To enable more
efficient drying, the slide codes should face toward the centri-
fuge rotor.
3.6 Scan the Slides Use a dual-channel scanner to excite the Cy3 and Cy5 hybridized
fluorophores and to record the emission signals as TIFF image files.
Use the green laser (532 nm wavelength) to excite and read the Cy3
signal, and the red laser (635 nm wavelength) to excite and read the
Cy5 signal. 24sure® and 24sure + ® slides should be scanned at a
10 μm resolution. To simplify the workflow and minimize errors,
we recommended that the slide barcode appears in the image file
name, with a TOP or BOTTOM suffix to identify which hybridiza-
tion area is included in the image file.
3.7 Data Analysis Use the specific array software to turn the scanned images into clear
and Interpretation profiles from which whole chromosome aneuploidy and
sub-chromosomal structural imbalances can be analyzed. The soft-
ware processes the TIFF image files and normalizes the fluorescent
intensities emitted by the Cy3 and Cy5 channels. The data are
processed and represented in a log2 ratio scale plot. The data
from each chromosome are plotted as green dots on a fused chart
display, representing the data from the sample tested versus the
reference DNA. For 24sure® microarrays the software compares
each sample with both male and female references whereas for
24sure + ® microarrays, comparisons are made only with male
reference DNA. The X chromosome log2 ratio (termed
X-separation) is compared with the log2 ratios of each chromosome
to highlight aneuploidy calls as gains [+] or losses [ ] [6]. Accord-
ing to these comparisons, we call a normal sample when there are
not any spot deviations from the reference DNA for any of the
chromosomes shown on the plot (Fig. 1). A sample is considered as
abnormal with specific chromosome aneuploidies (full chromo-
some gains or losses) when the plot shows all the dots for these
chromosomes within the upper or lower normal confidence lines
defined by the reference DNA. The presence of aneuploidies for
most of the chromosomes in the same specimen is interpreted as an
abnormal chaotic pattern. Partial gains or losses for specific seg-
ments of a chromosome are considered as segmental aneuploidies
when the size is higher than the resolution of the platform. Seg-
mental aneuploidies for the chromosomes involving the rearrange-
ment are the most commonly expected unbalances found in
chromosomal rearrangement carriers. In these cases, platforms
with a higher resolution are recommended (e.g., 24sure + ®, with
a higher number of BACs representing pericentromeric and sub-
telomeric regions, see Fig. 2). The samples resulting in amplification
failure show a typical pattern of XXY for sex chromosomes with a
chromosome 19 gain.
Fig. 1 PGS on a single cell using the 24sure array (20 Mb optimal resolution, but can achieve 10 Mb). (a)
Normal XX embryo vs male reference. (b) Abnormal embryo with single aneuploidy, monosomy 13. (c)
Complex abnormal embryo (between 2–5 aneuploid chromosomes). (d) Chaotic abnormal embryo (most of
the chromosomes are abnormal). (e) Profile for a sample with amplification failure
Fig. 2 PGS on a trophectoderm biopsy using the 24sure + array (10 Mb optimal resolution but can achieve
2 Mb) from a patient carrying a balanced reciprocal translocation, 46,XX,t(6;15)(p12;q14). The result shows an
embryo with an unbalanced translocation (partial gain of 6p, a partial loss of 15q) as well as a complete gain of
chromosome 16
4 Notes
1. Due to the low quantity of DNA we are working with for these
PGS and PGD experiments it is very important to work in
conditions of maximum sterility: i.e., a cap, lab coat, and gloves
should be worn; gloves should be alcohol sterilized before
starting any work. It is also important not to touch any surface
without gloves.
2. Work should be undertaken in a vertical flow cabinet. Treat the
cabinet with ultraviolet light for 10 min before starting the
process. Clean the hood, materials, and instruments with
alcohol.
3. Work in a clean area using a lab coat and gloves.
4. Store all of the buffers at room temperature.
5. We recommend that each step of the protocol in which the
samples are transferred to new tubes, plates, or slides is verified
by a second researcher.
6. We recommend working in a room separate from the IVF
laboratory; avoid manipulating DNA in this room, or storing
manipulated DNA samples in the freezer and fridge where the
reaction solutions are stored.
7. Before starting, wear a cap, lab coat, and gloves. Alcohol steril-
ize gloves before starting any work. Do not touch any surface
without gloves.
8. Work in a laminar flow cabinet and under the stereomicro-
scope. Treat the cabinet with ultraviolet light for 10 min before
starting the process. Clean the hood, materials, and instru-
ments with alcohol.
9. The pre-PCR lab for sample amplification should be DNA-free.

No caps, coats, or gloves used in other labs should be allowed
to enter this lab.
10. The thermo-cycler used in these steps should be used exclu-
sively for single- or low-cell number amplification.
11. It is important to check the sample amplification products
before hybridization because samples with amplification failure
can have the same profile as that of samples hybridized in the
same sub-array area.
References
1. Rodrigo L, Mateu E, Mercader A et al (2014) com/content/dam/illumina-support/docume

New tools for embryo selection: comprehensive nts/documentation/chemistry_documentation
chromosome screening by array comparative /arraykits/24sure/24sure-microarray-single-ch
genomic hybridization. Biomed Res Int annel-reference-guide-15056064-c.pdf
2014:517125. https://doi.org/10.1155/ 5. Illumina (2015) 24sure microarray dual channel
2014/517125 reference guide. https://support.illumina.com/
2. Handyside AH (2013) 24-chromosome copy content/dam/illumina-support/documents/
number analysis: a comparison of available tech- documentation/chemistry_documentation/
nologies. Fertil Steril 100(3):595–602. https:// arraykits/24sure/24sure-microarray-dual-chan
doi.org/10.1016/j.fertnstert.2013.07.1965 nel-reference-guide-15056062-c.pdf
3. Alfarawati S, Fragouli E, Colls P et al (2011) 6. Illumina (2014) A technical guide to aneuploidy
First births after preimplantation genetic diag- calling with 24sure single channel. https://sup
nosis of structural chromosome abnormalities port.illumina.com/content/dam/illumina-sup
using comparative genomic hybridization and port/documents/documentation/chemistry_
microarray analysis. Hum Reprod 26 documentation/arraykits/24sure/A-Technica
(6):1560–1574. https://doi.org/10.1093/ l-Guide-to-Aneuploidy-Calling-24sure-15056
humrep/der068 973-A.pdf
4. Illumina (2015) 24sure microarray single chan-
nel reference guide. https://support.illumina.
Chapter 6
Aneuploidy Screening using Next Generation Sequencing

Cengiz Cinnioglu, Refik Kayali, Tristan Darvin, Adedoyin Akinwole,
Milena Jakubowska, and Gary Harton
Abstract
Chromosomal aneuploidy is recognized to be a significant contributing factor in implantation failure and
spontaneous miscarriage Hellani et al. (Reprod Biomed Online 17:841–847, 2008), Vanneste et al. (Nat
Med 15:577–583, 2009) and is likely to be responsible for the majority of IVF failure [Baltaci et al. (Reprod
Biomed Online 12:77–82, 2006), Munne (Placenta 24:S70–76, 2003)]. Preimplantation genetic testing
for aneuploidy (PGT-A) screening, formerly termed preimplantation genetic screening (PGS), enables the
assessment of the numeric chromosomal constitution in blastomere and/or trophectoderm biopsy before
embryo transfer.
Preimplantation genetic testing for aneuploidy (PGT-A) has been proven to improve the selection of
embryos for transfer and therefore also assisted reproductive technology (ART) cycles. In this chapter we
describe the current gold standard platform for PGT-A, next generation sequencing (NGS) protocol used
in our laboratory.
Key words Preimplantation genetic testing for aneuploidy (PGT-A), Aneuploidy, Next generation
sequencing (NGS), Whole genome amplification (WGA)
1 Introduction
One of the most common genetic abnormalities in human embryos

is aneuploidy [1–3] which contributes to increased reproductive
issues of infertile couples including lack of implantation, recurrent
implantation failure, miscarriage, and birth of children affected
with various abnormalities associated with poor reproductive out-
comes [4–6]. To increase both the implantation and pregnancy
rates for these couples, performing PGT-A as a part of the fertility
treatment is recommended. There are several indications for
PGT-A, the most common being advanced maternal age
(AMA-typically female patients older than 35 years at the time of
IVF treatment). A study on polar body biopsies indicates that the
aneuploidy of chromosomes 13, 16, and 18 increases from 20% to
60% for women in age group 35–43 [7]. Among other frequent
85
86 Cengiz Cinnioglu et al.
indications for PGT-A are recurrent miscarriage, repetitive implan-

tation failure (RIF), male factor, and previous aneuploidy
conception [8].
Techniques used in PGT-A have changed significantly in the
last 20 years. Up until 2010, the most common way to diagnose
embryos with aneuploidy was through fluorescence in situ hybridi-
zation (FISH). The major limiting factor to this procedure is an
inability to test for the full chromosome complement (24 chromo-
somes in one test). In practice, only selected chromosomes are
tested using FISH: most commonly, chromosomes 15, 16,
17, 18, 21, 22, X and Y [9]. To increase the number of chromo-
somes that can be analyzed simultaneously, new techniques were
introduced including quantitative polymerase chain reaction
(qPCR), single nucleotide polymorphism arrays (SNP arrays), and
array comparative genomic hybridization (aCGH) [10]. All
24 chromosomes can be tested at the same time using these meth-
ods. The latest development in the aneuploidy screening field is the
development of next generation sequencing (NGS) platforms for
the analysis of the chromosome complement in early human
embryos. Early stages of NGS involved a number of manual pipet-
ting steps including whole genome amplification (WGA), library
preparation, and sequencing chip loading ahead of sequencing
which can be done using a number of NGS platforms commercially
available. In our laboratories, we began NGS using the personal
genome machine (PGM) from Thermo Fisher (San Francisco, CA).
Recently, our laboratories have begun using new technology
including robotics and machines such as Ion GeneStudio™ S5
with the Ion Chef™ system which allows for an automated chip
loading and ability to analyze up to 96 samples simultaneously in
less than 24 h. This makes this system the most costs effective, high
throughput NGS system currently available commercially [11].
The objective of this chapter is to describe our use of the NGS
platform from Thermo Fisher which utilizes semi-conductor
sequencer sequencing and proprietary algorithms to allow for
high throughput, high resolution NGS testing for PGT-A. All
PGT-A protocols begin with a whole genome amplification step
which allows for general amplification of the entire genome from
one to a few cells from early human embryos. The aim of this step is
twofold: to obtain sufficient amounts of deoxyribonucleic acid
(DNA) for further processing and to incorporate specific, unique
DNA barcodes to each embryo biopsy sample. By adding unique
barcodes to each embryo biopsy sample, multiple samples from
different embryos and/or different patients can be mixed together
prior to sequencing allowing for high throughput NGS. Following
WGA, 5 μL of each sample is taken and pooled or mixed with other
samples, followed by a purification step which separates impurities
from the double stranded DNA (dsDNA). The purified pool is
quantified using Qubit (Thermo Fisher Scientific, USA) and the
Aneuploidy Screening using Next Generation Sequencing 87
pool is diluted using nuclease-free water to obtain 18 pM concen-

tration of DNA. The pooled DNA is then ready for the next step
with is isothermal amplification (IA) (see Subheading 3.4). Follow-
ing IA, the samples can be loaded on the semi-conductor chip.
During the loading procedure, the DNA is fragmented, or cut, into
smaller segments that can be attached to tiny beads used in
sequencing. Each bead is the deposited into one well on the chip
[12]. The chip, loaded with beads covered with DNA fragments, is
then placed in the sequencer where a series of nucleotides are
flowed across each bead, one nucleotide at a time. When the next
nucleotide in the segment is incorporated, a hydrogen ion is
released, which changes the pH in the solution in the chip’s well.
Changes of pH are measured and converted into voltage. By mea-
suring the pH of each well during each nucleotide flow, incorpora-
tion of the next nucleotide is assessed and noted by the system.
Once each nucleotide has been incorporated, the sequence of each
unique DNA fragment can be cataloged and used later in the
process [13]. Once completed, the sequence of each sample is
aligned with a reference human genome. This is the followed by
use of the Ion Reporter™ software which calculates the number of
copies of each DNA fragment across the entire genome. This allows
for analysis of chromosome copy number as well as measurement of
small gains/losses [14]. Graphs are generated, and initial calls are
determined by the Ion Reporter™ software. This step is then
followed by analysis of the graphs by highly trained technologists
in our laboratory based on training and use of control cell lines in
each experiment with known chromosome abnormalities. These
controls were purchased by Coriell cell repositories (https://www.
coriell.org/) that were used initially for validation purposes and are
now used in each experiment as positive controls.
2 Materials
1. Stereo Microscope for Cell Manipulation.

2. Stripper micropipette (Vitrolife, Colorado, USA).
3. 96-Well Thermal Cycler with heated lid (0.2 mL block for
standard PCR amplification).
4. 96-Well Retainer for transferring tubes.
5. Microcentrifuge (capable of >15,500 g, fits 1.5 mL and
0.2 mL microcentrifuge tubes).
6. DynaMag™ -2 magnet (Thermo Fisher Scientific, Waltham,
MA).
7. Qubit™ 3.0 Fluorometer.
8. Heat block.
9. Ion OneTouch™ ES Instrument (Thermo Fisher Scientific,

Waltham, MA).
10. 18-MΩ water purification system.
11. Multistage (dual-stage) gas regulator (0-50 PSI, 2-3 Bar
output).
12. Ion Personal Genome Machine™ (PGM™) System (Thermo
Fisher Scientific, Waltham, MA).
13. Ion Chip™ Minifuge (Thermo Fisher Scientific, Waltham,
MA).
14. Vacuum filtration system.
15. 0.2 mL PCR tubes, Flat Cap (do not use polystyrene tubes).
16. 1.5 mL Eppendorf LoBind™ Tubes (DNA).
17. 2.0 mL Eppendorf LoBind™ Tubes (DNA).
18. Low TE (10 mM Tris pH 8.0, 0.1 mM EDTA).
19. Nuclease-free water.
20. NaOH (10 M), molecular biology grade.
21. 1 L Glass bottle.
22. 0.22 μm vacuum filter.
2.1 Cell Loading 1. 100–130 μm diameter micro glass capillaries.

2. 20 Phosphate Buffered Saline (PBS).
3. Double-distilled autoclaved water.
4. Mineral oil.
5. Polyvinylpyrrolidone (PVP).
2.2 Library 1. Ion SingleSeq™ Kit (Thermo Fisher Scientific, Waltham, MA):
Preparation, Contains: cell extraction buffer, extraction enzyme dilution
Purification, and buffer, extraction enzyme, pre-amplification buffer,
Quantification pre-amplification enzyme, amplification buffer, amplification
enzyme, nuclease-free water, Ion SingleSeq™ Barcode Set,
Human CEPH Genomic DNA Control 100 μg/mL.
2. Agencourt™ AMPure™ XP Reagent (Beckman Coulter,
Indianapolis, IN).
3. Qubit™ dsDNA High Sensitivity (HS) Assay Kit (Thermo
Fisher Scientific, Waltham, MA): Contains: Qubit™ dsDNA
HS Reagent (Component A), Qubit™ dsDNA HS Buffer
(Component B), Qubit™ Standards (Component C and D).
4. Qubit™ Assay Tubes (Thermo Fisher Scientific, Waltham,
MA).
2.3 Isothermal 1. Ion PGM™ Template IA Supplies (Thermo Fisher Scientific,

Amplification and Waltham, MA): Eppendorf™ LoRetention Dualfilter—300 μL
Enrichment PCR pipette tips, 8-well strips.
2. Ion PGM™ Template IA Reagents 500 (Thermo Fisher Scien-
tific, Waltham, MA): Ion PGM™ Template IA Pellets 500, Ion
PGM™ Template IA ISP Dilution Buffer, Ion PGM™ Tem-
plate IA Start Solution.
3. Ion PGM™ Template IA Reactions 500 (Thermo Fisher Sci-
entific, Waltham, MA): Ion PGM™ Template IA Ion Sphere
Particles™, Ion PGM™ Template IA Primer Mix S, Ion
PGM™ Template IA Rehydration Buffer.
4. Ion PGM™ Template IA Solutions 500 (Thermo Fisher Sci-
entific, Waltham, MA): Ion PGM™ Template IA Stop Solu-
tion, Ion PGM™ Template IA Recovery Solution, Ion PGM™
Template IA Wash Solution, MyOne™ Beads Wash Solution,
Neutralization Solution, Tween™ Solution.
5. Ion PGM™ Enrichment Beads (Thermo Fisher Scientific, Wal-
tham, MA): Contains Dynabeads™ MyOne™ Streptavidin C1
Beads.
2.4 Sequencing, 1. Ion PGM™ Sequencing Supplies (Thermo Fisher Scientific,

Initialization, and Waltham, MA): Contains: Wash 1 Bottle with label (250 mL),
Cleaning Wash 3 Bottle with label (250 mL), Ion PGM™ Reagent
Bottle Sipper Tubes, Ion PGM™ Wash Bottle Sipper Tubes,
Reagent Bottles with labels (50 mL).
2. Ion PGM™ Hi-Q™ View Sequencing Reagents (Thermo
Fisher Scientific, Waltham, MA): Contains: Ion PGM™
Hi-Q™ View Sequencing Polymerase, Sequencing Primer,
Control Ion Sphere™ Particles.
3. Ion PGM™ Hi-Q™ View Sequencing Solutions (Thermo
Fisher Scientific, Waltham, MA): Contains: Ion PGM™
Hi-Q™ View Sequencing W2 Solution, Ion PGM™ Cleaning
Tablet, Annealing Buffer, Ion PGM™ Hi-Q™ View Sequenc-
ing W3 Solution.
4. Ion PGM™ Hi-Q™ View Sequencing dNTPs (Thermo Fisher
Scientific, Waltham, MA): Contains: Ion PGM™ Hi-Q™ View
Sequencing dGTP, Ion PGM™ Hi-Q™ View Sequencing
dCTP, Ion PGM™ Hi-Q™ View Sequencing dATP, Ion
PGM™ Hi-Q™ View Sequencing dTTP.
5. Ion PGM™ Wash 2 Bottle Kit (Thermo Fisher Scientific,
Waltham, MA): Contains: Wash 2 Bottle with label (2 L),
Wash 2 bottle condition solution.
6. Ion Chip Kits (Thermo Fisher Scientific, Waltham, MA): Con-
tains: Ion 318™ Chip v2 BC.
2.5 Cell Loading 1. 1 PBS: Autoclave the 20 PBS. Prepare 1 mL of 1 PBS in a
sterile, autoclaved 1.5 mL tube by mixing 50 μL of the auto-
claved 20 PBS in 950 μL of the double-distilled autoclaved
water; vortex and briefly centrifuge it. Dispense 2.5 μL of 1
PBS into autoclaved 0.2 mL PCR tubes. Prepare as many tubes
as the number of embryo biopsies. Prepare two additional
tubes to be used as positive and negative PCR controls.
2. 1% Polyvinylpyrrolidone (PVP) Solution: In an autoclaved,
sterile 1.5 mL tube, dilute 0.01 g of polyvinylpyrrolidone in
1 mL of 1 PBS. Vortex and briefly centrifuge.
2.6 Purification and 1. 70% Ethanol: In a sterile 1.5 mL tube, dilute 210 μL of 100%
Quantification ethanol in 90 mL of nuclease-free water. Vortex and briefly
centrifuge.
2. Qubit™ Working Solution: In a sterile 1.5 mL tube, combine
1194 μL of Qubit™ dsDNA HS Buffer with 6 μL of Qubit™
dsDNA HS Reagent. Vortex, briefly centrifuge, and store in a
dark area.
2.7 Isothermal 1. Melt-Off Solution: In a sterile 1.5 mL tube, combine 280 μL of

Amplification and Tween™ Solution and 40 μL of 1 M NaOH. Vortex and briefly
Enrichment centrifuge.
2. 1 M NaOH: In a sterile 1.5 mL tube, dilute 100 μL of 10 M
NaOH in 900 μL of nuclease-free water. Vortex and briefly
centrifuge.
2.8 Sequencing, 1. 1 M NaOH: In a sterile 1.5 mL tube, dilute 100 μL of 10 M

Initialization, and NaOH in 900 μL of nuclease-free water. Vortex and briefly
Cleaning centrifuge.
2. Chlrorite Cleaning Solution: In a 1 L glass bottle, combine 1 L
of 18 MΩ water Ion Cleaning tablet (chlorite tablet). Allow the
tablet to dissolve completely. After the tablet has dissolved, add
1 mL of 1 M NaOH and filter the solution using a 0.22 μm
filter.
3. 100 nM NaOH: In a sterile 1.5 mL tube, dilute 50 μL of 1 M
NaOH in 450 μL of nuclease-free water.
4. Wash 2 Solution: Rinse the Wash 2 Bottle (2 L) 3 times with
200 mL of 18 MΩ water. Fill with 2 L of 18 MΩ water, an
entire bottle of Ion PGM™ Hi-Q™ View Sequencing W2
Solution, and 70 μL of 100 nM NaOH. Invert 5 times to mix
thoroughly.
3 Methods
The protocols described in this chapter are based on the protocols

recommended by the NGS manufacturer, Thermo Fisher Scientific.
3.1 Cell Loading The single cell, or 3–5 cells biopsied from the trophectoderm of
blastocyst stage embryos, should be washed before amplification to
minimize non-cellular DNA contamination. After this, the samples
are loaded into 0.2 mL PCR tubes to start the amplification
process.
1. Use one capillary Stripper tip for each individual cell or tro-
phectoderm biopsy.
2. Briefly centrifuge the 0.2 mL PCR sterile tubes containing 5 μL
1 PBS to avoid the formation of bubbles inside them.
3. Prepare a petri dish (one for each biopsied embryo) containing
a row with three droplets of 5 μL PBS/1% PVP solution and
cover with mineral oil. Capture the cell or trophectoderm
biopsy with the capillary and rinse it by passing it through
each of these droplets (release it in the first droplet, recapture
it, and transfer it into the second droplet; repeat for the third
droplet).
4. Deposit each biopsied cell, or trophectoderm cell sample, into
a 0.2 mL PCR sterile tube under the stereoscope. The cell
(s) release should be observed by the embryologist and done
with a minimal volume of washing media (maximum 0.5 μL).
Cap the PCR tube and place inside a cold rack.
5. Store at 20 C until amplification or maintain at 4 C if the
amplification procedure is immediate.
3.2 Sample Due to the low quantity of DNA we are working with for these
Amplification PGT-A experiments it is very important to work in conditions of
maximum sterility: i.e., a cap, lab coat, and gloves should be worn;
gloves should be alcohol sterilized before starting any work. It is
also important not to touch any surface without gloves. Work
should be done in a vertical flow hood. Sterilize the hood with
UV light for 10 min prior to starting the process. Clean the hood,
materials, and equipment with alcohol. Store all reagents in their
appropriate temperatures. We recommend that each step of the
protocol in which the samples are transferred to new tubes, plates,
or slides is verified by a second researcher. We recommend working
in Pre-PCR room separate from the IVF laboratory. It is crucial to
perform gel electrophoresis during the whole genome amplification
process to determine if amplification has been successful or not.
3.2.1 Cell Lysis and DNA 1. Defrost the positive control genomic DNA; vortex and briefly
Extraction centrifuge it. Store it in a rack.
2. Organize the batch of samples to be amplified and calculate the
buffer quantities for the different master mixes.
3. Defrost a vial of cell extraction buffer and extraction enzyme
dilution buffer, vortex and briefly centrifuge it.
4. Transfer the 0.2 mL PCR tubes containing the samples from
the batch into a cold rack. Prepare one tube containing 2.0 μL
CEPH DNA (10 pg DNA) and 5.0 μL washing buffer for the
positive control and one tube containing only 5.0 μL washing
buffer for the negative control. Briefly centrifuge all the tubes
before starting the amplification.
5. Add 2.5 μL cell extraction buffer to each tube and briefly
centrifuge to collect the contents.
6. Prepare a master mix in a 0.2 mL microcentrifuge tube as
follows: For each sample, use 4.8 μL extraction enzyme dilu-
tion buffer and 0.2 μL extraction enzyme. Scale the amounts
according to the number of samples being processed. Add the
extraction enzyme at the last minute and mix by inversion (or a
very short vortex burst). Briefly centrifuge and keep the master
mix in a cold rack.
7. Add 5 μL of freshly prepared master mix to each sample and the
controls. Briefly centrifuge.
8. Incubate the samples in a PCR thermal cycler with the lid
preheated to 95 C prior to starting the reaction and use the
following program settings:
l One Cycle: 75 C for 10 min.
l One Cycle: 95 C for 4 min.
l 22 C Hold.
3.2.2 Pre-amplification 1. Defrost a vial of pre-amplification buffer, vortex and briefly

centrifuge it.
2. Prepare a master mix in a 0.2 mL microcentrifuge tube as
follows: For each sample, use 4.8 μL pre-amplification buffer
and 0.2 μL pre-amplification enzyme. Scale the amounts
according to the number of samples being processed. Add the
pre-amplification enzyme at the last minute and mix by inver-
sion (or a very short vortex burst). Briefly centrifuge and keep
the master mix in a cold rack.
3. Under a pre-PCR hood, to add 5 μL of freshly prepared master
mix to each sample and the controls. Briefly centrifuge.
4. Incubate them in a PCR thermal cycler with the lid preheated

to 95 C and use the following program settings:
l Twelve cycles: 95 C for 15 s, 15 C for 50 s, 25 C for 40 s,
35 C for 30 s, 65 C for 40 s, 75 C for 40 s.
l 4 C Hold.
5. Place the PCR tubes into a cold rack (or in the fridge at 4 C)
and check for sample pre-amplification by agarose gel
electrophoresis.
3.2.3 Amplification 1. Defrost a vial of amplification buffer and a vial of nuclease-free

water, vortex and briefly centrifuge it.
For each sample, use 2.5 μL nuclease-free water, 27 μL ampli-
fication buffer, and 0.5 μL amplification enzyme. Scale the
amounts according to the number of samples being processed.
Add the amplification enzyme at the last minute and mix by
inversion (or a very short vortex burst). Briefly centrifuge and
keep the master mix in a cold rack.
3. Under a pre-PCR hood, add 30 μL of freshly prepared master
mix to each sample and the controls. Briefly centrifuge.
4. Retrieve the Ion SingleSeq™ Barcode Set, sterilize the surface
with 70% ethanol, and open carefully. Verify the position of the
samples in accordance with run workflow. Add 5.0 μL of the
respective barcode to their respective samples.
5. Mix the samples by inversion and incubate the samples in a
PCR thermal cycler with the lid preheated to 95 C and use
l Four cycles: 95 C for 20 s, 50 C for 25 s, 752 C for 40 s.
l Twelve cycles: 95 C for 20 sec, 72 C for 55 sec.
l 4 C Hold (see Note 1).
3.3 Library 1. Add 5 μL of each library to a new 200 μL microcentrifuge tube

Purification to create an equivolume pool. Vortex the tube to mix and
pulse-spin to collect contents at the bottom of the tube (see
Note 2).
2. Incubate the pool in thermal cycler with the lid preheated to
95 C and use following program settings:
l 22 C Hold.
3. Pulse-spin the tube to collect contents, then transfer the heated

library pool to a fresh 1.5 mL Eppendorf DNA
LoBind™ Tube.
4. Total the volumes of libraries added to the pool and add equal
volume of AMPure™ XP beads. Vortex briefly, pulse-spin the
tube to collect contents, then incubate for 5 min at room
temperature.
5. Place the tube on the DynaMag™-2 magnet, then wait 5 min
for beads to aggregate to the side of the tube.
6. At the end of the 5 min, aspirate the supernatant carefully and
discard. Wash beads with 150 μL of freshly prepared 70%
ethanol while the tube is still on the magnet and incubate for
30 s.
7. Aspirate and discard the wash solution. Repeat with a second
wash, incubate for 30 s, and aspirate the wash solution.
8. Allow the beads to dry at room temperature for 3–4 min with
the tube on the magnet. Then, remove the tube from the
magnet and resuspend beads in 200 μL of Low TE, vortex
thoroughly, and pulse-spin to collect the contents.
9. Incubate the tube at room temperature for 1 min.
10. Place the tube on the DynaMag™-2 magnet, then wait
2–3 min for beads to aggregate to the side of the tube.
11. Transfer 195 μL of the supernatant containing the purified
library pool to a fresh 1.5 mL Eppendorf DNA LoBind™
tube avoiding the carryover of beads. Label the tube accord-
ingly (see Note 3).
3.4 Library 1. Vortex the purified pool thoroughly, pulse-spin to collect con-
Quantification tents, and transfer 30 μL to a new 0.2 μL microcentrifuge tube.
2. Incubate the sample in a thermal cycler with the lid preheated
to 95 C and use following program settings:
l 4 C Hold.
3. Prepare the Qubit™ Working Solution by diluting the Qubit™
dsDNA HS Reagent (Component A) 1:200 in Qubit™
dsDNA HS Buffer (Component B) in a centrifuge tube enough
for the purified pool(s) plus two standards and vortex well.
4. Prepare the two standards by adding 190 μL Qubit™ Working
Solution to two labeled Qubit™ Assay Tubes. Add 10 μL of
each Qubit™ Standard #1 (Component C) and Qubit™ Stan-
dard #2 (Component D) to the appropriate tubes.
5. Prepare the unknowns by adding 198 μL Qubit™ Working
Solution to labeled Qubit™ Assay Tubes, 2 tubes per pool to
create an average. Add 2 μL of the heated library pool to the

appropriate sample tube.
6. Vortex each of the tubes for 2–3 s, avoid creating bubbles.
Incubate the tubes in the dark at room temperature for 2 min.
7. On a Qubit™ 3.0 Fluorometer, select “dsDNA Assay” and
then select “dsDNA High Sensitivity.” Choose “Read Stan-
dards” to measure standards that will generate a standard
curve.
8. Set the roller wheel to 2 μL and the units to ng/μL. Read the
standards and records the values. Calculate the average purified
pool and immediately proceed to isothermal amplification
reaction.
3.5 Isothermal 1. Turn on the heat block and set to 41 C. Fill each of the wells
Amplification with water.
(IA) Reaction 2. Thaw the Ion PGM™ Template IA Primer Mix S and Ion
3.5.1 Preparation of PGM™ Template IA Rehydration Buffer on cold block. Place
Template-Positive Ion
Ion PGM™ Template IA Pellet on cold block until needed.
PGM™ Template IA Ion 3. Dilute the library pool to 2.0 nM based on the recorded
Sphere Particles (ISPs) average during the quantification step. Based on dilution cal-
culation, use Low TE buffer to dilute.
4. Perform a 1:100 dilution using the freshly prepared 2.0 nM
library pool and nuclease-free water to generate a 20 pM
library pool.
5. Transfer 50 μL of 20 pM library pool to a new 0.2 mL PCR
tube. Heat the pool in the thermal cycler with the lid preheated
to 95 C and use following program settings:
l 4 C Hold.
6. Vortex the Ion PGM™ Template IA ISPs for 1 min.
7. In a new 2.0 mL Eppendorf DNA LoBind™ tube on a cold
block, prepare the templating solution by combining the fol-
lowing: 130 μL Ion PGM™ Template IA ISP Dilution Buffer,
8.0 μL Ion PGM™ Template IA Primer Mix S, 21 μL Ion
PGM™ Template IA ISPs, 10 μL of the preheated 20 pM
purified library. Vortex the tube, pulse-spin to collect the con-
tents, and place on cold block.
8. Invert the Ion PGM™ Template IA Rehydration Buffer
3 times to mix, the use 720 μL to rehydrate the Ion PGM™
Template IA Pellet. Vortex for 4 s at maximum setting, pulse-
spin to collect the contents, and place on cold block.
9. Transfer the rehydrated IA pellet to the templating solution,
vortex for 4 s, pulse-spin to collect contents, and store on cold
block.
10. Invert the Ion PGM™ Template IA Start Solution 3 times to

mix, then add 300 μL to the mixture using negative pipetting:
l Set a 1 mL pipette to 300 μL. Press the pipette to the second
stop and dip the tip into the Ion PGM™ Template IA Start
Solution. Slowly release the pipette and allow 10 s for the
contents to be fully drawn.
l Dispense the contents to the mixture by pressing the pipette
to the first stop and hold it. Allow 10 s for the contents to be
dispensed. Some liquid will remain in the tip.
11. Vortex the tube ten times in 1 s pulses at maximum settings.
Invert the tube and repeat the ten 1 s pulses. Pulse-spin to
collect the contents and place the tube on ice block.
12. Start the reaction by gently placing the tube in the preheated
heat block. Make certain the tube is immersed in water. Incu-
bate the IA reaction for 25 min at 41 C.
3.5.2 Recovery of 1. Stop the IA reaction by removing the tube from the heat block
Template-Positive Ion and adding 650 μL of the Ion PGM™ Template IA Stop
PGM™ Template IA Ion Solution. Vortex the tube thoroughly.
Sphere Particles (ISPs) 2. Centrifuge the tube at 7500 g for 6 min.
3. Aspirate and discard the supernatant, being careful not to
disturb the pellet. Leave approximately 100 μL in the tube.
4. Resuspend the pellet in 1 mL of the Ion PGM™ Template IA
Recovery Solution. Resuspend by pipetting up and down.
Vortex the mixture well, avoiding to create bubbles.
5. Add an additional 700 μL of the Ion PGM™ Template IA
Recovery Solution and vortex thoroughly. Incubate for 5 min
with 5 s of vortexing every minute.
6. Centrifuge the tube at 17,000 g for 6 min.
7. Immediately remove and discard all the supernatant without
disturbing the ISP pellet. Remove any bubbles prior to remov-
ing the bulk of the liquid to avoid frothing in subsequent steps.
8. Add 100 μL of the Ion PGM™ Template IA Wash Solution to
the ISP pellet. Resuspend the templated ISPs complete by
vortexing for 4 s at maximum speed, then pipetting the ISP
suspension up and down four times (see Note 4).
3.6 Enrichment 1. Prepare a melt-off solution as follows: In a sterile 1.5 mL

Eppendorf tube, combine 280 μL of the Tween™ Solution
3.6.1 Preparing the
and 40 μL of 1 M NaOH. Vortex well and briefly centrifuge.
Melt–Off Solution and the
Dynabeads™ MyOne™ 2. Vortex the tube of Dynabeads™ MyOne™ Streptavidin C1
Streptavidin C1 Beads Beads for 30 s to resuspend the beads thoroughly. Briefly
centrifuge the tube to collect the supernatant without getting
beads.
3. Transfer 13 μL of Dynabeads™ MyOne™ Streptavidin C1

Beads in a sterile 1.5 mL Eppendorf tube. Place the tube on
the DynaMag™-2 magnet for 2 min, then carefully remove,
and discard the supernatant without disturbing the pellet.
4. Add 130 μL of the MyOne™ Beads Wash Solution to the
pellet. Remove the tube from the magnet, vortex the tube for
30 s, then briefly centrifuge to collect the contents.
3.6.2 Filling the 8-Well 1. Vortex the template-positive ISPs (~100 μL) and transfer all
Strip contents in Well 1 in a sterile 8-well strip.
2. Fill the remaining wells with the following:
l Well 2: 130 μL of Dynabeads™ MyOne™ Streptavidin C1
Beads resuspended in MyOne™ Beads Wash Solution.
l Well 3: 300 μL of Ion PGM™ Template IA Wash Solution.
l Well 6: Empty.
l Well 7: 300 μL of freshly prepared Melt-Off Solution.
l Well 8: Empty.
3.6.3 Preparing the Ion 1. Place a new tip in the Tip Loader. Take the Tip Arm and push
OneTouch™ ES for down onto the tip and ensure that it is tightly secured. Once
Enrichment secure, return the Tip Arm back to its cradle ensuring it is in the
correct orientation.
2. In a sterile 0.2 mL PCR tube, add 10 μL of Neutralization
Solution and place it into the place holder on the device.
3. Place the 8-well strip in the slot, flushed all the way right. Pres
Start/Stop to begin the run (~36 min).
4. Immediately after the run, securely close and remove the PCR
tube containing the enriched ISPs. Invert the tube five times to
mix the contents.
5. Discard the used tip and 8-well strip (see Note 5).
3.7 Preparing the 1. Fill a brand-new Wash 2 Bottle to the mold line (~2 L) with
PGM for Initialization 18 MΩ water. Pour the entire contents of the Wash 2 Bottle
Condition Solution and invert the mix five times.
3.7.1 Condition the Wash
2 Bottle for Use 2. Allow the bottle to sit at room temperature for at least 8 h
before use.
3.7.2 Cleaning the PGM 1. If performing an 18 MΩ water cleaning, do the following:

l Empty and remaining solution from all bottle and rinse
them twice with 18 MΩ water.
l Add 250 mL of 18 MΩ water into the cleaning bottle and

securely attach this to the W1 position. Place the remaining
empty bottles in their respective W2 and W3 positions
without screwing the caps on.
l Remove any dNTP conical tubes and leave used sipper tips
attached. Place drip trays below.
l Ensure that a used chip is securely placed in the PGM.
l Press Clean on the PGM touchscreen and selection
18-MOhm water cleaning. Follow the onscreen instructions
and allow cleaning to begin.
l Empty all wastes once complete.
2. If performing a Chlorite cleaning, do the following:
l Empty and remaining solution from all bottle and rinse
them twice with 18 MΩ water.
l Fill a glass bottle with 1 L 18 MΩ water, then add an Ion
Cleaning tablet (chlorite tablet). Allow the tablet to dissolve
completely. When the tablet has completely dissolved, add
1 mL of 1 M NaOH and filter the solution using a 0.22 μm
filter.
l Add 250 mL of the filtered chlorite solution into the clean-
ing bottle and securely attach this to the W1 position. Place
the remaining empty bottles in their respective W2 and W3
positions without screwing the caps on.
l Remove any dNTP conical tubes and leave used sipper tips
attached. Place drip trays below.
l Ensure that a used chip is securely placed in the PGM.
l Press Clean on the PGM touchscreen and selection chlorite
cleaning. Follow the onscreen instructions and allow clean-
ing to begin.
l Empty all wastes once complete.
3.7.3 Initializing the PGM 1. Remove the dNTP stock solutions from the freezer and begin
thawing. Check the tank pressure for the nitrogen gas.
2. Rinse the conditioned Wash 2 Bottle 3 times with 200 mL of
18 MΩ water. Fill to the mold line with 18 MΩ water. Add the
entire bottle of Ion PGM™ Hi-Q™ View Sequencing W2
Solution to the Wash 2 bottle.
3. Prepare 500 μL of 100 mM NaOH by diluting 50 μL of 1 M
NaOH in 450 μL of nuclease-free water.
4. Add 70 μL of 100 nM NaOH to the Wash 2 Bottle. Cap the
bottle and invert it five times to mix contents thoroughly.
5. Rinse the Wash 1 and Wash 3 bottles with 18 MΩ water.
6. Add 350 μL of 100 nM NaOH to the Wash 1 Bottle.
7. Add Ion PGM™ Sequencing W3 Solution to the 50 mL line

marked on the Wash 3 Bottle.
8. Replace the sippers for the Wash Bottles, place the prepared
bottles in their respective positions, and securely fasten the caps
on each.
9. On the PGM, press Initialize and follow the instruction. Select
Ion PGM™ Hi-Q™ View Sequencing Kit when prompted,
then press next. Allow the PGM to complete the first portion
of the initialization process.
10. Using the thawed dNTP stocks, transfer 20 μL of each dNTP
stock solution to a new respective reagent bottle. Cap each
bottle until ready for use.
11. After the PGM completes the first portion of the initialization
process, replace the sippers for the dNTP slots, and securely
fasten each dNTP bottle into their respective positions. Press
next to allow the PGM to complete the final portion of the
initialization process.
12. If the initialization was successful, then a notification stating
Passed will appear on the screening. This will indicate that the
PGM is ready for sequencing.
3.8 Starting a 1. On the Ion Torrent Server, create a Planned Run using Ion
Sequencing Run ReproSeq™ Aneploidy.
3.8.1 Create a 2. Follow the on-screen instructions and confirm the plan run
Planned Run details.
3.8.2 Preparing the 1. Thaw the Sequencing Primer on ice.

Enriched, Template- 2. Vortex the Control Ion Sphere™ Particles (ISPs), then briefly
Positive ISPs centrifuge to collect the contents. Add 5 μL if the Control ISPs
to the enriched, template-positive ISPs. Mix well.
3. Centrifuge at 17,000 g for 3 min.
4. Aspirate the supernatant and avoid disturbing the pellet. Leave
~15 μL in the tube.
5. Vortex the thawed primer for 5 s and add 12 μL to the
enriched, template-positive ISPs. Vortex the mixture and
briefly centrifuge.
6. Incubate the samples in a PCR thermal cycler with the lid
preheated to 95 C prior to starting the reaction and use the
3.8.3 Performing a Chip 1. On the initialized PGM, press Run and follow the instructions
Check to perform chip check.
2. When prompted, place a new Ion 318™ Chip v2 BC into the
chip deck, securely fasten the chip, and allow the PGM to
calibrate the chip.
3.8.4 Binding the 1. After annealing the Sequencing Primer in the thermal cycler,
Sequencing Polymerase, add 3 μL of Ion PGM™ Hi-Q™ View Sequencing Polymerase
Loading the Chip, and to the ISPs. Mix by pipetting up and down, then allow the
Starting the Run mixture to incubate at room temperature for 5 min.
2. Following chip calibration, retrieve the new chip and aspirate
all liquid in the chip through the loading port. Using an Ion
Chip™ Minifuge, place the chip upside-down with the tab
pointing in. Centrifuge the chip for 10 s to remove of any
liquid.
3. Place the chip on a first surface. After the incubation is com-
plete, transfer 30 μL of ISPs to the chip by loading through the
loading port. Add the dial by dialing down ~1 μL per sec. Pay
attention to the dispersal of the liquid in the chip to ensure a
homogenous distribution is done.
4. Using an Ion Chip™ Minifuge, place the chip right-side up
with the tab point in and centrifuge for 1 min. Repeat the 1 min
centrifuge this time with the chip tab point out.
5. Remove the chip and hold it at a 45 angle. Without removing
the tip, slowly pipet the sample out and then back into the chip
one time. Pipet slowly to avoid creating bubbles. Then remove
as much liquid as possible.
6. Using an Ion Chip™ Minifuge, briefly centrifuge the chip
upside-down with the tab point in.
7. Load the chip onto the PGM and follow the onscreen
instructions.
8. Once all planned run details have been confirmed, begin the
run. Sequencing will take ~2 h.
3.9 Data Analysis Run data obtained by the PGT-A sequencer is processed and sent to
Ion Torrent Browser Suite™ version 5.6.0 for generation of data
files utilized for base calling and profile generation. Run quality
control (QC) parameters must be met to emit diagnosis. QC is
broken down into two components for run validation. (1) Run
performance (all samples are analyzed with identified control) and
(2) the analysis of each sample independently meets QC run
metrics. Optimal run performance metrics are as follows: Loading
>70%. Life Ion Sphere™ Particles (ISPS) >98%. Usable Reads
>30%. Polyclonality <50%. Optimal sample parameters are as fol-
lows: Duplicate Reads <30%. Median of the Absolute values of all
Pairwise Differences (MAPD) <0.35. Reads per sample >70,000.
If all QC parameters are met, diagnosis can begin. Runs should be

repeated if run parameter values are below the optimal QC values
determined by the manufacturer and internal validation protocol.
Sequencing data obtained by the PGT-A sequencer is processed
and sent to Ion Reporter™ version 5.4 software for data analysis.
This software uses the bioinformatic tool ReproSeq Low-pass
whole-genome aneuploidy workflow v1.0 to detect 24 chromo-
somes aneuploidies from a single whole-genome sample with low
coverage (minimum 0.01). Normalization is done using the bio-
informatics baseline ReproSeq Low-Coverage Whole-Genome
Baseline generated from multiple normal samples. Ion Reporter™
software generates a graph representing the copy number variation
(CNV) of the sample analyzed compared to the reference bioinfor-
matics baseline. An embryo is considered normal when it has no
deviations from the reference bioinformatics baseline for any of the
24 chromosomes. PGT-A test has different diagnostic results,
depending on which, the transference of the embryo would be
recommended or not. An embryo is considered normal when all
its chromosomes have two copies. An embryo is defined as abnor-
mal by the presence of aneuploidy (chromosome gains or losses)
when there are bins that are diverted into the upper (gain +) or
lower part (loss -) of the graph. The aneuploidies could be for a full
chromosome, when all the bins covering a chromosome are gained
or lost, or partial aneuploidies, when the bins gained or lost repre-
sent only part of the chromosome. These abnormalities are usually
called deletions or duplications. The presence of aneuploidies for
most of the chromosomes on the same specimen is interpreted as a
chaotic abnormal embryo. With this technique, it is not possible to
identify deletions and duplications smaller than 10 Mb (the limit of
resolution of this platform), balanced structural abnormalities, low
level mosaic aneuploidy, uni-parental disomy (UPD), haploidy, and
most polyploidies.
4 Notes
1. This is a safe stopping point. The amplified samples may be

stored at 20 C for up to 2 years.
2. When pooling fewer than 8 libraries, the pool volume drops
below 40 μL. Add nuclease-free water to bring the final volume
to 40 μL prior to library pool purification.
3. This is a safe stopping point. The purified pool may be stored at
20 C for up to 1 week.
4. This is a safe stopping point. The templated ISPs may be stored
at 20 C for up to 1 week.
5. This is a safe stopping point. The templated ISPs may be stored
at 4 C for up to 3 days.
References
1. Hellani A, Abu-Amero K, Azouri J et al (2008) technologies in reproductive medicine, 1st
Successful pregnancies after application of edn. CRC Press, Taylor and Francis Group,
array-comparative genomic hybridization in Boca Raton
PGS-aneuploidy screening. Reprod Biomed 9. Stephenson MD, Awartani KA, Robinson WP
Online 17(6):841–847 (2002) Cytogenetic analysis of miscarriages
2. Vanneste E, Voet T, Le Caignec C et al (2009) from couples with recurrent miscarriage: a
Chromosome instability is common in human case-control study. Hum Reprod 17
cleavage-stage embryos. Nat Med 15 (2):446–451
(5):577–583. https://doi.org/10.1038/nm. 10. Sermon K, Capalbo A, Cohen J et al (2016)
1924 The why, the how and the when of PGS 2.0:
3. Fragouli E, Alfarawati S, Spath K et al (2013) current practices and expert opinions of fertil-
The origin and impact of embryonic aneu- ity specialists, molecular biologists, and embry-
ploidy. Hum Genet 132(9):1001–1013. ologists. Mol Hum Reprod 22(8):845–857.
https://doi.org/10.1007/s00439-013-1309- https://doi.org/10.1093/molehr/gaw034
0 11. Yan L, Huang L, Xu L et al (2015) Live births
4. Baltaci V, Satiroglu H, Kabukçu C et al (2006) after simultaneous avoidance of monogenic
Relationship between embryo quality and diseases and chromosome abnormality by
aneuploidies. Reprod Biomed Online 12 next-generation sequencing with linkage ana-
(1):77–82 lyses. Proc Natl Acad Sci U S A 112
5. Munne S (2003) Preimplantation genetic diag- (52):15964–15969. https://doi.org/10.
nosis and human implantation--a review. Pla- 1073/pnas.1523297113
centa 24(Suppl B):S70–S76 12. Ion ReproSeq™ PGS Kits—Ion S5™ System
6. Rodrigo L, Mateu E, Mercader A et al (2014) USER GUIDE. (2017) Thermo Fisher Scien-
New tools for embryo selection: comprehen- tific, Inc. https://assets.thermofisher.com/
sive chromosome screening by array compara- TFS-Assets/LSG/manuals/MAN0016712_
tive genomic hybridization. Biomed Res Int IonReproSeqPGS_S5_UG.pdf. Accessed
2014:517125. https://doi.org/10.1155/ 4 March 2018
2014/517125 13. Merriman B, Ion TR, Team D et al (2012)
7. Kuliev A, Zlatopolsky Z, Kirillova I et al (2011) Progress in ion torrent semiconductor chip
Meiosis errors in over 20,000 oocytes studied based sequencing. Electrophoresis 33
in the practice of preimplantation aneuploidy (23):3397–3417. https://doi.org/10.1002/
testing. Reprod Biomed Online 22(1):2–8. elps.201200424
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014 Validation of next-generation sequencing for
8. Rubio C, Vera M, Lopez P et al (2018) Preim- comprehensive chromosome screening of
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(eds) Handbook of new genetic diagnostic rbmo.2015.09.002
Part III
Traditional Prenatal Diagnosis

Chapter 7
DNA Extraction from Various Types of Prenatal Specimens

Odelia Nahum, Amanda Thomas, and Brynn Levy
Abstract
A prenatal noninvasive genetic screening test that yields a positive result typically warrants further direct
assessment of fetal DNA following an invasive procedure. The precious nature of these invasively acquired
samples, combined with the time sensitive nature with which results should be reported, demands that the
methodologies used for analysis be quick, efficient, and dependable.
Prenatal diagnosis has been performed using DNA extracted from amniotic fluid and chorionic villi for
several decades, and more recently methodologies have been developed to extract cell free fetal DNA from
amniotic fluid. DNA extraction methodologies in these matrices should reliably and reproducibly isolate a
sufficient quality and quantity of DNA for the intended downstream application, and make it possible to
purify and concentrate samples that may arrive with suboptimal quality or quantity.
Phenol-Chloroform extraction followed by DNA precipitation in ethanol has historically been used for
prenatal samples, but this methodology is labor intensive, time consuming, and requires use of toxic
chemicals. There are now commercially available, solid phase-based kits for rapid and reproducible DNA
extraction and purification, enabling simultaneous extraction of a large number of samples. Commercial kits
are available for a variety of sample matrices including all prenatal specimen types, although other meth-
odologies including organic or inorganic liquid phase extraction may also be utilized.
Here, we describe extraction using both commercially available kits for direct amniocytes and chorionic
villi and cell free fetal DNA derived from amniotic fluid, as well as inorganic liquid phase extraction for tissue
culture of amniocytes, CVS, and products of conception.
Key words DNA extraction, Amniotic fluid, Chorionic villi, Cell free amniotic fluid, Cell culture,
DNA prep, Precipitation, cffDNA, Inorganic liquid phase extraction, DNA extraction kit
1 Introduction
Prenatal genetic diagnosis traditionally requires invasive procedures

to acquire fetal DNA for direct analysis. A critical feature of prenatal
diagnosis is the ability to return accurate results quickly and effi-
ciently, so that time may be afforded for decision-making or addi-
tional testing. In the laboratory, the first step of many diagnostic
techniques is the extraction of high quality DNA with sufficient
yield for the intended downstream methodology. DNA extraction
techniques should be robust and focused on maximizing the quality
and quantity of isolated DNA, as low yield or poor quality DNA
105
106 Odelia Nahum et al.
may require an additional invasive procedure. Extraction meth-

odologies should be fast, efficient, reliable, and reproducibly isolate
a sufficient quantity of DNA that is free from contaminating com-
pounds. All DNA extraction protocols should be optimized to
isolate DNA at both the quality and quantity sufficient for the
intended downstream application.
Historically, DNA from amniocytes and chorionic villi has been
isolated using phenol-chloroform extraction followed by ethanol
precipitation [1, 2]. While similar methodologies may still be uti-
lized for DNA extraction of prenatal samples, they are labor inten-
sive, time consuming, and require the use of toxic chemicals. Today,
a variety of commercial kits are available that quickly and efficiently
isolate DNA without compromising quantity or quality. The vol-
ume of sample required for sufficient DNA quantity will depend on
the sample matrix and the intended downstream application. Gen-
erally the quantity of DNA attainable from amniotic fluid is lower
than that of CVS [3], and this is reflected in the amount of sample
required for sufficient DNA yield.
Today, DNA extraction directly from chorionic villi and amnio-
cytes can generally be achieved with the same commercially avail-
able column-based kit, however, the initial sample preparation steps
vary depending on the sample type. Prior to DNA extraction,
chorionic villi samples should be cleaned and microdissection
employed to remove as much maternal tissue as possible. Further
disaggregation of the villi is achieved by treating with trypsin to
remove the trophoblastic layer, and collagenase to digest the inter-
cellular matrix. By contrast, extraction from direct amniocytes
requires only centrifugation to collect the cells, followed by wash-
ing and resuspension. Centrifugation of amniotic fluid allows for
separation of the amniocytes and supernatant. The amniotic super-
natant contains a high fraction of fetal cell free DNA and may also
be utilized for prenatal analysis.
Once amniocyte and CVS samples are prepared for extraction, a
broad spectrum protease such as Proteinase K, a ribonuclease, and
buffer containing a chaotrophic agent are added to the sample to
remove protein and RNA. DNA is bound to the column, washed,
and excess contaminates are filtered using a series of buffers with
varying stringencies to remove organic contaminants and salts.
DNA can be eluted using water. These commercially available
stationary phase kits typically produce high quality DNA free of
contaminants, with yield proportional to the volume of starting
material.
It is highly recommended that a backup culture be established
for every prenatal specimen being processed. Having backup cul-
tures provides multiple benefits including (1) having fetal material
available when the DNA quality/quantity from the uncultured
tissue (chorionic villi or amniocytes) is suboptimal, (2) the down-
stream assay fails and needs to be repeated and (3) additional
DNA Extraction from Various Types of Prenatal Specimens 107
confirmatory tests are required or ordered. In addition, establish-

ment of cell cultures may reduce the probability of maternal cell
contamination for amniotic fluid contaminated with peripheral
blood [4], and decrease the probability of DNA extraction failure
for suboptimal CVS samples [5]. DNA extraction from cultured
chorionic villi, cultured amniocytes, or cultured tissue derived from
products of conception can be attained with commercially available
kits, or inorganic liquid phase extraction as described here. Briefly,
when flasks are approximately 70–80% confluent, cells are trypsi-
nized, washed, and pelleted via centrifugation. Cells are lysed in a
detergent-based solution, typically 10% SDS, followed by a high
salt protein precipitation with ammonium acetate. DNA is precipi-
tated from the supernatant and resuspended in a Tris-EDTA buffer
solution. Spectrophotometric analysis can be used to confirm DNA
concentration and purity.
While prenatal diagnosis has been carried out in amniocytes and
chorionic villi for many decades, more recently the existence of cell
free fetal DNA (cffDNA) in amniotic fluid has been recognized as a
robust matrix for analysis of genetic information [6]. The amount
of cell free fetal DNA present in amniotic fluid seems to be a
function of both gestational age and ploidy status of the fetus (for
review see Hui [7]). Initial attempts at utilizing DNA extraction
methods similar to that of serum/plasma or body fluid modified for
large volumes met only partial success [6], and only 40% of the
samples extracted produced a sufficient yield for microarray analysis
[8]. Modification of protocols to resemble DNA extraction from
urine produced more sufficient yield of DNA [9], and this is the
basis for the current cffDNA methodologies. Further optimization
of the cffDNA purification protocol has led to the standardization
of this methodology, which is now commercially available in cell
free DNA extraction kits. Extraction of cffDNA from amniotic fluid
utilizes amniotic fluid remaining after the amniocytes have been
removed. Since this is typically >5 mL, DNA extraction methods
are adapted for a larger volume of starting material. Briefly, amni-
otic fluid samples are added to a binding optimization buffer con-
taining Proteinase K. Carrier RNA is added to increase the DNA
yield of low concentration samples [10]. DNA is concentrated by
passing the sample through a vacuum-based column. Bound DNA
is washed several times with a series of buffers with varying strin-
gencies, followed by a final wash in 100% ethanol. Elution is per-
formed with molecular biology grade water.
In our hands, the methodologies discussed in this chapter have
reliably and reproducibly extracted DNA at sufficient quantities,
with a 260/280 ratio between 1.8 and 2.0, and 260/230 ratio
greater than 1.5. All procedures should be independently opti-
mized to extract DNA of prenatal samples to a concentration
sufficient for the specific application for which it is intended.
2 Materials
2.1 DNA Extraction 1. AccuGENE 1 PBS (Lonza, Basel, Switzerland).

from Amniotic Fluid 2. QIAmp DNA Blood Mini Kit (QIAGEN, Germantown, MD,
USA).
3. Proteinase K (QIAGEN, Germantown, MD, USA).
4. RNase A Solution (100 mg/mL) (QIAGEN, Germantown,
MD, USA).
5. 100% Ethanol.
6. Molecular Biology Grade Water.
2.2 DNA Extraction 1. QIAmp DNA Blood Mini Kit (QIAGEN, Germantown, MD,
from Chorionic Villi USA).
2. Proteinase K (QIAGEN, Germantown, MD, USA).
3. RNase A Solution (100 mg/mL) (QIAGEN, Germantown,
MD, USA).
4. 100% Ethanol.
2.3 DNA Extraction DNA can be extracted from cultured chorionic villi, cultured amni-
from Cultured Cells otic fluid cells (amniocytes), or cultured tissue derived from pro-
ducts of conception.
1. Trypsin (TrypLE Express) (Gibco, Gaithersburg, MD).
2. Cell Lysis Solution (QIAGEN, Germantown, MD, USA).
3. Protein Precipitation Solution (Ammonium acetate 7.5 M).
4. RNase A Solution (QIAGEN, Germantown, MD, USA).
5. 100% isopropanol.
6. 70% ethanol.
7. TE Buffer.
2.4 DNA Extraction 1. QIAamp circulating nucleic acid kit (QIAGEN, Germantown,
from Cell Free MD, USA).
Amniotic Fluid 2. AccuGENE 1x PBS (Lonza, Basel, Switzerland).
3. 100% Ethanol.
5. Carrier RNA: Add 1550 μL buffer AVE from the kit to the tube
containing 310 μg lyophilized carrier RNA to obtain a solution
of 0.2 μg/μL. Dissolve it thoroughly and aliquot in 200 μL
tubes. Store at 20 C.
3 Methods
3.1 DNA Extraction 1. Spin the amniotic fluid (AF) specimen in a 15 mL tube at
from Amniotic Fluid 2500 g for 15 min at room temperature (see Note 1).
2. Remove the supernatant with a sterile transfer pipet to a new
tube (see Note 2).
3. Tap the tube to resuspend the pellet.
4. Add 900 μL of 1 PBS to the tube using the PBS to wash the
sides of the tube.
5. Spin the tube at 2500 g for 1 min at room temperature.
6. Remove the supernatant and add it to the original supernatant
tube, leaving the cell pellet with about 50 μL of supernatant.
7. Tap the tube to resuspend the pellet.
8. Add 475 μL of 1 PBS; tap the tube to mix ensuring that the
pellet is completely resuspended.
9. Add 50 μL of Qiagen proteinase K into the sample and mix well
by tapping.
10. Add 10 μL of RNAse A solution (100 mg/mL), mix by
tapping.
11. Add 600 μL of buffer AL (mix the buffer thoroughly before
adding).
12. Vortex the tube for at least a minute.
13. Incubate in a 70 C 2 C water bath for 10 min. Mix by
inverting it 20 times and incubate again at 70 C 2 C for
another 10 min (see Note 3).
14. Add 500 μL of room temperature 100% Ethanol. Mix well by
vortexing for at least 1 min.
15. Transfer 700 μL of solution to QIAmp mini column in a 2 mL
collection tube, being careful not to wet the rim of the column.
16. Microcentrifuge at 6000 g for 1 min.
17. Discard the filtrate and return the column to the tube.
18. Repeat steps 15–17 as necessary to filter remaining solution.
19. Add 500 μL of buffer AW1 (add ethanol to the buffer when
using for the first time according to the manufacturer’s instruc-
tions), being careful not to wet the rim of the column.
20. Centrifuge for 1 min at 6000 g.
21. Discard the filtrate and return the column to the tube.
22. Add 500 μL of buffer AW2 (add ethanol to the buffer when
tion), being careful not to wet the rim of the column.
23. Centrifuge for 3 min at maximum speed.
24. Transfer the column to a new 1.5 mL tube, and centrifuge for
another minute at maximum speed.
25. Place the column in a clean 1.5 mL tube and add 15 μL of H2O
(see Note 4).
26. Incubate at room temperature for 5 min.
27. Centrifuge for 1 min at 6000 g.
28. Remove and discard the column and measure the DNA using
any type of UV spectrophotometer (see Notes 5–8).
3.2 DNA Extraction 1. Add 180 μL of Buffer ATL to the cleaned and prepared CVS
from Chorionic Villi sample (If a precipitate has formed in Buffer ATL, dissolve by
(See Note 9) incubating at 56 C).
2. Add 20 μL proteinase K, mix by vortexing, and incubate at
56 C until the tissue is completely lysed. Incubation time will
vary depending on the size of the tissue sample, with larger
samples requiring longer incubation. Vortex occasionally dur-
ing incubation to assist in lysis.
3. Briefly centrifuge the tube to remove drops from the inside of
the lid.
4. If necessary, transfer the solution to a 1.5 mL tube.
5. Add 4 μL RNase A (100 mg/mL), mix by pulse-vortexing for
15 s, and incubate for 2 min at room temperature. Briefly
centrifuge the tube to remove drops from inside the lid, then
add 200 μL Buffer AL to the sample. Mix again by pulse-
vortexing for 15 s, and incubate at 70 C for 10 min. Briefly
centrifuge the tube to remove drops from inside the lid (It is
essential that the sample and Buffer AL are mixed thoroughly
to yield a homogeneous solution).
6. Add 200 μL ethanol (96–100%) to the sample, and mix by
pulse-vortexing for 15 s. After mixing, briefly centrifuge to
remove drops from inside the lid (It is essential that the sample,
Buffer AL, and the ethanol are mixed thoroughly to yield a
homogeneous solution).
7. Carefully apply the mixture (including the precipitate) to the
QIAamp Mini spin column (in a 2 mL collection tube) without
wetting the rim. Close the cap, and centrifuge at 6000 g for
1 min. Place the QIAamp mini spin column in a clean 2 mL
collection tube, and discard the tube containing the filtrate.
8. Repeat step 7 as necessary to filter remaining mixture.
9. Carefully open the QIAamp Mini spin column and add 500 μL
Buffer AW1 (add ethanol to the buffer when using for the first
time according to the manufacturer’s instructions) without
wetting the rim. Close the cap, and centrifuge at 6000 g
for 1 min. Place the QIAamp Mini spin column in a clean 2 mL
collection tube, and discard the collection tube containing the

filtrate.
10. Carefully open the QIAamp Mini spin column and add 500 μL
Buffer AW2 (add ethanol to the buffer when using for the first
time according to the manufacturer’s instructions) without
wetting the rim. Close the cap and centrifuge at full speed for
3 min.
11. Place the QIAamp Mini spin column in a new 2 mL collection
tube and discard the old collection tube with the filtrate.
Centrifuge at full speed for 1 min.
12. Place the QIAamp Mini spin column in a clean 1.5 mL micro-
centrifuge tube, and discard the collection tube containing the
filtrate. Carefully open the QIAamp Mini spin column and add
25 μL distilled water. Incubate at room temperature for 5 min,
and then centrifuge at 6000 g for 1 min.
13. Remove and discard the column and Measure the DNA using
any type of UV spectrophotometer (see Notes 5–8). If the yield
of DNA is insufficient, an additional 20 μL of distilled water
can be added to the QIAmp Mini spin column, incubated, and
centrifuged as in step 12.
3.3 DNA Extraction 1. Remove the supernatant from a confluent cell culture flask.
from Cultured Cells Save the supernatant in a 15 mL falcon tube.
2. Add 3 mL of Trypsin to flask and incubate for 3 min in the
incubator.
3. Tap flask to loosen cells, and check that cells are loose under the
microscope.
4. Add 1 mL of the supernatant back to the flask to wash it and
transfer the complete solution to the same 15 mL falcon tube
(which contains the remaining supernatant).
5. Centrifuge for 10 min at 600 g. Remove the supernatant
leaving behind 200 μL residual fluid. Thoroughly suspend the
pellet in the residual fluid by pipetting up and down.
6. Add 600 μL of Cell Lysis Solution and resuspend the cells by
pipetting up and down or vortex on high speed for 10 s to lyse
the cells.
7. Incubate for 10 min at 37 and transfer to a 1.5 mL Eppendorf
tube. Cool to room temperature.
8. Add 3 μL of RNase A Solution, and mix by inverting 25 times.
Incubate for 15 min at 37 C. Then incubate at room temper-
ature for 3 min to quickly cool the sample.
9. Add 200 μL of Protein Precipitation Solution, and vortex
vigorously for 20 s at high speed.
10. Centrifuge for 3 min at maximum speed and transfer the

supernatant by carefully pouring into a clean 1.5 mL tube
containing 600 μL of 100% isopropanol. Be sure not to dis-
lodge the protein pellet during pouring.
11. Mix by inverting gently 50 times until the DNA is visible as
threads or a clump (see Note 10).
12. Centrifuge for 1 min at maximum speed. The DNA may be
visible as a small white pellet.
13. Carefully discard the supernatant, and drain the tube by invert-
ing on a clean piece of absorbent paper, taking care that the
pellet remains in the tube.
14. Add 600 μL of 70% ethanol and invert several times to wash the
DNA pellet.
15. Centrifuge for 1 min at maximum speed.
16. Carefully discard the supernatant. The pellet might be loose
and easily dislodged. Drain the tube on a clean piece of absor-
bent paper, taking care that the pellet remains in the tube.
17. Air dry the pellet for at least 15 min. Avoid over-drying the
DNA pellet, as the DNA will be difficult to dissolve.
18. Add TE buffer and vortex for 5 s at medium speed to mix (see
Note 11).
19. Measure the DNA using any type of UV spectrophotometer
(see Notes 5–8).
3.4 DNA Extraction 1. Bring samples to room temperature.

from Cell Free 2. Bring total volume to 5 mL with PBS if necessary.
Amniotic Fluid
3. Set up the QIAvac 24 plus vacuum manifold as shown in Fig. 1.
4. Heat water bath to 60 C and Heat block to 56 C.
5. Bring buffer AVE to room temperature.
6. Add reconstituted carrier RNA to Buffer ACL (this will vary
depending on number of samples being processed—Follow the
manufacturer’s instructions).
7. Pipet 500 μL Proteinase K into a 50 mL tube.
8. Add 5 mL of amniotic fluid to the tube.
9. Add 4 mL buffer ACL containing the carrier RNA.
10. Pulse vortex for 30 s.
11. Immediately incubate the tube in the preheated water bath
(60 C) for 30 min.
12. Add 9 mL Buffer ACB (add isopropanol to the buffer when
tion) to the lysate in the tube.
13. Pulse vortex for 30 s.
Fig. 1 Setting up the QIAvac 24 plus with the QIAamp mini columns
14. Incubate the mixture on ice for 5 min.

15. Set up the vacuum with the columns according to the instruc-
tions with the 20 mL extension tubes (see Fig. 1).
16. With the vacuum in the off position, carefully free pour the
mixture into the extension tube, and then turn the value to
approximately 5/6 of the way. Adjust if the solution is draining
too quickly (should be around 10–15 min for the entire solu-
tion to drain).
17. Once the solution has entirely drained, switch off the vacuum
immediately so the membrane does not dry out.
18. Remove the extenders.

19. Apply 600 μL Buffer ACW1 (add ethanol to the buffer when
tions) to the column carefully—Not disturbing the membrane.
20. Switch the vacuum on until the volume has drained and return
to the off position.
21. Apply 750 μL Buffer ACW2 (add ethanol to the buffer when
tions) to the column carefully—Not disturbing the membrane.
23. Apply 750 μL of 100% ethanol to the column carefully—Not
disturbing the membrane.
25. Close the lid of the column and place inside an empty
1.5 mL tube.
26. Spin the column inside the tube at maximum speed for 3 min.
27. Discard the tube and place the column in a new 1.5 mL tube.
28. Open the lid of the column and place the centrifuge tube in the
hot block for 10 min to dry the membrane completely.
29. Place the column in a new 1.5 mL centrifuge tube and discard
the tube it was previously in.
30. Carefully apply 10 μL of nuclease free water to the membrane
to elute the DNA.
31. Incubate with the lid closed for 5 min at room temperature.
32. Centrifuge at maximum speed for 1.5 min to elute the nucleic
acids.
33. Measure the DNA using any type of UV spectrophotometer
(see Notes 5–8).
4 Notes
1. We recommend working with at least 10 mL of amniotic fluid.

The greater the volume of fluid, the higher the likelihood of
obtaining a sufficient DNA yield.
2. To maximize the quantity of fetal DNA, further DNA extrac-
tion from the supernatant of the cell free DNA can be per-
formed (see Subheadings 2.4 and 3.4). This is especially helpful
when a low volume of amniotic fluid is received or if the cell
pellet is very small. For those laboratories that store the super-
natant, the cell free DNA can be extracted at a later date if
necessary. We have had success extracting DNA from superna-

tant that was frozen for greater than 4 months.
3. If necessary, the extraction procedure can be stopped at this
point and continued the next day, keeping the tube at room
temperature overnight.
4. Some commercially available kits suggest elution in provided
buffer or other salt containing solution. We prefer to elute
using molecular biology grade H2O, as this enables further
concentration of DNA if necessary.
5. We recommend using the Nanodrop ONE to measure the
DNA concentration.
6. The 260/280 ratio should be between 1.8 and 2.0 and the
260/230 ratio should be above 1.5.
7. Given the precious nature of the DNA, it may still be worth
using the DNA even when the 260/280 ration is above 2.1
and/or if the 260/230 ratio is below 1.0. Quality control
metrics should be established for each downstream assay that
you perform.
8. We highly recommend performing maternal cell contamination
studies on all DNA extracted for prenatal studies (see
Chapter 8).
9. We recommend starting with at least 3 mg of tissue. However,
we have obtained successful results for SNP microarray analysis
using less than 3 mg. As with amniotic fluid, the greater the
starting quantity of villi, the higher the likelihood of obtaining
a sufficient DNA yield.
10. A DNA pellet is not always visible after the next centrifugation
step, so it is a good idea to position the tubes in the centrifuge
in such a way that the location of the pellet will be consistent.
We suggest placing all tubes with the hinge pointed out.
11. Adjust the TE volume according to the size of the pellet. If the
DNA pellet can be easily seen, we recommend adding at least
100 μL of TE buffer.
References
1. Rebello MT, Hackett G, Smith J et al (1991) (ed) Genetic disorders and the fetus: diagnosis,
Extraction of DNA from amniotic fluid cells for prevention, and treatment, 6th edn. John
the early prenatal diagnosis of genetic disease. Wiley & Sons, Ltd, Oxford, UK
Prenat Diagn 11(1):41–46 4. Winsor EJ, Silver MP, Theve R et al (1996)
2. Williamson R, Eskdale J, Coleman DV et al Maternal cell contamination in uncultured
(1981) Direct gene analysis of chorionic villi: amniotic fluid. Prenat Diagn 16(1):49–54.
a possible technique for first-trimester antena- https://doi.org/10.1002/(SICI)1097-0223(
tal diagnosis of haemoglobinopathies. Lancet 2 199601)16:1<49::AID-PD808>3.0.CO;2-U
(8256):1125–1127 5. Saura R, Roux D, Taine L et al (1994) Early
3. Old JM (2011) Prenatal diagnosis of the amniocentesis versus chorionic villus sampling
hemoglobinopathies. In: Milunsky AMJ
for fetal karyotyping. Lancet 344 karyotype. Am J Hum Genet 75(3):485–491.

(8925):825–826 https://doi.org/10.1086/423288
6. Bianchi DW, LeShane ES, Cowan JM (2001) 9. Lapaire O, Johnson KL, Bianchi DW (2008)
Large amounts of cell-free fetal DNA are pres- Method for extraction of high-quantity and
ent in amniotic fluid. Clin Chem 47 -quality cell-free DNA from amniotic fluid.
(10):1867–1869 Methods Mol Biol 444:303–309. https://doi.
7. Hui L, Bianchi DW (2011) Cell-free fetal org/10.1007/978-1-59745-066-9_24
nucleic acids in amniotic fluid. Hum Reprod 10. Shaw KJ, Thain L, Docker PT et al (2009) The
Update 17(3):362–371. https://doi.org/10. use of carrier RNA to enhance DNA extraction
1093/humupd/dmq049 from microfluidic-based silica monoliths. Anal
8. Larrabee PB, Johnson KL, Pestova E et al Chim Acta 652(1–2):231–233. https://doi.
(2004) Microarray analysis of cell-free fetal org/10.1016/j.aca.2009.03.038
DNA in amniotic fluid: a prenatal molecular
Chapter 8
Assessment of Maternal Cell Contamination in Prenatal

Samples by Quantitative Fluorescent PCR (QF-PCR)
Christie M. Buchovecky, Odelia Nahum, and Brynn Levy
Abstract
When biopsying a fetal tissue like chorionic villi or amniotic fluid, there is a chance of getting some maternal
material that could contaminate the fetal specimen and might lead to a misdiagnosis. Thus, all prenatal
samples should be subjected to testing for maternal cell contamination. This is done using quantitative
fluorescent PCR (QF-PCR) of short tandem repeat (STR) markers.
Key words Maternal cell contamination, Prenatal diagnosis, Microsatellite markers, Short tandem
repeats, Genotyping, Sample identity
1 Introduction
Prenatal sampling procedures pose a risk of including maternal cells

alongside the intended fetal sample. This risk has been empirically
calculated as approximately 0.5% in amniotic fluid sampling, 1–2%
in chorionic villi sampling, and has the potential to be much higher
in products of conception depending on the method by which the
sample was obtained [1, 2]. Although best practices for obtaining
and processing samples can limit the amount and frequency of
contamination, misdiagnosis due to maternal cell contamination
(MCC) remains a concern in all prenatal diagnosis arenas
[3–5]. For this reason, the American College of Medical Genet-
ics and Genomics (ACMG) and the Association for Molecular
Pathology (AMP) recommend MCC testing as the standard of
care for each prenatal specimen obtained for molecular diagnosis
[6, 7].
The method described herein employs DNA genotyping by
quantitative fluorescent PCR (QF-PCR), followed by capillary elec-
trophoresis, on 15 highly polymorphic microsatellite or short tan-
dem repeat (STR) markers, and an X–Y specific marker
(amelogenin). This allows distinction between maternal and fetal
117
118 Christie M. Buchovecky et al.
D19S433 vWA TPOX
80 120 160 200 240
6000 Fetal Sample informative marker

4000 S
stutter peak N
2000 C
0
sz 117.69 sz 172.15 sz 242.62
ar 18609 ar 22645 ar 47475
sz 121.63 sz 180.20 sz 246.76

ar 10634 ar 35516 ar 22042
sz 125.53 sz 184.10
ar 22300 ar 12905
80 120 160 200 240
5700
Maternal Sample
3800
1900
0
sz 121.61 sz 180.30 sz 242.63
ar 24783 ar 40966 ar 78113
sz 125.51 sz 184.19
ar 22945 ar 39603
Fig. 1 Prenatal Sample with 36.3% Maternal Cell Contamination. Marker D19S433 is not informative, as the
maternal allele that is not shared with the fetus overlaps a stutter peak from the fetal peak at size 125. Marker
TPOX is not informative because all maternal peaks are shared with the fetus. Marker vWA is informative, with
a shared peak (S) at 180, a non-shared, non-contaminating peak (N) at 172, and a contaminating peak (C) at
184 in the prenatal sample. Using the equation for determining MCC with heterozygous fetal alleles (see
Subheading 3.1), the MCC calculated from this marker is 36.3%. Note that the allele pattern in a prenatal
sample with 100% MCC would match the exact maternal allele pattern, with no unshared peaks
DNA to estimate the presence and level of MCC in prenatal sam-

ples. The markers present in the putative fetal DNA sample (derived
from cultured or uncultured cells from amniotic fluid, chorionic
villi, cord blood, or products of conception) are compared to those
in a maternal DNA sample (typically from a blood or buccal speci-
men). By examining these samples in parallel and comparing the
relative ratios of alleles at each marker, it is possible to estimate the
presence and level of contaminating maternal cells in the fetal
sample (see Fig. 1).
Prevention of misdiagnosis, especially of falsely reporting nor-
mal or wild-type results, is the primary reason for undertaking
MCC testing for each prenatal diagnostic sample obtained, includ-
ing for gene sequencing and microarray testing. If MCC is not
detected, the diagnostic test result can be considered accurate at
the level of detection of the MCC assay (which is 5% when
Assessment of Maternal Cell Contamination in Prenatal Samples by. . . 119
following this protocol). Because each instance of sampling

includes a risk for MCC, it is important that this testing be per-
formed on DNA extracted from the same sample or culture used for
diagnostic testing. In addition to this routine use in diagnostic
tests, ruling out MCC is vital when products of conception show
a normal female karyotype (46, XX), prenatal karyotyping or FISH
studies suggest XX/XY mosaicism, or cytogenetic specimens show
discordant results between direct and cultured samples.
Prenatal samples are considered precious as each sampling pro-
cedure is associated with a small but definitive risk of miscarriage
[8]. The finding of MCC does not automatically dictate that a
repeat invasive sampling procedure be recommended/performed.
The mere presence of MCC does not preclude diagnostic testing,
so long as the test employed is robust to the level of contamination
observed. Common cutoffs include 10% for SNP oligonucleotide
analysis (SOMA) and 15% for direct sequencing-based tests. For
other assays, the MCC cutoff to reject a sample for diagnostic
testing should be calculated based on the experimentally ascer-
tained effect of MCC on the sensitivity and specificity of the diag-
nostic test. If the level of MCC in a sample is anticipated to interfere
with diagnostic testing, the availability of a cultured specimen pre-
sents an opportunity for further analysis. Culturing of amniotic
fluid is especially amenable to reduction of MCC, as culture con-
ditions favor growth of amniocytes over maternal blood cells
[9]. On the other hand, culturing can increase levels of MCC in
chorionic villi or product of conception samples where maternal
decidua remained present in the culture setup, making careful
processing of these samples imperative [4]. In cases where clinically
significant MCC persists in cultured samples, a request for repeat
sampling may be warranted.
In rare instances, a prenatal sample may be negative for MCC,
but produce an unexpected result in this assay. Discordance of
alleles between the fetal and maternal samples at multiple loci
could represent an egg donor, surrogate pregnancy, or laboratory
error. If the origin of this discordance is confirmed to be either an
egg donor or a surrogate pregnancy by the clinical team, the MCC
assay should be used to check for contamination with alleles unique
to the gestational carrier. In another scenario, additional,
non-maternal contaminating peaks may be observed when there
are multiple gestational pregnancies (see Fig. 2). In multiple gesta-
tion pregnancies, the presence of fetus-to-fetus contamination
should be noted as potentially impacting the accuracy of results,
but typically does not prevent diagnostic testing. If this pattern
occurs without a clinical record of a multiple gestational pregnancy,
the possibilities of a vanishing twin or fetal chimerism should be
considered. Chromosomal aneuploidy is suggested when a pattern
of three fetal alleles at one or more loci is observed on a single
chromosome; When observed on multiple chromosomes, triploidy
AMEL D5S818 FGA

105 135 165 195 225 255
2400 Twin A Sample

twin-twin
1600
bleed-through contamination
800
sz 108.65 sz 157.22 sz 224.42 sz 246.42

ar 28190 ar 12130 ar 8517 ar 2242
sz 161.33 sz 233.30
ar 11314 ar 10552
sz 237.64
ar 2950
105 135 165 195 225 255
1200
Twin B Sample
800
bleed-through
400
sz 108.64 sz 157.22 sz 237.56

ar 12898 ar 6309 ar 6202
sz 161.34 sz 246.40
ar 5775 ar 4639
105 135 165 195 225 255
4800 Maternal Sample

3200
1600
sz 108.53 sz 157.14 sz 233.13

ar 56517 ar 26310 ar 26454
sz 161.20 sz 237.50
ar 24949 ar 20821
Fig. 2 Twin-Twin Contamination. Marker AMEL indicates the sex of each sample as female. Marker D5S818 is
not informative because both alleles are shared between all samples. Marker FGA shows Twin A contaminated
with cells from Twin B. The Twin B sample appears non-contaminated. Although the contaminating peak at
size 237 is shared with both the maternal sample and twin B, the peak at 246 is found only in twin B. From the
relative sizes of these two peaks and similar observations at other markers, we can conclude most of the
contamination is of co-twin, rather than maternal origin. Additionally, this figure shows an example of a
non-biological peak between the ranges for markers D5S818 and FGA; this is due to bleed-through of signal
from another fluorescent channel
D8S1179 D21S11 D7S820 CSF1PO

120 150 180 210 240 270 300 330
Fetal Sample - Triploid
sz 142.27 sz 201.66 sz 225.44 sz 278.78 sz 325.64

ar 36629 ar 49923 ar 20626 ar 44950 ar 53779
sz 146.99 sz 286.17 sz 329.77

ar 30768 ar 19494 ar 23599
sz 151.53
ar 31884
120 150 180 210 240 270 300 330
Maternal Sample
sz 142.17 sz 214.71 sz 278.84 sz 321.63

ar 37240 ar 26281 ar 53601 ar 30657
sz 146.86 sz 225.44 sz 325.76

ar 34350 ar 24339 ar 28025
Fig. 3 A Prenatal Sample with Triploidy. In the fetal sample, marker D8S1179 shows three peaks of roughly
equal area, while the other three markers each show two peaks with one double the area of the other,
indicating three alleles at each marker. A similar pattern could be seen with high MCC. However, this can be
excluded in this sample based on marker D21S11, where the larger peak is not shared with the maternal
sample, suggesting that the mechanism of triploidy in this fetus was likely caused by dispermy. If a similar
pattern of 3 equal peaks or a 2:1 ratio of peaks occurs at a single marker in a sample without MCC, the
possibility of trisomy should be considered
is the likely explanation (see Fig. 3). In instances where allele pat-
terns suggest a chromosome abnormality, appropriate diagnostic
testing should be performed to confirm this finding. Rarely, a fetal
sample may appear discordant to the maternal sample at a single
locus. Explanations for this phenomenon include mutation at the
primer site causing allelic dropout or de novo tandem repeat expan-
sion or contraction (see Fig. 4). Given a sufficient number of addi-
tional markers, the identity of the maternal specimen can still be
assured if there is concordance between the maternal fetal samples
at all other loci. Frequencies of null and de novo mutation rates
among many markers have been ascertained in the literature; data
for those used in this assay are provided with the PCR kit.
In conclusion, assessment of maternal cell contamination by
QF-PCR is an effective means of quickly assuring the identity and
sufficient purity of prenatal samples, thereby preventing a misdiag-
nosis that may otherwise occur. The permissible level of MCC for a
given diagnostic assay should be empirically determined in each
laboratory prior to implementation. Additionally, the data obtained
from MCC testing can suggest aneuploidy in a fetus, which should
D3S1358 TH01 D13S317 D16S539

90 120 150 180 210 240 270 300
6900 Fetal Sample discordant

4600 alleles
2300
0
sz 121.87 sz 172.47 sz 220.57 sz 246.25 sz 278.91
ar 48265 ar 42825 ar 46200 ar 39359 ar 48947
sz 126.04 sz 184.40 sz 286.26

ar 49638 ar 43485 ar 39860
90 120 150 180 210 240 270 300
6900 Maternal Sample

4600
2300
0
sz 121.73 sz 172.43 sz 224.88 sz 246.05 sz 275.27
ar 33176 ar 56462 ar 31308 ar 27703 ar 31817
sz 125.75 sz 282.61
ar 28817 ar 30180
Fig. 4 A prenatal Sample with a Single Discordant Marker. The first three markers (and all other markers from
the sample) are consistent with a matched maternal-fetal pair without MCC. However, the fetal sample does
not share any alleles with the maternal sample at marker D16S539. In this scenario, it is most likely that the
maternal STR allele assessed by this marker expanded or contracted by one repeat between generations as
the probability that a sample swap occurred with the mother still matching 14/15 loci is extremely low
then be confirmed by traditional diagnostic means. Lastly, use of

this assay prior to performing diagnostic testing reduces costs in a
clinical laboratory by allowing a director to avoid testing incorrect
or contaminated samples that would require reanalysis.
2 Materials
The protocol described here is for running the AmpFLSTR Identi-

filer PCR Kit PLUS on the ABI 3130 DNA analyzer (Applied
Biosystems, Foster City, CA, USA). The protocol is expected to
be similar but not identical using other PCR kits and/or DNA
analyzers. Please consider the manufacturers’ advice when adapting
and optimizing this protocol for use with different equipment and
reagents.
2.1 MCC Kit 1. AmpFLSTR Identifiler PCR Kit PLUS (Applied Biosystems,
Foster City, CA, USA).
2. HiDi Formamide (Applied Biosystems, Foster City, CA, USA).
3. GeneScan™ 500 LIZ™ dye Size Standard (Applied Biosys-
tems, Foster City, CA, USA).
2.2 DNA Extraction DNA can be extracted using any commercial kit (see Chapter 7).
2.3 ABI 3130 1. POP-7™ Polymer for 3130/3130xl Genetic Analyzers

(Applied Biosystems, Foster City, CA, USA).
2. 310 and 31xx Running Buffer, 10 (Applied Biosystems, Fos-
ter City, CA, USA).
3. 96-well optical reaction plate (Applied Biosystems, Foster City,
CA, USA).
3 Methods
Table 1 shows various troubleshooting solutions when problems

are encountered during PCR and electrophoresis.
3.1 Preparation The spiked positive controls are prepared from DNAs extracted
of 5% and 10% Spiked from a mother and fetus pair that is verified as being
Positive Controls non-contaminated.
Table 1
PCR & Electrophoresis troubleshooting
Problem Possible cause Solution

Weak or no signal at all loci, Absent/insufficient master mix Repeat PCR
including controls or primers
Master mix not mixed Vortex master mix thoroughly
thoroughly
Primer set exposed to too much Protect primer set from light
light
PCR system malfunction Check the system
Incorrect thermal cycler Check cycler for correct parameters
parameters
Insufficient PCR product Mix 1 μL PCR product, 20 μL
injected Formamide, and 0.5 μL standard
Degraded Formamide Avoid multiple freeze/thaw cycles
Positive signal from control Sample DNA is degraded Evaluate quality. If degraded,
DNA but not test samples re-amplify with more DNA
Quantity of sample DNA is low Quantitate DNA and repeat PCR
using 1 ng
Sample DNA contains high Purify DNA and repeat PCR
amount of PCR inhibitors
Not all loci visible Sample DNA is degraded Evaluate DNA quality. If degraded,
re-amplify with more DNA
Sample DNA contains high Purify the DNA and repeat PCR
amount of PCR inhibitors
Low peak heights Incorrect thermocycler Check parameters
parameters
1. Dilute the DNA of both maternal and fetal control samples

to approximately 1 ng/μL using molecular grade water (see
Note 1).
2. Prepare the 5% and 10% spiked positive control as follows (see
Note 2):
l 5%: Take 1.5 μL of maternal control DNA (1 ng/μL) and
add it to 28.5 μL of fetal control DNA (1 ng/μL).
l 10%: Take 3 μL of maternal control DNA (1 ng/μL) and
add it to 27 μL of fetal control DNA (1 ng/μL).
3. Store the original diluted maternal and original diluted
(non-spiked) fetal control samples as well as the spiked control
DNAs at 20 C.
3.2 DNA Extraction Extract DNA from maternal blood and fetal sample using your
standard prenatal DNA extraction methods (see Chapter 7).
3.3 PCR 1. Dilute the DNA of both maternal and fetal samples to approxi-
Amplification mately 1 ng/μL using molecular grade water.
2. Thaw the AmpFLSTR Identifiler Plus Kit Master Mix and the
AmpFLSTR Identifiler Plus Kit Primer Set, then vortex 3 s and
centrifuge briefly before opening the tubes (see Note 3).
3. Prepare the PCR master mix using reagents in the AmpFLSTR
Identifiler PCR Plus Kit:
For each sample use 10 μL of PCR mix and 5 μL of the
primer set.
4. Add 2 μL of the diluted DNAs to a well of the 96-well plate
(one well per sample) (see Notes 4 and 5).
5. Aliquot 15 μL of the PCR master mix into each well of the
plate.
6. Seal the plate, vortex for 3 s, and short spin (about 20 s) in a
tabletop centrifuge to remove bubbles.
7. Place the plate in a pre-warmed thermal cycler and run the
following program (approximately 3 h):
l Twenty nine cycles: 94 C for 20 sec, 59 C for 3 min.
l 4 C hold.
8. Short spin the plate.
3.4 Sizing the DNA 1. Prepare a master mix using 20 μL of formamide and 0.5 μL of
Fragments LIZ size standard per sample.
2. In a new 96-well optical reaction plate, add 20.5 μL of master
mix and 1 μL of PCR product (see Note 6).
3. Denature for 3 min at 95 C and put on ice for another 3 min.

4. Spin the plate and run on the ABI 3130 Genetic Analyzer.
3.5 Analysis and 1. Import data into a genotyping analysis software according to
Calculation that software’s microsatellite guidelines. Data for a sample will
of Maternal Cell include size, peak height, and area for each identified allele.
Contamination 2. Identify informative markers for each mother-fetus pair (see
Note 7).
l Informative marker: One maternal allele differs from both
fetal alleles for that marker and does not overlap with a fetal
stutter peak (stutter peaks occur one microsatellite repeat
less than the main allelic peak).
3. For each informative allele, determine contamination using the
appropriate formula below (see Note 8):
l Informative markers with heterozygous fetal allele: % con-
tamination ¼ area(C*)/[area(N**) + area(C*)].
l Informative markers with homozygous fetal allele: % contam-
ination ¼ [area(C*) 2] / [area(S***) + area(C*)].
*
Contaminating peak (C): peak at size of maternal allele
that is not shared with the fetus.
**
Non-shared, non-contaminating peak (N): fetal allele that
is not shared with mother.
***
Shared peak (S): fetal allele that is common to the mother
and fetus.
4. Calculate the mean contamination for all informative markers to
determine the maternal cell contamination for that sample (see
Fig. 1). Informative markers where no contaminating peak is
observed should be included as a marker with zero contamination.
3.6 Interpretation of l 100% MCC: Maternal and fetal samples are identical at all loci
Results tested, indicating the “fetal” specimen likely represents maternal
cells. This has been reported in samples from products of con-
ception. Possible diagnostic errors including sample mix-ups,
genotyping errors, rare genetic variants which interfere with
analysis, and other sources should be excluded.
l Mid-level MCC (>10%): Significant maternal cell contamination
is detected in DNA extracted from this fetal specimen. Mutation
analysis or SOMA of the fetal sample cannot be interpreted due
to the amount of maternal cells detected. Recommend retesting
fetal sample from cultured cells, or another direct sample if the
current fetal sample was cultured.
l Low-level MCC (5–10%): Low-level maternal cell contamination
was detected in DNA extracted from this fetal specimen. Some
tests may be amenable to testing despite low-level
contamination. If this is not the case, recommend retesting fetal

sample from cultured cells if the current MCC analysis was based
on a direct sample, or recommend another direct sample if the
current MCC analysis was based on cultured cells.
l Negative for MCC: No evidence of maternal cell contamination in
the fetal specimen at the level detectible by this assay (see Note 9).
4 Notes
1. The diluted maternal and non-spiked fetal control samples

(1 ng/μL) will also be utilized as control samples with every
MCC run (see Note 4).
2. The 5% and 10% spiked positive controls usually last for about
5 runs. They should be prepared fresh when the final calcula-
tions of a clinical run indicate drift from their expected percent-
age value, i.e., if the 5% appears more like 8% and the 10%
appears more like 14%, new spiked control batches should be
prepared.
3. Thawing is required only during first use of the kit. After first
use, reagents are stored at 2–8 C and, therefore, they do not
require subsequent thawing. Do not refreeze the reagents.
4. For each MCC run, you will be running 6 DNA control
samples as follows: (1) Fetal control DNA (see Note 1),
(2) maternal control DNA (see Note 1), (3) 5% spiked positive
control, (4) 10% spiked positive control, (5) non-template
control (nuclease-free molecular grade water), and (6) control
DNA that comes with the AmpFLSTR Identifiler PLUS
PCR Kit.
5. When the total concentration of the entire fetal DNA sample is
very low (3–10 ng/μL) it is better to use 3 μL of the diluted
fetal DNA sample instead of 2 μL.
6. When the total concentration of the entire fetal DNA sample is
very low (3–10 ng/μL) it is better to use 2 μL of the fetal PCR
product instead of 1 μL.
7. When the fluorescent signal in a sample is strong, peaks can be
seen that represent neither an allele from the individual tested,
nor a biological contaminant. These bleed-through peaks
occur in neighboring channels because the fluorescent filters
do not have perfect specificity for their fluorophores. When an
anomalous peak is observed in one channel that matches the
size of an intense biological peak in another channel, it is likely
a bleed-through peak rather than a sign of contamination.
8. When determining whether a peak is shared between the
maternal and fetal sample, the size of the maker may vary by
+/1 due to small differences in capillary electrophoresis runs
between samples.
9. At least 2 informative markers must be present to report

absence of maternal cell contamination. Peak heights of infor-
mative markers should be 1000rfu for heterozygous alleles and
2000rfu for homozygous alleles, to eliminate the possibility of
contamination at a 5% level (contaminating allele will then be
approximately 50rfu). This minimum does not apply when
maternal cell contamination is clearly detected.
References
1. Hsu LY (1992) Prenatal diagnosis of chromo- interpretation, and reporting of maternal cell
somal abnormalities through amniocentesis. In: contamination in prenatal analyses a report of
Milunsky A (ed) Genetic disorders and the fetus: the association for molecular pathology. J Mol
diagnosis, prevention and treatment, 3rd edn. Diagn 13(1):7–11. https://doi.org/10.1016/j.
Johns Hopkins University Press, Baltimore jmoldx.2010.11.013
2. Ledbetter DH, Zachary JM, Simpson JL et al 7. South ST, Lee C, Lamb AN et al (2013) ACMG
(1992) Cytogenetic results from the standards and guidelines for constitutional cyto-
U.S. collaborative study on CVS. Prenat Diagn genomic microarray analysis, including postnatal
12(5):317–345 and prenatal applications: revision 2013. Genet
3. Ledbetter DH (1993) Prenatal cytogenetics: Med 15(11):901–909. https://doi.org/10.
indications, accuracy and future directions. In: 1038/gim.2013.129
Simpson JL, Elias S (eds) Essentials of prenatal 8. Akolekar R, Beta J, Picciarelli G et al (2015)
diagnosis. Churchill Livingstone, New York Procedure-related risk of miscarriage following
4. Saura R, Roux D, Taine L et al (1994) Early amniocentesis and chorionic villus sampling: a
amniocentesis versus chorionic villus sampling systematic review and meta-analysis. Ultrasound
for fetal karyotyping. Lancet 344 Obstet Gynecol 45(1):16–26. https://doi.org/
(8925):825–826 10.1002/uog.14636
5. Weida J, Patil AS, Schubert FP et al (2017) 9. Winsor EJ, Silver MP, Theve R et al (1996)
Prevalence of maternal cell contamination in Maternal cell contamination in uncultured
amniotic fluid samples. J Matern Fetal Neonatal amniotic fluid. Prenat Diagn 16(1):49–54.
Med 30(17):2133–2137. https://doi.org/10. https://doi.org/10.1002/(SICI)1097-0223(
1080/14767058.2016.1240162 199601)16:1<49::AID-PD808>3.0.CO;2-U
6. Nagan N, Faulkner NE, Curtis C et al (2011)
Laboratory guidelines for detection,
Chapter 9
Rapid Prenatal Aneuploidy Screening by Fluorescence In

Situ Hybridization (FISH)
Anja Weise and Thomas Liehr
Abstract
The most common aneuploidies observed in prenatal diagnostics in the second trimester are trisomies of
the chromosomes 13, 18 or 21 and gonosomal abnormalities. Rapid detection of these aneuploidies after
amniocentesis is possible by fluorescence in situ hybridization (FISH) utilizing centromeric or locus-specific
probes. FISH aneuploidy screening results in uncultured amniocytes are available within 24 h or less.
Operators should be aware that there are possible pitfalls in connection with the commercially available
probe sets and in result interpretation in general and thus proceed with appropriate caution. Here, we
explain how rapid prenatal aneuploidy screening is performed using the Food and Drug Administration
(FDA-) approved Aneu Vysion kit (ABBOTT/Vysis) and a review is given of drawbacks and opportunities
of the method.
Key words Prenatal diagnosis, Molecular cytogenetics, Fluorescence in situ hybridization (FISH),
Interphase FISH, Pitfalls
1 Introduction
In the 1980s it took around 3–4 weeks to obtain a fetal karyotype

from amniotic fluid cells, mainly due to the necessity for cell cultur-
ing [1]. This timeframe was shortened to around 2 weeks [2], due
to progress in cytogenetic techniques, i.e., in cell growth media,
but also in cell preparation [3]. Even though this is a relatively short
time compared to the 3–4 weeks necessary in the 1980s, it was
recognized that such moderately long waiting times cause psycho-
logical distress for the pregnant women [1].
This was one of the main causes for the introduction of molec-
ular (cytogenetics) methods for prenatal diagnosis of the most
common chromosome disorders in the second trimester: trisomy
13, 18, and 21, monosomy X and other gonosomal aberrations, as
well as triploidy [4]. Molecular techniques can be performed
directly on uncultured fetal cells, obviating the time consuming
process of cell culture and allowing for results to be obtained within
129
130 Anja Weise and Thomas Liehr
24 h. The quantitative fluorescence polymerase chain reaction

(QF–PCR), Multiplex ligation-dependent probe amplification
(MLPA), or the fluorescence in situ hybridization (FISH) are
three popular methods utilized in clinical practice [1]. Despite
some key methodological advantages unique to FISH (like single
cell analysis and low level mosaicism detection), aneuploidy screen-
ing by FISH has virtually been replaced by q-PCR in many places in
Europe. However, in the USA, it remains the primary rapid aneu-
ploidy screening method of choice. Nowadays, where chromo-
somal microarray analysis is fast becoming the first line test in
invasive prenatal diagnostics, rapid molecular screening tests (like
FISH) still make sense in terms of saving time and resources for the
detection of frequent human aneuploidies.
The only Food and Drug Administration (FDA-) approved
FISH-test for rapid aneuploidy screening in uncultured amniocytes
cells is the Aneu Vysion kit (commercially available at ABBOTT/
Vysis). It consists of two locus-specific probes for 13q14 (LSI 13)
and 21q22.13~22.2 (LSI 21) (Fig. 1) and three alpha-satellite
DNA-probes for chromosomes X, Y, and 18 (cep X, cep Y, and
cep 18) (Fig. 2). The two locus specific and the three centromeric
probes are applied in two different hybridization experiments to the
samples.
One has to be aware of the fact, that other genetic disorders like
(un)balanced structural rearrangements, numerical aberrations
beside the five tested chromosomes, microdeletions/microduplica-
tions, uniparental disomy, or mutations detectable only by molecu-
lar genetics, are not excluded after obtaining a “normal” result for
the rapid aneuploidy screening by FISH. To obtain a reliable result
50–100 interphase nuclei have to be evaluated per probe mix; for all
five probes cut-off rates of ~10% were suggested recently [4].
Published studies on the use of FISH for rapid aneuploidy
screening include tens of thousands of cases and they all show
concordance rates in excess of 99% between the results obtained
by FISH and those seen by standard G-banded karyotyping
[4–10]. However, there are always single case reports of false-
positive or false-negative results that are due to centromeric poly-
morphisms, the presence of small supernumerary marker chromo-
somes, dicentric chromosomes not detectable in interphase nuclei,
or maternal contamination [4–14]. As recently suggested, all the
observed pitfalls and reported misdiagnoses (apart from maternal
contamination) could easily be prevented by the exclusive use of
locus-specific probes [13]. The advantage of such probes, like the
LSI 13 and LSI 21 probes in the FDA kit, was proven, e.g., by the
fact that the LSI 21 probe is better suited to distinguish between
free and translocation trisomy 21 in the interphase [15].
Rapid Aneuploidy Screening by FISH 131
Fig. 1 (a) The distribution of the probes LSI13 (SpectrumOrange) and LSI21
(SpectrumGreen) is depicted, which comprise probe set 1. (b) A nucleus with
normal signal distribution of LSI13 and LSI21 indicating—if present in at least
45 of 50 evaluated nuclei—for the absence of a numerical aberration of
chromosomes 13 and 21 within the fetus. The nucleus is counterstained in
blue by DAPI. (c) In this nucleus there are two signals for LSI13 and three for
LSI21 indicative of a (free or translocation) trisomy 21
In summary, the rapid prenatal FISH aneuploidy test is a pow-

erful tool for the clinician in the care for pregnant women. It should
be offered to patients with appropriate genetic counseling, discuss-
ing the benefits and limitations of the test. This robust rapid
prenatal FISH test may also be utilized on preimplantation
embryos [16] and prenatal chorionic villi samples [17].
Fig. 2 (a) The three centromeric probes cepX, cepY, and cep18 are labeled in
SpectrumGreen, SpectrumOrange, and SpectrumBlue, respectively, in probe set
2. (b) A nucleus with one signal, each, for cepX and cepY are seen in this cell of a
male fetus. Additionally, two signals only are present for the cep18 probe and
thus, with high probability a free trisomy 18 is excluded. (c) In this case three
signals for cep18, but as well two signals for cepX and one for cepY were
present in each of the analyzed nuclei. As also three signals, each, for LSI13 and
LSI21 were detected (results not shown), a clinically suspected triploidy could be
confirmed by FISH and the pregnancy was terminated
2 Materials
2.1 Preparation of 1. Trypsin/EDTA. Pre-warm 3 mL per case (37 C).

Amniotic Fluid Cells 2. Mix 1 PBS with fetal calf serum v/v 4/1. Pre-warm 5 mL per
case (37 C).
3. 0.075 M KCl.
4. Carnoy’s fixative: 3:1 Methanol:Glacial acetic acid.
2.2 Slide 1. 20 SSC stock solution: 3.0 M NaCl, 0.3 M Na-citrate; set up
Pretreatment with double distilled water, adjust to pH 7.0, autoclave and
store at room temperature.
2. Pepsin stock solution 10% (w/v): dissolve 100 mg pepsin in
1 mL of filtered double distilled water at 37 C; aliquot and
store at 20 C.
3. Pepsin-buffer: Add 1 mL of 1 M HCl to 99 mL of distilled
water and incubate at 37 C for about 20 min; then add 50 μL
of the pepsin stock solution 10% (w/v) and leave the coplin jar
at 37 C; make fresh as required.
4. 1 PBS/ MgCl2: 5% (v/v) 1 M MgCl2 in 1 PBS.
2.3 Slide 1. Denaturation-buffer: 70% (v/v) deionized formamide, 10%

Denaturation (v/v) filtered double distilled water, 10% (v/v) 20 SSC,
10% (v/v) phosphate buffer; make fresh as required.
2. Deionized formamide: Add 5 g of ion exchanger Serdolit
MB-1 (Serva, Heidelberg, Germany) to 100 mL of formamide,
stir for 2 h (room temperature) and filter twice through What-
mann no. 1 filter paper. Aliquot and store at 20 C.
3. Phosphate buffer: prepare 0.5 M Na2HPO4 and 0.5 M
NaH2PO4, mix these two solutions (1:1) to get pH 7.0, then
aliquot and store at 20 C.
2.4 Hybridization 1. Aneuvision Kit (Abbott Molecular, Des Plaines, IL, USA).
2.5 Posthy- 1. 20 SSC stock solution: 3.0 M NaCl, 0.3 M Na-citrate; set up
bridization Washing with double distilled water, adjust to pH 7.0, autoclave, and
store at room temperature.
2. DAPI-solution: Dissolve 5 μL of DAPI (4,6-diamidino-2-phe-
nylindol.2HCl stock-solution) in 100 mL 4 SSC/0.2%
Tween; make fresh as required.
3. Vectashield H-1000 Antifade Mounting Medium (Vector
Laboratories, Burlingame, CA, USA).
3 Methods
This section describes how amniotic fluid cells are prepared, how
the target of the hybridization, i.e., cytogenetic slides with inter-
phase cells, have to be pretreated and how FISH itself is performed
and evaluated.
3.1 Preparation of For the rapid aneuploidy screening it is necessary to prepare the
Amniotic Fluid uncultured amniocytes as recommended in the AneuVysion kit
(AF) Cells protocol.
1. Place 2–3 mL of AF into a 15 mL reaction tube and centrifuge

(175 g for 5 min); discard the supernatant (see Note 1).
2. Resuspend the pellet in 3 mL trypsine/EDTA and incubate for
15 min at 37 C.
3. Add 1 PBS/foetal calf serum (4/1), centrifuge (175 g for
5 min—discard the supernatant).
4. Resuspend the pellet in 5 mL 0.075 M KCl and incubate at
37 C for 20 min.
5. Add 2 mL Carnoy’s fixative, centrifuge (175 g for 5 min—
discard the supernatant).
6. Add 3 mL Carnoy’s fixative, resuspend and incubate at 20 C
for 5 min.
7. Finalize the preparation with one last centrifugation (175 g
for 5 min); discard the supernatant and dilute the cells in
200 μL of the remaining supernatant.
8. Dispense the entire remaining suspension on two small regions
of a single dry and clean slide and air dry for 10 min at room
temperature.
9. Dehydrate in an ethanol series (70%; 90%; 100%, for 3 min
each) and air dry.
3.2 Slide 1. Incubate slides in 2 SSC for 1 min at room temperature (in a
Pretreatment 100 mL coplin jar on a shaker).
2. Replace 1 PBS with 100 mL pre-warmed pepsin-buffer
(37 C) and incubate the slides for 5 min at 37 C, without
agitation (see Note 2).
3. Replace pepsin-buffer with 100 mL 1 PBS/MgCl2, incubate
at room temperature for 5 min with gentle agitation. MgCl2
will block the enzymatic activity of pepsin.
4. Postfix nuclei on the slide surface by replacing 1 PBS/MgCl2
with 100 mL of formalin-buffer for 10 min (room tempera-
ture, with gentle agitation).
5. Formalin-buffer is replaced by 100 mL 1 PBS for 2 min
(room temperature, with gentle agitation).
6. Finally, dehydrate slides through an ethanol series (70%, 90%,
100%, 3 min each) and then air dry (see Note 3).
3.3 Fluorescence In The FISH procedure is itself divided into several steps: denatur-
Situ Hybridization ation, hybridization, posthybridization washing, and evaluation.
(FISH) and Evaluation
3.3.1 Denaturation of 1. Add 100 μL denaturation-buffer to the slides and cover with
Target DNA (24 50 mm) coverslips.
2. Incubate the slides on a heating plate for 3 min at 75 C (see

Note 2).
3. Remove the coverslips immediately by forceps and place the
slides in a coplin jar filled with 70% ethanol (4 C) to conserve
target DNA as single strands.
4. Dehydrate the slides in ethanol (70%, 90%, 100%, 4 C, 3 min
each) and air dry.
3.3.2 Hybridization 1. For each slide to be hybridized add 8 μL of the probe solution
1 (LSI 13 and LSI 21) on one region of the slide and 8 μL of
the probe solution 2 (cep 18, cep X and cep Y) on the dry slide,
put 20 20 mm coverslips on the drops and seal with rubber
cement.
2. Incubate slides overnight at 37 C in a humid chamber (see
Note 4).
3.3.3 Posthybridization 1. Take the slides out of 37 C humid chamber and remove
Washing rubber cement with forceps.
2. Place slides in 4 SSC/0.2%Tween (room temperature,
100 mL coplin jar) and allow coverslips to slide off (see Note 5).
3. Postwash the slides 1 2 min in 0.4 SSC (56–62 C) fol-
lowed by 1 1 min in 4 SSC/0.2% Tween (100 mL, room
temperature).
4. Counterstain the slides with DAPI-solution (100 mL in a
coplin jar, room temperature) for 8 min.
5. Wash the slides in water for a few seconds and air dry.
6. Add 15 μL of Vectashield H-1000 antifade mounting medium,
cover with coverslips, and look at the results in a fluorescence
microscope.
3.3.4 Evaluation 1. Evaluate 50 interphase nuclei per case under the fluorescence
microscope. This is a semi-statistic evaluation counting 1, 2,
3, or 4 signals for each probe. In case of a questionable result
within the cut-off region, enhance evaluated nuclei to 100 or
more (see Note 6).
4 Notes
1. During the whole preparation avoid taking the cells up with a

pipette—this would only lead to undesired loss of cells; only
mix with supernatant by moving the tube.
2. Pepsin pretreatment conditions, as well as denaturation time of
the target DNA should be tested in each laboratory on a single
slide first. If pepsin concentration is too stringent, it can result
in clean slides without any remaining nuclei.
3. The pretreated slides can be hybridized immediately or stored

at room temperature for up to 3 weeks. If longer storage is
necessary, the slides are stable at 20 C for several months.
4. Incubation can be stopped, if necessary, after 3 h. However,
this may result in the signals being too weak for evaluation.
This is especially true of the LSI-probes.
5. During the washing steps, it is important to prevent the slide
surfaces from drying out, otherwise background problems may
arise.
6. As long as not more than 7%/10% of the studied cells present
with 3 specific signals, no trisomy of the corresponding chro-
mosomal region is suspected and a “normal” report can be
issued. Between 7%/10% and 20%, the evaluation can be
regarded as an “unclear result,” which is with high probability
a “normal” result. However, in such cases, try to evaluate
100 or more nuclei for the probe in question to come to a
final decision. A value of >20% is indicative at minimum of a
mosaic trisomy. We suggest different cut-off rates for female
and male fetuses for optimal interpretation of the result
[4]. This is due to the fact that maternal cell-contamination is
obvious in the amniotic fluid of a male fetus but is not discern-
ible in a female fetus. Thus, the results of female fetuses should
be interpreted with slightly more care and the cut-off rates
lowered. Similar suggestions are made for the handling of
XXX, XYY, or XXY results.
References
1. Hulten MA, Dhanjal S, Pertl B (2003) Rapid Lancet 366(9480):123–128. https://doi.org/

and simple prenatal diagnosis of common chro- 10.1016/S0140-6736(05)66790-6
mosome disorders: advantages and disadvan- 6. Eiben B, Trawicki W, Hammans W et al (1999)
tages of the molecular methods FISH and Rapid prenatal diagnosis of aneuploidies in
QF-PCR. Reproduction 126(3):279–297 uncultured amniocytes by fluorescence in situ
2. Held KR (2003) QS Zytogenetik Bericht hybridization. Evaluation of >3,000 cases.
2002/2003. Med Genet 15:420–421 Fetal Diagn Ther 14(4):193–197. https://
3. Claussen U, Ulmer R, Beinder E et al (1993) doi.org/10.1159/000020919
Rapid karyotyping in prenatal diagnosis: a 7. Leung WC, Waters JJ, Chitty L (2004) Prena-
comparative study of the ’pipette method’ and tal diagnosis by rapid aneuploidy detection and
the ’in situ’ technique for chromosome har- karyotyping: a prospective study of the role of
vesting. Prenat Diagn 13(12):1085–1093 ultrasound in 1589 second-trimester amnio-
4. Liehr T, Ziegler M (2005) Rapid prenatal diag- centeses. Prenat Diagn 24(10):790–795.
nostics in the interphase nucleus: procedure https://doi.org/10.1002/pd.985
and cut-off rates. J Histochem Cytochem 53 8. Tepperberg J, Pettenati MJ, Rao PN et al
(3):289–291. https://doi.org/10.1369/jhc. (2001) Prenatal diagnosis using interphase
4B6394.2005 fluorescence in situ hybridization (FISH):
5. Caine A, Maltby AE, Parkin CA et al (2005) 2-year multi-center retrospective study and
Prenatal detection of Down’s syndrome by review of the literature. Prenat Diagn 21
rapid aneuploidy testing for chromosomes (4):293–301
13, 18, and 21 by FISH or PCR without a 9. Weremowicz S, Sandstrom DJ, Morton CC
full karyotype: a cytogenetic risk assessment. et al (2001) Fluorescence in situ hybridization
(FISH) for rapid detection of aneuploidy: 14. Skinner JL, Govberg IJ, DePalma RT et al
experience in 911 prenatal cases. Prenat (2001) Heteromorphisms of chromosome
Diagn 21(4):262–269. https://doi.org/10. 18 can obscure detection of fetal aneuploidy
1002/pd.39 by interphase FISH. Prenat Diagn 21
10. Witters I, Devriendt K, Legius E et al (2002) (8):702–704
Rapid prenatal diagnosis of trisomy 21 in 5049 15. Liehr T, Starke H, Beensen V et al (1999)
consecutive uncultured amniotic fluid samples Translocation trisomy dup(21q) and free tri-
by fluorescence in situ hybridisation (FISH). somy 21 can be distinguished by interphase-
Prenat Diagn 22(1):29–33 FISH. Int J Mol Med 3(1):11–14
11. Estabrooks LL, Hanna JS, Lamb AN (1999) 16. Mir P, Rodrigo L, Mateu E et al (2010)
Overwhelming maternal cell contamination in Improving FISH diagnosis for preimplantation
amniotic fluid samples from patients with oli- genetic aneuploidy screening. Hum Reprod 25
gohydramnios can lead to false prenatal inter- (7):1812–1817. https://doi.org/10.1093/
phase FISH results. Prenat Diagn 19 humrep/deq122
(2):179–181 17. Toutain J, Epiney M, Begorre M et al (2010)
12. Liehr T, Beensen V, Hauschild R et al (2001) First-trimester prenatal diagnosis performed
Pitfalls of rapid prenatal diagnosis using the on pregnant women with fetal ultrasound
interphase nucleus. Prenat Diagn 21 abnormalities: the reliability of interphase fluo-
(5):419–421. https://doi.org/10.1002/pd. rescence in situ hybridization (FISH) on mes-
44 enchymal core for the main aneuploidies. Eur J
13. Liehr T, Schreyer I, Neumann A et al (2002) Obstet Gynecol Reprod Biol 149(2):143–146.
Two more possible pitfalls of rapid prenatal https://doi.org/10.1016/j.ejogrb.2009.12.
diagnostics using interphase nuclei. Prenat 015
Diagn 22(6):497–499. https://doi.org/10.
1002/pd.299
Chapter 10
Prenatal Detection of Chromosome Aneuploidy

by Quantitative Fluorescence PCR
Kathy Mann, Erwin Petek, and Barbara Pertl
Abstract
Autosomal chromosome aneuploid pregnancies that survive to term, namely, trisomies 13, 18, and
21, account for 89% of chromosome abnormalities with a severe phenotype identified in prenatal samples.
They are traditionally detected by full karyotype analysis of cultured cells. The average reporting time for a
prenatal karyotype analysis is approximately 14 days, and in recent years, there has been increasing demand
for more rapid prenatal results with respect to the common chromosome aneuploidies, to relieve maternal
anxiety and facilitate options in pregnancy. The rapid tests that have been developed negate the requirement
for cultured cells, instead directly testing cells from the amniotic fluid or chorionic villus sample, with the
aim of generating results within 48 h of sample receipt. Interphase fluorescence in situ hybridization is the
method of choice in some genetic laboratories, usually because the expertise and equipment are readily
available. However, a quantitative fluorescence (QF)-PCR-based approach is now widely used and reported
as a clinical diagnostic service in many studies. It may be used as a stand-alone test or as an adjunct test to full
karyotype or array CGH analysis, which scan for other chromosome abnormalities not detected by the
QF-PCR assay.
Key words Quantitative fluorescence-polymerase chain reaction (QF-PCR), Chromosome aneuploi-

dies, Rapid prenatal test, Down syndrome
1 Introduction
Autosomal chromosome aneuploid pregnancies that survive to

term, namely, trisomies 13, 18, and 21, account for 89% of chro-
mosome abnormalities with a severe phenotype identified in prena-
tal samples [1]. They are traditionally detected by full karyotype
analysis of cultured cells. The average UK reporting time for a
prenatal karyotype analysis is approximately 14 days [2], and in
recent years, there has been increasing demand for more rapid
prenatal results with respect to the common chromosome aneu-
ploidies, to relieve maternal anxiety and facilitate options in preg-
nancy. The rapid tests that have been developed negate the
requirement for cultured cells, instead directly testing cells from
the amniotic fluid (AF) or chorionic villus sample (CVS), with the
139
140 Kathy Mann et al.
aim of generating results within 48 h of sample receipt. Interphase-

fluorescence in situ hybridization (FISH) [3, 4] is the method of
choice in many genetic laboratories, usually because the expertise
and equipment are readily available. However, a quantitative fluo-
rescence polymerase chain reaction (QF-PCR)-based approach is
more suited to a high throughput diagnostic service. This approach
is now widely used and reported as a clinical diagnostic service
[5–14]. It may be used as a stand-alone test or as an adjunct test
to full karyotype or array CGH analysis, which scan for other
chromosome abnormalities not detected by the QF-PCR assay.
1.1 Principle QF-PCR refers to the amplification of chromosome-specific poly-

morphic microsatellite markers using fluorescence-labeled primers,
followed by quantitative analysis of the products on a genetic
analyzer to determine copy number of specific chromosomal mate-
rial. Tetranucleotide repeat markers are used to minimize
PCR-generated “stutter bands” (PCR artifacts that are one to
three repeat units smaller than the true allele size). Where a micro-
satellite marker is heterozygous, the ratio of its allele peak areas
represents a disomic (1:1) or trisomic (2:1, 1:2 or 1:1:1) chromo-
some complement (see Fig. 1). A marker is uninformative if only a
single peak is observed.
Due to allele size heterogeneity and differences in sample type
and quality, the amplification of a single marker relative to other
markers in the assay may vary greatly. Thus, a comparison of allele
peak areas between markers, as an indicator of chromosome copy
number, is not recommended. Furthermore, because only peak
areas within a single locus are compared, allele dosage ratios are
more resilient to the effects of the plateau phase of the PCR than
other dosage assays [15].
The procedure described here uses a “one-tube test,” where
17 markers are coamplified in one multiplex reaction (see Table 1
and Fig. 1) [9]. Five markers are used for both chromosomes
13 and 21, and seven markers are used for chromosome 18. A
separate sex chromosome multiplex can be used for sexing purposes
and to diagnose sex chromosome aneuploidy (see Table 2 and
Fig. 2) [9, 16]. The inclusion of a paralagous marker in the sex
chromosome assay is recommended as this adds confidence to a
diagnosis of monosomy X. Here, the TAF9L marker is used which
amplifies a chromosome 3 and X chromosome sequence with the
same primer pair. Comparison of the two peaks is used to determine
the copy number of the X chromosome sequence. Markers are
located along the length of each chromosome to increase the
chance of detecting unbalanced chromosome rearrangements.
Prenatal Detection of Chromosome Aneuploidy by Quantitative Fluorescence PCR 141
Fig. 1 Genotyper profile (Applied Biosystems CA) depicting the 17 polymorphic markers in the trisomy
multiplex. The seven chromosome 18 markers exhibit allele patterns consistent with three copies of
chromosome 18. All five chromosome 13 markers and four of five chromosome 21 markers show a normal
biallelic profile while the fifth chromosome 21 marker is uninformative. The results are consistent with trisomy
18 and a normal complement for chromosomes 13 and 21
1.2 Potential Evidence of a second genotype, as shown by inconsistent dosage

Problems ratios for each chromosome, extra allele peaks, or both, usually
indicates contamination of the sample by maternal cells (see
1.2.1 Maternal Cell
Fig. 3), although it may represent a twin or chimera. Maternal cell
Contamination (MCC)
contamination is usually associated with blood-stained AF samples,
although the degree of blood staining should not in itself be used as
an indicator of maternal cell contamination; blood cells may be fetal
or maternal in origin. Usually, samples are accompanied by some
degree of blood staining, although this ranges from a pale pink cell
pellet and clear liquor to a deep red color of the whole fluid. When
the majority genotype shows consistent normal or abnormal results
with no inconclusive allele ratios, then the result may be reported,
although it may be advisable to confirm the origin of the majority
genotype by analysis of a maternal blood sample. We have found the
level of blood staining in the AF cell pellet to broadly correlate with
the level of maternal genotype; the majority genotype from pellets
with fewer red blood cells is consistently fetal in origin. However,
when the presence of two genotypes causes allele ratios to skew
outside of the normal or abnormal range (see Subheading 3.5, step
1), it is recommended that the QF-PCR results are not interpreted
due to the increased risk of a misdiagnosis. In these cases, if a rapid
result is required and one of the genotypes is determined as fetal in
origin either by sexing or by genotype analysis of a maternal sample,
142
Table 1
Details of primers used in the trisomy multiplex. Size ranges given are those used in Genotyper, version 3.7 (Applied Biosystems). Heterozygosity
Kathy Mann et al.
values are based on our cohort and may vary in other populations. Primer concentrations will vary depending on supplier, batch and buffer. All
markers are tetranucleotide repeats
Final conc. (μM)

Marker Name Location Hetero-zygosity Allele size range (bp) Primer sequences 50 –30 (Forward, reverse) of each primer
D13S252 13q12.1 0.85 260–330 PET-GCAGATGTACTGTTTTCCTACCAA 0.75
AGATGGTATATTGTGGGACCTTGT
D13S305 13q13.3 0.75 418–482 VIC-GCCTGTTTGAGGACCTGTCGTTA 1
TGGTTATAGAGCAGTTAAGGCAC
D13S628 13q31.1 0.688 425–474 NED- TGGATGAATACGCCACTTTTC 1.5
TGGTTAAAAGATTGCCAAGGAG
D13S634 13q21.33 0.812 355–440 6FAM-GGCAGATTCAATAGGATAAATAGA 0.5
GTAACCCCTCAGGTTCTCAAGTCT
D13S325 13q12.12 0.75 235–315 VIC-CTGTGCTATCTCCTCCAACG 0.25
GTTTGAAAGATAGGCCATGCAG
D18S386 18q22.1 0.875 320–417 VIC-TGAGTCAGGAGAATCACTTGGAAC 0.75
CTCTTCCATGAAGTAGCTAAGCAG
D18S390 18q22.3 0.75 340–415 NED-GGTCAATAGTGAATATTTGGATAC 0.25
CTCCAACCTCACTTGAGAGTA
D18S391 18p11.31 0.75 190–235 VIC-GGACTTACCACAGGCAATGTGACT 0.1
CTGGCTAATTGAGTTAGATTACAA
D18S535 18q12.3 0.92 450–500 6FAM-CAGCAAACTTCATGTGACAAAAGC 0.5
CAATGGTAACCTACTATTTACGTC
D18S819 18q11.2 0.78 370–450 PET-CTTCTCACCTGAATTACTATGGT 1
TTTGTAATCGATCTACCACAGTT
D18S976 18p11.31 0.78 465–495 PET- GAGATCCTGAACATGGAGCAG 0.5
ACACTATTGGCATCCCTTGG
D18S978 18q12.3 0.667 180–230 NED-GTAGATCTTGGGACTTGTCAGA 0.25
GTCTCCCATGGTCACAATGCT
D21S11 21q21.1 0.9 220–283 6FAM-TTTCTCAGTCTCCATAAATATGTG 1
GATGTTGTATTAGTCAATGTTCTC
D21S1437 21q21.1 0.84 283–351 6FAM-CTACCACTGATGGACATTTAG 0.5
GTGGAGGGTGTACCTCCAGAA
D21S1409 21q21.2 0.81 205–250 PET-AAGCGAAGGATTTGGATCAG 0.5
TTTGCCTCTGAATATCCCTATC
D21S1411 21q22.3 0.933 256–345 ATAGGTAGATACATAAATATGATGA 0.75
NED-TATTAATGTGTGTCCTTCCAGGC
D21S1435 21q21.3 0.75 160–200 6FAM-CCCTCTCCAATTGTTTGTCTACC 0.25
ACAAAAGGAAAGCAAGAGATTTCA
Prenatal Detection of Chromosome Aneuploidy by Quantitative Fluorescence PCR
143
144
Table 2
Kathy Mann et al.
Details of primers used in the sex chromosome multiplex. Size ranges given are those used in Genotyper, version 3.7 (Applied Biosystems).
Heterozygosity (Het) values are based on our cohort and may vary in other populations. Primer concentrations will vary depending on supplier, batch
and buffer. DYS448 is a hexanucleotide repeat, DX6807, DXS981, DXS1187, XHPRT, DXS7423, DXS6803, DXS6809, DXYS267, DXYS218 are
tetranucleotide repeats and DXS1283E is a dinucleotide repeat. AMEL, SRY and TAF9L are not polymorphic. TAF9L is present on chromosome 3 and the
X chromosome and the ratio of both peaks is used to assess relative X chromosome copy number
Marker Final conc. (μM)

Name Location Hetero-zygosity Allele size range (bp) Primer sequences 50 –30 (Forward, reverse) of each primer
DXS6807 Xp22.3 0.7 300–380 6FAM- TCTCCCTTATTTGTGGTTTTGC 1.5
AGCAGTTCTCCCTTATCCAC
DXS1283E Xp22.3 0.89 295–340 NED-AGTTTAGGAGATTATCAAGCTG 0.75
CCCATACACAAGTCCTCAAAGTGA
DXS981 Xq13.1 0.86 225–260 6FAM-CTCCTTGTGGCCTTCCTTAAATG 0.25
TTCTCTCCACTTTTCAGAGTCA
DXS1187 Xq26.2 0.72 180–220 VIC-CAGCTACTCAATGAAAAGCC 0.25
ATGGGGTAGGGACCAAAAAT
XHPRT Xq26.2 0.78 265–300 VIC-ATGCCACAGATAATACACATCCCC 0.5
CTCTCCAGAATAGTTAGATGTAGG
DXS7423 Xq28 0.74 350–420 VIC- TACTGGAGGTGAGGGTTGTG 1.5
TGGGCTGCCCAGATACAACT
DXS6803 Xq21.31 0.86 135–152 6FAM-AAAATTTTCCTCAAAACAAAAAGG 0.5
AGAATATTCACCTAGAAATGTGC
DXS6809 Xq21.33 0.78 399–427 PET-TTGCTTTAGGCTGATGTGAGG 0.5
CAGGTTAATTCAAGATATTTGTCA
DXYS267 Xq21.31 0.87 240–280 PET-ATGTGGTCTTCTACTTGTGTCA 1.0
Yp11.31 GTG TGT GGA AGT GAA GGA TAG
AMEL Xp22.2 – 106 PET-CCCTGGGCTCTGTAAAGAATAGTG 0.25
Yp11.2 112 ATCAGAGCTTAAACTGGGAAGCTG
SRY Yp11.31 – 248 bp NED- AGTAAAGGCAACGTCCAGGAT 0.5
TTCCGACGAGGTCGATACTTA
DYS448 Yq11.223 – 323–370 PET- CAAGGATCCAAATAAAGAACAGAG A 0.5
GGTTATTTCTTGATTCCCTGTG
DXYS218 Xp22.33 0.74 383–411 6-FAM- AACTGAGGGGACCTGGAATG 0.5
Yp11.3 GAATCGATTCAACCCGGGAGA
TAF9L 3p24.2 – 116 AGCATCTCTGTTAAATTTAGAAATG 0.5
Xq21.1 125 PET-CAGGAAACAGCTATGACCTGC
TTTTGACAGGTAGTTTTGG
Prenatal Detection of Chromosome Aneuploidy by Quantitative Fluorescence PCR
145
Fig. 2 Three Genotyper Profiles (Applied Biosystems CA) depicting the 14 markers in the sex chromosome
multiplex. (a) A normal male sample is represented by DXYS267 and DXYS218 which indicate the presence of
two sex chromosomes, AMEL which exhibits equal quantities of the X and Y sequences, TAF9 which indicates
that there are half as many X chromosome sequences present as chromosome 3 sequences, the presence of Y
chromosome-specific sequences (SRY and DYS448) and eight apparently hemizygous polymorphic markers.
(b) A normal female sample is represented by eight biallelic polymorphic markers indicating two X chromo-
somes, TAF9 which indicates that X chromosome sequences are present in equal quantities to chromosome
3, and the absence of Y chromosome sequences (AMEL, SRY, DYS448). (c) A monosomy X sample is
represented by ten apparently hemizygous polymorphic markers, TAF9 which indicates that there are half
as many X chromosome sequences present as chromosome 3 sequences, and the absence of Y chromosome
sequences (AMEL, SRY, DYS448)
Fig. 3 Genotyper profile of a sample exhibiting a high level of maternal cell contamination. The characteristic
triallelic pattern, where the fetal-specific and maternal-specific allele peak areas combine to equal the shared
fetal/maternal allele, is observed in markers D21S11, D18S391, D18S386, D13S305, D21S1411, D13S628,
D13S252, D18S819
interphase-FISH may then be used if the analysis takes into account

the fetal to maternal ratio. The number of analyzed cells can be
increased to account for those that are maternal, or a sex chromo-
some probe can be cohybridized with an autosome probe and only
the male cells analyzed.
In our London sample set, approximately 10% of AF samples
are found to have two genotypes, although allele ratios vary con-
siderably, proportional to the relative contribution of each cell line.
The majority of these samples exhibit a very low level second
genotype and can be reported as normal, whereas approximately
2% of AF samples in both the London and Graz cohorts exhibit
second genotypes that prevent confident interpretation of allele
ratios; therefore, they are reported as unsuitable due to MCC.
The detection of maternal cells in an amniotic fluid sample should
not discredit the karyotype analysis of cultured cells. Subsequent
genotype analysis of cultured cells from samples showing MCC,
normally demonstrates a single genotype, consistent with the selec-
tion and growth of fetal cells, and loss of maternal cells during the
culture process [17]. In those samples with a mixed female/male
cell population evident on the direct analysis, a single male geno-
type is usually detected on the follow-up test. However, the pres-
ence of two genotypes in samples where no blood staining is
evident may indicate a maternal tissue plug. This may grow in
culture and therefore genotype analysis of cultured cells may be
useful.
Fig. 4 Genotyper profile of a sample exhibiting trisomy 21 mosaicism. All informative chromosome 21 markers
show either an unequal triallelic pattern (D21S11 and D21S1411) or unequal diallelic ratios (D21S1435,
D21S1437, and D21S1409). Chromosome 13 and 18 markers are normal. The presence of triallelic
chromosome 21 markers is consistent with a meiotic non-disjunction event followed by trisomy rescue to
generate the normal cell line
1.2.2 Mosaicism The problem of mosaic genotypes and karyotypes in prenatal sam-
ples is well documented, particularly in CVS (see Fig. 4). With
respect to QF-PCR, two issues are relevant; the levels of mosaicism
detectable by the QF-PCR technique, and the degree of concor-
dance between a direct test result and the fetal genotype.
The first of these can only be determined by the analysis of
samples (both postnatal and prenatal) exhibiting mosaicism for one
of the tested regions. The generation of “artificial mosaics” by the
mixing of two genotypes in known measures represents a chimera
rather than a mosaic genotype. The presence of a triallelic result is
consistent with a meiotic nondisjunction event generating the tri-
somy cell line, whereas the absence of a triallelic result is evidence,
although not diagnostic, of a normal conception followed by a
mitotic nondisjunction event. Analysis of mosaic cases in our sam-
ple set found that a minimum level of 15% trisomy mosaicism could
be detected if a triallelic allele pattern was observed and 20% tri-
somy mosaicism if only dialleic ratios were present [18]. Thus, a
mitotic error occurring in a disomic fetus may be harder to detect,
due to the absence of a third allele. Indeed, QF-PCR identified only
one of three trisomy 18 or 21 mosaics described by Pertl et al. [11];
this mosaic case was also triallelic, in this case for chromosome 21.
Discrepancies between the QF-PCR and karyotype result have
been described [19, 20]. These have been shown to be due to
mosaicism and confinement of cell lines to different regions of the
tested sample. To minimize such discrepant results it is recom-

mended that DNA is prepared from dissociated cells prepared
from 5 to 15 mg of cleaned villi. Although only a small aliquot of
this cell suspension is required, all major cell lines in both the
mesoderm and cytotrophoblast should be represented and detected
by QF-PCR. Karyotype analysis of cell populations that are subse-
quently cultured from this cell suspension should minimize discre-
pancies between the two techniques.
Chorionic villi consist of an outer cytotrophoblast and internal
mesoderm layers. The mesoderm layer is derived from a later fetal
cell lineage, whereas the cytotrophoblast is derived from a much
earlier lineage; thus, it is less representative of fetal tissue [21].
DNA prepared from dissociated cells represents cells from both
the cytotrophoblast layer and mesenchymal core [22]. Interphase-
FISH results also are thought to represent both cell layers. In
contrast, karyotype analysis of direct CVS preparations concerns
only the cytotrophoblastic line, whereas culture conditions primar-
ily lead to expansion of the mesoderm cell line, resulting in a final
karyotype that is more representative of the fetus. In summary, care
should be taken in the interpretation of trisomic prenatal results
derived from CVS material, in the absence of a triallelic result
demonstrating a meiotic origin to the trisomy cell line.
1.2.3 Submicroscopic Partial chromosome duplication may be identified by QF-PCR

Duplications analysis by the presence of both normal and abnormal marker
results on one chromosome. This pattern may indicate a cytogenet-
ically visible abnormality [8] or one that is submicroscopic. If the
most distal or proximal markers are duplicated, then this may
indicate the unbalanced product of a reciprocal translocation.
However, in our experience, the presence of a single abnormal
marker result, where all other informative results are normal, is
most likely to represent a submicroscopic duplication (SMD)
[6]. In the majority of cases, analysis of parental samples shows
these SMDs to be inherited. SMDs identified by a single marker
that have previously been reported and are flanked by normal
markers, require no further investigation and according to Best
Practice Guidelines do not have to be reported (http://www.acgs.
uk.com/media/765524/acc.cmgs_qfpcr_bp_jan2012_3.01.pdf).
A list of SMDs identified to date by markers used in QF-PCR assays
is available from Kathy.Mann@viapath.co.uk. For SMDs that have
not previously been described, it is necessary to establish the inher-
itance of the SMD; inherited submicroscopic duplications are
unlikely to be clinically significant [23].
1.2.4 Primer Site Primer site polymorphisms are a known phenomenon of PCR
Polymorphisms (PSPs) assays [24]. Sequence differences between the genomic DNA and
the primers can result in complete or partial allele dropout (ADO)
Fig. 5 Genotyper profile of a sample exhibiting a somatic microsatellite mutation (SMM) at marker D13S305.
The characteristic triallelic result is evident where the two lowest allele peaks representing the two mosaic
cell lines combine to equal the higher allele present in both cell lines. All other informative markers are normal
due to reduced or absent hybridization of the primers to genomic

DNA. Partial ADO in a normal sample can either give an abnormal
diallelic ratio consistent with trisomy for that region, or an incon-
clusive ratio (see Subheading 3.5). Complete ADO in an abnormal
sample can result in a normal diallelic ratio at that locus. In all cases
of suspected ADO caused by PSPs, it is recommended to repeat the
PCR at a lower annealing temperature (for example, 4 C lower
than the standard temperature). This provides a less stringent envi-
ronment for primer hybridization, resulting in reduced ADO, as
represented by a change in the allele ratio. If the follow-up tests are
consistent with the presence of a PSP, the marker result should be
failed and not used as part of the QF-PCR analysis, even if it shows a
normal ratio; a PSP may cause an abnormal diallelic result to seem
normal at a lower annealing temperature.
1.2.5 Somatic Somatic changes in the length of a microsatellite sequence, due to

Microsatellite Mutations DNA replication and proof-reading errors, may be visible as an
(SMMs) unequal triallelic result, where the areas of the two lowest alleles
combine to equal the highest allele (Fig. 5), or skewed diallelic
ratios. The characteristic triallelic pattern represents two cell lines
that have one common allele and a second allele of different
lengths. QF-PCR analysis of AF samples found <0.1% to have an
SMM at a single locus, whereas SMMs were observed at a much
higher frequency (4%) in individual whole villi [25]. This high
frequency is probably due to the small clonal cell population
being analyzed. QF-PCR analysis of digested/chopped villi con-
taining a more diverse cell population (see Subheading 3.1.2) there-
fore results in the identification of fewer SMMs. Interestingly, the
majority of SMMs are single repeat unit expansion/contraction
events. These somatic changes in repeat length are specific to the

microsatellite and are not clinically significant. Thus, a single
marker result, showing a characteristic triallelic pattern consistent
with a SMM, where all other markers on that chromosome are
normal, does not require further clarification. However, it is more
difficult to class skewed diallelic results as SMMs; PSPs (see Sub-
heading 1.2.4) and SMDs (see Subheading 1.2.3) may be alterna-
tive explanations. It is important to distinguish between these
events if a 1:2 or 2:1 allele ratio is observed, because these ratios
may represent a clinically significant imbalance. Once PSPs have
been excluded, analysis of cultured cell populations may help to
distinguish between an SMD and SMM. In the case of an SMM, the
proportion of the two cell lines may change between uncultured
and cultured cells thus altering the allele ratio.
2 Materials
2.1 DNA Preparation 1. InstaGene Matrix (Bio-Rad, Hercules, CA). Store at 4 C.
2.2 PCR 1. T.1E buffer: 10 mM Tris–HCl, pH 7.6, and 0.1 mM EDTA.

Filter, sterilizes, and store at room temperature.
2. 10 Multiplex PCR kit (MyTaq, Bioline, London, UK). Store
at 20 C. Once thawed store at 4 C. Kit contains Taq
polymerase and dNTPs.
3. 50 -labeled fluorescent oligonucleotide primers (Thermo Fisher
Scientfic, Waltham, MA, USA). We use primers, 50 -labeled with
6-FAM, VIC, NED, or PET. Fluorescence-labeled primers
should not be exposed to light for prolonged periods or repeat-
edly freeze-thawed (see data sheet). These are supplied as pre-
cipitates, and are resuspended in T.1E buffer to give a working
concentration of 100 μM. The primers are stored at 20 C.
Aliquots in use are stored at 4 C.
4. Unlabeled oligonucleotide primers (Sigma-Aldrich, Dorset,
UK). These primers are supplied as precipitates, and are resus-
pended in T.1E buffer to a working concentration of 100 μM
and stored at 20 C. Although unlabeled primers are more
stable than fluorescent-labeled primers, freeze-thawing should
be avoided. Aliquots in use are stored at 4 C.
2.3 Analysis 1. ABI PRISM 3100 capillary-based genetic analyzer

(Thermo Fisher Scientific). All consumables (10 EDTA
buffer, POP6 polymer, 36 cm capillaries, plate septa) are pur-
chased from Thermo Fisher Scientific.
2. Deionized formamide (Toxic: refer to the material safety data
sheet): HI-DI formamide (Thermo Fisher Scientific) is highly
deionized, to minimize the breakdown of the fluorescent label.
It is stored at 20 C in appropriate aliquots.
3. Genescan-500 LIZ size standard (Thermo Fisher Scientific).

Standard is designed for 100–500-bp fragment analysis.
4. 96-well plates: We use those recommended for the ABI
PRISM 3100 genetic analyzer (Thermo-Fast 96 Detection
Plates, ABgene, Epsom, Surrey, UK).
5. Software: For fragment analysis, GeneScan Analysis, version
3.7 was used. For allele size calling and labeling, Genotyper,
version 2.5 (Thermo Fisher Scientific) was used. Microsoft
Excel (Microsoft, Redmond, WA) is used to calculate and
tabulate allele dosage ratios. Other fragment analysis software
is available including GeneMapper IDX (Thermo Fisher Scien-
tific) and GeneMarker (SoftGenetics LLC, PA 16803, USA)
which include specific aneuploidy analysis modules.
3 Methods
The processing of a number of prenatal samples at one time, and

the risk of sample mixup, necessitates stringent quality control
procedures (see Note 1). In addition, care must be taken, as with
all PCR-based tests, to avoid contamination of tested material with
amplified products of previous reactions and external DNA (see
Note 2).
3.1 Sample Sample and DNA preparation procedures are carried out in a class
Preparation II biological containment cabinet, up to step 5, Subheading 3.2.
(see Note 3).
3.1.1 Amniotic Fluid (AF) Between 10 and 20 mL of AF is normally received in a 20 mL sterile

universal container.
1. Centrifuge the sample at 200 g for 10 min.
2. Carefully remove the supernatant fluid using a 20 mL syringe
and leave approximately 1 mL of fluid (equal to the top of the
conical section of the Universal) in the container.
3. Resuspend the cell pellet in the fluid by using a plastic Pasteur
pipette, and transfer approximately 100 μL (2–3 drops) to a
0.5 mL Eppendorf tube. This aliquot (approx 1/10 of the
original AF sample) can be stored at 4 C until processed.
The remainder can be cultured for subsequent karyotype anal-
ysis, as required.
3.1.2 Chorionic Villus 1. Remove the maternal decidua from 5 to 15 mg of chorionic

(See Note 4) and Tissue villi.
Samples (See Note 5) 2. Either finely chop the cleaned villi or digest with collagenase
(final concentration 2.5 mg/mL) at 37 C for 35 min, followed
by trypsin digestion (final concentration 2.5 mg/mL) at 37 C
for 25 min. The addition of 2 mL of medium prevents further

digestion.
3. Remove a single drop (approx 100 μL) of the dissociated cell
slurry for QF-PCR analysis. This may be stored at 4 C. The
rest of the sample is cultured for karyotype analysis or DNA is
extracted for array CGH analysis. For tissues, a small section of
whole tissue sample is isolated and can be stored at 4 C.
3.2 DNA Preparation This is a quick and simple procedure and it can be successfully
(See Note 6) applied to a number of sample types and different sample qualities
(see Note 7). Because it is based on a boiling lysis protocol, aided by
vigorous vortexing, the safety issues associated with phenol/chlo-
roform extractions also are avoided. After cell lysis, the InstaGene
Matrix commercial resin removes trace metal contaminants that
may inhibit the PCR.
Before the DNA extraction, place the InstaGene Matrix on a
magnetic stirrer and set at a medium speed for at least 5 min.
1. Pellet the cells/villus/tissue at 12,000 g for 1 min in a
microcentrifuge and carefully remove the liquid, leaving
enough to resuspend the pellet (approx 10–20 μL). For amni-
otic fluid samples, any blood staining in the cell pellet is noted
as a percentage of whole pellet. If >50% of the pellet is red,
follow steps 2 and 3. For all other samples, proceed to step 4.
2. Vortex to resuspend the cell pellet and add 200 μL of H2O to
wash the sample (see Note 8).
3. Vortex the sample, pellet the cells, and remove the wash solu-
tion as described above. Resuspend the cells in the remaining
wash solution by vortexing.
4. Add between 100 and 400 μL (see Note 9) of InstaGene Matrix
to the cells/villus by using a wide-bore pipette tip, e.g., a
Gilson p1000 tip and vortex.
5. Incubate at 100 C for 8 min.
6. Vortex again at high speed for 10 s, and pellet the InstaGene
Matrix at 12,000 g for 3 min in a microcentrifuge.
7. Place the samples on ice to cool.
8. The DNA preparation should be stored at 20 C (see Note
10).
3.3 PCR Setup Batches of PCR assays can be prepared in advance, tested and stored
at 20 C. These are 20 μL aliquots of a master mix that contains all
components except DNA, which is added immediately before tem-
perature cycling, to give a total volume of 25 μL. The final con-
centrations of the reaction components are 1 Multiplex PCR kit
and 2.5–42.5 pmol of each primer (see Tables 1 and 2), in a total
volume of 25 μL (Note: DNA is added according to volume rather

than concentration, as concentration is not measured).
1. To a thin-walled PCR tube containing 20 μL of the master mix,
add 5 μL of DNA solution (see Note 11), taking care not to
disturb the InstaGene Matrix pellet. Mix by pipetting.
2. Add 1 drop of mineral oil, if a heated lid is not being used, and
place in the PCR machine.
3. PCR cycling conditions: Taq polymerase activation and initial
denaturation at 95 C for 15 min followed by 25 cycles of
94 C for 30 s, 58 C for 90 s, 71 C for 90 s (see Note 12).
Final synthesis: 72 C for 20 min followed by storage at 10 C
(see Note 13).
3.4 Analysis Post-PCR cleanup to remove excess primers and free dye molecules
is not carried out (see Note 14). We use the Thermo Fisher Scien-
3.4.1 PCR Product
tific ABI 3100 PRISM genetic analyzers, and conditions specific to
Preparation
this instrument are described. Standard use of this analyzer is not
detailed here. Other genetic analyzers capable of fragment resolu-
tion, fluorescence detection, and quantification also can be used,
and they include Thermo Fisher Scientific ABI capillary-based ana-
lyzer model 3130, 3730, and 3500; and Spectrum Compact CE
system (Promega Corporation, Madison, WI, USA).
1. Prepare PCR products for analysis by the addition of 3 μL of
product to 15 μL of HI-DI formamide in 96-well plates (see
Note 15).
2. Denature at 95 C for 2 min and snap-chill on ice.
3.4.2 3100 Analysis Separate PCR products through a 36 cm capillary array filled with
POP6 (see Note 16). A 10-sec injection time is suitable for most
samples (see Note 17). The running conditions are 60 C for
3000 s.
3.4.3 Genotyper Analysis Macros are used to label allele peaks with marker name, size, and
peak area (Fig. 1) (see Note 18). The Genotyper table is transferred
to an Excel spreadsheet for allele ratio analysis.
3.5 Result The criteria listed below are based on >40,000 QF-PCR prenatal
Interpretation tests (see Note 19). For additional information, see the UK ACGS
Best Practice Guidelines for the Diagnosis of Aneuploidy at
(http://www.acgs.uk.com/media/765524/acc.cmgs_qfpcr_bp_
jan2012_3.01.pdf).
1. Normal allele dosage ratios range between 0.8 and 1.4 (see
Note 20). For alleles separated by >24 bp, ratios up to 1.5
are acceptable. Trisomy is indicated by an allele ratio of
between 1.8 and 2.4 or between 0.65 and 0.45 or by the
presence of three alleles of equal areas (see Note 21). All of

these results are described as informative.
2. Two informative markers per chromosome are required for
confident interpretation. This minimizes the risk of misdiagno-
sis due to primer-site polymorphisms and/or somatic repeat
instability (see Note 22). Single marker abnormal results
should not be reported.
3. If both normal and abnormal marker results are obtained for a
single chromosome, follow-up studies should be carried out.
Such results may represent polymorphisms or a clinically signif-
icant partial chromosome imbalance (see Subheadings
1.2.3–1.2.5). Single marker assays, additional markers, lower-
ing the PCR annealing temperature and analysis of cultured cell
populations and parental samples can clarify these results.
4 Notes
1. To prevent sample mix-up a minimal number of tube-to-tube

transfers should be used (three transfers are required for this
protocol). Each sample transfer and analysis should be checked
by another laboratory member, and the use of two identifiers
per tube, such as sample number and name, aids sample
tracking.
2. Contamination of a PCR by external DNA or PCR amplicons is
evident by the appearance of allele peaks in the negative
(no DNA) PCR control; a critical part of any PCR procedure
and a reaction that should be set up last in a series of samples.
Separation of the PCR setup and post-PCR analysis areas
should help to prevent contamination.
3. To ensure that DNA is prepared from the correct sample, it is
advisable to prepare the initial sample aliquot (see Subheadings
3.1.1 and 3.1.2) one sample at a time in a class II biological
containment cabinet, with only one sample in the cabinet
during the procedure. Because the subsequent DNA prepara-
tion (see Subheading 3.2) is carried out without further tube
transfers, DNA from a number of samples can be prepared
simultaneously. This DNA can then be used for subsequent
PCR tests.
4. It is recommended that DNA is prepared from dissociated cells
prepared from 5 to 15 mg of cleaned villi. Although only a
small aliquot of this cell suspension is required, all major cell
lines in both the mesoderm and cytotrophoblast should be
represented and detected by QF-PCR. The analysis of whole
villi has rarely been associated with discrepant results between
QF-PCR and karyotype analysis due to mosaic cell lines
confined to one region of the sample [19, 20]. Analysis of cell

populations that are subsequently cultured for karyotype or
array CGH analysis should minimize discrepancies between
the techniques.
5. The QF-PCR procedure can be applied effectively to solid
tissue samples (e.g., skin and cartilage). This is particularly
useful for confirming a prenatal diagnosis or for a poor-quality
sample where chromosome analysis of cultured cells may not be
possible.
6. DNA prepared using the protocols described here also may be
used for other molecular prenatal tests. The protocol has ben-
efits over the traditional phenol/chloroform-based approach in
terms of labor and time savings and reduced safety risks. How-
ever, as the extracted DNA may contain residual contaminants,
its suitability for use in other tests should be determined.
7. Although the procedure is generally successful in extracting
DNA of sufficient quality for use in the multiplex, in our
experience, DNA extracted from bloodstained or discolored
AF fluid may contain PCR inhibitors. These inhibitors can be
removed by a subsequent extraction (see Note 9).
8. Deionized water lyses red blood cells, and it also may aid lysis of
cells in villus and tissue samples. A deionized water wash is used
for blood-stained/discolored AF.
9. It is beneficial to adjust the volume of InstaGene Matrix to
balance removal of all cell lysis products with excessive dilution
of the DNA. A 300 μL volume of InstaGene Matrix is generally
used for CVS and tissue samples and the larger AF cell pellets
(those that cover the base of the 0.5 mL microcentrifuge tube).
Only 100 μL of InstaGene Matrix is required for average and
small AF pellets. It is important that the DNA extraction is not
overloaded with too much starting material. This leads to a
failure by the InstaGene Matrix to chelate all metal ions and can
result in inhibition of the PCR. In particular, the larger sized
markers in the multiplex may fail to amplify. If inhibition is
observed, a further extraction can be used to remove the con-
taminants; 100 μL of the DNA extract is added to 100 μL of
InstaGene Matrix and treated as per the extraction protocol (see
Subheading 3.2, steps 5–8).
10. Because the DNA prepared here is relatively crude, and cell
lysis products that damage DNA may remain, the DNA should
be stored at 20 C. However, the DNA is stable for at least
3 days at room temperature, which allows some flexibility, such
as transfer of the sample to another laboratory.
11. As well as the necessary inclusion of the negative (no DNA)
PCR control for the reasons given in Note 2, the use of a DNA
control trisomic for one of the chromosomes and exhibiting
2:1 or 1:2 dosage for at least one marker is recommended.

Normal allele dosage is exhibited by the other two nontrisomic
chromosomes. The control not only demonstrates that the
amplified DNA represents allele copy number for that proce-
dure but also can be used as a standard against which any
spurious background bands or free-dye peaks can be
compared.
12. Samples that do not generate sufficient amplified sequences for
analysis, due to low initial DNA concentration (usually evident
by a very small original cell pellet) can be amplified with a
greater number of cycles. Any change in PCR cycling condi-
tions should always be accompanied by a trisomy control, to
ensure the reaction is still quantitative.
13. The 20-min incubation at 72 C is necessary, because Taq
polymerases lacking exonuclease activity add a template-
independent dATP to the 30 end of amplified sequences
[26]. Without the 72 C incubation, a single base-pair size
difference in the amplified sequences can resolve as a “split
peak” on the profile and hinder analysis. This is a particular
problem here due to the size of the fragments generated
(100–500 bp). In this size range, the analysis systems efficiently
resolve single base-pair differences especially in smaller alleles.
14. Free-dye peaks are caused by the detachment of the fluorescent
molecule from the labeled primer. These molecules are
resolved as broad peaks, usually up to 180 bp (see Note 16);
as such, they can be distinguished from allele peaks. The break-
down of fluorescent primers can be minimized by the use of
deionized formamide stored at 20 C and reduced exposure
of labeled primers to temperatures above 20 C. Free-dye
molecules can be removed, along with unincorporated primers,
by standard post-PCR cleanup protocols if required.
15. Accurate transfer of samples to the wells of a 96-well plate can
be difficult. The risk of error can be minimized not only by the
use of a multichannel pipette but also by the addition of
loading buffer containing dextran blue, which is visible but
does not interfere with the fluorescent analysis. Transparent
piercable sheets also are available (ABGene) that can be sealed
onto the plate, or rubber septa can be placed over wells that are
not in use.
16. The POP6 polymer can be used on the 3100 genetic analyzer if
greater resolution is required, although this requires longer
run times. Resolution of free-dye molecules (see Note 14) is
not linear in respect to fragment size, but it is influenced by
both temperature and the separating matrix. A different poly-
mer may be used to resolve free-dye molecules that coincide
with allele peaks.
17. Allele peak heights >6000 fluorescent units on the 3100

genetic analyzer are not analyzed. The charge-coupled device
camera becomes saturated in this range, and peak fluorescence
may be underrepresented. If a sample is overloaded, injection
times can be reduced to accommodate differences in DNA
concentration and the corresponding amplification. This is
one of the advantages of the capillary-based genetic analyzers,
where a repeat injection does not require repeat sample
preparation.
18. Tetranucleotide alleles demonstrate few visible stutter bands
(see Subheading 1.2), and only the main allele peak is labeled.
However, some microsatellite markers contain a mix of both
tetranucleotide and dinucleotide repeats and generate signifi-
cant stutter bands. For dinucleotide alleles, the larger alleles
generally exhibit more significant stutter effects than smaller
alleles, due to the longer repeat. It is therefore necessary to
recognize and label at least the first stutter peak and include it
in the allele peak area measurement.
19. Although there are now several published studies describing
the use of QF-PCR as a diagnostic test [5–14], it is important
to validate the QF-PCR strategy in the laboratory in which it is
to be used. Control samples are required, and a pilot study is
recommended before the implementation of a QF-PCR-based
aneuploidy diagnostic service, especially if primer sets are used
that are not described in the published literature.
20. The large normal range is necessary due to the use of tetra-
nucleotide repeats. These can result in widely spaced alleles
(up to 50 bp apart), and marked preferential amplification of
the smaller allele, which in turn results in skewed allele dosage
ratios. However, closely spaced alleles should exhibit less allele
specific preferential amplification and would be expected to
have dosage ratios closer to 1.0.
21. As more than one sample is usually processed, the sample
identity of abnormal results should be confirmed. This can be
done by a repeat QF-PCR test. Alternatively, genotype analysis
of a maternal blood sample by using the same markers can be
used to confirm sample identification.
22. Polymorphisms in the primer-binding site can result in partial
or complete amplification failure of an allele (see Subheading
1.2.4). This can result in a misdiagnosis if the result is used in
isolation. In addition, SMMs (see Subheading 1.2.5) [25] and
submicroscopic imbalance (see Subheading 1.2.3) [6] also
could result in misdiagnosis if a single marker was used
independently.
References
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in situ hybridization (FISH). Am J Hum Genet pregnancies at risk. Mol Hum Reprod 6
51(1):55–65 (9):855–860
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Chapter 11
Multiplex Ligation-Dependent Probe Amplification (MLPA)

for Prenatal Diagnosis of Common Aneuploidies
Jan Schouten, Paul van Vught, and Robert-Jan Galjaard
Abstract
Multiplex Ligation-dependent Probe Amplification (MLPA) is a method to determine the copy number of
up to 60 genomic DNA sequences in a single multiplex PCR based reaction.
MLPA probes consist of two oligonucleotides that can hybridize next to each other on a certain DNA
sequence of interest, where they are ligated. All ligated probes are subsequently amplified by PCR using a
single set of primers. Each amplified MLPA probe has a unique length and can be visualized and quantified
by capillary electrophoresis. As the primers are almost 100% consumed in the PCR reaction, the quantity of
each PCR amplicon is proportional to the number of copies of each probe target sequence in the DNA
sample. A trisomy 21 can therefore be detected by an approximately 50% increased signal of each chromo-
some 21 specific probe relative to reference samples.
MLPA with the P095 Aneuploidy probemix for chromosomes 13, 18, 21, X and Y has been used as a
rapid detection method on large numbers of samples from uncultured amniotic fluid or from chorionic villi.
As compared to FISH and karyotyping, MLPA is more rapid, has a higher throughput, and is less expensive.
MLPA however cannot detect low grade mosaicism, female triploidies, and copy number neutral chromo-
some abnormalities such as inversions and translocations.
Key words Aneuploidy, Multiplex ligation-dependent probe amplification (MLPA), Trisomy, Multi-
plex polymerase chain reaction (PCR), Amniotic fluid, Chorionic villi, Gene dosage
1 Introduction
Multiplex Ligation-dependent Probe Amplification (MLPA) is a

multiplex method to detect abnormal copy numbers of up to
60 different genomic DNA sequences [1]. MLPA reactions are
easy to perform, require little hands-on time and results can be
obtained within 24 h. Furthermore, it requires only standard
equipment that is present in most DNA diagnostic laboratories.
In contrast to normal multiplex PCR, in MLPA not the sample
DNA is amplified, but the probes that are hybridized to the target
DNA (Fig. 1). Each single probe initially consists of two oligonu-
cleotides (left probe oligo [LPO] and right probe oligo [RPO]),
which are designed to hybridize immediate adjacent to each other
161
162 Jan Schouten et al.
Fig. 1 Outline of the MLPA technique. After hybridization to their target sequence
in the sample DNA, the probe oligonucleotides are enzymatically ligated. One
probe oligonucleotide contains a non-hybridizing stuffer sequence of variable
length. Ligation products can be amplified using PCR primer sequences X and Y,
amplification product of each probe has a unique length (90–500 nt). Amplifica-
tion products are separated by capillary electrophoresis. Relative amounts of
probe amplification products, as compared to a reference DNA sample, reflect
the relative copy number of target sequences. Adapted from www.mlpa.com
on the target DNA. After hybridization, the two oligonucleotides

can be ligated by a specific ligase enzyme, thereby creating a single
molecule, but only when both probe oligonucleotides are perfectly
hybridized to adjacent sites of the sample DNA. The ends of this
probe molecule contain two sequences recognized by a PCR primer
pair. After ligation, all probes are amplified by a single PCR primer
pair, of which one is fluorescently labeled. Since all probes have a
unique length, the resulting MLPA amplified products have a size
that ranges from 90 to up to 500 nucleotides, and can be visualized
by capillary electrophoresis.
Multiplex Ligation-Dependent Probe Amplification (MLPA) for Prenatal. . . 163
MLPA probes that do not find a target sequence cannot be

amplified by PCR, and do not have to be removed. The protocol for
an MLPA reaction is therefore very simple (Fig. 1):
1. Denature 20–500 ng DNA by heating to 98 C in a thermo-
cycler, followed by addition of the MLPA probes and buffer.
Leave overnight at 60 C for hybridization.
2. Add the Ligase and ligase buffers and ligate at 54 C for
15 min.
3. Inactivate the Ligase by heating to 98 C. Add PCR primers,
dNTPs and polymerase and start the PCR.
4. Analyze the products by capillary electrophoresis.
In MLPA, it is important that all fragments are amplified with
the use of only one pair of PCR primers that is present in limited
amounts. This abolishes differences in PCR efficiencies, which is
normally observed in multiplex PCRs, as well as the difference in
DNA input among multiple reactions. As a result, the relative signal
strength of each amplification product is determined primarily by
the copy number of the target sequence in a DNA sample. By
comparing these probe signals with those obtained from reference
samples, a decrease or increase in signal intensity can be observed,
reflecting a deletion or duplication, respectively (Fig. 2).
The MLPA technique allows discrimination of sequences that
differ only in a single nucleotide. MLPA can therefore also be used for
the detection of known mutations. A variation on the MLPA tech-
nique can be used to determine the methylation status of DNA
sequences [2]. MLPA products for more than 400 different
Fig. 2 Detection of trisomy 21 with MLPA. Arrows indicate alterations. An increase of the signal from the
chromosome 21 specific probes is seen
applications are commercially available from the MRC-Holland

company (www.mlpa.com).These include MLPA probemixes for
the detection of:
1. Aneuploidy of chromosomes 13, 18, 21, X, and Y (P095
probemix).
2. Deletions or duplications of specific chromosomal areas, e.g.,
the P250 probemix for the 22q11 region involved in DiGeorge
syndrome.
3. Copy number changes of all 41 unique subtelomeric regions
(P036 and P070 probemixes), all centromeric regions (P181,
P182 probemix) or 23 different microdeletion syndromes, in a
single reaction (P245 probemix).
4. Deletions or duplications involving only one or more exons of a
gene (e.g., DMD, BRCA1 or BRCA2). Small chromosomal
rearrangements can be detected as the sequences analyzed by
MLPA probes are only 60 nucleotides in length. Probemixes
for more than 100 different genes are available.
For aneuploidy testing on capillary electrophoresis systems, the
SALSA P095 probe mix is available. This probemix contains eight
independent probes for each of the chromosomes 13, 18, 21 and X,
as well as four Y-specific probes, and is used as a rapid test for
aneuploidies of these chromosomes. Extensive tests, several of
which included more than 4000 samples, indicated a diagnostic
accuracy of MLPA for the detection of common trisomies that is
statistically similar (non-inferior) to that of karyotyping or FISH
[3–10].
MLPA is not able to detect all chromosomal abnormalities seen
with karyotyping. It is designed for detection of specific copy
number changes of chromosomes 13, 18, 21 and the sex chromo-
somes. MLPA analysis is expected to detect high level of chromo-
somal mosaicism since it will give the average copy number per cell.
The P095 probemix is able to detect male triploidies but will not
detect cases of 69,XXX [3–6]. Cases of 69,XXY may be difficult to
discriminate from maternal DNA contamination as was shown for a
case of 69,XXY which was assigned as a sample contaminated by
maternal DNA [4]. However we have correctly diagnosed two cases
[6]. It should be noted, though, that many triploidies result in fetal
ultrasound abnormalities.
The MLPA aneuploidy kit is not designed for detection of
balanced chromosomal rearrangements like translocations and
inversions. MLPA however provides several opportunities that are
not possible with other techniques such as karyotyping. For exam-
ple, when a quick diagnostic result is warranted in case of fetal
abnormalities visualized by advanced ultrasound examination,
amniotic fluid samples can first be rapidly tested for the copy
number of the most common occurring aneuploidies. If normal,
further selective testing based on clinical preselection could be

done for using MLPA probemixes for all subtelomeric regions
and the most common microdeletion syndromes.
2 Materials
2.1 Contents of 1. SALSA Probemix: Mixture of up to 60 pairs of probe

SALSA MLPA Probemix oligonucleotides.
and Reagents 2. SALSA MLPA buffer: Contains 1.5 M salt + additives. Does
not always freeze at 20 C.
3. Ligase-65 enzyme solution. Does not freeze at 20 C.
4. Ligase-65 Buffer A: Contains cofactor NAD required for the
Ligase-65 enzyme.
5. Ligase-65 Buffer B: Contains the salts required by the Ligase-
65 enzyme.
6. SALSA PCR Primer mix: Contains one fluorescently labeled
and one unlabeled PCR primer + dNTPs.
7. SALSA Polymerase enzyme solution. Does not freeze at
20 C.
SALSA MLPA kits are stable for at least 1 year when stored in
the dark at 20 C.
All enzymes, nucleic acids, and buffer constituents are
non-hazardous.
In addition to standard lab equipment, such as pipettors and
water, a thermocycler with heated lid and capillary electrophoresis
instrument is required.
2.2 DNA Extraction 1. PBS .

from Amniotic Fluid 2. QIAamp DNA Mini Kit (QIAGEN, Hilden, Germany).
2.3 Separation and 1. Beckman D1-labeled 60–600 size standard (Beckman Coulter,
Quantification of the Brea, CA, USA).
MLPA Amplification 2. Deionized formamide.
Products by Capillary
Electrophoresis Using
the Beckman CEQ
Apparatus
3 Methods
3.1 Amniotic Fluid 1. Mix the amniotic fluid gently, just before a sample is removed.
Sample Preparation Most cells might be at the bottom of the tube.
Protocol: 2. Centrifuge a 2 mL sample of amniotic fluid for 5 min at
DNA-Isolation 10,000 g. Remove the supernatant carefully. Wash the pellet
with 1 mL of PBS and centrifuge for 5 min at 10,000 g.

Remove the supernatant carefully and resuspend the pellet in
200 μL of PBS. DNA-isolation is done according to the Qiagen
Blood and Body Fluid Spin Protocol. DNA is eluted with
50 μL buffer AE instead of 200 μL to increase the DNA
concentration. Transfer 5 μL DNA to a 0.2 mL vial tube for
the MLPA reaction. Store the remainder at 20 C.
3.2 DNA 1. Heat 5 μL DNA-sample (20–500 ng DNA) (see Notes 1 and 2)

Denaturation and for 5 min at 98 C in a 0.2 mL vial in a thermocycler with
Hybridization of the heated (105 C) lid.
SALSA Probes 2. Cool to 25 C before opening the thermocycler.
3. Prepare a mixture of equal volumes SALSA Probemix and
MLPA buffer at room temperature. Mix well.
4. Add 3 μL of this mixture to each sample. Mix with care by
repeated pipetting.
5. Incubate for 1 min at 95 C, followed by a 16 h incubation (see
Note 3) at 60 C (see Note 4).
3.3 Ligation Reaction 1. Prepare a Ligation master mix containing 3 μL Ligase-65 buf-
fer A, 3 μL Ligase-65 buffer B, 25 μL water, and 1 μL Ligase-65
enzyme for each reaction (see Note 5). Mix well by repeated
pipetting.
2. Reduce the temperature of the thermocycler to 54 C.
3. Add 32 μL Ligase master mix to the MLPA reaction, while the
samples are in the thermocycler, and mix by repeated pipetting.
4. Incubate for 15 min at 54 C, than heat 5 min at 98 C for
Ligase inactivation. Remove vials from the thermocycler.
3.4 PCR 1. Prepare a Polymerase master mix for each reaction containing
2 μL of SALSA PCR-primer mix, 7.5 μL of water, and 0.5 μL of
SALSA Polymerase. Mix well but do not vortex. Store on ice
until used (see Note 5).
2. While the vials are at room temperature, add 10 μL of Poly-
merase master mix to each vial. Mix by pipetting up and down,
place the vials in the thermocycler, and start the PCR reaction
(see Note 6).
3.4.1 PCR Conditions l 30 s 95 C.

l 30 s 60 C.
l 60 s 72 C; 35 cycles.
End with 20 min incubation at 72 C (see Note 7).
3.5 Separation and The amount of the MLPA PCR reaction required for analysis by
Quantification of the capillary electrophoresis depends on the apparatus and fluorescent
MLPA Amplification label used. As an example, conditions for the Beckman CEQ appa-
Products by Capillary ratus are shown:
Electrophoresis 1. Following the PCR reaction, mix 0.7 μL of the PCR reaction,
0.2 μL of the Beckman D1-labeled 60–600 size standard, 32 μl
deionized formamide.
l Settings: Capillary temperature 50 C. Denaturation 90 C
for 120 s. Injection time: 1.6 KV for 30 s. Runtime: 60 min
at 4.8 KV. Analysis settings: Include peaks >3%; Size
standard-600. Slope threshold 1.
3.6 Thermocycler l 5 min 98 C; 25 C pause.

Program for the l 1 min 95 C; 60 C pause.
Complete MLPA l 54 C pause; 15 min 54 C.
Reaction
l 5 min 98 C; 20 C pause.
l 30 s 95 C; 30 s 60 C; 60 s 72 C, 35 cycles.
l 20 min 72 C; 15 C pause.
N.B. Heated lid is at 105 C during all steps.
3.7 Data Analysis For data analysis of an MLPA experiment, Coffalyser.Net software
with the appropriate lot specific MLPA product sheet must be used
and is available free of charge on MRC-Holland’s website.
Although Coffalyser.Net uses a more sophisticated algorithm, this
section describes the basic principles.
Analysis of MLPA data consists of roughly three different parts:
First, Coffalyser.Net starts with raw data analysis (baseline cor-
rection, peak identification) and extensive quality control (e.g.,
DNA quantity used; complete DNA denaturation, degree of
sloping).
Next, MLPA data is normalized and peak signals are translated
to probe ratios. The absolute fluorescence measured by capillary
electrophoresis cannot be used directly for copy number calcula-
tions as it is affected by many variables. First, each probe’s measured
fluorescence must be normalized within each sample to get mean-
ingful data (intra-sample normalization).
In the third step, the relative probe signals are then used in the
inter-sample normalization; final probe ratios are determined by
comparing the relative probe peak in the DNA sample of interest to
all reference samples. Reference DNA samples are expected to have
a normal copy number for both the reference and target probes.
This final probe ratio is also called Dosage Quotient (DQ).
Coffalyser.Net calculates the DQ for each probe in each sample.
The MLPA peak pattern of a DNA sample without genomic
abnormalities will be identical to that of reference samples: final
Table 1
P095 Dosage Quotients expected in aneuploidy and normal cases when using 46, XY as reference
samples. Note that the P095 MLPA probemix cannot make distinction between 69, XXX and 46, XX
samples (*)
Sample type chr. 13 probes chr. 18 probes chr. 21 probes chr. X probes chr. Y probes
46, XY 1 1 1 1 1
46, XX * 1 1 1 2 0
47, XY,+13 1.5 1 1 1 1
47, XY,+18 1 1.5 1 1 1
47, XY,+21 1 1 1.5 1 1
45, X0 1 1 1 1 0
47, XXX 1 1 1 3 0
47, XXY 1 1 1 2 1
47, XYY 1 1 1 1 2
69, XXX * 1 1 1 2 0
69, XXY 1 1 1 1.33 0.67
69, XYY 1 1 1 0.67 1.33
probe ratios will be ~1.0, reflecting 2 copies for autosomal regions.

For heterozygous deletions, probe ratios of ~0.5 are expected,
while heterozygous duplications will have a DQ-value of ~1.5.
Probes should be arranged based on chromosomal location for
correct interpretation; this will also aid in detecting subtle changes
such as mosaicism.
3.8 Interpretation of Once the correct DQ values of each probe are established, the
Results relative copy number of each sample can be determined. In theory,
all probes located on a certain chromosome should give approxi-
mately the same DQ-value, also in cases of aneuploidy (Table 1). In
practice, however, certain variables such as SNPs, sample impurities
etc., can affect the DQ-value obtained for a certain probe. Also,
partial chromosome gains or losses may occur.
Based on Van Opstal and colleagues [6], the following rules are
recommended for determination of copy number status in aneu-
ploidy cases:
1. When the DQ-values of at least 4 of the 8 probes for a certain
chromosome are equal to or higher than 1.30, and the values
for the remaining four probes are close to 1.30, a trisomy for
that chromosome should be considered.
2. A monosomy X should be considered if the relative probe

signals of probes on the X chromosome are within the normal
ranges of those for normal males, and Y signals are absent.
3. A 47, XXY should be considered if the relative probe signals of
probes on the Y chromosome are within the normal ranges of
those for normal males, and those of the chromosome X probes
are within the normal range for normal females.
4. A 47, XYY should be considered if the relative probe signals of
probes on the Y chromosome are ~1.8 times those for normal
males, and those of the chromosome X probes are within the
normal range for normal males.
5. A 47, XXX should be considered if the relative probe signals of
probes on the X chromosome are ~2.5 times those for normal
males, and Y signals are absent.
6. We recommend follow-up study of samples in which several
probes for a certain chromosome are marked as having a statis-
tically abnormal probe value by Coffalyser.Net analysis. The use
of a 1.30 cut-off value to distinguish a normal from a trisomy
result does not allow detection of mosaic samples. As an exam-
ple, all chromosome 21 specific probes may have a ratio
between 1.10 and 1.20 in a mosaic sample with 30% trisomy
21 cells. Coffalyser.Net analysis may identify such mosaic sam-
ples when the experiment was performed well.
4 Notes
1. If necessary, dilute DNA with TE (10 mM Tris–HCl pH 8.0;

0.1 mM EDTA).
2. The volume of the reaction is important for the hybridization
speed, which is probe and salt concentration-dependent. Do
not use more than 5 μL sample DNA.
3. Minimum recommended hybridization period 14 h. Maximum
20 h.
4. Evaporation may occur during (A) overnight hybridization or
(B) pipetting the ligation reaction at 54 C. In case you suspect
evaporation problems, the following may help: (A): Test evap-
oration by incubating 8 μL H2O overnight at 60 C; >5 μL
H2O should remain, or (B): Reduce handling time by using
multi-channel pipettes. To reduce evaporation: (1) ensure
heated lid works well; (2) increase/decrease pressure of lid on
tubes; (3) try different tubes (e.g., Thermo Fischer ABgene
AB-0773, AB-0451); (4) use mineral oil (Vapor-lock, Qiagen
981611): add small drop of oil to DNA sample, just enough to
cover it. There is no need to remove oil. After addition of
MLPA buffer-probemix mixture or polymerase mix, centrifuge
very briefly. After addition of ligase mix, gently pipet up

and down.
5. Ligase and polymerase master mixes can be stored at 4 C for at
least 1 h.
6. Never use micro-pipettes for performing MLPA reactions that
have been used for handling MLPA PCR products! Following
PCR, the tubes should not be opened in the vicinity of the
thermocycler.
7. PCR products can be stored in the dark at 4 C for at least
4 days.
8. An extensive trouble shooting section is present on the www.
mlpa.com website.
References
1. Schouten JP, McElgunn CJ, Waaijer R et al ligation-dependent probe amplification: a pro-

(2002) Relative quantification of 40 nucleic spective study of 4000 amniotic fluid samples.
acid sequences by multiplex ligation- Eur J Hum Genet 17(1):112–121. https://
dependent probe amplification. Nucleic Acids doi.org/10.1038/ejhg.2008.161
Res 30(12):e57 7. Gerdes T, Kirchhoff M, Lind AM et al (2008)
2. Nygren AO, Ameziane N, Duarte HM et al Multiplex ligation-dependent probe amplifica-
(2005) Methylation-specific MLPA tion (MLPA) in prenatal diagnosis-experience
(MS-MLPA): simultaneous detection of CpG of a large series of rapid testing for aneuploidy
methylation and copy number changes of up to of chromosomes 13, 18, 21, X, and Y. Prenat
40 sequences. Nucleic Acids Res 33(14):e128. Diagn 28(12):1119–1125. https://doi.org/
https://doi.org/10.1093/nar/gni127 10.1002/pd.2137
3. Slater HR, Bruno DL, Ren H et al (2003) 8. Kooper AJ, Faas BH, Kater-Baats E et al
Rapid, high throughput prenatal detection of (2008) Multiplex ligation-dependent probe
aneuploidy using a novel quantitative method amplification (MLPA) as a stand-alone test for
(MLPA). J Med Genet 40(12):907–912 rapid aneuploidy detection in amniotic fluid
4. Gerdes T, Kirchhoff M, Lind AM et al (2005) cells. Prenat Diagn 28(11):1004–1010.
Computer-assisted prenatal aneuploidy screen- https://doi.org/10.1002/pd.2111
ing for chromosome 13, 18, 21, X and Y based 9. Kooper AJ, Faas BH, Feuth T et al (2009)
on multiplex ligation-dependent probe ampli- Detection of chromosome aneuploidies in cho-
fication (MLPA). Eur J Hum Genet 13 rionic villus samples by multiplex ligation-
(2):171–175. https://doi.org/10.1038/sj. dependent probe amplification. J Mol Diagn
ejhg.5201307 11(1):17–24. https://doi.org/10.2353/
5. Hochstenbach R, Meijer J, van de Brug J et al jmoldx.2009.070140
(2005) Rapid detection of chromosomal aneu- 10. Boormans EM, Birnie E, Oepkes D et al
ploidies in uncultured amniocytes by multiplex (2010) Comparison of multiplex ligation-
ligation-dependent probe amplification dependent probe amplification and karyotyp-
(MLPA). Prenat Diagn 25(11):1032–1039. ing in prenatal diagnosis. Obstet Gynecol 115
https://doi.org/10.1002/pd.1247 (2 Pt 1):297–303. https://doi.org/10.1097/
6. Van Opstal D, Boter M, de Jong D et al (2009) AOG.0b013e3181cbc652
Rapid aneuploidy detection with multiplex
Chapter 12
Chromosomal Microarray Analysis Using Array Comparative

Genomic Hybridization on DNA from Amniotic Fluid
and Chorionic Villus Sampling
Ankita Patel
Abstract
Chromosomal Microarray analysis offers an objective high resolution view of copy number changes in the
genome that contribute to genomic disorders in various clinical setting such as postnatal, prenatal, and
oncology. Here, we describe a fast and reliable method of using chromosomal microarray analysis in
detection of genomic imbalances that may be associated with congenital malformations in a prenatal
setting. Results can be obtained in 4–5 days using direct amniotic fluid (AF) or chorionic villus samples
(CVS).
Key words Array CGH, Microarray, Prenatal diagnosis, Comparative genomic hybridization
1 Introduction
Comparative genomic hybridization (CGH) was initially developed

for analysis of tumors with complex chromosomal imbalances and
low mitotic index [1]. Differentially labeled tumor and normal
DNA were applied to normal metaphase chromosomes and there-
fore the resolution of the analysis was still dependent on chromo-
some length. Crucial advancement in CGH technology came with
the replacement of normal metaphase chromosomes with arrays of
BAC or PAC (Bacterial or P1 artificial chromosome) clones immo-
bilized on glass slides for hybridization targets and later oligonu-
cleotide probes. This enabled the detection of copy number
changes throughout the genome at a higher resolution and there-
fore, provided a basis for high throughput analysis of genomic
imbalances for clinical diagnostics [2–6]. Clinical implementation
of Chromosomal Microarray Analysis (CMA) using BAC-based
comparative genomic hybridization was initially for the purpose of
diagnosing genomic imbalances primarily in individuals with devel-
opment delay, multiple congenital anomalies, and neuropsychiatric
171
172 Ankita Patel
disorders. The CMA arrays includes probes for virtually all known
microdeletion/duplication syndromes, pericentromeric and subte-
lomeric regions, as well as probes for some single gene disorders
that may occur due to gain or loss of large DNA segments. Probes
distributed randomly along all chromosome arms are also included
to identify any full trisomies. Therefore, with a single test, CMA will
detect almost all of the disorders detected by standard multiple
FISH tests and provides a major advance in the diagnosis of patients
in which a genetic cause of disability is strongly suspected but not
observed by traditional cytogenetic analysis. In an extensive review
of 33 studies including 22,698 patients overall, the International
Standard Cytogenomic Array Consortium found that CMA offered
a diagnostic yield of 15–20% as compared to 3% for G-banded
chromosome analysis in patients with intellectual disability or con-
genital anomalies [7]. Consequently, CMA is now recommended
by the American College of Medical Genetics as the first tier genetic
test for the evaluation of individuals with multiple anomalies and
nonsyndromic developmental delay/intellectual disability
[8]. CMA was then applied to prenatal studies and a large prospec-
tive NIHD study was initiated which showed the advantage of
CMA over karyotyping for detection of submicroscopic microdele-
tion and microduplication syndromes [9]. In this chapter, we
describe the methods for extracting DNA from amniotic fluid and
CVS samples for microarray analysis on oligonucleotide arrays
manufactured by Agilent Technologies. Included are methods of
whole genome amplification for when the DNA yield is low espe-
cially from lower gestation samples.
2 Materials
2.1 DNA Extraction 1. Phosphate Buffered Saline, 1.

of Direct Amniotic 2. Qiagen DNA extraction kit (Midi) (Qiagen, Germantown,
Fluid (AF) and CVS MD, USA).
Samples
3. Isopropanol (Molecular Grade, 99.5%).
4. Ethanol (Molecular Grade, 99.5%).
5. DNA Clean Concentrator ™-5 (Zymo Research, Irvine, CA,
USA).
2.2 Genomic DNA 1. NanoDrop (NanoDrop Technologies, Wilmington, DE,

Concentration USA).
and Integrity 2. Water (Molecular Grade).
Chromosomal Microarray Analysis Using Array Comparative Genomic. . . 173
2.3 Sample Digestion 1. RsaI (10.0 U/μl) (Promega, Madison, WI, USA).
for Hybridization 2. AluI (10.0 U/μl) (Promega, Madison, WI, USA).
(Optional Is Using
Agilent SNP Arrays)
2.4 Sample Labeling 1. SureTag complete DNA labeling kit (Agilent Technologies,
Santa Clara, CA, USA).
2. Human Cot-1 DNA (1.0 mg/ml) (Promega, Madison, WI,
USA).
2.5 Hybridization 1. Human Cot-1 DNA (1.0 mg/ml) (Promega, Madison, WI,
of Patient Samples USA).
2. 10 Blocking Agent (Agilent Technologies, Santa Clara, CA,
USA).
3. 2 Hybridization Buffer (Agilent Technologies, Santa Clara,
CA, USA).
4. SureHyb Chambers (Agilent Technologies, Santa Clara, CA,
USA).
5. CustomHD-CGH Microarray (Agilent Technologies, Santa
Clara, CA, USA).
6. Hybridization Chamber gasket slides (Agilent Technologies,
2.6 Array Washing 1. Distilled, deionized water (Millipore Sigma, Burlington, MA,
USA).
2. 100% Acetonitrile.
3. Agilent Oligo aCGH/ChIP-on-Chip Wash Buffer 2 (Agilent
Technologies, Santa Clara, CA, USA).
4. Agilent Oligo aCGH/ChIP-on-Chip Wash Buffer 1 (Agilent
Technologies, Santa Clara, CA, USA).
2.7 Scan Slides 1. Agilent Laser Scanner (Agilent Technologies, Santa Clara, CA,
USA).
2. Agilent Feature Extraction Software (Agilent Technologies,
3 Methods
3.1 DNA Extraction 1. Preparing the starting material.

from Direct Amniotic 2. Spin 3–10 cc amniotic fluid in a 15.0 ml tube in the Eppendorf
Fluid 5810 (swinging bucket table top), centrifuge at 3500 rpm
(2465 g) for 15 min at room temperature.
3. Remove the supernatant with a sterile transfer pipet, leaving the
cell pellet with approximately 50.0 μl of media.
174 Ankita Patel
4. Tap the tube to resuspend the pellet in the remaining media.

Ensure that the pellet is completely resuspended with no visible
clumps.
5. Add 500.0 μl 1 Phosphate Buffered Saline (PBS) to the cells
in the 15.0 ml tube with micropipettor taking care to wash
down the sides of the tube.
6. Transfer the sample using a Pasteur pipet to a new labeled
1.5 ml screw-top tube.
7. Add 500.0 μl PBS to the empty tube and transfer to the 1.5 ml
screw-top tube to transfer any remaining cells.
8. Centrifuge the tube at maximum speed in an Eppendorf
5415D benchtop microcentrifuge for 30 s.
9. Remove the PBS with an SL1000 micropipettor and transfer
into a second, new labeled 15.0 ml conical tube, leaving
approximately 20.0 μl. Vortex and resuspend the pellet.
10. Wash the cells with 1 ml of 1 PBS and centrifuge at maximum
speed for 30 s. Remove the PBS with a Pasteur pipet, leaving
~20 μl and repeat step 10.
11. Remove the PBS with a Pasteur pipet, leaving ~20 μl. Vortex
the tube vigorously to resuspend the pellet.
12. Add 600.0 μl of Cell Lysis Solution to the resuspended pellet.
13. Add 100.0 μl of Proteinase K (20.0 mg/ml) to the tube.
14. Screw on the top, place parafilm around the top, and invert
25 times to mix. Put the sample(s) in the “Shake and Bake”
oven at 37 C 2 C overnight with the rocker turned on.
15. Remove the tube(s) from the 37 C 2 C “Shake and
Bake” oven.
16. Add 3.0 μl of RNase A solution (100.0 mg/ml) to the cell
lysate. Screw on the top and place parafilm around it.
17. Invert the tube 25 times and then incubate at 37 C 2 C in
the “Shake and “Bake” oven for 15 min.
18. Remove the samples from the oven, briefly spin and place the
sample(s) on ice for 5 min. Add 200.0 μl of Protein Precipita-
tion Solution to the sample. Vortex vigorously for 20 s to
uniformly mix the sample.
19. Centrifuge at maximum speed in the Eppendorf 5415D table-
top centrifuge for 5 min.
20. Pipet 600.0 μl of 100% Isopropanol into a new 2.0 ml labeled
tube. Place on ice for at least 5 min to cool.
21. Transfer the supernatant to the tube containing the ice-cold
isopropanol.
22. Mix the sample by gently inverting 50 times.

24. Carefully pour off the isopropanol into a waste container, being
careful to leave the pellet undisturbed.
25. Spin for an additional 30 s. Pipet to remove residual
isopropanol.
26. Add 600.0 μl of 70% ethanol to the tube. Replace the top and
invert several times to wash the pellet.
28. Carefully pour off the ethanol into an ethanol waste container
being careful to leave the pellet undisturbed.
29. Spin for an additional 30 s. Pipet to remove residual ethanol.
30. Invert the tube on a clean absorbent paper for 10–15 min.
Make sure that all of the ethanol has evaporated before
proceeding.
31. Add 50.0 μl of DNA Hydration Buffer to the DNA pellet.
32. Incubate the tube at 55 C 2 C for 2 h to rehydrate the
pellet, tapping the tube periodically.
33. If possible, leave the tube overnight at room temperature to
further rehydrate the DNA.
34. Mix the sample by tapping the tube and briefly spin the sample.
35. Take and record the concentration using the NanoDrop (pro-
tocol Genomic DNA Concentration and Integrity).
36. Concentrate and clean up the DNA with Zymo column (Sub-
heading 3.3).
3.2 DNA Extraction 1. Centrifuge the tubes containing the cleaned CVS
of Direct Chorionic (3 mg–10 mg) at maximum speed for 2–5 s. Remove as much
Villus Sample (CVS) of the media as possible using a sterile transfer pipet.
2. The supernatant should be transferred to a new labeled 1.5 ml
screw top tube. It should be kept until you are sure that
sufficient DNA was isolated in this protocol and it should
then be immediately bleached and discarded.
3. If received in a 15 ml tube, transfer villi to a labeled 1.5 screw
top tube.
4. Add 1.0 ml 1 Phosphate Buffered Saline (PBS) to the villi in
the tube.
5. Centrifuge the tube at maximum speed in a benchtop micro-
centrifuge for 15 s.
6. Remove the PBS with an SL1000 micropipettor, leaving
approximately 20.0 μl.
176 Ankita Patel
7. Flick the tube gently to resuspend the pelleted villi and repeat
steps 4 and 5.
8. Add 600.0 μl of Cell Lysis Buffer and 20.0 μl of Proteinase K
(20 mg/ml) to the microfuge tube. Parafilm the tube and
invert 25 times to mix. Place on rocking platform at 37 C 2 C
overnight.
9. Remove the sample from the rocking platform (see Note 1).
10. Add 3.0 μl of RNaseA (100.0 mg/ml) to the cell lysate. Invert
the tube 25 times and incubate in a 37 C 2 C incubator for
15 min.
11. Add 600.0 μl of room-temperature 100% isopropanol to a new,
labeled 2.0 ml empty tube and place on ice for at least 2 min
before step 12.
12. Quickly spin the tube and place the sample on ice for 5–10 min.
13. Add 200 μl of protein precipitation solution and vortex vigor-
ously for 20 s.
14. Spin at maximum speed in a tabletop centrifuge for 5 min.
15. If the pellet is not tight, place on ice for another 5 min and
centrifuge again for 3 min.
16. Transfer the Supernatant to the tube containing ice-cold
isopropanol.
17. Mix the sample by gently inverting 50 times.
18. Centrifuge at maximum speed in the tabletop centrifuge for
5 min.
19. Pour off the isopropanol into a clean labeled waste tube, being
careful to leave the pellet undisturbed. Spin for an additional
30 s to remove any residual isopropanol and pipet out.
20. Add 600.0 μl of 70% ethanol to the tube. Replace the top and
invert several times to wash the pellet.
21. Centrifuge at maximum speed in the tabletop centrifuge for
5 min.
22. Pour off the ethanol into a labeled waste tube, being careful to
leave the pellet undisturbed. Spin for an additional 30 s to
remove any residual ethanol and pipet out.
23. Leave the tube to air dry for 10–15 min for the remaining
ethanol to evaporate off.
24. Add 50.0 μl of DNA Hydration Buffer (or more for a large
pellet) to the DNA pellet.
25. Incubate the tube at 55 C 2 C for 2 h to rehydrate the
pellet, tapping the tube periodically. If possible, leave the tube
overnight at room temperature to further rehydrate the DNA.
26. Mix the sample by tapping the tube and briefly spin the sample.
Measure the concentration using the Nanodrop and then clean
a maximum of 5 μg of DNA on a Zymo column.
27. If the concentration prior to cleanup is more than 500 ng/μl,
parafilm the sample and let it sit overnight at room
temperature.
3.3 DNA Clean Up 1. Add 100 μl of Binding buffer to 5 μg of DNA sample. Vortex
Using Zymo Columns and let it sit for 5 min at room temperature.
2. Add the sample to the zymo mini filter sitting in a collection
tube. Spin for 30 s. Discard the waste.
3. Add 200 μl of wash buffer to the zymo mini filter. Spin for 30 s.
Discard the waste. Repeat.
4. For elution A, transfer the mini filter to a new labeled
1.7 ml tube.
5. Add 18 μl of water to the filter. Let it sit for 10 min and spin for
30 s into the first labeled tube.
6. For elution B, transfer the mini column to another new labeled
1.7 ml tube.
7. Add 10 μl of water to the filter. Let it sit for 5 min. Spin for 30 s
into a second labeled tube.
8. Determine the concentration of each elution using the
Nanodrop.
3.4 Genomic DNA 1. Determine the DNA concentration and quality using any stan-
Concentration dard DNA measuring technique. 260/280 ration should be
and Integrity between 1.75 and 2.00 and the 260/230 ratio should be above
1.5.
2. If the 260/280 ratio is below 1.75 or above 2.00, or the
260/230 ratio is below 1.5, or the concentration is below
66 ng/μl, purify the sample by again by using the Zymo
column.
3. At this point an aliquot may be taken for maternal cell contam-
ination studies before proceeding further.
3.5 Sample Digestion 1. If using Agilent CGH + SNP arrays, then digestion of the DNA
for Hybridization with AluI and RsaI is necessary since the SNP probes on the
array are AluI/RsaI restricted. If the non-SNP Agilent arrays
are used or the SNP probes are not to be evaluated the DNA
can be processed for labeling (Go to DNA labeling method).
2. Place top labels on closed 1.7 ml microfuge tubes for both the
patients and the controls.
3. Add the appropriate amount of nuclease-free water and geno-
mic DNA to make 1.0 μg genomic DNA in 20.0 μl to a labeled
178 Ankita Patel
Table 1
Digestion master mix for each single tube reaction (prepare in order)
Reagent Volume (μl) Final concentration

1 Nuclease-free water 2.50
2 10 Buffer C 2.10
3 Acetylated BSA (10.0 μg/μl) 0.40 0.67 μg/μl
4 AluI (10.0 U/μl) 0.50 0.83 U/μl
5 RsaI (10.0 U/μl) 0.50 0.83 U/μl
Total volume 6.00
tube for each reaction, following the volumes stated on the

worksheet.
4. Add 1.0 μg final volume of 20 μl genomic gender-matched
control DNA to a separate labeled 1.7 ml microfuge tube.
5. The gender control should be thoroughly mixed by tapping or
inverting before use. DNA should not be vortexed regularly, as
this will shear it.
6. Prepare the digestion master mix on ice with the components
listed in Table 1, in the order indicated.
7. Add 6.0 μl of the Digestion Master Mix to each reaction tube.
8. Each tube should now have a total volume of 26.0 μl. Place the
tubes on ice as soon as you have added the Master Mix.
9. Mix well by flicking the bottom of the tubes and spin them at
maximum speed for 2–10 s.
10. Place the sample tubes at 37 C 2 C (either in a floating rack
in a water bath or in a standard rack in an oven) and incubate
from 2 to 17 h (approximately overnight).
11. Transfer the sample tubes to a floating rack in a water bath at
65 C 2 C and incubate for 20 min to inactivate the
enzymes. Place tubes on ice for at least 5 min.
12. Proceed to the Sample Labeling protocol.
3.6 Sample Labeling 1. If samples were stored at 20 C 2 C after enzyme digestion
(Sample Digestion for Hybridization protocol), thaw at room
3.6.1 Fluorescent
temperature for 5 min.
Labeling of Genomic DNA
2. Centrifuge the tubes at maximum speed for 2–10 s.
3. Add 5.0 μl of 10 Random Primers to each reaction tube
containing 26.0 μl of digested genomic DNA.
4. Flick to mix. Centrifuge the tubes at maximum speed for
2–10 s.
Table 2
Labeling master mix set up for Single Tube Reaction
Reagent Volume (μl)

1 Nuclease-free water 2.0
2 5 Reaction buffer 10.0
3 10 dNTP 5.0
4 dCTP labeled dye (Cy3—1.0 mM or Cy5—1.0 mM) 3.0
5 Klenow (Exo-) 1.0
Total volume 21.00
5. Denature the DNA in a 105 C 2 C heat block for 3 min.

6. Immediately place the tubes on ice for 5 min, and then spin the
tubes at maximum speed for 2–10 s. Return the tubes to ice.
7. Make a master mix for each dye (Cy5 for the sample DNA and
Cy3 for the control DNA) using the volumes in Table 2, which
list volumes for one labeling reaction. Add the components on
ice in the order indicated. Mix by tapping and spin at maximum
speed for 2–10 s.
8. Add 19.0 μl of Labeling Master Mix to each reaction tube. Flick
to mix. Centrifuge the tubes at maximum speed for 2–10 s.
9. Place the sample tubes in a floating rack in a water bath at
37 C 2 C and incubate for 2 h.
10. Transfer the sample tubes to a water bath at 65 C 2 C and
incubate for 10 min to inactivate the enzyme. Place tubes on
ice for at least 5 min (see Note 2).
3.6.2 Clean-Up 1. Unincorporated dyes and dNTPs are removed by using Milli-
of Labeled Genomic DNA pore Microcon Columns provided in the SureTag labeling kit.
2. Add 430.0 μl of 1 TE (pH 8.0) to each reaction tube,
bringing the total volume to 480.0 μl.
3. Place a Microcon YM-30 filter into the supplied labeled 1.7 ml
microfuge tube and load the correct labeled DNA sample onto
the filter. Spin for 10 min at 9300 RPM (8000 g) in a
microcentrifuge at room temperature. Discard the flow-
through in a labeled Cy-3 / Cy-5 waste container.
4. Add 480.0 μl of 1 TE (pH 8.0) to each filter. Spin for 10 min
at 9300 RPM (8000 g) in a microcentrifuge at room tem-
perature. Discard the flow-through in the labeled Cy-3 / Cy-5
waste container.
180 Ankita Patel
5. Invert the filter into a fresh supplied labeled 1.7 ml microfuge

tube. Spin for 1 min at 9300 RPM (8000 g) in a microcen-
trifuge at room temperature to collect the purified sample.
6. Measure and record the volume of each eluate in μl. If the
sample volume is more than 21 μl, return the sample to its
filter and spin for 1 min at 9300 RPM (8000 g) in a mir-
ocentrifuge at room temperature. Discard the flow-through in
a labeled Cy-3/Cy-5 waste.
7. Repeat steps 5 and 6 until each sample volume is less than or
equal to 21.0 μl.
8. Bring the total sample volume to 21.0 μl with 1 TE (pH 8.0)
if necessary.
9. Proceed to Subheading 3.6.3 (see Note 3).
3.6.3 Quantitate the Yield 1. Open program “ND-1000 v3.30” on the desktop of a com-
and Specific Activity puter attached to a NanoDrop Spectrophotometer.
of the Labeling 2. Select “MicroArray” from the first screen.
3. Place 1.5 μl of Nuclease-Free water on the pedestal. Lower the
arm. Click OK. Wait for reading.
4. Select “DNA-50” for “Sample Type” from the next screen.
5. Wipe off pedestal with a Kimwipe.
6. Place 1.5 μl of 1 TE (pH 8.0) on the pedestal.
7. Click “Blank” or press F3. All readings should go to 0.0.
8. Wipe off the sample with a Kimwipe and replace with a fresh
1.5 μl of 1 TE (pH 8.0).
9. Click “Measure” or press F1.
10. If the concentration reading is between 0.0 ng and 1.0 ng,
proceed to the next step.
11. Measure the absorbance at A260nm (DNA), A550nm (cyanine
3), and A650nm (cyanine 5).
12. Calculate DNA (μg) ¼ (A260–A320) 50 μg/ml volume in ml
Dye Incorporation:
Cy3™ (pmole) ¼ (A550–A650)/0.15 volume in μl
Cy5™ (pmole) ¼ (A650–A750)/0.25 volume in μl
13. Match each patient sample with the gender-matched control
that most closely matches its concentration. The concentration
must be above 4 μg and the pmole >100.
14. Proceed to Hybridization of Patient Samples to Agilent Arrays
protocol (see Note 4).
3.7 Hybridization 1. Add the components listed in Table 3, in the order listed, to a
nuclease-free labeled tube.
Table 3
Hybridization volumes for different Agilent array formats
2400K array 4180K array 2105K array 860K array

Component (μl) (μl) (μl) (μl)
1 Cyanine 5-labeled sample 19.5 19.5 19.5 19.5
2 Cyanine 3-labeled sample 19.5 19.5 19.5 19.5
3 1 TE 40.0 0.0 40.0 0.0
4 CoT-1 DNA (1.0 mg/ml) 25.0 5.0 25.0 2.5
5 Agilent 10 blocking agent 26.0 11.0 26.0 4.5
6 Agilent 2 hybridization 130.0 55.0 130.0 22.5
buffer
Final hybridization sample 260.0 110.0 260.0 68.5
volume
2. Mix the sample by pipetting it up and down several times. Flick

the bottom of the tube to mix. Centrifuge the tubes at
8000 g for 2–10 s.
3. Place sample tubes in a heat block at 105 2 C and incubate
for 3 min.
4. Immediately transfer sample tubes to a water bath at 37 2 C
and incubate for 30 min.
5. Remove sample tubes from the water bath. Spin for 1 min at
8000 g in a microcentrifuge.
6. Load a clean gasket slide into the Agilent SureHyb chamber
base with the gasket label facing up and aligned with the
rectangular section of the chamber base. Ensure that the gasket
slide is level and seated properly within the chamber base.
7. Slowly dispense the hybridization sample mixture into the
gasket well. Load all gasket wells, being extremely careful to
dispense each sample into the correct well.
8. After checking the Hybridization Worksheet to be sure that
you have the correct slide, place the slide microarray-side down
onto the SureHyb gasket slide, so the numeric barcode side is
facing up and the “Agilent”-labeled barcode is facing down.
The label on the gasket slide should be lined up with the label
on the array slide.
9. Gently place the SureHyb chamber cover onto the sandwiched
slides and slide the clamp assembly onto both pieces.
10. Hand-tighten the clamp onto the chamber. Experience will
show how to get the chambers tight enough, but not too
tight to break the slide.
182 Ankita Patel
11. Vertically rotate the assembled chamber to wet the slides and
assess the mobility of the bubbles. Tap the assembly on the
palm of your hand if necessary to move bubbles.
12. Place the assembled slide chamber in the rotator rack in a
hybridization oven set to 65 C 2 C. Be sure that the rotator
is balanced both side-to-side and front-to-back. Set the hybri-
dization rotator to 20 rpm.
13. Hybridize at 65 2 C for 20–68 h.
14. Proceed to Array Washing protocol.
3.8 Array Washing 1. Place at least 250 ml of Agilent Oligo aCGH/ChIP-on-Chip

Wash Buffer 2 in a 37 2 C water bath and a dish labeled
“Wash BF2” in a 37 2 C incubator overnight, or at least 3 h
before washing.
2. Fill a slide-staining dish labeled “Wash BF1” with approxi-
mately 350 ml of room-temperature Agilent Oligo aCGH/
ChIP-on-Chip Wash Buffer 1 and place it in a fume hood.
3. Place a slide rack into another slide-staining dish labeled “Wash
BF1.” Add a magnetic stir bar. Fill this dish with enough room-
temperature Oligo aCGH Wash Buffer 1 to cover the slide rack
(~250 ml). Place this dish on a magnetic stir plate in the hood.
4. Place the dish labeled “Acetonitrile” in the fume hood and fill
with approximately 300 ml with room-temperature acetonitrile
(see Note 5).
5. The procedure of washing is conducted as depicted in Table 4.
6. Remove the hybridization chambers to be washed from the
65 2 C incubator.
7. Place the first slide into the wash buffer. Place the hybridization
chamber assembly on a flat surface and disassemble it. Loosen
the thumbscrew, turning counter-clockwise. Slide off the
clamp assembly and remove the chamber cover. Remove the
array-gasket sandwich from the chamber base by grabbing the
slides from their ends.
8. Keep the microarray slide numeric barcode facing up and sub-
merge the array-gasket sandwich into the first dish containing
Oligo aCGH Wash Buffer 1. Do not let go of the slides.
9. With the sandwich completely submerged in Oligo aCGH
Wash Buffer 1, pry the sandwich open from the barcode end.
Insert one end of the plastic forceps between the slides and
gently turn the forceps to separate the slides. Let the gasket
slide drop to the bottom of the staining dish. Remove the
microarray slide and place it into the slide rack in the second
dish containing Oligo aCGH Wash Buffer 1, being very careful
not to touch the array. Minimize exposure of the slide to air.
Table 4
Procedure of microarray slide washing
Procedure Wash Buffer Temperature Time

Disassembly Oligo aCGH Wash Buffer 1 Room temperature Depends on the total number of slides
1 Oligo aCGH Wash Buffer 1 Room temperature 5 min
2 Oligo aCGH Wash Buffer 2 37 2 C 1 min
3 Acetonitrile Room temperature 1 min
10. Repeat steps 7 and 8 for up to four additional slides, leaving a

blank space between each pair of slides. No more than five
slides can be washed in each wash batch. Larger dishes can be
used for a maximum of 10 slides.
11. When all slides in the batch are placed into the slide rack in the
second dish, stir using a setting between 110 and 130 for 5 min
(Table 4).
12. When there is 1 min left on the previous wash, remove a
pre-warmed dish and the Oligo aCGH Wash Buffer 2 from
the 37 C 2 C incubator and water bath. Place the dish on a
stir plate, add a magnetic stir bar, and fill the dish with Oligo
aCGH Wash Buffer 2 to the top of the label.
13. When the time is up, transfer the slide rack to the dish contain-
ing the Oligo aCGH Wash Buffer 2 and stir on a setting
between 110 and 130 for 1 min (Table 4).
14. Remove the slide rack from the dish and tilt the rack slightly to
minimize wash buffer carry-over. Transfer the slide rack to the
dish containing Acetonitrile and leave for 1 min (Table 4).
15. Remove the slide rack very slowly to minimize droplets on the
slides. It should take about 10 s to remove the slide rack.
16. Scan slides immediately to minimize the impact of air oxidation
on signal intensities, or store slides in slide boxes in a
desiccator.
17. If necessary, repeat steps 6–16 for the next group of five slides
using fresh Oligo aCGH Wash Buffer 1 and pre-warmed Oligo
aCGH Wash Buffer 2. The Oligo aCGH Wash Buffer 1 in the
dish used to open the array-gasket assembly can be used
throughout the day and should be discarded after all of the
day’s washing has been completed. The Oligo aCGH Wash
Buffer 1 (on the stir plate) and Oligo aCGH Wash Buffer 2 can
be used for up to 5 slides (see Notes 5 and 6).
18. When all washing for the day has been completed, wash all of
the dishes, the slide rack, and the stir bars. Pour used Oligo
aCGH Wash Buffer 1 and Oligo aCGH Wash Buffer 2 down
184 Ankita Patel
the lab sink. Rinse the dish and lid with tap water, and then fill
the dish several times with Millipore water and empty. Rinse
the lid with Millipore water. Air dry the Oligo aCGH Wash
Buffer 1 dishes. The Oligo aCGH Wash Buffer 2 dish can be
placed directly in the incubator. Discard the acetonitrile in the
acetonitrile waste container in the hood. Air dry the dishes in
the hood.
3.9 Scanning Slides 1. Open the “Feature Extraction” software by double-clicking its
and Analysis icon on the desktop. Select the appropriate project. Start
Extracting, or click the “start extracting” icon. Minimize the
“Feature Extraction” window.
2. Open the “Scan Control” software by double-clicking its
shortcut on the desktop.
3. Place the slides in the Agilent slide carriers.
4. Place the slide carrier on a flat surface. Use your thumb to slide
the ridged end of the cover toward the open rectangle at one
end. Open the hinged cover.
5. Holding a slide with the array side up and with the word
“Agilent” toward the open rectangle. Place the one end of
the slide on the ledge inside the slide holder. Gently lower
the slide into place. Make sure that it is seated flush on the
support ledges.
6. Gently lower the hinged cover. If it does not close easily, make
sure that the slide is seated properly on the ledge. Push down
the cover gently and slide the locking tabs in place.
7. Place the slide carriers into the carousel. Open the hinged
chamber of the scanner.
8. Place each slide carrier into the carousel.
9. Do not place a carrier in the slot marked “H.” Place the first
carrier in position one and continue to load all of the carriers.
Do not skip any slots otherwise this will cause the run to stop.
The slides should be placed with the word “Agilent” at the
center and facing toward lower numbers, i.e., the open back of
the slide showing the barcode should be facing out.
10. Close the chamber door.
11. Enter the position of the first slide to be scanned in the “Start
slot:” field of the software. This will normally be “1.”
12. Enter the position of the last slide to be scanned in the “End
slot:” field.
13. Select the appropriate scanner profile.
14. Select the proper resolution. The current settings are 5 μm for
standard resolution arrays and 3 μm for high resolution arrays.
15. Press the “Scan slot 1—X” button, where “X” is the number
you entered for the last slide to be scanned.
16. Scanning will take approximately 10–15 min per slide depend-
ing on the resolution.
17. The .tiff images will be automatically output to the folder “D:
\Scanner Output Location.”
18. The .tiff files will be automatically extracted as they are com-
pleted. The software will automatically detect new .tiff files as
they are created.
19. The software will remain active until it has not detected a new
file generation in X min (X can be a number you can preset on
the Feature Extraction program). The program can be stopped
by the user by selecting the “Project/Stop” icon from the
menu bar.
20. Only the slides with barcodes will be automatically extracted.
21. After scanning the text files can be uploaded into the Agilent
CytoGenomics software. After normalization and subtraction
of background noise the data are to log ratios and plotted.
4 Notes
1. DNA isolation can stop here for continuation the next day.
Samples may also be left at this point over the weekend. Keep
the cell lysate at room temperature if extraction is to be
continued later.
2. Samples can be stored at 20 C for a day before proceeding to
the clean-up step.
3. Cleaned, labeled DNA can be stored overnight or over the
weekend at 20 C 2 C, either before or after quantitating.
4. Subheading 3.7 is adapted with modification from Agilent
manual “Agilent Oligonucleotide Array-Based CGH for Geno-
mic DNA Analysis”.
5. If fewer than 5 slides are washed and more are to be washed
later the same day, leave the Oligo aCGH Wash Buffer 1 in the
hood and place the Oligo aCGH Wash Buffer 2 in its tray in the
37 2 C incubator until ready for use.
6. Acetonitrile can be used for up to 20 slides. Leave in the hood if
it will be used later for another wash.
186 Ankita Patel
References
1. Kallioniemi A, Kallioniemi OP, Sudar D et al 6. Cheung SW, Shaw CA, Yu W et al (2005) Devel-
(1992) Comparative genomic hybridization for opment and validation of a CGH microarray for
molecular cytogenetic analysis of solid tumors. clinical cytogenetic diagnosis. Genet Med 7
Science 258(5083):818–821 (6):422–432. https://doi.org/10.109701.
2. Solinas-Toldo S, Lampel S, Stilgenbauer S et al GIM.0000170992.63691.32
(1997) Matrix-based comparative genomic 7. Miller DT, Adam MP, Aradhya S et al (2010)
hybridization: biochips to screen for genomic Consensus statement: chromosomal microarray
imbalances. Genes Chromosomes Cancer 20 is a first-tier clinical diagnostic test for indivi-
(4):399–407 duals with developmental disabilities or congen-
3. Snijders AM, Nowak N, Segraves R et al (2001) ital anomalies. Am J Hum Genet 86
Assembly of microarrays for genome-wide mea- (5):749–764. https://doi.org/10.1016/j.ajhg.
surement of DNA copy number. Nat Genet 29 2010.04.006
(3):263–264. https://doi.org/10.1038/ng754 8. Manning M, Hudgins L, Professional P et al
4. Cai WW, Mao JH, Chow CW et al (2002) (2010) Array-based technology and recommen-
Genome-wide detection of chromosomal imbal- dations for utilization in medical genetics prac-
ances in tumors using BAC microarrays. Nat tice for detection of chromosomal
Biotechnol 20(4):393–396 abnormalities. Genet Med 12(11):742–745.
5. Ylstra B, van den Ijssel P, Carvalho B et al (2006) https://doi.org/10.1097/GIM.
BAC to the future! or oligonucleotides: a per- 0b013e3181f8baad
spective for micro array comparative genomic 9. Wapner RJ, Martin CL, Levy B, et al (2012)
hybridization (array CGH). Nucleic Acids Res Chromosomal microarray versus karyotyping for
34(2):445–450. https://doi.org/10.1093/ prenatal diagnosis. The New England journal
nar/gkj456 of medicine. 367(23):2175–2184. https://doi.
org/10.1056/NEJMoa1203382
Chapter 13
Prenatal Diagnosis Using Chromosomal SNP Microarrays

Mythily Ganapathi, Odelia Nahum, and Brynn Levy
Abstract
Chromosomal microarray is a high resolution genomic technology to diagnose genetic conditions asso-
ciated with losses or gains of the human genome. This technology is currently routinely used in numerous
clinical settings, including postnatal diagnosis of disorders with genetic etiologies such as intellectual
disability, developmental delay, neurocognitive phenotypes, congenital anomalies, and prenatal diagnosis
wherein the referral could be ultrasound anomalies, advanced maternal age, and normal course of preg-
nancy. We describe the use of Chromosomal SNP microarrays for prenatal diagnosis of genetic disorders
which result from both copy number or copy neutral changes in the genome.
Key words Chromosomal microarray analysis, Chromosomal SNP microarray, Prenatal diagnosis,
Ultrasound abnormalities
1 Introduction
Chromosomal microarray analysis offers a high resolution approach

to diagnose disease conditions which occur due to losses and gains
in the human genome which are denoted as copy number variants
(CNVs). They include whole chromosomal aneuploidies as well as
submicroscopic gains or losses that are too small to be detected by
conventional karyotyping. For chromosomal microarray analysis,
patient’s DNA is fluorescently labeled and hybridized to a solid
support containing thousands of oligonucleotide-based DNA
probes. The copy number of a probe on the microarray is deter-
mined by comparative hybridization of the labeled patient DNA to
an in silico reference set. Oligonucleotide probes can be designed to
detect copy number changes in a sequence compared with a con-
trol, additionally, they may be designed to identify a specific geno-
type or allele of a single nucleotide polymorphism (SNP).
Chromosomal microarray platforms with copy number as well as
SNP probes additionally provide clinically relevant information
about copy neutral changes such as long continuous stretches of
187
188 Mythily Ganapathi et al.
homozygosity, uniparental disomy and will also identify ploidy

changes.
Chromosomal microarray analysis is particularly useful in the
diagnosis of genetic abnormalities in the prenatal setting as a direct
CVS/amniocentesis/POC sample can be assayed obviating the
need for a cultured specimen, resulting in faster turnaround
times. However, if DNA amounts from a direct sample are insuffi-
cient, then a cultured sample can be used for the assay. Targeted
chromosomal microarrays wherein known regions of interest are
analyzed are the preferred choice of most laboratories offering
prenatal testing to avoid the calling of VOUS variants which can
be a source of anxiety and distress for the parents. The American
College and the Society for Maternal-Fetal Medicine in 2013
recommended chromosomal microarray analysis as the First-tier
prenatal test in case of abnormal ultrasound findings [1]. Further,
CNVs identified by chromosomal microarray analysis such as
microdeletions and microduplications are not associated with
advanced maternal age [2]; hence it was recommended to offer
this test to all pregnant women and not restrict it to those aged
35 years and older. In cases of intrauterine fetal demise or stillbirth
chromosomal microarray analysis on fetal tissue (i.e., amniotic
fluid, placenta, or products of conception) was recommended
because of its improved detection of causative abnormalities.
The microarray platform we currently use is the Affymetrix
Cytoscan HD array which has a total of 2.67 million markers and
includes 1.9 million non-polymorphic copy number probes
and 750,000 SNPs for genotype calls. The described method can
be applied to direct or cultured prenatal specimens (amniotic fluid
or chorionic villi) as well as for products of conception from mis-
carriages. On the day of sample receipt, DNA can be extracted,
restriction enzyme digested and ligated with adaptors. Extraction is
performed by a lysis procedure using a commercial extraction kit
(Qiagen DNA mini kit cat, QIAGEN, Germantown, MD, USA).
The DNA is then PCR amplified, fragmented by DNAse digestion,
and labeled by Terminal Deoxynucleotidyl transferase reaction.
DNAse digestion is performed to fragment the genomic DNA
into segments approx 25 bp to 125 bp. After labeling, the DNA
is denatured and then loaded on to the array. The arrays are hybri-
dized overnight, washed to remove any unbound genomic DNA,
stained, scanned and analyzed. Herein, we report common chro-
mosomal SNP microarray procedure used to take a sample from
receipt to results in a clinical diagnostic setting.
Prenatal Diagnosis Using Chromosomal SNP Microarrays 189
2 Materials
Unless otherwise specified, reagents are provided with the Affyme-

trix Cytoscan HD kit (Thermo Fisher Scientific, Waltham, MA,
USA) (see Note 1).
2.1 Restriction 1. 100 BSA.

Enzyme Digestion 2. 10 NspI Buffer.
3. NspI Enzyme.
4. Chilled Affymetrix Nuclease-free water.
2.2 Ligation Reaction 1. 10 T4 DNA Ligase Buffer.

2. 50 μM Adaptor, NspI.
3. T4 DNA Ligase.
2.3 PCR 1. Affymetrix Nuclease-free water.

2. PCR Primer 002 (100 μM).
3. dNTPs (2.5 mM each).
4. Titanium DNA Amplification Kit (Clontech Laboratories,
Mountain View, CA, USA): Contains GC-Melt (5 M), TITA-
NIUM™ Taq DNA Polymerase (50), TITANIUM™ Taq
PCR Buffer (10).
2.4 Checking the 1. Affymetrix Nuclease-free water.

PCR Reaction Results 2. DNA Marker (USB/Thermo Fisher Scientific, Waltham, MA,
USA): PCR Markers, 50–2000 bp.
3. E-Gel EX 2% Agarose Gels (Invitrogen, Carlsbad, CA, USA).
4. 6 DNA loading buffer (USB/Thermo Fisher Scientific, Wal-
tham, MA, USA).
5. Strip Tubes (Bio-Rad Laboratories, Hercules, CA, USA).
2.5 PCR Product 1. Elution Buffer.

Purification 2. Purification Wash Buffer.
3. Magnetic beads.
2.6 PCR Product 1. Affymetrix nuclease-free water.

Quantitation of
Purified PCR Products
2.7 Fragmentation 1. Fragmentation Buffer.

2. Fragmentation Reagent (enzyme; DNase I).
3. Affymetrix Nuclease-free water.
2.8 Checking the 1. 4% E-Gel EX agarose gel (Invitrogen, Carlsbad, CA, USA).
Fragmentation 2. DNA Marker (TrackIt-25 bp) (Invitrogen, Carlsbad, CA,
Reaction by Running a USA).
Gel
3. 6 DNA loading buffer (USB/Thermo Fisher Scientific, Wal-
tham, MA, USA).
4. Strip Tubes (Bio-Rad Laboratories, Hercules, CA, USA).
2.9 Labeling 1. DNA Labeling Reagent (30 mM).

2. Terminal Deoxynucleotidyl Transferase (TdT; 30 U/μL).
3. Terminal Deoxynucleotidyl Transferase Buffer (TdT Buffer;
5).
2.10 Hybridization 1. Hybridization buffer Part 1.

2. Hybridization buffer Part 2.
5. Oligo Control Reagent (OCR), 0100.
2.11 Washing, 1. Stain Buffer 1.

Staining and Scanning 2. Stain Buffer 1.
the Arrays
3. Affymetrix GeneChip Array holding buffer.
4. Affymetrix GeneChip Wash A.
5. Affymetrix GeneChip Wash B.
3 Methods (See Notes 2–5)
1. Include Affymetrix Nuclease-free water as a negative control

and REF 103 DNA (supplied in the kit) in every batch of
samples to be set up for microarray analysis.
2. The duration of the assay can take between 3 and 4 days. We
usually do it in 4 days as described below, however it can be
shortened to 3 days:
l Day 1: Digestion + Ligation.
l Day 2: PCR + Purification.
l Day 3: Fragmentation + Labeling + Hybridization
(overnight).
l Day 4: Washing and Staining + Scanning.
3.1 DNA Dilution A starting concentration of 50 ng/μL (range: 45–64 ng/μL) of

DNA is needed for samples that have to be run on the Affymetrix
Cytoscan HD protocol (see Note 6).
1. Thaw the genomic DNA (gDNA).
2. Vortex the gDNA samples at high speed for 3 s and short spin
it.
3. If sample concentration is unknown, take an OD measurement
of the sample.
4. Based on OD measurements, dilute each sample to 50 ng/μL
using molecular biology grade water/TE (depending upon the
original solution that the DNA was eluted in) (see Note 6).
3.2 Restriction During this stage, one aliquot of each sample is digested by the
Enzyme Digestion NspI restriction enzyme. This assay has to be conducted in a Pre-
PCR clean area (preferably in a hood).
1. Thaw the Buffers and BSA on ice.
2. Leave the enzymes at 20 C until ready to use.
3. Using a marker, label one 1.5 mL Eppendorf tube as NspI.
4. Put tube on ice. Vortex the gDNA and controls at high speed for
3 s, spin down the DNA, and place in the cooling chamber on ice.
5. Vortex the buffers and BSA three times, 1 s each time.
6. Pulse spin for 3 s and place in the cooling chamber on ice.
7. Power on the thermal cycler to preheat the lid.
8. Make a master mix by adding the following to the labeled
Eppendorf tubes (see Notes 7 and 8): For each sample add
5 μL of the diluted genomic DNA (250 ng), 11.55 μL Affyme-
trix Nuclease-free water, 2 μL 10 NspI Buffer, 0.2 μL 100
BSA and 1 μL NspI. Scale quantities according to the number
of samples to be processed. Place the master mix on the cooling
chamber between adding reagents.
9. Vortex the master mix at high speed three times, 1 s each time.
10. Pulse spin for 3 s and place in the cooling chamber.
11. Return any remaining enzyme to the freezer.
12. Using a single channel P20 pipette, add 14.75 μL of the
Digestion Master mix to each well containing the genomic
DNA. The total volume in each well is now 19.75 μL.
13. Seal the plate tightly with a microseal film.
14. Vortex the center of the plate and all corners at 75% speed for
1 s. Spin the plate at 650 g for 1 min.
15. Ensure that the lid of thermal cycler is preheated.
16. Load the plate onto the thermal cycler and run the CytoScan
Digest program as follows: 37 C for 120 min, 65 C for
20 min, 4 C hold (see Note 9).
17. When the program is finished, remove the plate and spin it
down at 650 g for 1 min.
18. If proceeding directly to the ligation assay, place the plate on a
cooling chamber on ice. Otherwise, store the plate at 20 C.
3.3 Ligation Reaction During this stage, the NspI digested samples are ligated using the
NspI Adaptor.
This assay should be conducted in a Pre-PCR clean area (pref-
erably in a hood).
1. Allow the following reagents to thaw on ice: Adaptor NspI
(50 μM), T4 ligase buffer (10)—mix it thoroughly before
use to ensure precipitate is resuspended.
2. Turn the Power on the thermal cycler to preheat the lid.
3. Make a master mix by adding the following to a 500 μL
Eppendorf tube (see Notes 7 and 8): For each sample add
0.75 μL Adaptor NspI/Sty (50 μM), 2.5 μL T4 DNA Ligase
buffer (10), 2 μL T4 DNA ligase (400 U/μL). Scale quan-
tities according to the number of samples to be processed.
Place the master mix on the cooling chamber between adding
reagents.
4. Vortex the master mix at high speed three times, 1 s each time
then spin for 3 s and place in the cooling chamber.
5. Using a single channel P100 pipette, add 5.25 μL of the
Ligation Master mix to each well containing the digested
genomic DNA. The total volume in each well is now 25 μL.
6. Seal the plate tightly with microseal film.
7. Vortex the center and the corners of the plate at 75% speed for
3 s.
8. Spin the plate at 650 g for 1 min.
10. Load the plate onto the thermal cycler and run the Ligation
program as follows: 16 C for 180 min, 70 C for 20 min, 4 C
hold (see Note 9).
11. When the program is finished, remove the plate and spin at
650 g for 1 min.
12. Place the plate on a cooling chamber if PCR assay is going to be
set up immediately. Otherwise, store the plate at 20 C.
3.4 PCR The assay should be set up in a Pre-PCR clean area (preferably in a
hood). The PCR should be run on the thermal cycler in a post PCR
area.
1. Add 75 μL of Affymetrix Nuclease-free water to each DNA
ligation reaction, in order to dilute the DNA (see Note 10).
2. Tightly seal the plate. Vortex at 75% speed power for 1 s; then
spin the plate at 650 g for 1 min.
3. Allow the following reagents to thaw on ice: G-C Melt (5 M),
dNTP (2.5 mM each), PCR Primer (100 μM), Taq buffer
(10).
4. Turn the Power on the thermal cycler to preheat the lid.

5. Label a fresh 96-well plate in such way that each sample will be
added in 4 wells. In a case where you process 8 samples (a
whole column) the new plate should have 4 columns labeled,
so each sample will now have 4 wells. To each well, add 10 μL
of the diluted ligated DNA.
6. Make a master mix by adding the following to a 1.5 mL
Eppendorf tube (see Notes 7, 8, and 11): For each reaction add
39.5 μL Affymetrix nuclease-free water, 10 μL Taq buffer
(10), 20 μL, G-C melt (5 M), 14 μL dNTP, 4.5 μL PCR
primer, 2 μL Taq polymerase. Scale quantities according to the
number of reactions to be processed. Place the master mix on
the cooling chamber between adding reagents.
7. Vortex the master mix at 75% speed power three times, 1 s each
time.
8. Using a multi-channel P100 pipette, add 90 μL of the PCR
Master mix to each well containing the diluted genomic DNA
to obtain a total volume of 100 μL (see Note 10).
9. Seal the plate tightly with microseal film.
10. Vortex the center of the plate at high speed for 3 s.
13. Load the plate onto the thermal cycler and run the Cyoscan
PCR program as follows:
l Thirty cycles: 94 C for 30 s, 60 C for 45 s, 68 C for 15 s.
l Once cycle: 68 C for 7 min.
l 4 C hold (see Note 9).
14. When the program is finished, remove the plate and perform a
short spin.
15. If proceeding immediately to the purification step, put the
plate in a cooling chamber on ice. Otherwise, store the samples
at 20 C.
3.5 Checking the This step serves as the first quality control (QC) checkpoint. Gel
PCR Reaction by should be run in the post PCR area. Never bring in amplified PCR
Running a Gel products into the Pre-PCR Clean room.
1. Label a fresh set of strip tubes.
2. Aliquot 0.6 μL of 6 Gel Loading Dye and 5 μL water to each
well in the strip to be used.
3. Load 5 μL DNA marker to the first lane of the gel.
Fig. 1 Example of PCR products run on 2% agarose gel at 120 V for 10 min
4. Using a multichannel pipette transfer 3 μL of NspI PCR prod-

uct from each well in one column only (no need to run all 4
duplicates) to the corresponding wells of the gel plate.
5. Load 8 μL from each well of the gel plate onto a 2% agarose gel.
Load all 4 wells of the negative control.
6. Run the gel program: “E-Gel EX 1-2%” for 10 min.
7. Verify that the majority of the PCR product is between ~150 bp
to 200 bp (compare to Fig. 1).
8. The lanes with the negative control should be empty as shown
in Fig. 1 (see Note 12).
3.6 PCR Product This assay should be performed in post PCR area.
Purification
1. If frozen, thaw the PCR products in a plate holder on the
bench top to room temperature.
3. Add 45 mL of absolute Ethanol to the Purification wash buffer
prior to use. Cap the bottle tightly and shake. Enter the prepara-
tion date on the bottle label and put a check mark in the check box.
4. Using a P200 single channel pipet, transfer all 4 aliquots of
each sample to a 2.0 mL round bottom tube: Mark each
2.0 mL microcentrifuge tube with the appropriate sample
number (such as A, B, C, D) after the sample was transferred.
5. Cut the seal of only one row at a time so only one sample will be
transferred at a time. Do not pool the negative control. Be very
careful to avoid cross-contamination while pooling the PCR
products.
6. When finished, examine the PCR plate and ensure that the total
volume in each well has been transferred and pooled.
7. Thoroughly mix the magnetic bead stock by vigorously shaking
the bottle. Examine the bottom and the sides of the bottle and
ensure that the solution appears homogenous.
8. Aliquot 720 mL of magnetic beads to each pooled sample.
Securely cap each tube and mix well by inverting 10 times.
9. Incubate at room temperature for 10 min. During incubation,
the DNA binds to the magnetic beads.
10. Load the tubes—cap hinge facing out—onto the microcentri-
fuge and spin for 3 min at maximum speed.
11. Place the tubes on the magnetic stand. Leaving the tubes in the
rack, pipet off the supernatant without disturbing the bead
pellet and discard.
12. Using a P1000 pipet, add 1 mL of washing buffer to each tube
(be sure to add 45 mL of absolute ethanol to the purification
wash buffer prior to use).
13. Cap the tubes and load them into the foam tube adaptor. Fully
insert tubes into the foam to ensure they are secure. Space
tubes adequately to balance. Vortex at maximum settings for
2 min.
14. Centrifuge the tubes for 3 min at maximum speed (hinges
facing out).
15. Place the tubes on the magnetic stand. Leaving the tubes in the
rack, pipet off the supernatant without disturbing the bead
pellet and discard.
16. Spin the tubes for 30 s at maximum speed (hinges facing out).
17. Place the tubes back on the magnetic stand. Using a P20 pipet,
remove remaining drops of purification wash buffer from the
bottom of each tube. Transfer the tubes to a different rack and
allow the remaining purification wash buffer to evaporate by
leaving the tubes uncapped at room temperature for10 min.
18. Using a P100 pipet, add 52 μL of Elution buffer to each tube.
19. Cap the tubes and load them into the foam tube adaptor.
Vortex at maximum power for 10 min. Vortexing will resus-
pend the magnetic beads. Examine each tube to ensure that the
beads are resuspended in homogeneous slurry. If needed you
can flick the pellet to ensure full resuspension.
20. Centrifuge the tubes for 3 min at maximum speed (hinges
facing out).
21. Place the tubes on the magnetic stand for 10 min. The mag-
netic beads are pulled to the side of the tube. Check that all of
the beads have been pulled to the side in each tube. If all of the
beads have not been pulled to the side of the tubes, leave the
tubes on the stand an additional 3 min.
22. Transfer 47 μL of eluted sample to the appropriate well on a
fresh 96-well plate. Open one tube at each time of transfer, and
make sure the order of the samples matches the one in the
plate. Presence of some brown residue on pipet tips is fine.
Tightly seal the plate.
23. Vortex the plate at high speed in all corners and in the center,
and spin down at 650 g for 1 min.
3.7 Quantitation of This assay should be performed in the post PCR area.
Purified PCR Products
1. Label 500 μL tubes as A, B, C, D according to your plate
on a Nanodrop layout.
2. Using a P20 pipet, aliquot 18 μL of water to the corresponding
500 μL tube. Place the dilution tube in a separate rack.
3. Using a P2 pipet, transfer 2 μL of each purified sample to the
corresponding tube. Make sure to open only the tube of the
sample that is being diluted. Dilute one sample at a time.
4. Vortex and spin the diluted samples. The result is a tenfold
dilution.
5. Blank the NanoDrop with water.
6. Take 1.5 μL of the diluted sample and measure the OD of each
sample at 260 and 280 nm.
7. The average purification yield for seven or more samples should
be 3 μg/μL (see Notes 13–15).
8. PCR products can be stored at 20 C if not proceeding to the
fragmentation assay.
3.8 Fragmentation This assay should be performed in the post PCR area on ice.
1. Thaw the Fragmentation Buffer (10) on ice.
2. Vortex the buffer three times, 1 s each time. Pulse spin for 3 s
and place in the cooling chamber.
3. Preheat the thermal cycler.
4. The Fragmentation Reagent must be diluted to 0.1 U/μL in
the master mix. Read the Fragmentation Reagent tube label
and record the concentration.
5. Prepare fragmentation mastermix in an eppendorf tube on ice,
as per Table 1 (see Note 16).
6. Vortex the master mix at high speed for 5 s. Pulse spin for 3 s
and immediately place on ice.
7. Add 10 μL of the fragmentation mastermix to each sample. The
total volume of each sample should be 55 μL.
Table 1
Mastermix volumes for Fragmentation
Reagent Volume (μL)

Chilled nuclease free water 271.2
Fragmenation buffer 343.8
Fragmentation reagent 10
8. Cover the plate with microseal and pulse vortex five times 1 s
each.
9. Spin down at 650 g for 1 min in a prechilled centrifuge.
10. Place in the preheated thermal cycler (at 37 C) and run the
Fragmentation program as follows: 37 C for 350 , 95 C for
150 , 4 C hold.
11. Take the plate out and spin briefly.
3.9 Check the This step serves as the second QC checkpoint.

Fragmentation
1. Add 28 μL of affymetrix nuclease-free water, to the appropriate
Reaction by Running a
wells in the strip.
Gel
2. Add 4.0 μL of each sample to the wells containing the water.
3. Take an 8 μL aliquot out and add to a strip tube.
4. Add 0.6 μL gel loading dye to each sample.
5. Load 8 μL of each sample onto the gel.
6. Dilute the DNA marker as follows: 1 μL of marker with 9 μL of
water.
7. Load 10 μL of DNA marker to the first lane.
8. Run the gel program: “E-Gel Ex 4%” for 15 min.
9. Inspect the gel and compare it against the Fig. 2. Average
fragment size should be between 25 and 125 bp (see Note 17).
3.10 Labeling 1. Thaw the buffer and labeling reagent on ice.

2. Vortex each reagent three times on high speed, 1 s each time.
Pulse spin for 3 s, then place on cooling chamber on ice.
4. Make a master mix by adding the following to a 1.5 mL
Eppendorf tube (see Notes 7 and 8): For each sample add
14 μL 5 TdT buffer, 2 μL DNA Labeling reagent (30 mM),
3.5 μL TdT (30 U/μL). Scale quantities according to the
number of samples to be processed. Place the master mix on
the cooling chamber between adding reagents.
Fig. 2 Typical example of fragmented PCR products
Pulse spin for 3 s.
6. Using a P20 pipet, add 19.5 μL of the master mix to each
fragmented sample. Pipet up and down to ensure that all of the
mix is added to the samples.
7. Tightly seal the plate. Vortex at high speed for 3 s; then spin the
plate at 650 g for 1 min.
8. Place on the preheated thermal cycler block and run the Label-
ing program as follows: 37 C for 4 h, 95 C for 15 min, 4 C
hold (see Note 9).
9. When the Labeling program is finished, remove the plate from
the thermal cycler and spin down for 30 s.
10. If not proceeding with the hybridization assay, freeze the
labeled samples at 20 C.
3.11 Preparation of 1. Unwrap the arrays and place on the bench top, septa-side up.
Arrays for 2. Mark the front of each array with the sample number (A, B, C).
Hybridization
3. Allow the arrays to warm to room temperature on the bench
top 10–15 min.
4. Log in the arrays in AGCC. The sample information is stored in
a Sample file with an ARR extension. The arrays used in analysis
and the data files produced by analysis are linked to this sample
file.
Fig. 3 Arrays prepared for sample loading
5. Insert a 200 μL pipet tip into the upper right septum of each
array (see Fig. 3).
7. Preheat the hybridization oven to 50 C at least 1 h before
hybridization.
8. Make a master mix by adding the following to a tube (15 mL
falcon tube or 1.5 mL/2 mL amber tube. The choice of the
tube depends on the number of samples to be hybridized): For
each sample add 165 μL Hyb Buffer part 1, 15 μL Hyb Buffer
part 2, 7 μL Hyb Buffer part 3, 1 μL Hyb Buffer part 4, 2 μL
Oligo Control Reagent. Scale quantities according to the num-
ber of samples to be processed.
Pulse spin for 3 s and place in the cooling chamber.
10. Using a single channel P200 pipette, add 190 μL of the Master
mix to each well containing the labeled DNA, pipette up and
down to mix. The total volume in each well is now 260 μL.
11. Seal the plate tightly with microseal film. Vortex and spin the
plate at 650 g for 1 min.
13. Load the plate onto the thermal cycler and run the Hybridiza-
tion program as follows: 95 C for 10 min, 49 C hold.
14. Allow the samples to incubate at 49 C for at least 1 min before
loading.
15. Open the thermocycler lid, and load 200 μL of each sample in
the bottom left septa of the corresponding array. Make sure
that another technician is watching so that all the samples are
hybridized to the right labeled array. It is critical that the
Fig. 4 Covering the array septa with Tough-Spots
samples remain on the thermal cycler at 49 C after denatur-

ation and while being loaded onto arrays.
16. Take out the tip, and apply Tough-Spot on both septae of the
array (Fig. 4).
17. Load the arrays into hybridization oven (50 C and 60 rpm) for
16–18 h. Ensure that the oven is balanced as the trays with the
arrays are loaded.
3.12 Priming the Priming ensures the lines of the fluidics station are filled with the
Fluidics Station for appropriate buffers and the fluidics station is ready to run fluidics
Washing and Staining station protocols. Priming should be done:
l When the fluidics station is first started.
l When wash solutions are changed.
l Before washing, if a shutdown has been performed.
l If the LCD window instructs the user to prime.
1. Turn on the Fluidics station using the switch on the lower left
side of the machine.
2. Pour wash solutions A and B into the appropriate bottles, and
fill the water bottle with DI water.
3. Place all the bottles including the waste in the right place on the
side of the machine, and insert the corresponding tubing.
4. Turn on the computer.
5. Click on Affy Launcher icon, and hit Fluidics from the menu
bar.
6. In the experiment name field choose “no probe array.”
7. In the protocol field choose “Prime_450.”
8. Check “all modules” when using all four modules, otherwise

choose the Prime_450 protocol for the respective modules.
9. Click run for each module to begin priming, and follow the
LCD instructions (see Note 18).
3.13 Washing and The staining protocol is a three-stage process: (1) A Streptavidin
Staining the Arrays Phycoerythin (SAPE) stain, (2) An antibody amplification step, and
(3) A final stain with SAPE. Once stained, each array is filled with
Array Holding Buffer prior to scanning.
1. After 16–18 h of hybridization, remove the arrays from the
oven.
2. Remove the Tough-Spot from both septae of the array (see
Note 19).
3. Select your sample name on the computer. The Probe Array
Type and the corresponding protocol appear automatically
(i.e., “CytoScanHD_Array_450” protocol for Cytoscan
arrays).
4. Start the protocol and follow the instructions in the LCD on
the fluidics station.
5. Insert an array into the designated module of the fluidics
station while the cartridge lever is in the Down or Eject
position.
6. When finished, verify that the cartridge lever is returned to the
Up or Engaged position.
7. Remove any vials remaining in the positions of the fluidics
station module(s) being used.
8. When prompted to “Load Vials 1-2-3,” place the three vials
into positions 1, 2, and 3 on the fluidics station.
l Place one amber tube containing 500 μL stain buffer 1 in
position 1.
l Place one clear tube containing 500 μL stain buffer 2 in
position 2.
l Place one amber tube containing 800 μL Array Holding
Buffer in position 3.
l Press down on the needle lever to snap needles into position
and to start the run.
9. Once these steps are complete, the fluidics protocol begins.
The Fluidics Station dialog box at the workstation terminal and
the LCD window displays the status of the washing and stain-
ing steps (see Notes 20 and 21).
10. When staining is finished, Remove the arrays from the fluidics
station by first pressing down the cartridge lever to the eject
position.
11. Check the array window for large bubbles or air pockets. If
large bubbles are present, insert the array back into the module
and follow the instructions on the LCD panel.
12. Remove all 3 microcentrifuge vials containing stain and buffer
and replace with three empty vials as prompted.
13. Check the array window for dust or glue spots and wipe it off
with kimwipes wet with DI water or non-abrasive towel.
14. If the array has no large bubbles, it is ready for scanning. Pull
up on the cartridge lever to engage wash block and proceed to
scanning.
15. If the arrays cannot be scanned promptly, store the arrays at
4 C in the dark until ready for scanning. Scan must be per-
formed within 24 h.
16. When finished washing and staining, insert all wash lines
(except the waste one) to the DI bottle.
17. In the fluidics station dialog box (on the computer), select “no
probe array.”
18. In the protocol field choose “Shutdown_450.”
19. Check “all modules” when using all four modules, otherwise
choose the “Shutdown_450” protocol for the respective
modules.
20. Hit run to begin shutdown.
21. Shutdown takes about 10–15 min. After the protocol is over,
shut down the machine with the switch located on the lower
left side of the machine (see Note 22).
3.14 Scanning the 1. Turn on the scanner at least 10 min before use.
Arrays 2. If the arrays were stored at 4 C, allow them to warm to room
temperature before scanning.
3. If necessary, clean the glass surface of the array with a
non-abrasive towel or tissue before scanning. Do not use alco-
hol to clean the glass.
4. On the back of the array cartridge, clean excess fluid from
around the septa.
5. Carefully cover both septa with Tough Spots. Press to ensure
the spots remain flat.
6. Open the scanner lid and place the arrays in the carousel start-
ing with position # 1.
7. Close the lid.
8. Click on Affymetrix Launcher icon on the computer, and hit
Scanner control from the menu bar.
9. Hit “Start” from the scanner menu bar.
10. If you re-scan some of the chips, check the “allow rescan” box
and then hit ok. Otherwise, uncheck it and just hit ok.
11. If you need to add additional arrays to the scanner in the
middle of a run, hit “add chips”-> “add arrays after scan.”
Once scanning of the current array is completed, the lid of
the scanner will become “unlocked” and more arrays can be
added. After you close the lid hit” resume.”
12. In case there is an error during scan and the array wasn’t
scanned (usually “fousuing error”) cleaning again the array’s
glass might help (see Note 23).
13. When scanning, the green light is flashing. The light is stable
when it’s switching between arrays.
14. After scanning is done (green light is stable) take the arrays out
of the carousel, and turn off the scanner.
4 Notes
1. Some of the Affymetrix reagents can be substituted with the

following:
l NspI enzyme and NspI 10 buffer (New England Biolabs,
Ipswich, MA, USA).
l T4 DNA Ligase enzyme and buffer (New England Biolabs,
Ipswich, MA, USA).
l Agencourt AMPure XP magnetic beads (Beckman Coulter,
Brea, CA, USA).
l Qiagen elution (EB) buffer (Qiagen, Germantown, MD,
USA).
l 75% Ethanol solution can be used instead of the Affymetrix
wash buffer in the purification step.
2. Perform assays in designated areas (DNA extraction, Pre- PCR
and Post- PCR areas).
3. Never bring amplified products into the Pre-PCR Clean Area.
4. Keep dedicated equipment in each room or area used for this
protocol. To avoid contamination, do not move pipets between
the Pre-PCR Clean Area and the Post-PCR Area.
5. It is essential to set up the digestion, ligation, and PCR reac-
tions in the Pre-PCR clean room. This helps prevent
contamination.
6. Prenatal samples, particularly, uncultured amniotic fluid, usu-
ally do not yield much DNA. We found that for these samples
we can use a lower concentration than what is recommended by
the manufacturer. We have used DNA concentrations as low as
2 ng/μL and the data was still good.
7. Preparing master mixes with a 15% excess ensures consistency

in reagent preparation by minimizing pipetting errors and
reducing handling time of temperature sensitive reagents. The
success of this assay depends on the accurate pipetting and
subsequent thorough mixing of small volumes of reagents.
8. Before making the mastermix, remove the enzyme from the
freezer and immediately place in a cooler. Spin the enzyme for
3 s before adding to mastermix and then immediately add the
enzyme to the master mix. Place the remaining enzyme back in
the cooler.
9. If necessary, sample may be left in the PCR machine at 4 C
overnight.
10. When processing more than four samples, you should use a
reservoir to aliquot the water and later the PCR mastermix to
the samples, using a multichannel pipette.
11. 1 sample in this table represents one well of this sample. Since
each sample is being multiplied by 4, the calculation should
be 4 plus the 15% excess.
12. In case where all four lanes of the negative show some DNA
contamination, it is better to start the assay from the digestion
step using new vials of reagents. However, if only one lane of
the four shows contamination, it is most likely due to a splash
during vortexing and it does not necessarily mean that there is
an actual contamination.
13. Remember that each sample is diluted 10, so the measure-
ments should be multiplied by 10 in order to get the right yield.
14. If the sample yield is between 2.5 and 3 μg, it is still worth
continuing with the sample to see if it passes the second QC
point (which is the fragmentation gel electrophoresis). In situa-
tions where the sample is lower than 2.5 μg, it is better to
re-PCR the sample.
15. If the sample fails any of the QC checks after PCR, before
repeating the entire assay from scratch, you could try re-PCR
the sample using the remaining diluted ligated DNA and then
recheck the QC metrics.
16. Before making the mastermix, remove the fragmentation
enzyme from the freezer and vortex it vigorously for 10 s.
Immediately place in a cooler. Spin the enzyme for 3 s before
adding to mastermix and then immediately add the enzyme to
the master mix. Place the remaining enzyme back in the cooler.
17. It is very important to get the right sizing after fragmentation.
If the samples show a “tail” as seen in the first three samples in
the image, it is ok to proceed as long as most of the DNA is
between 25 and 125 bp. Samples that look too fragmented or
under fragmented should not be taken to the next step and
should be re-amplified by PCR. Also samples that look faint
compared to the other samples on the gel should not be taken

to the next step and should be re-amplified by PCR.
18. The priming takes about 10–15 min. In the meantime, prepare
the staining solutions.
19. Forgetting to do so might not interfere with the washing
process, but can damage the needle of the fluidics that needs
to go through the septae.
20. The washing and staining takes about 1.5 h. Ten minutes
before it ends, turn on the scanner for it to warm up (orange
light is on). Once the scanner is ready the green light will be on.
21. It is very important to check the computer every once in a
while during the wash for an error message. When there is fluid
missing, an error message pops up. In this case, check the level
of wash A and B in the bottle and make sure the tube is in the
fluid. Make sure to hit “resume” so the wash will continue from
where it stopped.
22. A cleaning protocol using sodium hypochlorite bleach should
be performed on the Fluidics Stations biweekly or as needed.
This is designed to eliminate any residual SAPE- antibody
complex that may be present in the fluidics station tubing and
needles. This protocol runs a bleach solution through the
system followed by a rinse cycle with DI water.
23. Sometimes there are glue stains on the array glass. Simply use
wet kimwipes with DI water to try and remove the stain.
Acknowledgments
We thank the technical staff at Affymetrix, who developed and

standardized these protocols for use in a clinical laboratory. The
members of clinical chromosomal array laboratory at Columbia
University Medical Center are also acknowledged for setting up
and fine-tuning these protocols.
References
1. American College of O., Gynecologists Com- 2. Wapner RJ, Martin CL, Levy B et al (2012)
mittee on G (2013) Committee Opinion No. Chromosomal microarray versus karyotyping
581: the use of chromosomal microarray analysis for prenatal diagnosis. N Engl J Med 367
in prenatal diagnosis. Obstet Gynecol 122 (23):2175–2184. https://doi.org/10.1056/
(6):1374–1377. https://doi.org/10.1097/01. NEJMoa1203382
AOG.0000438962.16108.d1
Chapter 14
Rapid Detection of Fetal Mendelian Disorders: Thalassemia

and Sickle Cell Syndromes
Joanne Traeger-Synodinos, Christina Vrettou, and Emmanuel Kanavakis
Abstract
The inherited disorders of hemoglobin synthesis constitute the most common monogenic diseases world-
wide. The clinical severity of β-thalassemia major and the sickle cell syndromes targets them as priority
genetic diseases for prevention programs, which incorporates population screening to identify heterozy-
gotes, with the option of prenatal diagnosis for carrier couples. Rapid genotype characterization is funda-
mental in the diagnostic laboratory, especially when offering prenatal diagnosis. The application of real-time
PCR provides a means for rapid and potentially high throughput assays, without compromising accuracy. It
has several advantages over end-point PCR analysis, including the elimination of post-PCR processing steps
and a wide dynamic range of detection with a high degree of sensitivity. Although there are over 200 muta-
tions associated with the β-thalassemia and sickle cell syndromes, the relatively small size of the β-, HBB
gene (less than 2000 base-pairs) and the close proximity of most mutations facilitates the design of a
minimal number of real-time PCR assays using the LightCycler™ system, which are capable of detecting
the majority of most common β-gene mutations world-wide. These assays are highly appropriate for rapid
genotyping of parental and fetal DNA samples with respect to β-thalassemia and sickle cell syndromes.
Key words Prenatal diagnosis, β-thalassemia and sickle cell syndromes, Real-time PCR
1 Introduction
Prenatal diagnosis (PND) aims to provide an accurate, rapid result

as early in pregnancy as possible. A prerequisite involves obtaining
fetal material promptly and safely. In addition, for monogenic dis-
eases the parental mutation(s) have to be characterized prior to
analysis of the fetal sample. The majority of methods currently used
for genotyping parental samples and performing prenatal diagnosis
are based on the polymerase chain reaction (PCR) (e.g., allele-
specific oligonucleotide (ASO) hybridization analysis of PCR
amplicons, amplification refractory mutation system (ARMS)
PCR, restriction endonuclease analysis of PCR amplicons, and
direct DNA sequencing). Most of these techniques are relatively
simple, fairly quick and inexpensive (with the possible exception of
207
208 Joanne Traeger-Synodinos et al.
sequencing) but may not provide a rapid final genotype nor facili-
tate the processing of many samples simultaneously (“high
through-put”). This can be a disadvantage with respect to identify-
ing the parental mutations if the disease-associated gene has a wide
spectrum of potential mutations. The application of real-time PCR
offers a means for rapid and potentially high throughput assays,
without compromising accuracy, making it an ideal approach for
genotyping parental and fetal DNA samples in the context of
prenatal diagnosis.
1.1 An Introduction Real-time PCR integrates microvolume rapid-cycle PCR with fluo-
to Real-Time PCR rometry, allowing real-time fluorescent monitoring of the amplifi-
for Genotyping cation reaction for quantitative PCR and/or qualitative
Applications characterization of PCR products. The latter application provides
a means for rapid genotyping, precluding any post-PCR sample
manipulation. Several real-time PCR instruments are available on
the market, and in addition there are a number of detection che-
mistries, some of which can be used on any instrument and others
which are instrument-specific. Real-time PCR machines can be
classified as either “flexible” or high throughput. The “flexible”
instruments, usually faster and with a wider choice of running
parameters, are more suitable for smaller batches of samples,
whereas the high throughput instruments are more appropriate
for running large batches of samples requiring a smaller repertoire
of assays. Flexible instruments are probably more appropriate for
PND applications since the processing of samples in small batches
precludes potential errors, e.g., through the occurrence of a tube-
switch. However, high throughput instruments may be advanta-
geous when screening parental samples to characterize mutations
before performing the prenatal diagnosis.
There are a number of detection chemistries suitable for geno-
typing but those more commonly described for use in genotyping
monogenic diseases include hybridization probes, Taqman probes
and molecular beacons [1].
The protocol that we describe in this chapter employs the use of
a flexible instrument (the LightCycler™ 1.0, LightCycler™ 1.5, or
LightCycler™ 480 II, Roche Molecular Biochemicals) with “hybri-
dization probes.”
Hybridization probes [2] involve a dual probe system. The two
fluorescently labeled probes hybridize to adjacent sequences within
the amplified target DNA, one of which covers the region expected
to contain the mutation(s). Close proximity of annealed probes
facilitates fluorescence resonance energy transfer (FRET) between
them, and a fluorescent signal is only generated when both of the
probes are hybridized to the target amplicon (Fig. 1a). The two
probes of the pair are designed to have different melting tempera-
tures (Tm), whereby the probe with the lower Tm lies over the
mutation site. Monitoring the emitted fluorescent signal under
conditions of increasing temperature will detect a loss of
Rapid Detection of Fetal Mendelian Disorders: Thalassemia and Sickle Cell. . . 209
Fig. 1 (a) Principle of hybridization probes for allele discrimination. (1) One probe
is labeled with a donor fluorophor (D) and the other with an acceptor fluorophore
(A). The probe with the acceptor fluorophore is complimentary to the region
including the expected mutation(s). (2) Close proximity of annealed probes
facilitates fluorescence resonance energy transfer (FRET) and the emission of
a fluorescent signal. (b) Plots of fluorescence versus temperature from melting
curve analysis using hybridization probes. Top plot shows the raw melting peak
data with fluorescence (F) versus temperature (T); Bottom plot shows the melting
peaks displayed when the computer software automatically converts and dis-
plays the first negative derivative of fluorescence to temperature vs. temperature
(-dF/dT vs. T). The latter facilitates easy discrimination between wild type and
mutant alleles
LC1F LC1R
LC2F LC2R
Set III
III.1A IIID III.2A LC3F LC3F
5’UTR 3’UTR
EXON 1 Intron 1 EXON 2 Intron 2 EXON 3
I.1A ID I.2A IID IIA IVD IVA VD VA
Set I SetII Set IV Set V
Lightcycler primers, LC1 (for probe sets I, II, III)
Lightcycler primers, LC2 (for probe sets II, III, IV)
Lightcycler primers, LC3 (for probe set V)
Allele-specific acceptor probes
Donor probes
Fig. 2 The position of the β-globin gene primers and LightCycler™ hybridization probe sets appropriate for
prenatal diagnosis and preimplantation genetic diagnosis protocols. Three PCR primer sets can be used (either
LC1 or LC2, which can both be multiplexed with LC3 if required). F ¼ forward primer; R ¼ reverse primer.
Probe sets I-V are used according to the mutations under investigation (see Table 1)
fluorescence (F) as the lower Tm probe melts off the template. A

single base mismatch under this probe results in a Tm shift of
5–10 C, allowing easy distinction between wild type and mutant
alleles (Fig. 1b). The ability to detect any nucleotide mismatches
under the low Tm probe (mutation detection probe) and the use of
different coloured probes (according to the properties of the real-
time PCR instrument in use) can allow more than one mutation to
be assayed in a single PCR reaction (Fig. 2).
1.2 Molecular Basis The β-globin gene (HBB: OMIM 141900) is a relatively small gene
of the (<2000 bp) located in the short arm of chromosome 11. (11p15.5,
β-Hemoglobinopathies GenBank: NM_000518). Although more than 200 causative muta-
and Design of Real- tions have been identified for β-thalassemia syndromes (HbVar
Time PCR Mutation http://globin.bx.psu.edu/hbvar/menu.html), the spectrum of
Detection Assays mutations and their frequency in most populations usually includes
a limited number of common mutations (e.g., up to approximately
six) plus a slightly larger number of rare mutations (e.g., around
10). In most populations worldwide the majority of the most
common mutations tend to cluster within small distances within

HBB. The probes sets described in this chapter were designed based
on those mutations most commonly found in the Mediterranean
populations, although they are also suitable for detecting several of
the common mutations found in other world populations (e.g.,
Asian Indian, SE Asian, and Chinese) [3, 4]. Overall they facilitate
mutation detection with the use of a small number of probe sets
within a minimal number of amplicons.
All the allele-specific (mutation-detection) probes described in
this chapter were designed to be complementary to the wild-type
sequence of HBB, rather than complementary to each specific
mutation. This potentially allows the distinction of normal alleles
from any allele with a nucleotide variation under the length of the
probe, minimizing the number of assays that need to be performed,
and potentially reducing costs and time required to identify the
mutations in parental samples, and subsequently perform the PND.
Although most mutations have a distinct melting profile and can be
implicated by comparison with controls, the definitive characteri-
zation of each mutation should be achieved by a second method
such as an ARMS-PCR assay [5, 6]. This is recommended for best-
practice, and any laboratory performing DNA diagnostics and
PND should have more than one available mutation detection
method [7, 8].
The LightCycler™ PCR primers (sets LC1, LC2 and LC3)
were designed with the aid of computer software (Amplify version
2.0, Bill Engels, 1992–1995). LC1 and LC2 amplify two over-
lapping regions of HBB surrounding the majority of the most
common β-thalassemia mutations in all world populations, along
with the HbS mutation, and when multiplexing mutation detection
they should be selected according to the mutations under investi-
gation (Table 1 and Fig. 2). LC3 amplifies a 30 region of HBB which
contains some rare β-thalassemia mutations and if necessary can be
multiplexed with either set LC1 or LC2.
Design of LightCycler™ mutation detection probe sets took
into account secondary structure properties (e.g., hairpin-loops)
and potential primer-dimer formation between the probes them-
selves and the PCR primers, evaluated by computation prior to
synthesis (TIB MOLBIOL, Berlin, Germany). In addition, design
of mutation detection probes avoided the regions of HBB known to
contain common polymorphic point mutations such as codon
2 (CAC > CAT). The LightCyclerTM systems 1.0 and 1.5 can
detect 2 fluorescent labels (LightCyclerTM Red 640 [LC Red 640]
and LightCyclerTM Red 705 [LC Red 705]), as well as SYBR®
green. The choice of fluorescent label used, for each probe will
depend upon the relative frequency of mutations in the population
under study and the potential requirement for multiplexed assays
when more than one mutation is investigated within a single sam-
ple. For example in the Greek population, HBB:c.93-21G>A
212
Table 1
Lightcycler mutation detection probe sets for the most common β-thalassemias mutations worldwide (and HbS mutation)
Probe Beta-gene mutation

set Acceptor probe name and sequence Donor probe name and sequence (s) detected HGVS*** nomenclature
*
Set I I.1A: ID: CAP +20 (C>T) HBB:c.31C>T
50 -ttc tga cac aac tgt gtt cac tag ca-30 LC FITC 50 -cct caa aca gac acc atg gtg cac c-30 CAP+22 (G>A) HBB:c.29G>A
Joanne Traeger-Synodinos et al.
Red** FITC
I.2A: HbS (Cd 6 A>T) HBB:c.20A>T
LC Red** 50 -gac tcc tga gga gaa gtc tgc-30 Cd5 (CT) HBB:c.17_18delCT
P Cd6 (A) HBB:c.20delA
Cd8 (AA) HBB:c.25_26delAA
Cd 8/9 (+G) HBB:c.27_28insG
Set II IIA: IID: IVSI-1 (G>A) HBB:c.92+1G>A
LC Red** 50 -tgt aac ctt gat acc aac ctg ccc 50 -tgc cca gtt tct att ggt ctc ctt aaa cct gtc-30 IVSI-1 (G>T) HBB:c.92+1G>T
a-30 P FITC IVSI-2 (T>G) HBB:c.92+2T>G
IVSI-2 (T>C) HBB:c.92+2T>C
IVSI-2 (T>A) HBB:c.92+2T>A
IVSI-5 (G>A) HBB:c.92+5G>A
IVSI-5 (G>C) HBB:c.92+5G>C
IVSI-5 (G>T) HBB:c.92+5G>T
IVSI-6 (T>C) HBB:c.92+6T>C
Set III III.1A: IIID: IVSI-110 (G>A) HBB:c.9321G>A
50 -tct gcc tat tgg tct att ttc cc-30 LC Red** FITC 50 -ccc tta ggc tgc tgg tgg tc-30 FITC IVSI-116 (T>G) HBB:c.9315T>G
III.2A: Cd39 (C>T) HBB:c.118C>T
LC Red** 50 -acc ctt gga ccc aga ggt tct t-30 Cd37 (TGG>TGA) HBB:c.114G>A
P Cd41/42 (delTTCT) HBB:
c.126_129delCTTT
Set IV IVA: VID: IVSII-1(G>A) HBB:c.315+1G>A
LC Red** 50 -tct cag gat cca cgt gca gct 50 -gtc cca tag act cac cct gaa g-30 FITC.
tg-30 P
SetV VA: VD: polyA signal mutation HBB:c.*110T>C
LC Red** 50 gct caa ggc cct ttc ata ata tcc cc 50 ttt ttc att agg cag aat cca ga-30 FITC. AATAAA>AACAAA HBB:c.*111A>G
AATAAA>AATGAA HBB:c.*112A>G
AATAAA>AATAGA HBB:c.*113A>G
AATAAA>AATAAG HBB:c.
AATAAA>AA- - AA [*109_*110delAT
AATAAA>A- - - - - or *110_*111delTA]
HBB:c.
*108_*112delAATAA
The β-globin gene specific PCR primers include (see Fig. 2):
For probes sets I, II, III: LC1F: 50 -GCT GTC ATC ACT TAG ACC TCA-30 ; LC1R 50 -CAC AGT GCA GCT CAC TCA G-30 ;
For probes sets II, III,IV: LC2F 50 -CAA CTG TGT TCA CTA GCA AC-30 ; LC2R 50 -AAA CGA TCC TGA GAC TTC CA-30 ;
For probe set V: LC3F 50 -ATT TCT GAG TCC AAG CTG GGC -30 ; LC3R 50 -AAA TGC ACT GAC CTC CCA-30
FITC: Fluorescein; P ¼ Phosphorylated
* ¼ Polymorphism linked with the IVSII-745 (C>G) mutation ** LC Red: The fluorescent label used for each probe will depend upon the relative frequency of mutations in the
population under study and the potential requirement of multiplexed assays. ***HGVS ¼ Human Genome Variation Society
Rapid Detection of Fetal Mendelian Disorders: Thalassemia and Sickle Cell. . .
213
(IVSI-110 G>A) is the most common mutation, so the probes for

most other mutations encountered in Greece are labeled with the
opposite fluorescent marker to that used for HBB:c.93-21G>A.
More specifically, two of the probe combinations (named set I and
set III) include 2 acceptor (mutation detection) probes with one
central donor probe, (Fig. 2), foreseeing the use of one or both
acceptor probes of the set according to the needs of any genotyping
assay. Each of the acceptor probes in sets I and III are labeled with
different acceptor fluorophores and the central donor probe,
designed to span the distance between the two acceptor probes, is
labeled with a fluorescein (F) molecule at both 50 and 30 ends. Sets
II and V were designed to screen for several neighboring mutations
with use of a single mutation detection (acceptor) probe. Set IV was
designed to detect a single mutation each and involved a donor
probe which was labeled with fluorescein only at the end adjacent to
the acceptor probe (Table 1, Fig. 2). In all sets the mutation-
screening (acceptor) probes were designed to have a lower Tm
relative to the donor probes, thereby ensuring that the fluorescent
signal generated during the melting curve is determined only by the
specificity of mutation-screening probe.
2 Materials
2.1 Equipment The method described was set up using a LightCycler™ system
instrument version 1.0 or 1.5 (Roche Diagnostics Corporation,
Indianapolis, IN, USA) (see Subheading 4.1 Notes 1 and 2).
1. Aluminum cooling block, which holds 32 LightCycler™ Cen-
trifuge Adapters (Roche Diagnostics Corporation,
Indianapolis, IN, USA), in which the real-time PCR reactions
are set up.
2. LightCycler™ glass capillary tubes (20 μl) (Roche Diagnostics
Corporation, Indianapolis, IN, USA), in which the real-time
PCR reactions are run in the LightCycler™ instrument.
3. A bench centrifuge with a well-depth of approximately 4.5 cm,
for centrifugation (maximum 3000 g) to pull the reaction
volume (20 μl) to the base of the glass capillary, prior to loading
in the LightCycler™ instrument.
2.2 Reagents 1. QIAMP DNA mini kit (Qiagen, Hilden, Germany) for extract-
ing DNA from chorionic villi samples or amniocytes.
2. Pair(s)of PCR primers selected according to mutations under
study (either LC1F/LC1R or LC2F/LC2R, if necessary with
LC3F/LC3R, as shown in Table 1 and Fig. 2).
3. Fluorescently labeled mutation detection probe sets, appropri-
ate for mutations under study (see Table 1 and Fig. 2).
4. LightCycler™ DNA Master Hybridization probes Kit (Roche

Diagnostics Corporation, Indianapolis, IN, USA), which also
includes MgCl2 (25 mM) and PCR-grade water.
2.3 Handling 1. All PCR primers to be used on the LightCycler™ are diluted as
and Storage of PCR stock solutions of 100 μM, divided into aliquots of convenient
Reagents volume (e.g., 25 μl) and stored at 20 C. For primer working
solutions the stock solutions are diluted to 10 μM and can be
subsequently stored at 4 C for up to 3 months.
2. The LightCycler™ hybridization probes are diluted to 3 μM
and stored in aliquots of relatively small volume (e.g., 20 μl) at
20 C. A thawed aliquot should not be refrozen, but can be
used up to 1 month when stored at 4 C.
3. The “Master Mix” from the LightCycler™ -DNA Master
Hybridization probes Kit should not be refrozen once thawed
but can be used for up to 1 month when stored at 4 C.
3 Methods
3.1 DNA Extraction The real-time PCR genotyping method described below assumes
from Chorionic Villi that assays are performed on good quality genomic DNA samples
Samples or (parental or fetal) with a concentration of 30–50 ng/μl. Extract
Amniocytes DNA using the QIAMP DNA mini kit according to the “Blood and
Body Fluid Spin Protocol” outlined in the manual supplied with
the kit.
3.2 Real-Time PCR 1. In an eppendorf tube make a premix for the amplification
Reaction Setup reactions for a total reaction volume of 20 μl (or 10 μl) per
for the Lightcyclertm sample. Each reaction should contain the ready-to-use reaction
1.0 and 1.5 (See mix provided by the manufacturer (LightCycler™ DNA Mas-
Subheading 4.2 Notes ter Hybridization Probes) plus MgCl2, a β-globin gene PCR
1 and 2) primer pair (i.e., LC1, LC2, LCR3) and LightCycler™ fluo-
rescent probe sets for the relevant mutations. A typical PCR
reaction for single color detection for one sample is shown in
Table 2.
2. When calculating the premix volume, make premix enough for
the number of samples being genotyped, a PCR premix blank
plus controls for the mutation(s) under investigation. The
controls should include a homozygous wild-type sample
(N/N), a sample heterozygous for the mutation (M/N) and
a sample homozygous for the mutation (M/M).
3. Place the appropriate number of LightCyclerTM glass capillary
tubes in the centrifuge adapters in an aluminum-cooling block.
4. Distribute accurately 18 μl (or 9 μl) of premix in all the
capillaries.
Table 2
A typical PCR reaction for single color detection for one sample
Stock Conca Final Conca b

μl/sample
H2O (PCR grade) 9.6 μl
MgCl2 25 mM 4 mM 2.4 μl
Beta globin forward primer
LC1(F) or LC2(F) or LC3(F) 10 μM 0.5 μM 1 μl
Beta globin reverse primer
LC1(R) or LC2(R) or LC3(R) 10 μM 0.5 μM 1 μl
LC Red allele-specific
probe (640 or 705) 3 μM 0.15 μM 1 μl
FITC donor 3 μM 0.15 μM 1 μl
Master Mix 2 μl
Premix volume 18 μl
DNA sample volume 2 μl
Total reaction volume 20 μlb
The volume of water is always adjusted to give final reaction volume of 20 μl/sample even when more than one primer or
probe set is included in the reaction. For example a PCR reaction with dual color detection using 2 allele-specific LC
probes (one labeled with Red 640 and the other with Red 705) and a common (central) doubly labeled FITC probe, or
even two sets of LC donor-acceptor probes (i.e., 4 probes)
a
Conc ¼ concentration
b
All volumes in the reaction may be halved without compromising the result
5. Add 2 μl (or 1 μl) genomic DNA (approximately 50 ng) per

sample and controls, and finally add the same volume of
double-distilled water to the PCR blank.
6. Once the PCR reactions have been set up in the capillaries at
4 C place the caps carefully on each capillary without pressing
down completely yet.
7. Remove the aluminum centrifuge adaptors containing the
capillaries from the cooling block and place in a bench centri-
fuge with wells deep enough to hold the aluminum centrifuge
adaptors (approximately 4.5 cm).
8. Spin at a maximum speed of 3000 g for 10 s to pull the
reaction volume to the base of the glass capillary.
9. Place each glass capillary carefully into the LightCycler™ car-
ousel by letting it simply “slip” in place. Then gently press the
cap completely in to the capillary and simultaneously the glass
capillary fully down into position in the LightCycler™
carousel.
10. Put the loaded carousel in to the LightCycler™ and initiate the
PCR cycles and melting curve protocols using the LightCycler
software version 3.5.1, according to the manufacturer’s
specifications.
3.3 Amplification 1. Preprogram the LightCycler™ software (according to the

and Melting Curve instrument model and the manufacturer’s guidelines) for the
Analysis following amplification steps: a first denaturation step of 30 s at
95 C followed by 40 cycles of 95 C for 3 s, 58 C for 5 s and
72 C for 20 s with a temperature ramp of 20 C/s. During the
PCR, emitted fluorescence can be measured at the end of the
annealing step of each amplification cycle to monitor progres-
sion of amplification.
2. Immediately after the amplification step, the LightCycler™ is
programmed to perform melting curve analysis to determine
the genotypes. This involves a momentary rise of temperature
to 95 C, cooling to 45 C for 2 min to achieve maximum
probe hybridization, and then heating to 85 C with a rate of
0.4 C/s during which time the melting curve is recorded.
3. Emitted fluorescence is measured continuously (by both chan-
nels F2 (640 nm) and F3 (705 nm) if necessary) to monitor the
dissociation of the fluorophore-labeled detection probes from
the complementary single-stranded DNA (F/T) (F: Fluores-
cence emitted, T: Temperature). The computer software auto-
matically converts and displays the first negative derivative of
fluorescence to temperature versus temperature (-dF/dT vs. T)
and the resulting melting peaks allow easy discrimination
between wild type and mutant alleles (Fig. 1b).
4 Notes
4.1 LightCycler 1. The assay reactions described have also been used on a Light-
CyclerTM 480 II instrument without any modifications.
2. The Real-time PCR reaction set up for the LightCyclerTM
480 II is done in 96-well plates, sealed with Lightcycler
480 Sealing Foil, which precludes the use of adaptors, capil-
laries and caps as described in Subheading 3.2.
4.2 Melting Curves 1. Trouble with melting curve analysis usually occurs when a
relatively high number of samples are included in a single run.
Under these circumstances wide melting peaks are observed
with only minor differences in the central Tm peak of the
melting curve for all genotypes, e.g., M/M, N/N, and M/N;
furthermore heterozygous DNAs will not give a double peak
following melting curve analysis but a single peak which lies
between that of the N/N and M/M controls/ samples.
2. The LightCyclerTM (system 1.0 and 1.5, software version 3.5)

is designed to run a maximum of 32 samples in a single run, but
our experience indicates that analysis of more than approxi-
mately 20 samples produces wide and “flattened” melting
curves. The problem is probably due to the way that the Light-
cyclerTM performs the melting curve analysis. During the
LightCyclerTM melting curve analysis the temperature
increases from 45 C to 90 C, which takes about 150 s when
using the “continuous acquisition” mode and a temperature
increment of 0.3 C/s. In “continuous acquisition” mode the
temperature increases continuously, and does not take in to
account that the measurement of fluorescence between one
capillary and the next takes a certain time. If a small number
of samples are analyzed, the fluorescence of each will be read
more often during the 150 s of melting curve data acquisition,
compared to a run with more samples. In the latter situation
there are too few measurement points to calculate a detailed
melting curve. There are three possible solutions to this
problem:
(a) For high sample numbers it is recommended that the
temperature increment used for the melting curve is
decreased, e.g., to 0.1 C/s when using “continuous
acquisition” mode, or to 0.4 C/s when using “stepwise
acquisition” mode.
(b) If the melting curve still fails to give a satisfactory result,
we have found that additional melting curves can be per-
formed using other temperature increments, although it
must be noted that the quality of the melting curves is
reduced each time an analysis is performed (and this is not
recommended by the manufacturer).
(c) In cases when the melting curve still fails to give satisfac-
tory result when analyzing more than about 20 samples,
pause the LightCycler™ program following the amplifica-
tion step and perform melting curve analyses in batches
(including the appropriate controls with each melting
curve analysis).
References
1. Wilhelm J, Pingoud A (2003) Real-time poly- of multiple beta-globin gene mutations by real-
merase chain reaction. Chembiochem 4 time PCR on the LightCycler: application to
(11):1120–1128. https://doi.org/10.1002/ carrier screening and prenatal diagnosis of thal-
cbic.200300662 assemia syndromes. Clin Chem 49(5):769–776
2. Lyon E (2001) Mutation detection using fluo- 4. Vrettou C, Traeger-Synodinos J, Tzetis M,
rescent hybridization probes and melting curve Palmer G, Sofocleous C, Kanavakis E (2004)
analysis. Expert Rev Mol Diagn 1(1):92–101. Real-time PCR for single-cell genotyping in
https://doi.org/10.1586/14737159.1.1.92 sickle cell and thalassemia syndromes as a rapid,
3. Vrettou C, Traeger-Synodinos J, Tzetis M, accurate, reliable, and widely applicable protocol
Malamis G, Kanavakis E (2003) Rapid screening for preimplantation genetic diagnosis. Hum
Mutat 23(5):513–521. https://doi.org/10. 7. Old J, Petrou M, Varnavides L, Layton M, Mod-

1002/humu.20022 ell B (2000) Accuracy of prenatal diagnosis for
5. Old JM, Varawalla NY, Weatherall DJ (1990) haemoglobin disorders in the UK: 25 years’
Rapid detection and prenatal diagnosis of beta- experience. Prenat Diagn 20(12):986–991
thalassaemia: studies in Indian and Cypriot 8. Traeger-Synodinos J, Harteveld CL, Old JM,
populations in the UK. Lancet 336 Petrou M, Galanello R, Giordano P,
(8719):834–837 Angastioniotis M, De la Salle B, Henderson S,
6. Kanavakis E, Traeger-Synodinos J, Vrettou C, May A, Ehbp m (2015) EMQN Best Practice
Maragoudaki E, Tzetis M, Kattamis C (1997) Guidelines for molecular and haematology
Prenatal diagnosis of the thalassaemia syndromes methods for carrier identification and prenatal
by rapid DNA analytical methods. Mol Hum diagnosis of the haemoglobinopathies. Eur J
Reprod 3(6):523–528 Hum Genet 23(4):426–437. https://doi.org/
10.1038/ejhg.2014.131
Chapter 15
Prenatal Diagnosis of Cystic Fibrosis

Anastasia M. Fedick, Jinglan Zhang, Lisa Edelmann, and Ruth Kornreich
Abstract
Cystic fibrosis (CF) is an inherited disease characterized by the accumulation of thick, sticky mucus which
damages epithelia in organs such as the lungs, pancreas, liver, intestines, and other parts of the body. The
most common symptoms are sinopulmonary disease and chronic gastrointestinal tract problems resulting
from decreased mucociliary clearance and inflammation. CF is the most common life-limiting autosomal
recessive disorder in people of northern European ancestry and it affects other populations with different
prevalence. CF can be diagnosed by many methods including testing for blood immunoreactive trypsin,
sweat chloride, transepithelial nasal potential difference, and molecular genetic testing.
Key words Cystic fibrosis, Recessive, Mutation, Chronic pulmonary disease, Congenital bilateral
absence or atrophy of the vas deferens
1 Introduction
Cystic fibrosis (CF) is a severe autosomal recessive disease with its

highest prevalence found in Caucasian populations at an estimated
one in 2000–3000 births. The major clinical symptoms of CF
include chronic pulmonary disease leading to progressive lung
failure, pancreatic exocrine insufficiency, and male infertility due
to congenital bilateral absence or atrophy of the vas deferens
(CBAVD) or obstructive azoospermia. Respiratory failure is
responsible for 80–95% of CF mortalities, which usually occur
around the age of 37. Symptoms of CF usually present in early
childhood, however there are rare instances of adult diagnoses.
Although multidisciplinary care for CF is available, its morbidity
and mortality often results from airway obstruction and pulmonary
defense system impairment associated with chronic infection and
neutrophil inflammation of the small and large airways. Persistent
infections, especially with Pseudomonas aeruginosa and Staphylo-
coccus aureus, cause chronic sputum production, and eventually
bronchiectasis and lung destruction.
221
222 Anastasia M. Fedick et al.
1.1 Molecular CF is a monogenic disorder caused by mutations in the CFTR

Genetics of Cystic gene. The CF disease gene, cystic fibrosis transmembrane conduc-
Fibrosis tance regulator (CFTR), was identified in 1989 [1, 2]. CFTR is a
transmembrane protein that consists of five domains: two
membrane-spanning segments each consisting of six regions
(TM1-TM12) that form the chloride channel pore, two
ATP-binding domains (NBF1 and NBF2) that control channel
activity by interacting with cytosolic nucleotides, and a regulatory
domain that also controls channel activity via phosphorylation
[3]. The CFTR gene is ~215 kb in size and located on chromosome
7q31.2 with 26 introns and 27 exons. Over 1980 mutations have
been reported to date in the Cystic Fibrosis Mutation Database
(www.genet.sickkids.on.ca/cftr/app) including missense, non-
sense, frameshift, splicing, and small and large insertions or dele-
tions, located in both the promoter region and throughout the
entire gene. Mutations in CF patients are usually inherited from
both of their carrier parents although de novo CFTR mutations
occur very rarely. Uniparental disomy (UPD) of chromosome
7 resulting in homozygosity of a CFTR mutation has also rarely
been causative of CF [4–6], as well as mosaicism for UPD7 [7].
CFTR mutations can be categorized into five different groups
based on the impact that they have on gene function. Class I
mutations cause defective protein production due to premature
stop codons, Class II mutations block protein processing due to
mislocalization, Class III mutations effect the nucleotide binding
domains causing errors in regulation and non-functional chloride
channels, Class IV mutations effect the membrane spanning
domains and impact conductance and/or gating properties, and
Class V mutations include those that reduce the synthesis of the
CFTR protein [3, 8]. While phenotype-genotype correlations have
not been established using the above categorization for most CF
phenotypes, there has been a correlation observed for most
category I, II, and III mutations with severe pancreatic-
insufficiency [3, 8].
The three base pair deletion of phenylalanine at position
508 (c.1521_1523delCTT, p.Phe508del) is the most common
CF mutation, accounting for approximately two-thirds of all cases
[9]. While CF is pan-ethnic, various ethnic populations have differ-
ent risks of being carriers for the disease [10]. CF is most common
in Ashkenazi Jews (carrier frequency of 1 in 25) and Caucasians
(carrier frequency of 1 in 26) and least common in Asians (carrier
frequency of 1 in 94). Certain mutations are also more prevalent in
specific ethnic populations due to founder effects, such as the
c.3846G>A, p.Trp1282X mutation in the Ashkenazi Jewish popu-
lation which accounts for around 60% of all mutations [11] and the
c.3302T>A, p.Met1101Lys mutation in the Hutterites of South
Dakota which accounts for around 69% of all mutations [12].
Prenatal Diagnosis of Cystic Fibrosis 223
When conducting molecular genetic tests, especially targeted

genotyping assays, false positive results can occur if there is a variant
at or near the same location of a known mutation. For example,
three variants: c.1516A>G, p.Ile506Val; c.1519A>G, p.Ile507Val;
and c.1523T>G, p.Phe508Cys have been known to cause errone-
ous results when testing for the c.1519_1521delATC, p.Ile507del
or c.1521_1523delCTT deletions. Depending on the method
used, it may be necessary to test for the additional variants when-
ever a c.1521_1523delCTT mutation is identified. It is particularly
evident when an individual who is a carrier for the deletion appears
to be homozygous because he or she is also a carrier for one of the
above non-pathogenic variants. In such instances where the patient
does not show any symptoms of CF, follow-up tests should be done
for those specific variants to ensure that the results are
accurate [13].
1.2 Genetic Testing The American College of Obstetricians and Gynecologists (ACOG)
for Cystic Fibrosis: and The American College of Medical Genetics (ACMG) both
Carrier and Diagnosis published guidelines for prenatal and preconception carrier screen-
ing for CF in 2001. Both committees originally recommended
conducting carrier screening for 25 common mutations that caused
classic CF and had mutant allele frequencies of 0.1% or greater in
the general United States population. The ACMG then updated
their recommendation in 2004 and removed two mutations:
c.948delT, p.Phe316LeufsX12 due to an observed frequency of
less than 0.1% and c.443T>C, p.Ile148Thr because it is not a
true pathogenic variant (it can occur in cis with the severe CF
mutation c.3067_3072delATAGTG, p.Ile1023_Val1024del),
reducing the recommended panel to 23 mutations. By screening
for the 23 mutations, it is estimated that the approximate carrier
risk after a negative test drops from 1:24 to 1:380 for Ashkenazi
Jews, 1:25 to 1:200 for non-Hispanic whites, 1:58 to 1:200 for
Hispanic whites, 1:61 to 1:170 for African Americans, and 1:94 to
1:180 for Asian Americans [14]. Genetic counseling is recom-
mended both pre- and post-testing so that patients understand
their remaining residual risk of having an affected child, especially
based on ethnic background and test detection rates [15].
Since the publication of the guidelines in 2001, it has been
routine for most pregnant women to have CF testing. When under-
going carrier screening, couples can choose to either be tested at
the same time or in tiers, where one partner is screened first and the
second is only screened if the first is found to be a carrier. In the
event that one partner is determined to be a carrier of a common
mutation, the second partner may want to be screened for more
than just the mutations recommended by ACMG or ACOG due to
the high heterogeneity of CFTR. For targeted mutations, low
throughput genotyping methods were originally used to diagnose
individuals, which included: the classic dot blot involving target
DNA immobilized on a membrane and then hybridized to a labeled

probe, the reverse dot blot which had an unlabeled probe immo-
bilized on a membrane and then hybridized to a labeled PCR
product, and gel-based restriction enzyme analysis which involved
DNA being cut at specific restriction sites and then run on gels to
genotype based on band sizes and patterns. Now with the advent of
high throughput genotyping, new methods involving fluorescent-
based or molecular weight-based allelic discrimination are utilized
which has allowed for many commercial laboratories to offer tar-
geted mutation panels that extend beyond those specifically recom-
mended by ACOG or ACMG. Sequencing can also be used for
diagnostic purposes or even for screening [16], which would
involve either Sanger or next generation sequencing, or a combina-
tion of both methods.
The ACMG has also recommended that reflex testing be done
if carrier screening indicates that an individual has the c.350G>A,
p.Arg117His mutation. Different phenotypes have been associated
with the c.350G>A mutation depending on whether it is found in
cis or trans with the 5T/7T/9T variant. If the 5T variant is found
in trans with c.350G>A, then the CBAVD phenotype occurs with-
out any other symptoms of CF, while if it is found in cis, classical CF
is expressed. Since the frequency of the 5T variant alone is high in
the general population, however, it is only recommended to screen
for this variant if the c.350G>A mutation is detected first [17].
While the diagnosis of CF is typically performed by molecular
genetic testing, confirmation is usually done via a sweat test which
measures the amount of salt in a patient’s sweat, with high salt levels
confirming the diagnosis. Once a diagnosis of CF has been made,
additional tests can include chest X-rays as well as lung function
tests to monitor the pulmonary phenotype, a sinus X-ray to look for
sinusitis, which can be a complication of CF, and sputum cultures
to look for pulmonary infection with Pseudomonas aeruginosa,
which is a bacteria that indicates advanced CF [18].
Currently, preconception carrier screening for couples is only
recommended, however newborn screening for CF is required in all
states in the United States. Testing can involve a two-tier system
where immunoreactive trypsinogen (IRT) analysis is done first on
dried neonatal blood spots, followed by DNA analysis for either the
common delta c.1521_1523delCTT mutation or other CF muta-
tions if the IRT levels are high [19].
1.3 Prenatal Testing Preconception screening is very important for couples so that they
and Counseling can make informed decisions about their reproductive options. If a
couple is found to be at an increased-risk of having an affected child
due to both parents carrying a mutation, the couple can consider
preimplantation genetic diagnosis (PGD) on embryos at a fertility
clinic so that only embryos known not to have CF will be candidates
for implantation. Couples can also choose to conceive naturally but
then undergo prenatal testing with the option of elective abortion if

the fetus is found to be affected. Using donor sperm or eggs is
another option, especially when the fathers are affected with CF
since these men have high rates of infertility, as well as adoption.
Prenatal testing for CF should be offered in instances of
increased risk either through direct or cultured chorionic villus
sampling (CVS) at 10–12 weeks or amniocentesis at
15–18 weeks. Since CF screening is recommended for all pregnant
women followed by partner screening in the case of carrier status,
targeted mutation analysis can be done on the prenatal sample for
the mutation(s) known to be present in the parents. In addition to
known carrier status of the parents or a family history of CF, fetal
intestinal obstruction, congenital viral infection, and intra-amniotic
bleeding are other prenatal symptoms that can be associated with
CF and indicate that molecular testing should be performed.
Meconium ileus, which is a bowel obstruction that occurs when a
newborn’s stool is thicker and sticker than normal, is observed in
10–20% of newborns that have CF, and since it can be seen in
second-trimester ultrasounds as an echogenic bowel, it can also
be used as a prenatal predictor of CF [20]. In instances of ultra-
sound indications, molecular testing of the parents to identify the
known mutation(s) is preferred, followed by testing the fetus in the
same laboratory. If only one mutation is found in the fetus, its risk
of being affected with CF can range from 13 to 43% based on the
prior empirical risk, while the detection of two mutations would be
diagnostic and the detection of no mutations decreases the risk to
less than 1 in 645 for Caucasian fetuses [21].
Whenever prenatal testing is being performed, it is always
important to test both the maternal blood sample and the fetal
sample to ensure that there is no maternal cell contamination
(MCC). Cytogenetic data has indicated that MCC can occur in
0.6–1.0% of cultured amniocytes, 0.1–0.9% of direct CVS, and
1.8–12.6% of cultured CVS [22]. Testing for MCC can be done
by comparing the maternal and fetal samples using microsatellite or
other polymorphic markers.
2 Materials
2.1 DNA Extraction 1. PureGene™ Genomic DNA purification kit (Qiagen, German-
town, MD, USA).
2.2 DNA-Based CFTR PCR primers must span all exons and at least 20 nt of
Molecular Diagnosis: intronic sequences must be M13 tagged (for standardization of
Sanger Sequencing assays—not necessary for all laboratories) and synthesized by an
established facility.
1. Primer working solutions: Prepare a 10 μM primer working
solution.
Table 1
Sequences for CFTR coding exon gene specific primers
Primer Sequence (50 ! 30 )
Product Length
Exon Forward Reverse (bp)
1 CCCAGAGTAGTAGGTCTTTGGC AAACCCAACCCATACACACG 204
2 GTGCATAATTTTCCATATGCC ttagccaccatacttggctC 341
3 GGGTTAATCTCCTTGGATATAC TTCACCAGATTTCGTAGTC 305
TTG TTTTC
4 TCTTGTGTTGAAATTCTCAGGG AAAACTACAACAGAGGCAG 525
TTTACAG
5 GAACCTGAGAAGATAGTAAGC GAAAACTCCGCCTTTCCAG 321
TAGATG
6 TGATCATATAAGCTCCTTTTAC TCCTGGTTTTACTAAAGTGGGC 343
TTGC
7 TGCCCATCTGTTGAATAAAAG CAAACATCAAATATGAGG 340
TGGAAG
8 cttccattccaAGATCCCTG TGAACATTCCTAGTATTAGC 476
TGGC
9 TGCTTGGCAAATTAAC gcACCTGGCCATTCCTCTAC 440
TTTAGAAC
10 CAGTGTAATGGATCATGGGC TGGAGAAGAGGATGACCACTG 853
11 cccttgtatcttttgtgcatagc AACCGATTGAATATGGAGCC 465
12 GGAAGATGTGCCTTTCAAATTC CCCACTAGCCATAAAACCCC 301
13 TGCATGTAGTGAACTG TGCAATCTATGATGGGACAG 255
TTTAAGG
14a AAATGCTAAAATACGAGACATA TCTTCGATGCCATTCATTTG 485
TTGC
14b GAAGGAGATGCTCCTGTCTCC CTACTCAATTGCATTCTGTGGG 529
15 ACAATGGTGGCATGAAACTG TGAGCTTTCGAATCTCTTAACC 547
16 AATTTAGATGTGGGCATGGG GGATTACAATACATACAAACA 201
TAGTGG
17 GGTTAAGGGTGCATGCTCTTC AAAGCCAGCACTGCCATTAG 473
18 GAGAAATTGGTCGTTACTTGAA GCAATAGACAGGACTTCAACCC 457
TC
19 GACTAGGAATAGAA CATTTGGGAACCCAGAGAAA 1055
TGGGGAGAGTA
20 TCTATTCAAAGAATGGCACCAG CAATGGAAATTCAAAGAAA 549
TCAC
(continued)
Table 1
(continued)
Primer Sequence (50 ! 30 )
Product Length
Exon Forward Reverse (bp)
21 TGGTTGAATACTTACTATA TGACAGATACACAGTGACCCTC 449
TGCAGAGC
22 AGCAAGTGTTGCATTTTACAAG GCTAACACATTGCTTCAGGC 428
23 GGTGACAGGATAAAATATTCCAA TTGCAGAGTAATATGAATTTC 362
TG TTGAG
24 TGATGGTAAGTACATGGGTG TTGTGCACACACATACATGC 334
TTTC
25 TCAAATGGTGGCAGGTAGTG GTGTCACCATGAAGCAGGC 385
26 CTACCCCATGGTTGAAAAGC TGAGTAAAGCTGGATGGCTG 421
27 CAAAATGCAAGGCTCTGGAC TCCTCAATTCCCCTTACCAA 494
CFTR PCR primers were M13 tagged (not shown on the above table)
2. Molecular biology grade water.

3. dNTPs.
4. Platinum Taq DNA Polymerase.
5. 10 PCR Buffer.
6. 50 mM MgCl2.
7. Shrimp Alkaline Phosphatase.
8. Exonuclease (USB, Thermo Fisher Scientific, Waltham, MA,
USA).
9. Primer sequences are described in Table 1.
2.3 DNA-Based 1. xTAG Cystic Fibrosis 60 kit v2 (Luminex Corporation, Austin,

Molecular Diagnosis: TX, USA).
CF60 Luminex
3 Methods
3.1 CFTR Gene 1. Extract DNA from prenatal cells using the PureGene™ Geno-
Sequence Analysis by mic DNA purification kit (see Note 1).
Sanger Sequencing 2. Dilute purified DNAs to a concentration of 50 ng/μL.
3.1.1 DNA Preparation, 3. Prepare PCR master mix for the number of samples to be tested
PCR, and Sequencing including a reagent blank and two extra for pipetting.
4. Distribute a 1.0 μL aliquot of the prepared DNAs (50 ng/μL)
to thin-walled PCR (0.2 mL) tubes with 1.2 μL of exon F/R
primer mix (10 μM working solution), 15.4 μL distilled water,

2.5 μL 10 PCR buffer, 0.75 mM MgCl2, 4.0 μL 0.2 μM
dNTP, and 0.2 μL Platinum Taq (5 U/μL).
5. Run the following PCR profile: 95 C for 5 min (95 C for 30 s,
60 C for 30 s, 72 C for 30 s) 35, 72 C for 7 min and
4 C hold.
6. Perform Exo/SAP treatment to clean up the PCR products
(2.5 μL Shrimp Alkaline Phosphatase and 1 μL Exonuclease).
Process the reactions in a thermal cycler programmed as fol-
lows: 37 C for 30 min, 99 C for 15 min, 4 C hold.
7. Perform bi-directional DNA sequencing for CFTR exon
(s) with 8–20 ng of the purified PCR product using procedures
recommended by the manufacturer.
3.1.2 Sanger Sequencing 1. Separate PCR products by electrophoresis in agarose gels to

Analysis ensure proper amplification, which should demonstrate a single
strong band with the expected size for each exon to be ana-
lyzed. The blank must not contain any amplification products.
If there is contamination in blank, all PCR reagents should be
discarded and new amplification reactions should be set up.
2. The chromatograms containing the sequencing data should
have unique, nonoverlapping peaks for homozygous samples.
A heterozygous missense or nonsense mutation will produce an
overlapped peak at the mutant position and deletion or inser-
tion in one strand will produce overlapped peaks at all positions
after the change. Sequencing results must show the variation in
both the forward and reverse directions.
3.2 CFTR Targeted 1. Extract DNAs for CFTR genotyping from blood samples or
Mutation Analysis by direct or cultured prenatal cells using the PureGene™ Geno-
Luminex Beads-Based mic DNA Purification Kit (see Note 2).
Genotyping 2. For multiplex PCR, add the following into each specimen tube:
3.2.1 CF60: DNA
9.75 μL DNase and RNase Free Distilled Water, 5.0 μL 5
Preparation, PCR and ASPE
Platinum Tfi Reaction Buffer, 1.75 μL Tfi 50 mM MgCl2,
2.5 μL xTAG CFTR PCR Primer Mix V2, 1 μL Platinum Tfi
exo() DNA Polymerase and 5.0 μL of appropriate DNA
sample.
3. Run the following PCR profile: 94 C for 2 min, (94 C for
15 s, 58 C for 30 s, 72 C for 30 s) 30, 72 C for 50 and 4 C
hold. Set the thermal cycler temperature as BLOCK Tempera-
ture with the heated lid enabled.
4. Prepare Enzyme Mix as follows: 2.5 μL Shrimp Alkaline Phos-
phatase and 1.0 μL Exonuclease I. Add 3.7 μL of the Enzyme
Mix into each of the PCR tubes. Incubate the tubes in a
thermal cycler programmed as follows: 37 C for 30 min and
99 C for 5 min and 4 C hold.
5. For multiplex ASPE, there will be two different ASPE Master

Mixes (A and B). Add the reagents in the order listed below to
prepare the ASPE A Master Mix: 6.8 μL DNase and RNase Free
Distilled Water, 4.0 μL 5 Platinum Tfi Reaction Buffer,
1.2 μL Tfi 50 mM MgCl2, 2.0 μL xTAG ASPE Primer A Mix
v2, and 1.0 μL Platinum Tfi exo() DNA Polymerase. For the
ASPE B Master Mix, repeat the above steps, but instead use
xTAG ASPE Primer B Mix v2. Add 5.0 μL of the same treated
PCR product to both ASPE Master Mixes. Cap each tube
immediately after addition of sample.
6. Place tubes in thermal cycler and cycle under the following
conditions: 94 C for 2 min, (94 C for 15 s, 56 C for 30 s,
74 C for 30 s) 30, 99 C for 5 min and 4 C hold. Set the
thermal cycler temperature as BLOCK temperature with the
heated lid enabled.
7. Thaw and bring both the xTAG CFTR Bead Mix A v2 and the
xTAG CFTR Bead Mix B v2to room temperature, limiting its
exposure to light.
8. Vortex the A and B Bead Mixes for 10 s and then sonicate for
10 s to disperse the beads. Repeat this step.
9. Vortex for several seconds and aliquot 22.5 μL of the A Bead
Mix into one set of eight labeled tubes, and aliquot 22.5 μL of
the B Bead Mix into a different set of eight labeled tubes.
10. Aliquot 2.5 μL of the A ASPE product into the A Bead Mix
tube and aliquot 2.5 μL of the B ASPE product into the B Bead
Mix tube.
11. Place tubes in a thermocycler programmed as follows: 96 C
for 2 min, 37 C for 30 min, and 37 C hold.
12. Before proceeding to the next step (about 5 min before com-
pletion of the half hour incubation) prepare the Reporter
Solution. Vortex the tube of Streptavidin, R-Phycoerythrin
(SA-PE) conjugate for 2–5 s. For one sample, add 1.25 μL of
SA-PE to 123.75 μL of xTAG 1 Wash Buffer.
13. Add 100 μL reporter (made from above step) to each well,
briefly pipette up and down and transfer into Costar plate.
There will be both an “A” plate or column and a “B” plate or
column.
14. Incubate the plate(s) at room temperature for 15 min. Run
samples on Luminex machine according to manufacturer’s
instructions.
3.2.2 CF60: Results 1. Data interpretation is accomplished using the TDAS CFTR
Analysis for CFTR Analysis Software.
Genotyping Using Luminex 2. Select both the “A” and “B” output data files that you want to
Bead Technology analyze and analyze them using the xTAG Cystic Fibrosis v2
assay with 60 variations detected.
3. A negative control (blank) must be included in every run.

Identify your negative control in the software.
4. In the Mask Editor dialog box, click on both checkboxes next
to the ACMG Panel column header and the Full Panel column
header.
5. Positive control samples are rotated on each run. If the signals
obtained for the blank are too high, the run is invalid and must
be repeated. Samples with insufficient signals or low bead
counts are repeated.
4 Notes
1. Any equivalent DNA extraction method may be used.

2. The QIASymphony DNA extraction Technology or any equiv-
alent DNA extraction method may be used.
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https://doi.org/10.1586/14737159.4.1.49
Chapter 16
Prenatal Diagnosis of Tay-Sachs Disease

Jinglan Zhang, Hongjie Chen, Ruth Kornreich, and Chunli Yu
Abstract
Tay-Sachs disease (TSD) is an autosomal recessive lysosomal storage disorder caused by mutations of the
HEXA gene resulting in the deficiency of hexosaminidase A (Hex A) and subsequent neuronal accumula-
tion of GM2 gangliosides. Infantile TSD is a devastating and fetal neurodegenerative disease with death
before the age of 3–5 years. A small proportion of TSD patients carry milder mutations and may present
juvenile or adult onset milder disease. TSD is more prevalent among Ashkenazi Jewish (AJ) individuals and
some other genetically isolated populations with carrier frequencies of approximately ~1:27 which is much
higher than that of 1:300 in the general population. Carrier screening and prenatal testing for TSD are
effective in preventing the birth of affected fetuses greatly diminishing the incidence of TSD. Testing of
targeted HEXA mutations by genotyping or sequencing can detect 98% of carriers in AJ individuals;
however, the detection rate is much lower for most other ethnic groups. When combined with enzyme
analysis, above 98% of carriers can be reliably identified regardless of ethnic background. Multiplex PCR
followed by allele-specific primer extension is one method to test for known and common mutations.
Sanger sequencing or other sequencing methods are useful to identify private mutations. For prenatal
testing, only predefined parental mutations need to be tested. In the event of unknown mutational status or
the presence of variants of unknown significance (VUS), enzyme analysis must be performed in conjunction
with DNA-based assays to enhance the diagnostic accuracy. Enzymatic assays involve the use of synthetic
substrates 4-methylumbelliferyl-N-acetyl-β-glucosamine (4-MUG) and 4-methylumbelliferyl-6-sulfo-2-
acetamido-2-deoxy-β-D-glucopyranoside (4-MUGS) to measure the percentage Hex A activity (Hex A%)
and specific Hex A activity respectively. These biochemical and molecular tests can be performed in both
direct specimens and cultured cells from chorionic villi sampling or amniocentesis.
Key words Tay-Sachs disease, Hexosaminidase A (Hex A) deficiency, HEXA gene, Targeted muta-
tion, HEXA sequencing, 4-methylumbelliferyl-N-acetyl-β-glucosamine (4-MUG), 4-methylumbelli-
feryl-6-sulfo-2-acetamido-2-deoxy-β-D-glucopyranoside (4-MUGS), Percentage Hex A activity (Hex
A%), Specific Hex A activity, Prenatal diagnosis
1 Introduction
The GM2 gangliosidoses are a group of related lysosomal storage

disorders with a deficiency of either β-hexosaminidase or an activa-
tor protein causing the inability to break down the gangliosides
GM2. As a result, the GM2 gangliosides and related substrates are
accumulated in the lysosomes of neuronal cells and lead to neuronal
233
234 Jinglan Zhang et al.
swelling and neurological degeneration [1–4]. Infantile Tay-Sachs

disease (TSD) is the prototype GM2 gangliosidoses which is char-
acterized by hypotonia, loss of motor skills, decreased attentiveness,
and increased startle response with onset from 3 to 6 months of age
followed by progressive neurological deterioration including blind-
ness, dementia, seizures, and subsequent death before the age of
3–5 years. Juvenile (subacute), chronic and adult-onset variants of
TSD have later onset, milder and variable neurological findings
with slower progression [2–4]. The other two GM2 gangliosidoses,
Sandhoff disease and GM2 activator deficiency, have clinical presen-
tations indistinguishable from TSD, except that there are also other
symptoms involved in Sandhoff disease such as organomegaly,
skeletal abnormalities, and oligosacchariduria [3].
Two lysosomal β hexosaminidase isoenzymes (Hex A and Hex
B) are critical for the degradation of gangliosides GM2 in the central
nervous system and other substrates such as glycolipids and glyco-
proteins throughout the body. The enzymatic functions are specific
toward the substrates’ terminal nonreducing sugars N-acetylgluco-
samine (GlcNAc) or N-acetylgalactosamine (GalNAc) in β-linkage
[2, 5]. Hex A (αβ) and Hex B (ββ) enzymes consist of α- and
β-subunits, which are encoded by the HEXA and HEXB genes,
respectively [6]. Neutral water soluble substrate such as artificial
4-methylumbelliferyl-N-acetyl-β-glucosamine (4-MUG) is specific
for the β-subunit and therefore can be hydrolyzed by both Hex A
and Hex B. The negatively charged hydrophobic gangliosides GM2
can only be hydrolyzed by the α-subunit and hence Hex A in the
presence of GM2 activator protein which binds the membrane
bound GM2 gangliosides to form soluble complex for Hex A
cleavage [5]. The synthetic 4-methylumbelliferyl-6-sulfo-2-aceta-
mido-2-deoxy-β-D-glucopyranoside (4-MUGS) is also specific
toward the α-subunit of hexosaminidase and is used to measure
the specific Hex A activity in vitro without the requirement of the
activator protein [7]. Three forms of GM2 gangliosidoses are auto-
somal recessive disorders with deficiencies of Hex A enzyme. TSD is
caused by mutations of the HEXA gene and profound deficiency of
Hex A, while Sandhoff disease is caused by mutations of the HEXB
gene and deficiencies of both Hex A and Hex B. GM2 activator
protein deficiency results from mutations of the G2MA gene lead-
ing to nonfunctional Hex A activity in vivo [2–4]. Due to the severe
clinical outcome and lack of effective therapy, carrier screening and
prenatal testing for at risk pregnancies are important to prevent the
birth of affected fetuses. The prenatal testing strategy and biochem-
ical and molecular methods used for carrier and prenatal testing for
TSD are the focus of this chapter.
1.1 Molecular TSD is an autosomal recessive disease caused by mutations in the

Genetics of TSD HEXA gene. The HEXA gene is 35 kb in size and located on
chromosome 15q23-24 with 13 introns and 14 exons. Before
Prenatal Diagnosis of Tay-Sachs Disease 235
population-based carrier screening programs started in the 1970s,

TSD was most commonly found in the Ashkenazi Jewish
(AJ) population with an incidence of 1:3600 births and a carrier
frequency of ~1:28 [8, 9]. In some genetically isolated populations
such as French Canadians in Quebec, Cajuns from Louisiana and
the Old Order Amish in Pennsylvanian, TSD carrier frequency is
also increased and similar to that of AJ individuals due to the
founder effects. Overall, TSD incidence is 100 times lower in the
general population with a carrier frequency of approximately 1:300
[2, 4]. Three mutations c.1274_1277dupTATC (1278+TATC),
c.1421+1G>C (IVS12+1G>C), and c.805G>A (p.G269S)
account for more than 98% disease alleles in AJ individuals
[9, 10]. Founder mutations c.1073+1G>A (IVS9+1G>A) in the
Irish [11] and g.2644_10588del (del7.6 kb) in French Canadians
have also been identified [12]. More than 170 mutations in the
HEXA gene have been reported to date in Human Gene Mutation
Database (http://www.hgmd.cf.ac.uk/). Most of the mutations are
associated with infantile TSD. The genotype and phenotype corre-
lation has been well studied for TSD. The residual Hex A activity
correlates inversely with the severity of the disease. Individuals with
acute infantile TSD usually have two null alleles and no Hex A
enzymatic activity (e.g., c.1274_1277dupTATC, c.1421+1G>C,
c.1073+1G>A, and g.2644_10588del). Individuals with juvenile
or chronic and adult onset TSD are usually compound heterozy-
gotes for a null allele and an allele that results in low residual Hex A
activity, or compound heterozygotes of two alleles that result in low
residual Hex A activity [2, 4, 13]. Missense mutations at two
codons (178 and 258), c.532C>T (p.R178C), c.533G>A (p.
R178H), c.533G>T (p.R178L), and c.772G>C (p.D258H), are
associated with B1 variant of TSD, accounting for a small propor-
tion of TSD patients [14]. The B1 variants have normal enzyme
activities toward the 4-MUG substrate; however are inactive
toward natural substrates in vivo or the Hex A-specific 4-MUGS
substrate [15]. For this reason, some B1 variant carriers might be
falsely negative by the routine heat inactivation assay with the
4-MUG substrate. Individuals with a null allele and a B1 variant
present with juvenile TSD. Individuals with two B1 alleles usually
have chronic and adult onset disease [2, 14]. False positive carrier
screening results for TSD by enzyme analysis are caused by the
presence of one of two known non-pathogenic pseudo-deficiency
alleles, c.739C>T (p.R247W) or c.745C>T (p.R249W). These
two variants are not associated with the disease, but result in the
production of a Hex A enzyme with decreased activity toward the
4-MUG substrate used in enzyme assays [16]. It has been reported
about 35% of non-Jewish and 2% of Jewish individuals who are
carriers by enzyme analysis are carriers for one of these two pseudo-
deficiency alleles [16].
1.2 Biochemical The biochemical diagnosis of TSD is established when Hex A

Enzyme Assays activity is absent or extremely low with normal or even elevated
Hex B activity. Enzyme analysis has also been the primary method
for carrier testing of TSD in non-Jewish populations. The natural
substrate GM2 gangliosides is most specific for testing Hex A activ-
ity but it is expensive, and assays using it are time-consuming and
difficult to perform in clinical laboratories. Synthetic artificial sub-
strates have been widely used for enzyme testing for TSD [17]. The
enzyme assay can be performed using serum (or plasma), white
blood cells, chorionic villi, amniotic fluid, and other tissues. In
women who are either pregnant [18] or using birth control medi-
cation [19], interfering Hex P in serum (or plasma) may impact the
percentage Hex A activity (Hex A%) and cause false positive results;
therefore, the enzyme assay for these individuals should not be
performed in serum or plasma. 4-MUG is the most sensitive and
commonly used artificial substrate in TSD biochemical carrier
screening tests. The Hex A and Hex B isoenzymes are both reactive
toward 4-MUG with similar kinetics; however they have different
thermal stabilities. At pH 4.4, Hex B is stable up to 55 C, whereas
Hex A is inactivated under heat. The half-life of Hex A is approxi-
mately 10 min at 50 C and 3 min at 55 C. The differential thermal
stabilities of Hex A and Hex B are the basis for the routinely used
heat-inactivation hexosaminidase activity assay. The total hexosa-
minidase activities (Hex A and Hex B) are measured with 4-MUG
before and after the denaturation of the Hex A at 50–55 C for
2–3 hour. The Hex A% is calculated from the difference and divided
by the total activity to reflect the proportion of Hex A from total
hexosaminidase [2]. Both total hexosaminidase activity and Hex A
% are routinely reported for the 4-MUG-based enzyme assay;
however, only Hex A% is used for the clinical diagnosis of TSD
and TSD carriers. TSD is characterized by a very low Hex A%
(<10%) and normal total activity (Hex B). Carriers of TSD have
an intermediate Hex A% (20–50%) that can be discriminated from
non-carriers (60–65% 5%). When testing for Tay-Sachs carriers
with a 4-MUG-based enzyme assay, Sandhoff carriers can also be
revealed by a characteristic high Hex A% activity (75–80%) and
relatively low total activity. In the individuals affected with Sandhoff
disease, the total activity is absent or extremely low with an
extremely high Hex A% (>80–90%). Another commonly used syn-
thetic sulfated substrate 4-MUGS is used to measure the specific
Hex A activity as it is almost exclusively hydrolyzed by Hex A.
4-MUGS substrate has been used in the diagnosis of TSD, and
particularly the B1 variant [15]. TSD and Sandhoff disease patients
demonstrate absent or extremely low specific Hex A activity
(0–15 nmol/hour/mg protein) compared to normal individuals
(>200 nmol/hour/mg protein). The B1 variant could be falsely
normal or intermediate when tested with 4-MUG substrate, there-
fore should be tested with 4-MUGS substrate. The 4-MUGS sub-
strate is not appropriate for carrier testing of TSD.
1.3 Molecular Molecular testing for TSD includes targeted DNA assays for
Genetic Testing founder mutations in various populations at higher risk for TSD.
Five HEXA mutations c.1274_1277dupTATC, c.1421+1G>C,
c.805G>A, c.1073+1G>A, and g.2644_10588del are included in
many DNA-based TSD carrier screening programs together with
two non-pathogenic pseudo-deficiency alleles c.739C>T and
c.745C>T [9]. One commonly used genotyping technology to
test these HEXA mutations is based on allele specific primer exten-
sion (ASPE) and is deployed on many different platforms. ASPE
usually involves an enzymatic reaction to determine the target
genotype by specific nucleotide incorporation followed by a
platform-specific detection step to quantify the allele-specific nucle-
otide incorporation. However, the detection rate of this five-
mutation panel is much lower in the general population and thus
makes targeted mutation testing less effective. Therefore, it is
recommended to always perform the enzymatic test with targeted
mutation analysis and reflex to full gene sequencing to screen for
the pathogenic mutation when the enzyme test results are positive
or inconclusive. Generally, only targeted genotyping or sequencing
of the predefined pathogenic mutations identified in parents should
be performed for TSD prenatal testing. When a fetus is negative for
at least one of parental pathogenic alleles, the likelihood this fetus is
affected by TSD is greatly diminished. Maternal cell contamination
studies (e.g., testing maternal and fetal microsatellite DNA
sequences) should always be performed to ensure that negative
results such as a heterozygote call in the fetus are not caused by
the presence of maternal DNA. Microsatellites (short tandem
repeats, STRs) are small arrays of tandem repeats (1–4 bp) inter-
spersed throughout the genome occurring on average once every
50 Kb. Because of the highly variable number of repeats, micro-
satellites are widely used in linkage studies and paternity tests.
Microsatellite markers can be used to determine the status of
maternal cell contamination in prenatal samples. Usually, the fluo-
rescently labeled PCR products are run by capillary electrophoresis
on a genetic analyzer.
1.4 Prenatal Testing Prenatal testing for TSD should be offered for at-risk pregnancies.
and Counseling Chorionic villus sampling (CVS) at 10–12 weeks or amniocentesis at
15–18 weeks can be performed to obtain fetal cells. DNA-based
testing can be performed with either direct or cultured CVS cells or
amniocytes. Enzyme analysis can be performed using direct or
cultured CVS and cultured amniocytes. In addition, enzyme analysis
can also be performed with cell-free amniotic fluid specimens. If
both parental disease-causing alleles are identified, DNA-based
assays for the parental mutations in the fetus are sufficient and
accurate. If the risk for TSD of a pregnancy is determined solely by
parental enzyme analysis and the underlying HEXA mutation(s) are
unknown, enzyme testing in combination with specific mutation
analysis can be pursued with formal genetic counseling. With the
increasing trend of pan-ethnic carrier screening and utilization of

sequencing-based DNA assays in clinical testing, identifications of
novel mutations and variants of unknown significance (VUS) have
increased. It is important that both enzyme and DNA-based assay
are performed concurrently in these situations. Both 4-MUG and
4-MUGS assays should be performed to increase test sensitivity as
well as testing a reference enzyme β-galactosidase to ensure sample
integrity. Normal and positive controls of the appropriate specimen
type are always tested with the fetal specimen. Maternal cell contam-
ination should be excluded by molecular marker studies.
2 Materials
2.1 Enzymatic 1. 0.9% saline (normal saline).

Assays 2. 0.05% trypsin-EDTA solution.
3. Serum Buffer: 12 mM citrate-20 mM Na2HPO4, pH 4.4.
Dissolve 461 mg citric acid (MW ¼ 192.1) and 568 mg
Na2HPO4 (MW ¼ 142.0) in 150 mL deionized water. Adjust
pH to 4.4, and then bring volume to 200 mL. Store at 20 C
for up to 2 years.
4. WBC buffer: 6 mM citrate–10 mM Na2HPO4, pH 4.4/
0.6% HAS.
Dilute 50 mL serum buffer to a final volume of 100 mL with
deionized water. Add 0.6 g of Human Serum Albumin (HSA).
Store at 20 C for up to 2 years.
5. Citrate–Na2HPO4 Buffer: 0.1 M citrate–0.2 M Na2HPO4
(0.1/0.2 M C–P), pH 4.2.
0.2 M citrate: dissolve 19.21 g of citric acid in 500 mL deio-
nized water.
0.4 M Na2HPO4: dissolve 28.4 g of Na2HPO4 in 500 mL
deionized water.
Mix 58.8 mL of 0.2 M citrate with 41.2 mL of 0.4 M
Na2HPO4, adjust pH to 4.2 by adding either 0.2 M citrate or
0.4 M Na2HPO4. Bring volume to 200 mL with deionized
water. Discard after making the 4-MUGS substrate.
6. Sodium acetate Buffer: 100 mM sodium acetate/sodium chlo-
ride, pH 4.0.
Dissolve 1.64 g sodium acetate (MW ¼ 82.04) and 1.17 g
sodium chloride (MW ¼ 58.44) in 180 mL deionized water,
adjust pH to 4.0 with acetic acid, and bring the volume to
200 mL. Discard after making the β-Galactosidase substrate.
7. 4-MUG substrate: 3.0 mM 4-MUG.
Dissolve 16 mg of 4-MUG (MW ¼ 379.36) in 14 mL of either
serum or WBC buffer for a batch of 20 samples. This substrate
is freshly made for use before the enzyme reaction.
8. 4-MUGS substrate: 3.0 mM 4-MUGS.

Dissolve 14.9 mg 4-MUGS potassium salt (MW ¼ 497.5) in
10 mL of 0.1/0.2 M C-P buffer, pH 4.2. Store at 20 C for
2 years.
9. β-Galactosidase substrate: 0.5 mM β-Galactosidase.
Dissolve 1.7 mg 4-methylumbelliferyl-β-D-galactopyranoside
(MW ¼ 338.3) in 10 mL 100 mM sodium acetate/sodium
chloride buffer, pH 4.0. Store at 20 C for 2 years.
10. Stop solution (for stopping enzyme reaction): 0.1 M ethylene
diamine.
Dilute 6.75 mL anhydrous ethylene diamine (MW ¼ 60.1,
density ¼ 0.899 g/mL) with deionized water to a final volume
of 1000 mL. Store at room temperature in amber glass dis-
pense bottle for up to 1 year.
11. 4-MU stock solution: 25 mM 4-MU.
Dissolve 44 mg of 4-MU (MW ¼ 176.2) in 10 mL DMSO.
12. 4-MU intermediate stock solution: 100 μM 4-MU.
Dilute 50 μL of the 25 mM 4-MU stock solution with DMSO
to a final volume of 12.50 mL.
13. 4-MU working solution: 10 μM 4-MU.
Dilute 0.5 mL of the 100 μM 4-MU intermediate stock solu-
tion with DMSO to a final volume of 5 mL.
2.2 DNA Extraction 1. PureGeneTM Genomic DNA purification kit (Qiagen, Ger-
mantown, MD, USA).
2.3 DNA-Based HEXA PCR primers must span all exons and at least 20 nt of
Molecular Diagnosis: intronic sequences must be M13 tagged (for standardization of
Sanger Sequencing assays—not necessary for all laboratories) and synthesized by an
established facility. 10 μM working solutions of the primers are
prepared.
1. Primer working solutions: Prepare a 10 μM primer working
solution.
2. DNase and RNase-free molecular biology grade distilled water.
3. dNTPs.
4. Platinum Taq DNA Polymerase.
5. 10 PCR Buffer.
6. 50 mM MgCl2 (Invitrogen, Carlsbad, CA, USA).
USA).
9. Primer sequences are described in Table 1.
Table 1
Sequences for HEXA coding exon gene-specific primers
Primer sequence (50 !30 )

Product
Exon Forward Reverse length (bp)
1 CCAGGCCGGAAGTGAAAG CTCCTGATTGAACCGTAGTCC 638
TA
2 TAGGGTCTTGGTTTTGCCTG AGGCCATCCAGAGTTACAGC 267
3 GTCCAGTGATTTATATAGAATATC AACACCAACCTTCCCACATC 249
TGGTC
4 TGCTCTGCTACATTGAGAACC CAATATTGGGATCCAACCCC 230
5 TTGTCTTCATCTCCCTGTGC GGAACTTGGTCTGTCCGTTG 292
6 CCAACATCGCAAGTTTGAGG GCCACAGCCAGATTCAGAC 268
7 TGTGGGCATTTTGAGTATCTTC AGCCAGTGCCCTGAAGC 315
8 TTACGTGTAGGACTGTGCGTG CCTCGGGTGCTAACTTCTA 357
TTC
9–10 TAATCCCCAGGCATTAGGC TCTGTAGAGGCAGGGAGGAG 629
11–12 GACATACTTTGCTGCTGGGG CTTCAGAAGGCTCGTTGCAC 782
13 GGTAGCAGCCTGTGGATGTC CTCTCTAAGGGGTTCCCCAG 286
14 GTGTGAAAAGTGTTGCTGGG TGCCACATTACTCTTTATTGAA 345
TG
HEXA PCR primers were M13 tagged (not shown on the above table)
2.4 DNA-Based 1. Primer sequences provided by Luminex (Luminex Corporation

Molecular Diagnosis: Austin, TX, USA).
Luminex 2. 10 PCR Primer Mix (Luminex Corporation Austin, TX,
USA).
3. ASPE Primer Mix (Luminex Corporation Austin, TX, USA).
4. Bead Mix (Luminex Corporation Austin, TX, USA).
5. DNase and RNase-free molecular biology grade distilled water.
6. dNTPs.
7. HotStarTaq® DNA Polymerase (Qiagen, Germantown, MD,
USA).
8. 10 Qiagen HotStar® PCR Buffer (Qiagen, Germantown,
MD, USA).
9. 25 mM MgCl2 (Qiagen, Germantown, MD, USA).
Table 2
Sequences for microsatellite marker primers used in maternal cell contamination studies
Primer sequence (50 !30 )

Product
Maker Forward Reverse length (bp)
D7S1817 [6-FAM]CAAATTAATGGCAAAAACTGC CCCCCCATTGAGG 122
TTATTAC
D2S406 [5-HEX]GTGATGGTAAATAATTTCTGAGACC GACAACTGAC 184
TTTCCCAGGA
DxS981 [6-FAM]TCAGAGGAAAAGAAGTAGACATACT TTCTCTCCAC 187
TTTTCAGAGTCA
D7S821 [5-HEX]ACAAAACCCCAAGTACGTGA TATGACAGGCATC 248
TGGGAGT
D11S1392 [6-FAM]GCAGGTATATTGCATCCATACG AGAAGGCC 196
TTGAGACATCCA
CSF1PO [6-FAM]AACCTGAGTCTGCCAAGGACTAGC TTCCACACACCAC 319
TGGCCATCTTC
TPOX [5-HEX]GCACAGAACAGGCACTTAGG CGCTCAAACG 270
TGAGGTTG
VWA [5-HEX]GCCCTAGTGGATGATAAGAATAATC GGACAGATGA 151
AGTATGTG TAAATACA
TAGGATGGATGG
D7S820 [6-FAM]ATGTTGGTCAGGCTGACTATG GATTCCACATTTA 243
TCCTCATTGAC
Fluorescent dyes were conjugated to the 50 -end of forward PCR primers

USA).
12. 10 Wash Buffer (Luminex Corporation Austin, TX, USA).
13. Streptavidin, R-Phycoerythrin conjugate (Invitrogen, Carls-
bad, CA, USA).
2.5 DNA-Based 1. PCR primers for microsatellite markers are diluted to a working
Marker Studies for concentration of 20 μM.
Maternal Cell 2. Primer sequences are described in Table 2.
Contamination
Analysis
3 Methods
3.1 Methods for Normal and affected controls must be performed with the prenatal
Enzymatic Assays specimen. Affected controls are aliquots of previously diagnosed
affected samples kept at 80 C. Concurrent normal controls are
3.1.1 Quality Control
provided from the prenatal sampling facilities or tissue culture
facilities.
3.1.2 Blanks and For 4-MUG assays, the WBC buffer is used as blank for direct and
Standards cultured CVS and cultured amniocytes; and serum buffer is used as
blank for cell-free amniotic fluid. For 4-MUGS and β-galactosidase
assays, water is used as blank for all specimen types. In addition, a
self-blank is used for 4-MUGS assay in amniotic fluid, where the
stopping solution is added before the substrate. One point 4-MU
calibration containing 1 nmol 4-MU standard (100 μL of 10 μM
4-MU standard) is used in every prenatal assay for calculation of
enzyme activities. Full range 4-MU calibration is performed every
6 months.
3.1.3 Preparation of 1. Clean and wash the direct CVS samples with normal saline
Direct CVS Sample (performed by a cell culture laboratory).
2. Add 300–1000 μL of normal saline to the freshly prepared or
frozen CVS sample according to the size of the sample.
3. Freeze and thaw the CVS sample in a dry ice/ethanol bath and
a 37 C water batch for five times to lyse the cells.
4. Centrifuge at 4600 g for 1 min.
5. Use the supernatant for enzyme assay or store at 20 C until
assay.
3.1.4 Preparation of 1. Decant the culture media and wash with 10 mL normal saline.
Cultured CVS Cells or 2. Add 0.75 mL 0.05% trypsin-EDTA to T25 flask and incubate
Amniocytes at 37 C incubator for 5 min. Check under a microscope to
make sure the cells are detached then add 0.75 mL saline.
3. Transfer the cell suspension into a 1.5 mL Eppendorf micro-
fuge tube and centrifuge at 4600 g for 3 min.
4. Discard the supernatant and wash the cell pellet with 1.5 mL
normal saline twice.
5. Remove the normal saline by inverting the tube over a paper
towel.
6. Add 300–1000 μL of chilled water depending on the size of the
pellet. Vortex to resuspend the pellet.
7. Sonicate the sample for 10 s using Fisher F60 sonic dismem-
brator with output power setting of 4.
8. Centrifuge at 4600 g for 1 min and the resultant supernatant
is ready for enzyme assay or store at 20 C until analysis.
3.1.5 Total The general procedures described below apply to all prenatal speci-
Hexosaminidase and Hex men types with minor differences in buffers, dilution factor, and
A% Activity with 4-MUG reaction times.
Substrate
1. Dilute cell lysates from direct CVS, cultured CVS, and cultured
amniocytes with WBC buffer and a 1:20 dilution, e.g., 570 μL
WBC buffer +30 μL lysates. Dilute amniotic fluid sample with
serum buffer and a 1:10 dilution, e.g., 540 μL serum buffer
+60 μL of amniotic fluid sample.
2. Label four sets of 12 75 mm glass tubes for blank, normal
control (NC), affected control (AC), and prenatal sample in
duplicates (set-0, 1h, 2h, and 3h).
3. Place 50 μL of diluted controls and patient sample in each tube
and cover with parafilm to avoid evaporation of the samples
during heat inactivation. 50 μL WBC buffer is used as blank for
lysates. 50 μL serum buffer is used as blank for amniotic fluid
sample.
4. Place “0” set of tubes in ice-water bath for non-heat or total
hexosaminidase.
5. Place the other rack containing 1h, 2h, and 3h sets in 50 C
water bath for heat-inactivation.
6. Take out sample tubes labeled 1h, 2h, and 3h after 1 hour,
2 hour, and 3 hour heat-inactivation respectively. Place on
ice-water bath immediately.
7. Add 100 μL of the freshly prepared 3.0 mM 4-MUG substrate
in WBC buffer to all samples. 3.0 mM 4-MUG substrate in
serum buffer is used for amniotic fluid samples.
8. Incubate in 37 C water bath for 15 min (30 min for amniotic
fluid sample).
9. Add 2.35 mL of stop solution to each tube to stop enzyme
reaction.
10. Read fluorescence at 360 nm excitation and 450 nm emission.
11. Determine protein concentration in mg/mL by method of
preference (e.g., Lowry or Bradford methods).
12. Total β-hexosaminidase (Hex) activity (nmol/hour/mg pro-
tein or nmol/hour/mL) is expressed as nmol of 4-MU pro-
duced per hour per mg of protein in the cell lysates or per mL
of amniotic fluid, where 4-MU turnover is calculated from
comparing fluorescence intensity of sample to the 1 nmol
4-MU standard.
13. The Hex A% or [Hex A/(Hex A + Hex B)]% is calculated from
the fluorescence difference of the non-heated and heated sam-
ples divided by the fluorescence of the non-heated samples and
multiplied by 100 (see Note 1). Hex A% of 1h is reported.
3.1.6 Specific Hex A 1. Label 12 75 mm glass test tubes in duplicate for blank,
Activity with 4-MUGS normal control, affected control, and sample.
Substrate 2. Place 20 μL of lysates for 4-MUGS assay. Add 30 μL of water to
controls and sample. Pipette 50 μL of water for blank.
3. Add 50 μL of 4-MUGS substrate to each sample. Quickly
vortex or shake all the samples for 10 s and incubate in 37 C
water bath for 30 min.
4. After incubation, remove tubes from the water bath and imme-
diately place in ice-water bath.
5. Add 2.4 mL of stop solution to all tubes to stop enzyme
reaction.
6. Read the fluorescence density at 365 nm excitation and 450 nm
emission.
7. The specific Hex A activity is expressed as nmol of 4-MU
produced per hour per mg of protein in the cell lysates or per
mL of amniotic fluid.
3.1.7 Reference Enzyme 1. Label 12 75 mm glass test tubes for blank, normal control,
β-Galactosidase Activity affected control, and sample.
2. Place 10 μL of lysates for β-galactosidase activity assay. Add
40 μL of water to controls and patient samples. Pipette 50 μL
of water for the blank.
3. Add 50 μL of 3.0 mM β-galactosidase substrate to each sample.
Quickly vortex or shake all the samples for 10 s and incubate in
37 C water bath for 30 min.
4. After incubation, remove tubes from the water bath and imme-
diately place in ice-water bath.
5. Add 2.4 mL of stop solution to all tubes to stop enzyme
reaction.
6. Read the fluorescence density at 365 nm excitation and 450 nm
emission.
7. The β-galactosidase activity is expressed as nmol of 4-MU
produced per hour per mg of protein.
3.1.8 Result The Hex A% after 1 hour heat-inactivation and specific Hex A
Interpretation activity are reported for the prenatal sample along with the normal
and affected controls. It is important for the prenatal testing labo-
ratory to establish normal and affected ranges for Hex A% and
specific Hex A activity in different specimen types. The affected
ranges of Hex A% in all specimen types are <5–10%. The affected
ranges of specific Hex A activity are less than 5–10% of the normal
mean of tested specimen type. The enzyme results are not valid if
there are significant maternal cell contaminations or sample integ-
rity is questionable reflected by low reference enzyme activity.
3.2 Methods for 1. Extract DNA from prenatal cells using the PureGene™ Geno-
HEXA Gene Sequence mic DNA purification kit (see Note 2).
Analysis by Sanger 2. Dilute purified DNA to a concentration of 50 ng/μL.
Sequencing
3. Prepare PCR master mix for the number of samples to be tested
3.2.1 DNA Preparation, including a reagent blank and two extra for pipetting loss.
PCR, and Sequencing Distribute a 1.0 μL aliquot of the prepared DNA (50 ng/μL)
to thin-walled PCR (0.2 mL) tubes with 1.2 μL of exon F/R
primer mix (10 μM working solution), 15.4 μL distilled water,
2.5 μL 10 PCR buffer, 0.75 mM MgCl2, 4.0 μL 0.2 μM
dNTP, and 0.2 μL Platinum Taq (5 U/μL).
4. Run the following PCR profile:
(a) One cycle: 95 C for 5 min.
(b) Thirty-five cycles: 95 C for 30 s, 60 C for 30 s, 72 C for
30 s.
(c) One cycle: 72 C for 7 min.
(d) 4 C hold.
5. Perform Exo/SAP treatment to clean up the PCR products
(2.5 μL Shrimp Alkaline Phosphatase and 1 μL Exonuclease).
Process the reactions in a thermal cycler programmed as
follows:
(b) One cycle: 99 C for 15 min.
(c) 4 C hold.
6. Perform bi-directional DNA sequencing for HEXA exon
(s) with 8–20 ng of the purified PCR product using procedures
recommended by the manufacturer.
3.2.2 Sanger Sequencing 1. Separate PCR products by electrophoresis in agarose gels to

Analysis ensure proper amplification, which should demonstrate a single
strong band with the expected size for each exon to be ana-
lyzed. The blank must not contain any amplification products.
If there is contamination in blank, all PCR reagents should be
discarded and new amplification reactions should be set up.
2. The chromatograms containing the sequencing data should
have unique, non-overlapping peaks for homozygous samples.
A heterozygous missense or nonsense mutation will produce an
overlapped peak at the mutant position and deletion or inser-
tion in one strand will produce overlapped peaks at all positions
after the change. Sequencing results must show the variation in
both the forward and reverse directions.
3.3 HEXA Targeted 1. Extract DNA for HEXA genotyping from blood samples or
Mutation Analysis by direct or cultured prenatal cells using the PureGene™ Geno-
Luminex Beads-Based mic DNA Purification Kit (see Note 3).
Genotyping 2. For multiplex PCR, add the following into each specimen tube:
3.3.1 DNA Preparation,
4.0 μL DNase and RNase Free Distilled Water, 1.9 μL 10
PCR, and ASPE
Qiagen HotStar® PCR Buffer, 1.0 μL Qiagen 25 mM MgCl2,
2.5 μL PCR Primer Mix (with dNTPs), 0.6 μL HotStarTaq®
DNA Polymerase and 2.5 μL of appropriate DNA sample.
3. Set the thermal cycler temperature as BLOCK Temperature
with the heated lid enabled and run the following PCR profile:
(b) Thirty-three cycles: 95 C for 30 s, 58 C for 30 s, 72 C
for 2 min.
(d) 4 C hold.
4. Prepare Enzyme Mix as follows: 1.3 μL Shrimp Alkaline Phos-
phatase and 0.5 μL Exonuclease I. Add 1.8 μL of the Enzyme
Mix into each of the PCR tubes. Incubate the tubes in a
thermal cycler programmed as follows:
(c) 4 C hold.
5. For multiplex ASPE, add the reagents in the order listed below
to prepare the ASPE Master Mix. 10.5 μL DNase and RNase
Free Distilled Water, 3.0 μL 10 Qiagen HotStar® PCR
Buffer, 1.6 μL Qiagen 25 mM MgCl2, 2.0 μL ASPE Primer
Mix (with dNTPs), 0.4 μL HotStarTaq® DNA Polymerase.
Add 2.5 μL of treated PCR product to the appropriately
labeled ASPE tube. Cap each tube immediately after addition
of sample.
6. Set the thermal cycler temperature as BLOCK Temperature
with the heated lid enabled and place tubes in thermal cycler.
Run the following program:
(b) Forty cycles: 95 C for 30 s, 56 C for 30 s, 72 C for
1 min.
(d) 4 C hold.
7. Thaw and bring the 2 Beads Mix to room temperature,
limiting its exposure to light, and prepare a 1.1 Wash Buffer
solution from the 10 Wash Buffer stock.
8. Vortex the 2 Bead Mix tube for 10 s and then sonicate for 10 s
to disperse the beads. Repeat this step.
9. Use black tube (1.5 mL or 5 mL) to dilute 2 beads mix with
1.1 wash buffer at 1:1 volume ratio. The final concentration
for beads mix should be 1.
10. Vortex for several seconds and aliquot 45 μL of the Bead Mix
into the eight labeled tubes.
11. Aliquot 5 μL of the ASPE reaction into the corresponding
labeled tube or well.
12. Place tubes in a thermal cycler programmed as follows:
(c) 37 C hold.
13. Add 100 μL 1 wash buffer to each well, then place the strips/
plate on the Magnetic Particle Concentrator (The concentrator
has 13 columns, which enables the 96-well plate to shift its
position). Hold 10 s and shift strips/plate position by one
column. Hold another 10 s and then shift the strips/plate
back by one column. Hold at least 1 min for beads precipita-
tion. Brown palates should be seen along one side of the well
inner wall.
14. Add 120 μL 1 wash buffer to each well and repeat the shifting
and pouring procedures once.
15. Before proceeding to the next step (about 5 min before com-
pletion of the 1 hour incubation) prepare the Reporter Solu-
tion. Vortex the tube of Streptavidin, R-Phycoerythrin
(SA-PE) conjugate for 2–5 s. For eight samples, add 6 μL of
SA-PE (1 mg/mL) to 2 mL of 1 Wash Buffer
(Tm Bioscience) in a 15 mL sterile centrifuge tube.
16. Add 100 μL reporter (made from above step) to each well,
briefly pipette up and down, and transfer into Costar plate.
17. Incubate the plate at room temperature for 15 min. Run
samples on Luminex machine according to the manufacturer’s
instructions.
3.3.2 Results Analysis for 1. Data interpretation is accomplished by obtaining the raw data
HEXA Genotyping Using as a comma delimited file (csv file) which subtracts the back-
Luminex Bead Technology ground from each variation in each sample, calculates the allelic
ratio for each variation, and ultimately defines a “SNP Call” for
each sample and variation.
2. The calls are made using a combination of thresholds that
require each signal to be significantly higher than the back-
ground and that the allelic ratios fall within empirically derived
ranges. The following QC cutoff is used to accept a genotyping
result: minimum MFI of 300, minimum bead count of 100.

Genotype is called based on allelic ratios: if 0.85: homozy-
gous mutation; if >0.7 and <0.85 or >0.15 and <0.3: No
Call; if 0.3 and 0.7: heterozygous; if 0.0 and 0.15:
wild type.
3. A negative control (blank) must be included in every run.
Positive control samples are rotated on each run. If the signals
obtained for the blank are too high, the run is invalid and must
be repeated. Samples with insufficient signals or low bead
counts are repeated.
3.4 Maternal Cell 1. Extract DNA from maternal blood and prenatal samples.
Contamination Study 2. PCR Amplification. Add the reagents in the order listed below
3.4.1 Method for Marker to prepare the PCR Master Mixes. 9.8 μL DNase and RNase
Studies Using PCR and Free Distilled Water, 2.6 μL 10 Qiagen HotStar® PCR
Capillary Electrophoresis Buffer, 1.5 μL Qiagen 25 mM MgCl2, 4.0 μL 1.25 mM
dNTP and 2.5 μL R/F primers, 1 U platinum Taq polymerase,
and 2.5 μL of appropriate DNA sample into each
specimen tube.
3. Run the following PCR profile:
(b) Twenty-five cycles: 94 C for 15 s, 58 C for 20 s, 72 C
for 20 s.
(d) 4 C hold.
4. After thermal cycling, store the fluorescent PCR products in
the dark at 4 C until ready to proceed, and store at 20 C for
longer-term storage.
5. Prepare the samples for loading in the Genetic Analyzer by
denaturing the samples at 95 C for 3 min and then placing
the plate on ice for 3 min.
6. Run the samples on the Genetic Analyzer according to the
manufacturer’s instructions.
7. Analyze the data according to the Genemapper instructions.
3.4.2 Analysis for the 1. Most samples will be heterozygous and show two major bands
Marker Study due to the high heterozygosity of the markers. Homozygous
samples will show one major band.
2. The stutter bands and nonspecific bands should be minimal
and distinguishable from real products. The maternal and fetal
patterns are compared.
3. At least one band should be in common between the maternal
and fetal samples. A marker is considered informative if the
patterns are different between the maternal and fetal samples.
4. If none of the initial five markers tested are informative, addi-

tional markers are tested.
5. If maternal cell contamination is not present, there will be a
band present in the maternal sample that is not observed in the
fetal sample.
4 Notes
1. Multiple heat-inactivation time points are the experience of this

testing laboratory.
2. Any equivalent DNA extraction method may be used.
3. The QIASymphony DNA extraction Technology or any equiv-
alent DNA extraction method may be used.
References
1. Moriwaki S, Takashima S, Yoshida H, population: the first 1000 cases at New York
Kawano N, Goto M (1977) Histological obser- University Medical Center, New York,
vation of the brain of Tay-Sachs disease with NY. Arch Intern Med 158(7):777–781
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of an autopsy case. Acta Pathol Jpn 27 RJ, Kornreich R (2010) Experience with car-
(3):387–407 rier screening and prenatal diagnosis for
2. Gravel RA, Clarke JTR, Kaback MM, 16 Ashkenazi Jewish genetic diseases. Hum
Mahuran D, Sandhoff K, Suzuki K (1995) Mutat 31(11):1240–1250. https://doi.org/
The GM2 gangliosidoses. In: Scriver CR, 10.1002/humu.21327
Beaudet AL, Sly WS et al (eds) The metabolic 10. Fernandes MJ, Kaplan F, Clow CL,
and molecular basis of inherited disease, 7th Hechtman P, Scriver CR (1992) Specificity
edn. McGraw-Hill, New York, pp 2839–2879 and sensitivity of hexosaminidase assays and
3. Sandhoff K, Harzer K (2013) Gangliosides and DNA analysis for the detection of Tay-Sachs
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(25):10195–10208. https://doi.org/10. org/10.1002/gepi.1370090303
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4. Kaback MM, Desnick RJ (1993) Hexosamini- ner EE, Prence EM (1996) Heterozygosity for
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University of Washington, Seattle, Seattle, WA Med Genet 33(10):829–832
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Isoenzymes of N-acetyl-beta-hexosaminidase. Kaplan F (1992) The French Canadian
Acta Biochim Pol 46(3):739–751 Tay-Sachs disease deletion mutation: identifi-
6. Mahuran DJ, Tsui F, Gravel RA, Lowden JA cation of probable founders. Hum Genet 89
(1982) Evidence for two dissimilar polypeptide (1):83–87
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Proc Natl Acad Sci U S A 79(5):1602–1605 Banwell B, Blaser S, Kok F, Giugliani R,
7. Bayleran J, Hechtman P, Saray W (1984) Syn- Mahuran D, Clarke JT (2006) The natural his-
thesis of 4-methylumbelliferyl-beta-D-N-acet- tory of juvenile or subacute GM2 gangliosido-
ylglucosamine-6-sulfate and its use in sis: 21 new cases and literature review of
classification of GM2 gangliosidosis genotypes. 134 previously reported. Pediatrics 118(5):
Clin Chim Acta 143(2):73–89 e1550–e1562. https://doi.org/10.1542/
8. Kronn D, Jansen V, Ostrer H (1998) Carrier peds.2006-0588
screening for cystic fibrosis, Gaucher disease, 14. Mahuran DJ (1999) Biochemical conse-
and Tay-Sachs disease in the Ashkenazi Jewish quences of mutations causing the GM2
gangliosidoses. Biochim Biophys Acta 1455 17. Lowden JA, Zuker S, Wilensky AJ, Skomor-
(2–3):105–138 owski MA (1974) Screening for carriers of
15. Peleg L, Meltzer F, Karpati M, Goldman B Tay-Sachs disease: a community project. Can
(1995) GM2 gangliosidosis B1 variant: bio- Med Assoc J 111(3):229–233
chemical and molecular characterization of 18. Navon R, Mark Z, Mashiah S, Padeh B (1973)
hexosaminidase A. Biochem Mol Med 54 Determination of Tay-Sachs genotypes in preg-
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JS, Prence EM, Brown D, Chabot T, Triggs- (1979) Human hexosaminidase isozymes.
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tion. Am J Hum Genet 53(6):1198–1205
Chapter 17
Next Generation Sequencing of Prenatal Structural

Chromosomal Rearrangements Using Large-Insert Libraries
Benjamin B. Currall, Caroline W. Antolik,
Ryan L. Collins, and Michael E. Talkowski
Abstract
Precise tests for genomic structural variation (SV) are essential for accurate diagnosis of prenatal genome
abnormalities. The two most ubiquitous traditional methods for prenatal SV assessment, karyotyping and
chromosomal microarrays, do not provide sufficient resolution for some clinically actionable SVs. Standard
whole-genome sequencing (WGS) overcomes shortcomings of traditional techniques by providing base-
pair resolution of the entire accessible genome. However, while sequencing costs have continued to decline
in recent years, conventional WGS costs remain high for most routine clinical applications. Here, we
describe a specialized WGS technique using large inserts (liWGS; also known as “jumping libraries”) to
resolve large (>5000–10,000 nucleotides) SVs at kilobase-resolution in prenatal samples, and at a fraction
of the cost of standard WGS. We explicate the protocols for generating liWGS libraries and supplement with
an overview for processing and analyzing liWGS data.
Key words Jumping libraries, Whole-genome sequencing, Prenatal diagnosis, Structural variation,
Chromosomal abnormalities, Copy-number variation
1 Introduction
Structural variation (SV), including copy-number variations

(CNVs; deletions and duplications) and balanced chromosomal
abnormalities (BCAs; SV absent gross gain or loss of DNA, such
as translocations or inversions), represents a highly penetrant class
of deleterious genetic mutations that can arise sporadically in off-
spring and perturb genes essential in human development. Tradi-
tional cytogenetic techniques revolutionized genetic analysis of
large genomic structural variation (SV) and are a cornerstone of
prenatal diagnosis for many genetic disorders; however, these main-
stay techniques have substantial limitations [1–3]. Conventional
G-banded karyotyping, while sensitive to all forms of extremely
large SV, cannot provide breakpoint resolution below the size of
individual chromosome banding patterns (~3–10 million
251
252 Benjamin B. Currall et al.
nucleotides) [4–7]. Fluorescence in situ hybridization (FISH) pro-

vides improved resolution over karyotyping, but requires prior
knowledge of the approximate genomic location of an SV, and
thus is only a tenable approach when specific SV are already sus-
pected [8]. Array-based comparative genomic hybridization
(aCGH) has far higher resolution, but is only sensitive to CNVs
and therefore blind to all BCAs.
Whole-genome sequencing (WGS) represents a major advance
beyond traditional methods by providing comprehensive data
about the accessible genome in its entirety, thus in theory permit-
ting delineation of all forms and sizes of SV to base-pair resolution
[9, 10]. Standard WGS preparation involves acoustic or enzymatic
fragmentation of genomic DNA, attaching adapters that are com-
patible with the selected sequencing technology, and identifying
indices to isolate these fragments [11]. The resulting DNA
“libraries” are often further amplified by polymerase chain reaction
(PCR) before high throughput sequencing. At present, Illumina
sequencing instruments are the most widely used, such as the
HiSeq 2000/2500 and MiSeq, and employ Solexa “sequencing-
by-synthesis” chemistry. Other approaches are also viable, and each
offers technical advantages and disadvantages. High throughput
sequencing yields an abundance of short (generally ~150 nucleo-
tide) reads from the original genomic DNA. These reads are com-
putationally mapped back against a reference genome, and
unexpected read mappings can be used to flag sites of divergence
versus the reference genome (e.g., sites of genetic variation in the
sequenced sample). Standard WGS approaches, however, can be
clinically cumbersome due to high sequencing costs and require-
ments of computational infrastructure and expert knowledge [12].
An alternative WGS method, known as long-insert WGS
(liWGS; also known as “jumping libraries”), can provide a cost-
effective alternative to conventional WGS while still capturing
nearly all large SV in the genome [13–16]. Importantly, the resolu-
tion of liWGS is roughly four orders of magnitude (1000-fold)
higher than karyotyping and one order of magnitude (tenfold)
higher than most clinical aCGH platforms. In contrast to standard
WGS, liWGS involves an initial fragmentation of genomic DNA to
a target size of typically 3–5 thousand nucleotides. These fragments
are circularized in a manner that allows specific retention of the
junction sites following fragmentation of the circle. DNA libraries
are prepared from the smaller junction fragments, which contain
the ends of the initial, larger fragments. Paired sequencing reads
from such libraries effectively span the sample genome in “jumps”
proportional to the initial fragment size, permitting liWGS to infer
~10-fold more covered genomic bases per sequenced fragment as
compared to standard WGS. The resultant sequencing data can be
processed and analyzed analogously to standard WGS. At current
sequencing costs, this liWGS approach results in an approximate
Next Generation Sequencing of Prenatal Structural Chromosomal. . . 253
eightfold cost reduction to achieve comparable nucleotide coverage

of the genome by conventional short-insert WGS compared to the
physical coverage of the inserts produced by liWGS. However, this
improved efficiency comes at a significant tradeoff in the genomic
information that can be obtained. Given the low nucleotide cover-
age achieved by liWGS, only relatively large SVs are accessible, and
the lower size threshold is directly proportional to the size of the
insert. The method cannot detect small SVs (~5000 nucleotides)
nor single nucleotide variants (SNVs) and small insertion/deletions
(1–49 nucleotides). The demonstrated value of liWGS has been in
the delineation of large SVs with high confidence and the genomic
features that they disrupt, providing clinically relevant information
that is otherwise inaccessible to conventional technologies such as
karyotyping and aCGH [17, 18].
Herein, we describe a liWGS preparation method we developed
in our laboratory for detecting chromosomal abnormalities in pre-
natal samples, which we have applied to samples received within our
hospital and collaborating hospitals [19]. Notably, the method is
also applicable to pediatric and adult populations, as has been
demonstrated in numerous research studies [13, 18, 20–25].
2 Materials
2.1 Fragmentation of 1. miniTUBE Red 5.0 kb (Covaris, Woburn, MA).

Human Genomic DNA 2. 1 Tris-EDTA buffer.
3. Covaris focused ultrasonicator such as E220evolution (Covaris,
Woburn, MA).
2.2 End-Repair of 1. End-It DNA End-Repair Kit (Epicentre, Madison, WI).

Sheared DNA 2. QIAquick PCR Purification Kit (Qiagen, Germany).
2.3 EcoP15I Cap 1. Cap Adapter 1: /5Phos/ACAGCAG (Integrated DNA Tech-

Adapter Ligation nologies, Coralville, IA).
2. Cap Adapter 2: /5Phos/CTGCTGTAC (Integrated DNA
Technologies, Coralville, IA).
3. Nuclease-Free Duplex Buffer (Integrated DNA Technologies,
Coralville, IA).
4. Quick Ligation Kit (New England Biolabs, Inc., Ipswich, MA).
5. QIAquick PCR Purification Kit (Qiagen, Germany).
2.4 Gel Size 1. 1 Kb Plus DNA Ladder (Thermo Fisher Scientific, Cambridge,
Selection MA).
2. QIAquick Gel Extraction Kit (Qiagen, Germany).
2.5 T4 1. Internal adapter 1: /5Phos/CGTTC/iBiodT/CCGT

Circularization (Integrated DNA Technologies, Coralville, IA).
2. Internal adapter 2: /5Phos/GGAGAACGGT (Integrated
DNA Technologies, Coralville, IA).
3. Nuclease-Free Duplex Buffer (Integrated DNA Technologies,
Coralville, IA).
4. T4 DNA Ligase (New England Biolabs, Inc., Ipswich, MA).
2.6 DNase Treatment 1. Plasmid-Safe ATP-Dependent DNase (Epicentre, Madison,

WI).
2.7 EcoP15I 1. EcoP15I (New England Biolabs, Inc., Ipswich, MA).

Digestion 2. InSolution sinefungin (EMD Millipore, Billerica, MA). Make
up solution for protocol at 10 mM.
2.8 End-Repair of 1. DNA Polymerase I, Large (Klenow) Fragment (New England

Digested DNA Biolabs, Inc., Ipswich, MA).
2. dNTPs, 25 mM.
2.9 Bead Binding 1. 1 wash buffer:

(a) 60 μL Tris–HCl, 1 M pH 7.5.
(b) 2.4 mL NaCl, 5 M.
(c) 12 μL EDTA, 0.5 M.
(d) 0.5 μL Tween 20.
(e) QS to 12 mL with sterile water.
2. 2 binding buffer:
(a) 60 μL Tris–HCl, 1 M pH 7.5.
(b) 2.4 mL NaCl, 5 M.
(c) 12 μL EDTA, 0.5 M.
(d) QS to 6 mL with sterile water.
3. Dynabeads MyOne Streptavidin C1 (Thermo Fisher Scientific,
Waltham, MA).
4. Magnetic rack.
2.10 dA-Tailing 1. NEBNext dA-Tailing Module (New England Biolabs, Inc.,

Ipswich, MA).
2.11 Adapter 1. Universal adapter: ACACTCTTTCCCTACAC-

Ligation GACGCTCTTCCGATC*T (Integrated DNA Technologies,
Coralville, IA).
2. Indexed adapter: /5Phos/GATCGGAAGAGCACACGTCT-

GAACTCCAGTCAC(6BPindex) (Integrated DNA Technolo-
gies, Coralville, IA).
3. Quick Ligation Kit (New England Biolabs, Inc., Ipswich, MA).
2.12 PCR on Beads 1. Universal forward primer: AATGATACGGCGACCACCGA-

GATCTACACTCTTTCCCTACACGACGCTCTTCC-
GATC*T (Integrated DNA Technologies, Coralville, IA).
2. Custom reverse indexed primer: CAAGCAGAAGACGGCA-
TACGAGAT(+6BPprimerindex)GTGACTGGAGTTC
(reverse primer index is the reverse complement of the barcode
adapter index) (Integrated DNA Technologies, Coralville, IA).
3. Phusion High-Fidelity PCR Master Mix with HF Buffer (New
England Biolabs, Inc., Ipswich, MA).
2.13 Final Gel 1. 1 Kb Plus DNA Ladder (Thermo Fisher Scientific, Cambridge,
Selection MA).
2. QIAquick Gel Extraction Kit (Qiagen, Germany).
3 Methods
3.1 Fragmentation of 1. Load 5–10 μg of DNA into a red miniTUBE and combine with
Human Genomic DNA 1 TE for a total volume of 200 μL (see Note 1).
2. Shear DNA to a target size of 3 kb using a Covaris focused-
ultrasonicator with red miniTUBEs and the 5k shear protocol
as specified by the manufacturer.
3. Transfer fragmented DNA (200 μL) to clean 1.5 mL tube.
4. Use the QIAquick PCR Purification Kit to purify sample as
specified by the manufacturer’s protocol, eluting in 35 μL
Buffer EB.
3.2 End-Repair of 1. Use the End-It End-Repair Kit to end-repair fragmented

Sheared DNA DNA: to the fragmented and purified DNA, add 5 μL End-It
Buffer (10), 5 μL End-It dNTPs (2.5 nM), 5 μL End-It ATP
(10 nM), and 1 μL enzyme mix (see Note 2).
2. Mix sample, spin down, and incubate at room temperature for
30–40 min.
3. Purify end-repaired DNA using QIAquick PCR Purification
Kit, eluting in 52 μL Buffer EB.
3.3 EcoP15I Cap 1. Resuspend cap adapters 1 and 2 together in Nuclease-Free

Adapter Ligation Duplex Buffer to a final concentration of 50 μM.
3.3.1 Prepare Cap 2. Incubate at 94 C for 2 min, then allow to cool on ice.
Adapters
3.3.2 Ligate Cap 1. Determine concentration of DNA and calculate the volume of
Adapters duplexed cap adapters required for an adapter:fragment ratio of
10:1 (see Notes 3 and 4).
2. To each sample, add: the volume of cap adapters calculated for
that sample, 55 μL Quick Ligase Buffer (2), and 2 μL Quick
Ligase (2,000,000 U/mL).
15 min.
4. Purify samples using QIAquick PCR Purification Kit, eluting in
30 μL Buffer EB.
3.4 Gel Size 1. Prepare a 1% agarose gel with ~0.17 μg of ethidium bromide
Selection per milliliter.
3.4.1 Run Agarose Gel 2. Add an appropriate loading dye to DNA ladder and samples,
and Select Band from Gel mix, and load ladder and samples on gel. Load 1 Kb Plus
Ladder each in the right- and leftmost lanes of each row of
gel; load samples so that empty wells are left between the ladder
and sample lanes, as well as between lanes containing different
samples, to reduce the risk of contamination.
3. Run gel at 100 V for approximately 1 h, until a bromophenol
dye indicator has migrated about 2 cm (see Note 5).
4. Use a razor blade or scalpel to select a DNA band in the 3–5 kb
range from each sample on the gel (see Notes 6 and 7).
3.4.2 Extract DNA Using 1. Weigh gel slice and add 3 volumes of Buffer QG (see Note 8).
Qiagen’s QIAquick Gel 2. Let it sit at room temperature until gel is completely dissolved.
Extraction Kit
3. Use columns to purify sample as directed in the manufacturer’s
protocol, but wash sample two times with 600 μL Buffer PE
during the wash step.
4. Elute in 100 μL Buffer EB (see Note 9).
3.5 T4 DNA 1. Resuspend internal adapters 1 and 2 together to a final con-

Circularization centration of 2 μM in Nuclease-Free Duplex Buffer.
3.5.1 Prepare Internal 2. Incubate at 94 C for 2 min, then allow to cool on ice.
Adapters
3.5.2 Circularize DNA 1. Determine concentration of DNA using Nanodrop instrument

and calculate the volume of duplexed internal adapters to add
for an adapter:fragment ratio of 3:1 (see Note 10).
2. To each sample, add 78 μL water, 20 μL T4 Ligation Buffer

(10), the calculated amount of internal adapters, and 2 μL T4
DNA Ligase (see Note 11).
at least 3 h.
4. Purify with QIAquick PCR Purification Kit, eluting in 60 μL
Buffer EB.
3.6 DNase Treatment 1. Using the Plasmid-Safe DNase Kit, add the following to each
sample: 24.5 μL water, 10 μL Plasmid-Safe Buffer (10), 5 μL
ATP (25 mM), 2 μL Plasmid-Safe DNase (100 U/μL) (see
Note 12).
2. Mix, spin down, and incubate at 37 C for 40 min.
Buffer EB (see Note 13).
3.7 EcoP15I Digest 1. Add the following to each sample: 10 μL NEB Buffer 3.1
(10), 20 μL ATP (10), 1 μL sinefungin (10 mM), and
6 μL EcoP15I enzyme (10,000 U/mL).
2. Mix samples by pipetting and digest at 37 C overnight.
3. After digestion is complete, inactivate enzyme by heating for
20 min at 65 C; cool on ice for 5 min after heat inactivation.
3.8 End Repair of 1. Add 1.5 μL dNTPs (25 mM) and 1.5 μL of DNA Polymerase I,
Digested DNA Large (Klenow) Fragment (5000 U/mL) to each sample. Mix
samples by pipetting and incubate at room temperature for
30 min.
2. Inactivate by heating for 20 min at 65 C, then cool on ice for
5 min.
3.9 Streptavidin 1. Aliquot 30 μL beads (10 mg/mL) per sample into a 1.5 mL
Bead Binding tube (see Note 14). Separate beads from solution on magnetic
rack and discard supernatant.
2. Add 500 μL 1 wash buffer to sample, remove from magnet,
and gently mix. Separate beads from solution on magnet and
discard supernatant.
3. Repeat step 3 twice for a total of three washes.
4. Add 500 μL 1 binding buffer, remove from magnet, and
gently mix. Separate beads from solution on magnet and dis-
card supernatant (see Note 15).
5. Resuspend beads in the original volume using 1 binding
buffer.
6. Add 105 μL 2 binding buffer and 30 μL beads to each sample
and mix by pipetting (see Note 16).
7. Bind for 30 min at room temperature, mixing by pipetting

every 10 min.
8. Separate beads from solution on magnet and discard
supernatant.
9. Wash beads four times with 200 μL wash buffer (see Note 17).
3.10 dA-Tailing 1. Using the NEBNext dA-Tailing Module, add 42 μL water,

5 μL dA-Tailing Buffer (10), and 3 μL Klenow (exo-) frag-
ment (5000 U/mL) to each sample. Mix by pipetting and
incubate for 30 min at 37 C.
supernatant.
3. Wash beads four times with 200 μL wash buffer, then wash
once with ~50 μL Quick Ligase Buffer (1) (see Note 18).
3.11 Adapter 1. Resuspend universal adapter with individual barcoded adapters

Ligation in Nuclease-Free Duplex Buffer to a final concentration of
15 μM.
3.11.1 Prepare Adapters
2. Incubate for 2 min at 94 C, then allow to cool on ice.
3.11.2 Ligate Adapters to 1. Using the Quick Ligation Kit, add 24.5 μL water, 25 μL Quick
Samples Ligase Buffer (2), 1 μL duplexed adapter (15 μM), and 1.3 μL
Quick Ligase (2,000,000 U/mL) to each sample, using a
different barcoded adapter for each sample.
2. Incubate at room temperature for 45 min.
supernatant.
4. Wash beads four times with 200 μL wash buffer, then wash
once with 200 μL Buffer EB.
5. Resuspend beads in 30 μL Buffer EB.
3.12 Amplify 1. Add 1 μL universal forward primer (25 μM), 1 μL sample-

Samples on Beads to specific reverse indexed primer (25 μM), and 75 μL Phusion
Add Illumina Adapters Master Mix (2) to each sample (see Note 19).
Using Phusion HF PCR 2. Mix samples by pipetting and split individual samples into three
Master Mix with HF reactions of 50 μL each for PCR.
Buffer 3. PCR amplify using the following conditions (see Note 20):
(a) One cycle: 98 C for 30 s.
(b) Eleven cycles: 98 C for 10 s, 65 C for 30 s, 72 C for
30 s.
(d) 10 C hold.
4. After PCR is complete, combine replicates for each sample into

single 1.5 mL tube.
5. Separate beads from solution on magnet and transfer superna-
tant to a new 1.5 mL tube.
Buffer EB.
3.13 Gel Purification 1. Prepare a 1.5–2% agarose gel with ~0.17 μg of ethidium bro-
of Final Product mide per mL.
2. Add an appropriate loading dye to DNA ladder and samples,
mix, and load 1 Kb Plus Ladder (1 μg/μL) and samples on gel.
Load one ladder each in the right- and leftmost lanes of each
row of gel, then load samples onto gel, leaving an empty well
between ladder and sample.
3. Run gel at 100 V for approximately one and a half hours.
4. Use a razor blade or scalpel cut the band ~200 bp from the gel
for each sample (see Notes 21 and 22).
5. Extract sample using QIAquick Gel Extraction kit as described
in Subheading 3.4.2, but elute in 20 μL EB.
3.14 Quantify and 1. Quantify using Agilent Bioanalyzer, Agilent Tapestation, or

Pool Libraries qPCR.
2. Pool libraries in desired ratio for Illumina sequencing (see
Note 23).
3.15 Sequencing and 1. Sequence libraries with paired 25 bp read chemistry and a 6 bp
Data Processing barcode read on an Illumina sequencing platform.
2. Demultiplex reads corresponding to the sample-specific 6 bp
barcode index attached to the prepared library, in accordance
with Illumina’s standard recommended protocols.
3. Reverse-complement the raw reads; this can be done with tools
such as fastx or seqtk (https://github.com/lh3/seqtk) [26].
4. Align reads using a pairwise-aware aligner appropriate for short
(25 bp) reads, such as BWA-backtrack (see Note 24), against a
human genome reference assembly (see Note 25) [27].
5. Mark duplicate sequenced fragments; this can be done with
tools such as Picard MarkDuplicates (http://broadinstitute.
github.io/picard/) or SAMBLASTER [28].
6. Sort aligned reads by numerical coordinate ordering; this can
be done with tools such as samtools or sambamba [29, 30].
7. Evaluate alignment metrics of the processed library; this can be
done with PicardTools (recommended; http://broadinstitute.
github.io/picard/), sambamba/samtools flagstat, bamtools
stats, and numerous other programs (see Note 26 and
Table 1) [29–31].
Table 1
liWGS library alignment metrics on nine multiethnic samples sequenced to deep coverage as part of
the 1000 Genomes Project: Human Genome Structural Variation Consortium
Alignment Duplication
rate Proper rate Median Haploid
Raw read pair Chimeric insert physical
Sample pairs Read Pair rate Read Pair pair rate size coverage
HG00512 183,274,642 97.2% 95.1% 88.0% 12.4% 10.3% 7.4% 3401 161.4
HG00513 195,273,608 96.8% 94.4% 88.1% 11.9% 10.2% 6.6% 3315 166.9
HG00514 178,875,771 96.0% 93.5% 86.9% 14.1% 11.6% 7.0% 3325 147.5
HG00731 187,721,771 95.8% 93.2% 88.0% 15.5% 12.6% 5.5% 3439 159.9
HG00732 180,471,734 95.4% 92.9% 85.0% 18.4% 16.5% 8.4% 3525 144.6
HG00733 211,630,514 96.9% 94.6% 80.8% 8.1% 7.0% 14.4% 3736 193.8
GM19238 198,430,973 96.2% 93.6% 79.1% 12.9% 10.7% 15.4% 3493 158.1
GM19239 182,791,861 97.0% 94.7% 87.2% 13.0% 11.2% 7.8% 3420 158.3
GM19240 208,381,715 97.2% 95.2% 85.3% 11.9% 9.8% 10.3% 3497 184.1
Mean 191,872,510 96.5% 94.1% 85.4% 13.1% 11.1% 9.2% 3461 163.9
8. Perform visual quality assurance on the distribution of insert

sizes as determined by Picard CollectInsertSizeMetrics (see
Fig. 1, Table 1, and Note 27; http://broadinstitute.github.
io/picard/).
3.16 Data Analysis 1. Isolate anomalous read-pairs using samtools or sambamba

[29, 30].
2. Search across anomalous read-pairs for unexpected aggrega-
tions of anomalous pairs in close proximity (<5 kb) with high
alignment and read qualities.
4 Notes
1. 3 μg of DNA is recommended as a minimum starting input, but

the protocol can also be done with less DNA input if
optimized.
2. Aliquot ATP from kit into smaller, single use tubes to reduce
freeze-thaw cycles. Always keep ATP-containing solutions on
ice during reaction preparation.
3. The amount of DNA used in this step and the subsequent gel
purification can affect the percentage of chimeras present in the
final sequencing reads. To greatly reduce the amount of chi-
meras, limit the DNA used in this step to no more than 4 μg
Fig. 1 Example insert size distributions of sequenced fragments from nine

libraries generated by the Talkowski Laboratory with this protocol for the
Human Genome Structural Variation project. Given an original input of 5 μg
genomic DNA and a targeted insert size of 3.5 kb, the below distribution of
proper sequenced fragments were obtained when sequenced to a yield of 180 M
individual fragments with 25 bp reads on an Illumina HiSeq 2500. Reads were
processed as described in Subheading 3.14 and the below distribution was
generated from aligned BAM files by PicardTools (http://broadinstitute.github.io/
picard/). These data are available freely through the 1000 Genomcs Project ftp
server, accessible here: http://www.1000genomes.org/human-genome-
structural-variation-consortium/. Notably, for detection of known structural
variation, coverage can be greatly reduced as desired; 30–50 physical
coverage is typically sufficient for confirmation of known rearrangements
(80 ng/μL in 50 μL volume), though reactions can be split into

multiple samples if more complex libraries are desired.
4. The volume of cap adapter added to a sample should not
exceed 4 μL; samples that require higher volumes should be
diluted and total sample should be reduced.
5. Exceeding the recommended gel run time will likely require
multiple Qiagen columns for purification to retain the desired
size range.
6. Capturing pre- and post-size selection images of the gel is
recommended.
7. A straight edge is recommended to ensure accurate and
uniform size selection of bands, and selection should be
below the 6 kb threshold to minimize chimera rates (expected
size of concatemerization of two 3 kb fragments).
8. The gel slice should weigh ~0.4 g. If exceeded, two spin
columns may be required.
9. To maximize yield during extraction, two elution steps of

50 μL each (total vol ¼ 100 μL) should be used.
10. A maximum of 1 μg DNA (10 ng/μL in 100 μL volume) is
recommended for use in this step. If DNA concentration is
higher, increase volume of total reaction or split into multiple
tubes to obtain ~1 μg of DNA per reaction. Higher amounts of
DNA can increase chimera rates.
11. Generally, adding 0.6–0.7 μL of internal adapters per sample is
recommended.
12. Aliquot ATP from kit into smaller, single-use tubes to reduce
freeze-thaw cycles.
13. If working with more than eight samples at once, transferring
samples to a 96-well plate at this time is suggested.
14. Beads should be at room temperature prior to use; mix beads
thoroughly before use to ensure solution is uniform.
15. A brief spin in a centrifuge is recommended at this stage to
ensure complete removal of any residual wash buffer.
16. Mix samples slowly on beads to prevent excessive air bubble
formation, which can interfere with wash buffer removal.
17. Take care to remove all wash buffer after this step before
proceeding. Residual wash buffer can interfere with later
reactions.
18. Washing with a small amount of Quick Ligase Buffer (1) aids
in removing any remaining traces of wash buffer.
19. For each sample, use the specific reverse primer that corre-
sponds to the adapter added in the previous step.
20. The number of cycles can be adjusted based on anticipated
library complexity and tolerance of duplications for a specific
experiment. Increasing cycles will increase final yield but may
increase duplication rates.
21. Product should be visible as a ~200 bp band on the gel. If the
size is lower than ~200 bp, run the gel longer to avoid possible
contamination with adapter/primer dimers during gel
extraction.
22. Capturing pre- and post-size selection images of the gel is
recommended.
23. It is recommended to save an aliquot of the final pool.
24. Many common aligners are not appropriate for 25 bp reads
generated by this protocol. Ensure your choice of aligner is
appropriate. BWA-Backtrack is the recommended aligner [27].
25. The most current human reference genome assembly is avail-
able directly from the NCBI Genome Reference Consortium
(GRC; http://www.ncbi.nlm.nih.gov/projects/genome/
assembly/grc/).
26. Given average read quality and sufficient sequencing depth for
minimal analysis of a human whole-genome (>40 M
sequenced read-pairs), desirable alignment quality metrics
might be: pairwise alignment rate 90%; chimeric pair
rate 10%; pairwise duplication rate 10%; median insert
size ¼ 3500 bp. An example of typical metrics for a high
coverage library (~150 haploid coverage) such as those gen-
erated for the 1000 Genomes Project/Human Genome Struc-
tural Variation Consortium is provided in Table 1.
27. Desirable insert size distributions of sequenced libraries will
feature a sharp left tail (right skew) leading to a peak at the
approximate desired library insert size. The right tail can be
elongated, but generally is not advised to extend significantly
beyond ~8 kb for a 3.5 kb median insert library, at which point
the distribution kurtosis will adversely impact read alignment
algorithms. See Fig. 1 and Table 1 for an example of desirable
insert size distributions and related alignment statistics.
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Chapter 18
Prenatal Diagnosis by Whole Exome Sequencing in Fetuses

with Ultrasound Abnormalities
Vanessa Felice, Avinash Abhyankar, and Vaidehi Jobanputra
Abstract
Whole-exome sequencing (WES) has been used as a standard of care for postnatal diagnosis in the clinical
setting in the past few years for children and adults with undiagnosed disease. Many rare disorders have been
diagnosed through WES, which is less expensive than the traditional serial genetic testing where patients
had previously spent years on an uninformative diagnostic odyssey. Seeking a diagnosis often entails
enduring time consuming, and sometimes invasive procedures which may be associated with medical
risks that are stressful for families and impose a heavy burden on the health-care system. However, the
use of WES is considered impractical in the prenatal and neonatal testing period because of the technical and
computational challenges of performing genomic sequencing from small amounts of genetic material, and
the need for faster turnaround time (TAT) than the current 6–8 weeks TAT provided by most clinical labs
offering postnatal testing. With the rapidly evolving methods of sequence analysis, there are clinical
challenges such as the constantly increasing number of genes being identified which are not yet fully
phenotypically characterized, especially when ascertained prenatally or neonatally before all the clinical
features may be evident. Despite these challenges, there are many clinical benefits to acquiring genomic
information in the prenatal and neonatal period. These include superior prognostic information which
allows for prenatal planning of mode of delivery and hospital for delivery and optimized neonatal manage-
ment. We have developed a clinical WES assay using small amounts of DNA with a TAT of 10 days for use in
the prenatal or neonatal setting. This test is used to detect small nucleotide variants and indels in fetuses and
neonates with structural abnormalities.
Key words Prenatal diagnosis, Whole-exome sequencing, Fetal anomalies
1 Introduction
The risk of major structural birth defects among live births in the
United States is approximately 3% and is associated with inherited
or de novo mutations as well as with maternal factors, such as
advanced age and exposure to teratogens [1, 2]. With advances in
imaging, the ability to detect birth defects prenatally and neonatally
and subsequently use this information to optimize perinatal and
neonatal management has increased tremendously. Simultaneously,
molecular genetic diagnostics have improved the ability to more
267
268 Vanessa Felice et al.
precisely identify the underlying cause of these birth defects and

provide additional prognostic information to improve prenatal and
postnatal management. The first major improvement in genomic
molecular prenatal diagnosis was chromosome microarray analysis
(CMA) [3]. The introduction of prenatal CMA has provided an
invaluable opportunity to evaluate the clinical value of new geno-
mic technologies in prenatal care and to understand the challenges
of interpretation of data in an evolving field.
Generally a combination of karyotype, Fluorescence In Situ
Hybridization (FISH), and CMA are used to investigate the etiol-
ogy of fetal anomaly identified by an ultrasound scan in the second
trimester. Of cases that undergo diagnostic testing, a karyotype
abnormality is found in 8% to 10% of cases, whereas a microdele-
tion/duplication is identified in another 6%, leaving most abnormal
fetuses without a specific genetic diagnosis [3, 4]. Advances in
throughput and decreased cost of next generation sequencing
technology have enabled whole-exome sequencing (WES) which
is ideal for diagnosing disorders that are genetically heterogeneous
and for which gene discovery is incomplete. The use of clinical
exome sequencing for the diagnosis of a wide range of indications
including birth defects has been described in several studies
[5–7]. The diagnostic yield of WES in pediatric patients with
undiagnosed disease is approximately 25% [5], suggesting that it
might complement genetic diagnosis in other settings. A case
report [8] and a study of 30 fetuses and neonates with structural
anomalies identified by ultrasound, illustrate the utility of identify-
ing variants that potentially cause abnormal fetal development
[9]. While these studies performed sequencing on prenatal speci-
mens, the data were not produced within a timeframe that would
have allowed the results to be used until after the birth of the baby.
Since then, additional smaller studies show that prenatal WES,
when the standard genetic testing is normal, can elucidate the
pathogenic variants in approximately 10–50% of fetal structural
anomalies [10–13].
There are many challenges to implementing genomic prenatal
and neonatal testing. These include the technical and computa-
tional challenges of performing genomic sequencing from small
amounts of genetic material, the need for faster turnaround time
(TAT) than the current 6-8 weeks TAT in postnatal cases, and the
rapidly evolving methods of sequence analysis. Clinical challenges
include the constantly increasing number of genes being identified
which are not yet fully phenotypically characterized, especially
when ascertained prenatally or neonatally before all the clinical
features may be evident. Despite these challenges, there are many
clinical benefits with proper utilization of prenatally and neonatally
acquired genomic information including increasingly precise prog-
nostic information allowing prenatal planning of mode of delivery
and hospital for delivery and optimized neonatal management. A
Prenatal Diagnosis by Whole Exome Sequencing in Fetuses with Ultrasound. . . 269
few large studies are currently ongoing to determine the clinical

utility of WES in prenatal setting.
At present, the American College of Medical Genetics and
Genomics (ACMGG) and the Society for Maternal Fetal Medicine
(SMFM) do not recommend routine use of WES for prenatal
diagnosis outside of the context of clinical trials [14]. In select
cases in which other approaches to diagnosis have been uninforma-
tive, it may be appropriate to offer WES. Examples of such cases
include recurrent or multiple congenital anomalies, heterotaxy, and
undiagnosed skeletal dysplasias. Prenatal WES also has a role in
cases in which a fetus has structural abnormalities with reported
consanguinity or homozygosity indicating relatedness on microar-
ray [4]. We describe below our laboratory method for WES with
rapid turnaround time which is emerging as a valuable tool for
genetic diagnosis in the prenatal and neonatal setting.
2 Materials
2.1 DNA Library 1. SureSelectXT Library Prep Kit (Agilent Technologies, Santa
Preparation Clara, CA, USA).
2. SureSelectXT Clinical Research Exome V2 (Agilent Technolo-
gies, Santa Clara, CA, USA).
3. Agencourt AMPure XP Kit (Beckman Coulter, Indianapolis,
IN).
4. Dynabeads MyOne Streptavidin T1 (Thermo Fisher Scientific,
Waltham, MA).
5. Ethanol, 200 proof for molecular biology.
6. Sterile, nuclease-free water.
7. DNA 1000 CHiP Kit (Agilent Technologies, Santa Clara, CA,
USA).
8. Dynal DynaMag-2 magnetic stand (Thermo Fisher Scientific,
Waltham, MA).
9. Covaris microTUBE plate (Covaris, Woburn, MA).
10. 96-Well Deep Well Plates.
11. 96-Well skirted PCR plates.
2.2 Illumina DNA 1. HiSeq Paired End Flow Cell v4 (Illumina, Inc., San Diego, CA,
Sequencing USA).
2. HiSeq Paired End Cluster Kit v4 (Illumina, Inc., San Diego,
CA, USA).
3. HiSeq Rapid Paired End Flow Cell v2 (Illumina, Inc., San
Diego, CA, USA).
4. cBot Manifold (Illumina, Inc., San Diego, CA, USA).
5. 1.0 N Sodium Hydroxide, JT Baker 200 mM Tris–HCl,

pH 8.0.
6. 200 mM Tris–HCl, pH 8.0.
7. Laboratory-grade water.
2.3 Laboratory 1. Thermo Mixer C 5382 (Eppendorf, Hauppauge, New York,

Equipment USA).
2. Agilent Technologies 2100 Bioanalyzer (Agilent Technologies,
3. PCR Thermal cycler.
4. Covaris LE-220 Sample Preparation System (Covaris, Woburn,
MA).
5. Vacuum DNA concentrator.
6. Microsample Incubator.
7. High Speed Microplate Shaker.
8. HiSeq 2500 (Illumina, Inc., San Diego, CA, USA).
9. cBot (Illumina, Inc., San Diego, CA, USA).
3 Methods
The protocols described in this chapter require high molecular

weight DNA of good quality. Any commercial DNA extraction kit
may be utilized (see Chapter 7). In addition, maternal cell contami-
nation studies should be performed (see Chapter 8) to ensure that
WES results accurately reflect the genomic status of the fetus.
3.1 DNA Library The library preparation process can be completed within 48 hours.
Preparation This process includes shearing genomic DNA, repairing the ends of
the fragments, adding an A-base to the 30 ends, ligating Illumina
adapters, and amplifying the DNA. The exome is then captured
through a hybridization to RNA baits and enriched with PCR to
prepare the samples for sequencing on Illumina next generation
sequencing instruments.
3.1.1 DNA Shearing The shearing process uses sonication to break up DNA into smaller
fragments which will be selected out based on size in the
subsequent steps.
1. Normalize 1500 ng of DNA in 130 μL of nuclease-free water
and transfer to a Covaris plate.
2. Cover the plate with Covaris foil tape and spin down briefly (see
Note 1).
3. Turn on the Covaris LE220 instrument.
4. Chill the water to 7 C and degas the instrument for at least

45 min prior to starting fragmentation.
5. Select load Samples to bring the sample tray to the front of the
instrument.
6. When the green “Open” button is illuminated, press the but-
ton and gently pull on the handle to open the instrument door.
7. Place the Covaris plate containing the samples to be sheared
onto the sample platform.
8. Close the door and select the following conditions:
Time ¼ 380 s, Duty Cycle ¼ 30%, PIP ¼ 450, Cycles Per
Burst ¼ 200.
9. Ensure that the positions of the samples on the software setup
page are correct and press Start to begin the fragmentation.
10. Once the fragmentation is complete, open the door and
remove the plate.
11. Transfer the sheared samples from the Covaris plate to a new
96-well plate.
12. Be sure the AMPure beads have been removed from the refrig-
erator and have acclimated to room temperature for at least
30 min prior to proceeding.
13. Make 80% Ethanol just prior to starting this procedure.
14. Vortex the AMPure XP beads until they are well dispersed.
15. Add 180 μL of the AMPure XP beads to each well of the plate
containing sample and pipette up and down 15 times.
16. Incubate the plate at room temperature for 10 min.
17. Place the plate on the appropriate magnetic stand and let it
incubate at room temperature for an additional 5 min.
18. Remove and discard all the solution of the supernatant from
each well.
19. With the plate still on the magnetic stand, add 200 μL of
freshly prepared 80% ethanol to each well without disturbing
the beads.
20. Incubate the plate at room temperature for 30 s, then remove
and discard all the supernatant from each well.
21. Repeat steps 19 and 20 once for a total of two ethanol washes.
22. Following the last ethanol wash, use a p10 to remove any
residual ethanol from the bottom of the wells while keeping
the plate on the magnetic stand.
23. Keep the plate on the magnetic stand for no more than 5 min
to dry the beads (see Note 2).
24. Remove the plate from the magnet and add 52 μL of nuclease-
free water to resuspend the beads by gently pipetting up and
down ten times.
25. Incubate the plate with the resuspended samples for 2 min at
room temperature.
26. Place the plate on the magnetic stand and let the plate incubate
at room temperature for an additional 5 min.
27. With the plate still on the magnetic stand, transfer 48 μL of the
supernatant to a new plate.
3.1.2 End Repair Following shearing, the ends of the DNA are of varied lengths and
have 30 and 50 overhangs. For adapter ligation to be efficient, the
overhands must be repaired. The end repair step extends the over-
hangs to create blunt-ended DNA fragments.
1. Create an End repair master mix using the guidelines in
Table 1.
2. Add 52 μL of End Repair/A-Tail Mix to each sample of the
plate and pipette up and down to mix.
3. Seal the plate and spin down briefly.
4. Place the plate in the thermal cycler and run for 30 min at
20 C.
5. Proceed immediately to the next step.
3.1.3 AMPure Bead 1. Be sure the AMPure beads have been removed from the refrig-
Cleanup erator and have been acclimated to room temperature for at
least 30 min prior to proceeding.
2. Make 80% Ethanol just prior to starting this procedure, unless
it was already prepared during the DNA shearing cleanup.
3. Vortex the AMPure XP beads until they are well dispersed.
4. Add 180 μL of the AMPure XP beads to each well of the plate
containing sample and pipette up and down to mix.
Table 1
End repair master mix setup
Reagent Vol. for 1 library (μL) Vol. for N library (μL)

Nuclease-free water 35.2 (N + 1) 35.2
10 end repair buffer 10 (N + 1) 10
dNTP mix 1.6 (N + 1) 1.6
T4 DNA polymerase 1.0 (N + 1) 1.0
Klenow DNA polymerase 2.0 (N + 1) 2.0
T4 polynucleotide kinase 2.2 (N + 1) 2.2
Total volume 52.0 (N + 1) 52.0

7. Remove and discard all the solution of the supernatant from
each well without disturbing the beads.
8. Use a p10 multichannel pipette to ensure that as much of the
solution is removed as possible.
9. With the plate on the magnetic stand, add 200 μL of freshly
prepared 80% ethanol to each well without disturbing the
beads.
11. Be careful not to disturb the beads.
the wells on the magnetic stand.
14. Keep the plate on the magnetic stand for 5 min to dry.
15. Add 32 μL of nuclease-free water to each well and resuspend
the beads by pipetting up and down.
room temperature.
18. With the plate on the magnetic stand, transfer 30 μL of the
supernatant from the plate to a new PCR plate and proceed to
the next steps.
3.1.4 Adenylation The adapter ligation requires the presence of a 30 A-base on the
of the 30 Ends double stranded DNA fragments. The adenylation step uses dATPs
and Exo() Klenow to adenylate the DNA fragments
1. Create an A-tail master mix using the guidelines in Table 2.
2. Obtain the post end repair plate from the thermocycler and add
20 μL of the master mix to each sample.
3. Incubate the plate for 30 min at 37 C.
3.1.5 AMPure Bead 1. Vortex the AMPure XP beads until they are well dispersed.
Cleanup 2. Add 90 μL of the AMPure XP beads to each sample.
5. Remove and discard all of the supernatant from each well.
Table 2
A-tail master mix setup
Reagent Vol. for 1 library (μL) Vol. for N libraries (μL)

10 Klenow polymerase buffer 5.0 (N + 1) 5.0
dATP 1.0 (N + 1) 1.0
Exo() Klenow 3.0 (N + 1) 3.0
6. With the plate still on the magnetic stand, add 200 μL of freshly
beads.
10. Keep the plate on the magnetic stand for not more than 5 min
to dry the beads.
11. Add 15 μL of nuclease-free water to each well of the plate using
a multichannel pipette. Resuspend the beads gently by pipet-
ting 10 times.
room temperature.
supernatant to a new PCR plate and proceed immediately to
the next step.
3.1.6 Adapter Ligation 1. Create a master mix using the guidelines in Table 3.
2. Add 37 μL of Adapter Ligation Mix to each sample of the plate
and pipette up and down to mix.
4. Place the plate in the thermal cycler and run for 15 min at
20 C.
Table 3
Adapter ligation master mix setup

5 T4 DNA ligase 10.0 (N + 1) 10.0
SureSelect adapter oligo mix 10.0 (N + 1) 10.0
T4 DNA ligase 1.5 (N + 1) 1.5
5. Remove and discard all of the solution of the supernatant from
each well.
beads.
to dry the beads.
ting ten times.
room temperature.
supernatant to a new PCR plate and proceed immediately to
the next step.
Table 4
DNA enrichment master mix setup

SureSelect primer 1.25 (N + 1) 1.25
ILM indexing PCR reverse primer 1.25 (N + 1) 1.25
5 Herculase II reation buffer 10.0 (N + 1) 10.0
100 nM dNTP 0.5 (N + 1) 0.5
Herculase II fusion DNA polymerase 1.0 (N + 1) 1.0
3.1.8 PCR Enrichment 1. Create a PCR master mix for the samples using the guidelines
in Table 4.
2. Pipette 35 μL of PCR master mix to the wells of a new PCR
plate.
3. Add 15 μL of ligated DNA and mix by pipetting up and down
at least ten times.
5. Place the plate in the thermal cycler and run the Enrichment
program as follows:
(b) Six cycles: 98 C for 30 s, 65 C for 30 s, 72 C for 1 min.
each well.
beads.

to dry the beads.
ting ten times.
room temperature.
supernatant from the plate to a new PCR plate.
15. Run the whole genome libraries on the Agilent 2100 Bioana-
lyzer using the DNA 1000 chip.
16. Libraries that pass QC have a concentration >25 ng/μL and a
library size >200 bp with and average peak >400 bp.
3.1.10 Capture 1. Normalize 750 ng of library in 20 μL of nuclease-free water in a

Hybridization PCR plate.
2. Place the plate in a speed vacuum on medium heat until the
samples have dried down completely.
3. While the samples are drying down, create three individual
master mixes: Hybridization Mix, Capture Mix, and
Block Mix.
4. Create a Capture master mix for the samples using the guide-
lines in Table 5 and keep the master mix on ice until ready
to use.
5. Create a Hybridization master mix using the guidelines in
Table 6 and keep at room temperature until ready touse.
6. Create a Blocking master mix using the guidelines in Table 7
and keep and keep on ice until ready to use.
7. After the samples have dried down, add 3.4 μL of nuclease-free
water and resuspend the samples.
8. Add 5.6 μL of the Block master mix to each sample and mix by
pipetting up and down.
9. Seal the plate and run the hybridization program as follows:
(b) 65 C hold.
10. While the thermal cycler is cooling down to the 65 C hold,
prepare the capture/hybridization mix by combing the capture
and hybridization master mixes as shown in Table 8.
Table 5
Capture master mix setup

SureSelect capture reagent 5 (N + 1) 5.0
SureSelect RNAse block 0.5 (N + 1) 0.5
Table 6
Hybridization master mix setup

SureSelect Hyb #1 6.63 (N + 1) 6.63
Table 7
Blocking master mix setup

Indexing block #1 2.5 (N + 1) 2.5
Block #2 2.5 (N + 1) 2.5
Indexing block #3 0.6 (N + 1) 0.6
Table 8
Capture/hybridization master mix setup

Capture mix 7 (N + 1) 7
Hybridization mix 13 (N + 1) 13
11. Once the cycler has reached 65 C, keep the plate on and
remove the seal.
12. With the plate on the thermal cycler, quickly add 20 μL of the
capture/hybridization mix to all the samples and pipette up
and down several times.
13. Seal the plate and incubate at 65 C for 18–24 h.
3.1.11 Preparation 1. Remove a vial of DynaBeads MyOne Streptavidin T1 beads

of Streptavidin T1 Magnetic from the 4 C refrigerator and vortex vigorously to resuspend
Beads the beads in their storage buffer (see Note 3).
2. Obtain a new deep well plate and label it with “Wash Buffer 2.”
3. Add 280 μL of SureSelect Wash Buffer 2 to six wells of the
“Wash Buffer 2” plate for each sample being processed.
4. Seal the plate and place in the Hybex incubator set at 65 C.
Incubate at 65 C until needed.
5. Aliquot 50 μL of Dynabeads to each well of a deep well plate
that will contain sample.
6. Add 200 μL of SureSelect Binding Buffer to each well contain-
ing beads.
7. Mix the beads by pipetting up and down at least ten times.
8. Place the plate on a magnetic block and allow the beads to
migrate to the magnet for at least 5 min.
9. Remove and discard the supernatant without disturbing the
beads.
10. Repeat steps 6–10 two times, for a total of three washes.
11. Add 200 μL of SureSelect Binding Buffer to each of the wells.
Resuspend the beads by pipetting up and down.
12. The beads are now washed and ready to use.
3.1.12 Wash and Target 1. With the thermal cycler still holding at 65 C, open the lid and
Capture remove the seal.
2. Transfer the hybridization mixture to the associated wells in the
deep well plate containing the washed beads.
3. Place the sealed plate on a plate shaker set on low speed for
30 min.
4. Following incubation, spin the plate briefly in a centrifuge.
5. Place the plate on a magnetic block and allow beads to migrate
for 5 min.
6. Once the beads have bound to the magnet and the supernatant
is clear, remove the supernatant and discard.
7. Remove the plate from the magnetic block and resuspend the
beads in 500 μL of SureSelect Wash Buffer 1 by pipetting up
and down.
8. Seal the plate and plate the place on a plate shaker.

9. Mix the samples on the plate shaker for 15 min at room
temperature.
10. After the incubation, briefly spin the plate down.
11. Place the plate on a magnetic block for 5 min while the beads
migrate to the magnet.
12. Once the beads have bound to the side of the well and the
solution appears clear, remove the supernatant and discard.
13. Remove the plate from the magnetic block.
14. Resuspend the beads in 500 μL of the 65 C pre-warmed
SureSelect Wash Buffer 2.
15. Mix by pipetting up and down at least five times (see Note 4).
16. Seal the plate and incubate the samples for 10 min at 65 C.
17. Following the incubation, briefly spin the plate down in a
centrifuge.
18. Place the plate on the magnetic block and allow the beads to
migrate for 5 min.
19. Once the beads have bound to the sides of the well, remove and
discard the clear supernatant. Remove the plate from the mag-
netic block.
20. Repeat steps 14–19 twice, for a total of three washes.
21. After the final wash has been completed, use a 10 μL pipette to
make sure all the wash buffer has been removed.
22. Resuspend the beads in 30 μL of nuclease-free water.
23. The hybridized libraries are now bound to the Dynabeads.
3.1.13 DNA Fragment 1. Create a PCR master mix for the samples using the guidelines
Enrichment in Table 9.
2. Pipette 35 μL of PCR master mix to the wells of a new PCR
plate for each sample needed.
3. Add 14 μL of the library on bead to each well.
4. Add 1 μL of the index to each sample and mix by pipetting up
and down (see Note 5).
6. Place the plate in the thermal cycler and run the Enrichment
program as follows:
(b) Ten cycles: 98 C for 30 s, 57 C for 30 s, 72 C for 1 min.
7. Proceed immediately to next step.
Table 9
DNA enrichment master mix setup

ILM post capture forward primer 1.0 (N + 1) 1.0
5 Herculase II reation buffer 10.0 (N + 1) 10.0
100 nM dNTP 0.5 (N + 1) 0.5
Herculase II fusion DNA polymerase 1.0 (N + 1) 1.0
each well.
beads.
to dry the beads.
ting ten times.
room temperature.
supernatant from the plate to a new PCR plate.
15. Run the final libraries on the Agilent 2100 Bioanalyzer using
the DNA 1000 chip.
16. Final libraries that pass QC have a concentration >2 ng/μL
and a library size >200 bp with and average peak >400 bp.
3.2 DNA Sequencing The final DNA library is diluted, denatured, and introduced into
on the Illumina HiSeq the lanes of the flow cell using the cBot according to the manufac-
2500 tures protocol. The libraries are loaded at a coverage of 100 for
the proband sample and 60 for the parent samples. The DNA
library templates are captured by the oligonucleotides that are
affixed to the surface of the flow cell. Templates bound to the
oligonucleotides on the flow cell are 30 extended, producing
covalently-attached discrete single molecules. The double-stranded
molecule is denatured, and the original template is washed away.
The free ends of the bound templates hybridize to the adjacent
lawn primers to form U-shaped bridges. The DNA bridge is then
copied from the primer to create a double-stranded DNA bridge.
The resulting dsDNA is denatured, hybridized to lawn-primers to
form new bridges and extended again. This process of iso-thermal
bridge amplification is repeated 35 times to create a dense cluster of
over 2000 molecules. The reverse strands in the cluster are removed
by cleavage at the reverse strand-specific lawn primers, leaving a
cluster with forward strands only. The free 30 -OH ends are blocked
to prevent nonspecific priming. Sequencing primers are hybridized
to the free ends of the DNA templates. The flow cell is now ready to
be sequenced on the Illumina HiSeq 2500 and is loaded onto the
sequencer according to the manufacturer’s protocol (see Note 6).
3.3 Data Processing 1. Demultiplexing: Once the sequencing chemistry is complete

the raw sequencing data is written in binary base call (BCL)
format. Illumina’s bcl2fastq software is used to demultiplex the
raw sequencing data and convert it into standard FASTQ for-
mat for downstream analysis.
2. Alignment: Individual sequencing reads in the FASTQ files are
mapped to reference human genome using Borrows-Wheeler
Aligner (BWA) [15]. This process generates a BAM format file
which is a compressed binary representation of aligned reads in
Sequence Alignment Map (SAM) format [16].
3. Duplicate marking: Duplicate reads are defined as sequence
reads originating from a single fragment of DNA. They can
arise due to PCR-based library preparation or as a result of
single amplification clusters on the flowcell being incorrectly
detected as multiple clusters by the sequencing instrument
optics. The BAM file generated in step 2 is processed through
Picard tools [http://broadinstitute.github.io/picard] which
locates and tags duplicate reads.
4. Base Quality Score Recalibration (BQSR): Base quality scores
are important quality indicators and are used by variant calling
algorithms for accuracy. Multiple factors like library prepara-
tion and sequencing instrumentation can introduce biases in
the sequencer-assigned base qualities. To correct any systematic

bias observed in the data, the duplicate marked BAM file is
processed using BaseRecalibrator tool of the Genome Analysis
Toolkit (GATK) [17]. At this stage the aligned BAM file is
ready for variant discovery.
5. Variant discovery: Identifying short variants (Single Nucleotide
Variants—SNVs and insertion/deletions—Indels) is typically a
two-step process. GATK HaplotypeCaller tool is run on each
sample separately in GVCF mode for scalable variant calling.
This produces an intermediate file format called gVCF (for
genomic VCF). GVCFs of multiple samples are then run
through a joint genotyping step to produce a multi-sample
VCF using GATK GenotypeGVCFs tool. Variant quality
score recalibration (VQSR) using GATK VariantRecalibrator
is then performed to filter low quality variants.
6. Variant annotation: To simplify and accelerate variant prioriti-
zation, the filtered VCF is processed through Ensembl Variant
Effect Predictor (VEP) [18]. Relevant variant-level and gene-
level annotations are added to each variant in this step. These
annotations include, but are not limited to, variant conse-
quence (missense, nonsense, frameshift, etc.), allele frequency
in gnomAD database [http://gnomad.broadinstitute.org/
about], in-silico damaging predictions from SIFT [19] and
PolyPhen [20], association with human phenotypes from
OMIM [https://omim.org] and previously reported variant
clinical significance from ClinVar [21].
7. Variant prioritization: To identify candidate function-
impacting variant(s) the annotations are used to perform
step-wise variant filteration. The variants are first filtered
based on the allele frequency in the population (gnomAD).
Typically, for rare disorders any variant observed at a frequency
of 1% or above is filtered out. It should be noted that this
frequency cutoff should be determined based on the pheno-
type being investigated and ancestry of the case. When parental
genotypes are available variants are further filtered based on the
expected mode of inheritance. For example, if recessive mode
of inheritance is being considered for a fully penetrant pheno-
type and the parents are unaffected, only homozygous variants
in the proband are retained with both parents heterozygous for
the variant allele. Similar filtration is done for other modes of
inheritance. At this stage, retained variants and associated genes
are manually evaluated for association with the phenotype
being investigated using information from OMIM, ClinVar,
and PubMed. Once variants possibly associated with the phe-
notype are identified, their clinical significance is assessed using
ACMG standards and guidelines for interpretation of sequence
variants [22]. According to the assessment the variants are
categorized into the following categories—Benign, Likely

Benign, Uncertain Significance, Likely Pathogenic, and
Pathogenic.
8. Pathogenic and likely pathogenic reportable variants may be
confirmed by using standard Sanger sequencing.
4 Notes
1. Visually check each sample well to ensure that there are no air
bubbles present prior to shearing and the metal rod is in the
center of the Covaris tube. Air bubbles can cause variable
shearing. If air bubbles are present, briefly centrifuge the plate
and then check again.
2. Over drying the beads can lead to significant sample loss.
3. This step should be performed not more than 1 h prior to
removal of the Hybridization plate from the thermal cycler.
The Hybridization plate should remain incubating at 65 C
during this process.
4. When mixing the beads with the buffer incubated at 65 C, this
should be done as quickly as possible. If the temperature drops
much below 65 C, nonspecific binding can occur.
5. The list of Agilent adapters can be obtained at: https://www.
agilent.com/cs/library/usermanuals/Public/G7530-90000.
pdf
6. Introduction to Next Generation Sequencing technology can
be found at: https://www.illumina.com/content/dam/
illumina-marketing/documents/products/illumina_sequenc
ing_introduction.pdf
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Chapter 19
Isolation and Characterization of Amniotic Fluid-Derived

Extracellular Vesicles for Biomarker Discovery
Blake Ebert and Alex J. Rai
Abstract
Amniotic fluid, the fetal-protective liquid that fills the amniotic sac, represents a rich source of biomarkers.
The diagnostic utility of amniotic fluid relies on the highly abundant maternal and fetal nucleic acid and
proteomic content, which allows for the simultaneous determination of mother and fetal health status.
Extracellular vesicles (ECVs) that are released by all cells and found in amniotic fluid could be harnessed to
provide substantial clinically actionable data. ECVs are mediators of critical biological functions and reflect
the health of the parent cell. Thus, ECVs released from cells in distress may provide important diagnostic
information. Here, we describe a straightforward and optimized method for isolating ECVs from amniotic
fluid. In addition, we validate our procedure through western blotting using antibodies targeting canonical
ECV protein markers and via direct visualization using transmission electron microscopy.
Key words Amniotic fluid, Apoptotic vesicles, Exosomes, Extracellular vesicles, Microvesicles,
Biomarkers
1 Introduction
Amniotic fluid (AF) is a dynamic and complex biological fluid

comprised of fetal secretions from the respiratory and excretory
systems, amniocytes, and maternal plasma [1]. AF changes in vol-
ume and composition throughout pregnancy in response to fetal
development and is crucial for the healthy development of the fetus
[1]. The embryo-containing amniotic sac forms 12 days after con-
ception and is immediately filled with AF [2]. At this point, AF is
composed primarily of water derived from the maternal plasma. By
12–14 weeks, fetal urine is the most abundant component. From
14–32 weeks, volume steadily increases to a maximum of approxi-
mately 1000 mL [2].
Changes in the AF cells, nucleic acids, proteins and extracellular
vesicles may reflect fetal abnormalities [2]. In current clinical prac-
tice, AF status is assessed in vivo via ultrasound for detection of fetal
malformations and is typically collected in the second trimester via
287
288 Blake Ebert and Alex J. Rai
amniocentesis to assess certain pregnancy-related complications in

high-risk mothers, including chromosomal abnormalities, fetal
infection, and encephalopathies [3]. While the ultimate goal is to
eliminate the need for amniocentesis by noninvasive fetal testing
through maternal serum or urine, AF as a diagnostic resource has
many benefits [4]. Namely, AF contains a higher abundance of cell-
free fetal/maternal DNA, RNA, and proteins than maternal serum
and urine [4]. In addition, AF reflects the health of both the
mother and the fetus simultaneously [4]. Taken together, these
findings suggest the utility of AF as a diagnostic tool for a wide
spectrum of diseases and abnormalities.
In particular, AF-derived extracellular vesicles (ECVs) may rep-
resent a rich trove of biomarkers. Released from all cells and found
ubiquitously in every biological fluid, ECVs carry genetic material
and proteins throughout the body that reflect the original parent
cell [5]. ECVs are primarily classified by size, although there are
discrepancies in the literature with regard to size limits. In general,
exosomes are the smallest, ranging from ~50–150 nm; microvesi-
cles are larger, ranging up to 1000 nm, and apoptotic vesicles are
the largest of the three, and can be up to 5000 nm. In recent years,
evidence has accumulated suggesting that ECVs, and exosomes
specifically, may be important regulators of key biological functions
[5]. Thus, in an abnormal state, an assessment of vesicular contents
may yield valuable clinically actionable information.
ECV-based biomarkers in amniotic fluid have the potential to
significantly improve clinical risk assessment and maternal/fetal
patient management [4]. Although AF is collected invasively
through amniocentesis, the high concentration of cell-free fetal
and maternal DNA, RNA, and proteins in AF relative to maternal
circulation suggests its utility in discovery-based analysis. We report
here an optimized methodology for isolating AF-derived ECVs and
validate this procedure via western blotting and negative staining
based transmission electron microscopy.
2 Materials
2.1 ECV Isolation 1. Centrifuge capable of 20,000 g speed.

2. D-Dithiothreitol (DTT) (100 mg/mL in ddH2O).
2.2 SDS-PAGE 1. 2 LDS-reducing agent buffer.

and Western Blotting (a) 4 LDS Stock sample buffer (Thermo Fisher Scientific,
Carlsbad, CA).
(b) 10 stock sample reducing agent (Thermo Fisher Scien-
tific, Carlsbad, CA).
Prepare by adding 500 μL of 4 LDS Stock sample buffer,
200 μL 10 stock sample reducing agent, 300 μL ddH2O.
Isolation and Characterization of Amniotic Fluid-Derived Extracellular. . . 289
2. 4–12% Bis-Tris 15 well gel (Thermo Fisher Scientific, Carlsbad,

CA).
3. 1 MES SDS running buffer: Prepare by adding 50 mL 20
MES and 950 mL ddH2O.
4. iBlot™ Transfer Stack, nitrocellulose, regular size (Thermo
Fisher Scientific, Carlsbad, CA).
5. Ponceau S solution (Sigma Aldrich, St. Louis, MO). 0.1% in 5%
acetic acid.
6. 1 PBS-Tween 20 (PBS-T): Prepare by adding 50 mL 20
PBS-T and 950 mL ddH2O.
7. 5% milk in PBS-T: Prepare by dissolving 5 g nonfat dry milk in
100 mL PBS-T.
8. SuperSignal™ West Pico PLUS Chemiluminescent Substrate
(Thermo Fisher Scientific, Carlsbad, CA): Contains Super-
Signal west pico and luminol and peroxide chemiluminescence
solutions.
9. XCell SureLock Mini-cell (Thermo Fisher Scientific, Carlsbad,
CA).
10. 600 V Power Supply.
11. iBlot™ 2 Gel Transfer Device (Thermo Fisher Scientific, Carls-
bad, CA).
2.3 Primary 1. PDCD6IP Antibody—C-terminal region (ARP76945_P050)

Antibodies (Aviva Systems Biology, San Diego, CA).
2. CD81 Antibody—C-terminal region (ARP63231_P050)
(Aviva Systems Biology, San Diego, CA).
3. CD9 antibody—N-terminal region (ARP61171_P050) (Aviva
Systems Biology, San Diego, CA).
4. HSP90B1 antibody—C-terminal region (ARP40463_P050)
5. ACTN4 antibody—N-terminal region (ARP42202_T100)
6. Purified Mouse Anti-Flotillin-1 (Clone 18/Flotillin-1)
(BD Biosciences, San Jose, CA).
2.4 Secondary 1. Anti-rabbit IgG, HRP-linked antibody (Cell Signaling Tech-

Antibodies nology, Danvers MA): 1:5000 in 5% milk
2. Anti-mouse IgG, HRP-linked antibody (Cell Signaling Tech-
nology, Danvers MA): 1:5000 in 5% milk.
2.5 Transmission 1. 2% paraformaldehyde (PFA) in 0.1 M PBS.

Electron Microscopy, 2. Formvar-carbon-coated EM grids (Electron Microscopy
Negative Staining Sciences, Hatfield, PA).
3. Uranyl-oxalate, pH 7 (prepare from stock solution of 4% uranyl

acetate in ddH2O and 0.15 M oxalic acid, brought to pH 7 by
titrating with 25% NH4OH).
4. JEM-1200 EXII transmission electron microscope (JEOL
Ltd., Tokyo, Japan) (see Note 1).
3 Method
The method described herein is straightforward, inexpensive, and

does not require complex equipment. These characteristics allow
for its incorporation into a standard operating procedure and use in
a CLIA-certified clinical laboratory setting. The first two steps
involve low-speed centrifugation spins to remove dead cells and
cellular debris. Successful isolation requires the addition of dithio-
threitol (DTT), which removes high-abundance components and
allows for the detection of lower-abundance proteins. See Fig. 1 for
protocol details.
Fig. 1 Schematic diagram of ECV isolation from amniotic fluid. Starting material
is subjected to two centrifugation spins. The pellet is subsequently treated with
DTT and centrifuged again at 20,000 g. The final pellet can then be subjected
to downstream analyses
3.1 Purification 1. Centrifuge amniotic fluid sample at 3000 g for 10 min at

of ECVs 4 C (see Note 2).
2. Transfer supernatant to fresh Eppendorf tubes.
3. Centrifuge sample at 20,000 g for 20 min at 4 C.
4. Decant and discard supernatant and add 50 μL D-Dithiothrei-
tol (see Notes 3 and 4).
5. Pipette up and down to resuspend the pellet and centrifuge at
20,000 g for 20 min at 4 C.
6. Decant and discard supernatant. Store pellet at 80 C until
ready for further processing.
3.2 Preparation 1. Prepare fresh working 2 LDS-reducing agent sample buffer.

of ECVs and SDS-PAGE 2. Resuspend pellet in 30 μL 2 LDS-reducing agent sample
Analysis buffer.
3. Boil samples on heat block at 95 C for 3 min.
4. Cool immediately on ice.
5. Load 15 μL of sample in 4–12% Bis-Tris gel secured in XCell
Sure Lock electrophoresis cell, and run at 150 mV for 1 h or
until dye front reaches the bottom of the gel.
6. Transfer gel to nitrocellulose membrane using the iBlot Trans-
fer Device.
3.3 Western Blotting 1. Incubate the nitrocellulose membrane in Ponceau S stain on

After Gel the rocker for 5 min at room temperature.
Electrophoresis 2. Decant stain, rinse several times with dH2O, and acquire an
image of the stained blot.
3. De-stain in PBS-T on the rocker for 5 min at room
temperature.
4. Prepare primary antibody in 5% milk/PBS-T at 1:1000 dilution
(see Note 5).
5. Incubate membrane in primary antibody solution overnight at
4 C.
6. Wash three times in PBS-T for 5 min each.
7. Prepare fresh HRP-conjugated secondary antibody in 5%
milk/PBS-T at 1:5000 dilution.
8. Incubate membrane in secondary antibody solution for 45 min
at room temperature.
9. Wash three times in PBS-T for 5 min each, on rocker.
10. Add 1 mL of each of the chemiluminescent substrates (Super-
Signal west pico and luminol and peroxide chemiluminescence
solutions), being careful to use separate pipette tips for each.
The solution is activated immediately after mixing.
11. Process image on chemiluminescent developer. Representative
western blot results are shown in Figs. 2 and 3.
1 2 3 4 5 6
260
140
1: anti-CD9
100 2: anti-CD81
70 3: anti-ALIX
4: anti-hsp90b1
50
5: anti-ACTN4
40 6: anti-FLOT1
35
25
Fig. 2 Antibody optimization. Primary antibodies (Aviva Systems Biology, San Diego, CA) were tested at various
dilutions using cultured melanoma cells (92.1) whole cell lysate. For each antibody, a 1:500 and 1:1000
dilution were tested (shown above); 1:1000 was deemed to be optimal
Fig. 3 Validation of ECV isolation procedure by western blotting using antibodies to canonical ECV protein
markers, see panel on RIGHT. Multiple canonical protein markers for extracellular vesicles were detected in
ECV fractions from AF including: CD9, CD81, ALIX, hsp90β1, ACTN4, and flotillin-1. Ponceau S staining reveals
the requirement for DTT treatment to remove high-abundance proteins from the ECV pellet in order to visualize
lower-abundance proteins and increase ECV yield (compare lanes 3, 4, and 5 in Ponceau S Stain). Lanes on
gel correspond to steps from ECV isolation protocol, as denoted on left panel
Fig. 4 Transmission electron microscopy imaging of amniotic fluid-derived ECVs. Samples were prepared for
negative staining and were viewed at 25,000 magnification (left panel). We viewed an individual exosome
and microvesicle at 200,000 magnification (right panel). The inner diameter of exosome is 123 nm in
diameter and that of the microvesicle is 180 nm
3.4 Transmission 1. Resuspend ECV pellet in 2% PFA in 0.1 M sodium phosphate

Electron Microscopy buffer.
2. Deposit 10 μL of sample onto Formvar-carbon-coated EM
grids and let sit for 20 min at room temperature.
3. Blot off sample with filter paper, being careful to not
completely dry out the grid.
4. Add 20 μL uranyl-oxalate solution and leave at room tempera-
ture for 5 min.
5. Remove stain with filter paper and let grids dry at room
temperature.
6. Examine grids using transmission electron microscopy (see
Fig. 4).
4 Notes
1. Any transmission electron microscope can be used. Our expe-

rience is with the JEM-1200 EXII transmission electron
microscope.
2. We have used 1 mL starting volume of amniotic fluid sample
for the exosome isolation and analysis, as described above
(results shown in Fig. 3). However, we note that results can
be obtained (and confirmed by western blotting and transmis-
sion electron microscopy) using <200 μL amniotic fluid sam-
ple, once conditions are optimized.
3. Pellets are usually light yellow in color, but some samples may
not have visible pellets. We typically add 100 μL DTT to
resuspend. For smaller pellets and/or smaller volumes, we
suggest using lower volumes, e.g., 25–50 μL DTT, and test
sample by western blotting.
4. If starting sample volume is high, sample may be split into

multiple tubes for convenience (this may also be necessitated
by rotor size and maximum volume/number of tubes). In such
cases, final pellets from multiple tubes can be combined into
one tube to increase ECV recovery.
5. Antibodies should be optimized by testing at various dilutions
and conditions using the appropriate sample of interest. Rou-
tinely, we test all antibodies initially using cell lysate, and once
optimal western blotting conditions are delineated, we subse-
quently test for the presence of target proteins using experi-
mental samples of interest (see Fig. 2).
References
1. Palmas F, Fattuoni C, Noto A et al (2016) The Obstet Gynecol Clin N Am 42(2):193–208.
choice of amniotic fluid in metabolomics for the https://doi.org/10.1016/j.ogc.2015.01.011
monitoring of fetus health. Expert Rev Mol 4. Kamath-Rayne BD, Smith HC, Muglia LJ et al
Diagn 16(4):473–486. https://doi.org/10. (2014) Amniotic fluid: the use of high-
1586/14737159.2016.1139456 dimensional biology to understand fetal well-
2. Kolialexi A, Tounta G, Mavrou A et al (2011) being. Reprod Sci 21(1):6–19. https://doi.
Proteomic analysis of amniotic fluid for the diag- org/10.1177/1933719113485292
nosis of fetal aneuploidies. Expert Rev Proteo- 5. Urbanelli L, Buratta S, Sagini K et al (2015)
mics 8(2):175–185. https://doi.org/10.1586/ Exosome-based strategies for Diagnosis and
epr.10.112 Therapy. Recent Pat CNS Drug Discov 10
3. Evans MI, Andriole S, Evans SM (2015) Genet- (1):10–27
ics: update on prenatal screening and diagnosis.
Part IV
Non-Invasive Prenatal Testing

Chapter 20
Quad Screen Test, A Multiplexed Biomarker Assay

for Prenatal Screening to Assess Birth Defects: The
Columbia University Experience Using the Beckman
Access2 Immunoassay Analyzer and Benetech PRA
Awet Tecleab, Alex K. Lyashchenko, and Alex J. Rai
Abstract
In the prenatal quad screen, the levels of four analytes in maternal serum are used to calculate the risk of
serious birth defects. The Beckman Access2 Immunoassay System is an automated analyzer that enables
rapid measurement of alpha-fetoprotein, unconjugated estriol, human chorionic gonadotropin, and
dimeric inhibin A. The Benetech PRA software package is used to convert maternal serum analyte
concentrations to multiples of the median (MoM) and calculates the risks of particular birth defects. The
results from this simple and minimally invasive screen determine the need for more sensitive, specific, and
usually riskier diagnostic procedures. We present herein some recent data from our experience at Columbia
University Medical Center in New York, NY, using the Beckman Access2 immunoassay analyzer and
Benetech PRA software package.
Key words Fetal defect markers, Prenatal diagnosis, Maternal screening, Quad screen
1 Introduction
Prenatal screening tests commonly use maternal blood or amniotic

fluid for the detection of pregnancies with high risk of serious birth
defects. The quad screen test measures the maternal serum levels of
four biomarkers: alpha-fetoprotein (AFP), unconjugated estriol
(uE3), human chorionic gonadotropin (β-hCG), and dimeric
inhibin A (DIA). It is performed in the second trimester, with
optimal sample collection time between 14 and 22 weeks [1], and
is approved for screening of trisomy 18, trisomy 21 (Down syn-
drome), and open neural tube defects. It has a sensitivity of 80% for
the detection of Down syndrome at a 5% false positive rate [2, 3].
Awet Tecleab and Alex K. Lyashchenko contributed equally to this work.
297
298 Awet Tecleab et al.
In order to accurately calculate the risks of birth defects using

the quad screen, each laboratory must first establish the normal
reference ranges of maternal serum AFP, uE3, β-hCG, and DIA
levels using its own patient population to account for regional and
demographic variation.
AFP is the most abundant fetal serum protein and is an impor-
tant marker of open neural tube defects (ONTDs), which occur at a
rate of 1–2 per 1000 live births in the United States [4]. It is
synthesized by the fetal liver and the yolk sack, passes into the
amniotic fluid in fetal urine, and reaches the maternal circulation
through the placenta or by diffusion through the fetal membranes.
In a fetus with an open neural tube defect such as anencephaly or
spina bifida, the defective closure of the developing nervous system
allows AFP to leak out and increases its concentration in the amni-
otic fluid and in the maternal serum [5].
In addition to ONTDs, AFP is also used as a marker for Down
syndrome and trisomy 18, both of which are associated with
decreased maternal serum AFP levels [6–9]. Prior to the develop-
ment of this screening method, pregnant women aged 35 and
above were routinely assessed for Down syndrome by the more
invasive procedures of amniocentesis and karyotyping [6–8].
AFP is first detected in the maternal serum at about tenth week
of gestation, its concentration starts to rise around week 16 and
subsequently declines after about week 32 [ref. 10, and Table 1].
Thus, for prenatal screening, the maternal serum AFP concentra-
tion is typically measured in the second trimester, as the first tri-
mester levels are too low to reliably differentiate between an
affected and an unaffected fetus.
The Beckman Access2 Immunoassay System can quickly and
accurately measure the levels of AFP and the other quad screen
analytes as mass per unit volume of maternal serum (amniotic fluid
can also be used for the measurement of AFP levels, but there are
caveats- see Notes 1–6). However, in order to be able to interpret
these results, each laboratory must establish its own normal reference
AFP ranges in unaffected singleton pregnancies; this is routinely
done using 50–100 maternal serum samples for each gestational
week. In addition to gestational age, other factors such as maternal
weight, race, and diabetes mellitus status also significantly affect
maternal serum AFP concentration and must be taken into account
[11, 12]. For this reason, each measured maternal serum AFP level is
normalized to the appropriate median level in the reference popula-
tion to derive the multiples of the median (MoM) value, which is
then further adjusted for additional factors mentioned above.
Table 1 shows the mean maternal serum AFP concentrations as
well as the calculated MoMs for each gestational week 15–19 for
2359 samples analyzed at Columbia University Medical Center.
Estriol is the predominant form of estrogen in pregnant
women. It is synthesized by the placenta, is made water soluble by
Quad Screen Test, A Multiplexed Biomarker Assay for Prenatal. . . 299
Table 1
Median maternal serum AFP levels by week of gestation
Multiples of median concentration (ng/mL)

Number Median concentration
Week of Gestation of samples (ng/mL) 2.0 2.5 3.0
15 435 31.1 62.2 77.8 93.4
16 506 36.0 72.0 90.0 108.0
17 452 41.6 83.2 104.1 124.9
18 425 48.1 96.3 120.3 144.4
19 413 55.7 111.3 139.2 167.0
20 308 64.4 128.8 161.0 193.2
Table 2
The range in uE3 levels by gestational age week
GA week Range of ue3 results (ng/mL)

13 0.42–0.50
14 0.52–0.62
15 0.64–0.76
16 0.79–0.94
17 0.97–1.17
18 1.20–1.44
19 1.49–1.78
20 1.84–2.20
21 2.27–2.72
conjugation with sulfate and glucuronate in the maternal liver, and

is excreted by the kidney. However, approximately 10% of estriol
circulates unconjugated [13]. Low maternal serum unconjugated
estriol (uE3) levels are associated with Down syndrome and trisomy
18 [9, 14].
Table 2 shows the range in uE3 levels by gestational age week,
derived from patients presenting at Columbia University Medical
Center.
Human chorionic gonadotropin (hCG) is a glycoprotein hor-
mone synthesized by the placenta that is used to screen for Down
syndrome and trisomy 18. It is composed of alpha and beta sub-
units, with the beta subunit (β-hCG) being specific to hCG
[12, 15]. The serum levels of β-hCG in non-pregnant women are
very low, usually less than 5 IU/L, but soon after implantation of
Table 3
Median serum β-hCG levels in non-pregnant women
Reference population Median Range 95th Percentile

(non-pregnant females) N (mIU/mL) (mIU/mL) (mIU/mL), [95% CI]
18 and < 40 years 132 0 0–0.6 0.3[0.2–0.4]
40 years 141 0 0–3.1 1.5[1.1–2.9]
Post-menopausal 134 2.8 0.1–11.6 7.7[6.4–10.4]
the zygote, β-hCG levels increase rapidly and are used for early
detection of pregnancy. The level of β-hCG doubles every 1.5 to
3 days for the first 6 weeks of gestation, continues to increase until
the end of the first trimester, and then declines for the remainder of
the pregnancy [16]. Maternal serum β-hCG levels are typically
increased approximately two-fold in pregnancies with Down syn-
drome but much decreased in pregnancies with trisomy 18 [9].
The median, the range, and the 95th percentile values of
maternal serum β-hCG concentrations in healthy non-pregnant
women of several age groups measured at Columbia University
Medical Center are shown in Table 3.
Inhibin A is a gonadal protein hormone secreted by the Sertoli
cells of the testis or the granulosa cells of the ovary. It inhibits the
secretion of follicle stimulating hormone (FSH) by the pituitary
gland and regulates gamete production, as well as embryonic and
fetal development [17]. Increased maternal serum dimeric inhibin
A levels are associated with Down syndrome [18].
The median, the 2.5th, and the 97.5th percentiles of maternal
serum dimeric inhibin A levels measured at Columbia University
Medical Center in women at various stages of the menstrual cycle,
as well as in postmenopausal women and normal men are shown in
Table 4.
Once the appropriate reference ranges are established, the
maternal serum concentrations for each analyte measured by the
Beckman Access 2 Immunoassay System can be converted into
multiples of the median by dividing the sample concentration by
the median of the appropriate reference range, for that gestational
age. The multiple of median (MoM) is further adjusted by the
variables that are known to affect the MoM such as weight, race,
prior pregnancies and deliveries, and age at the time of delivery.
The patient-specific risk for a specific abnormality is then calcu-
lated by multiplying the pretest probability, obtained from large
epidemiological studies, by likelihood ratio based on the
adjusted MoM.
Finally, the quad screen test results are reported as the maternal
serum concentrations and the adjusted MoM values for each ana-
lyte, the risks for specific birth defects, and the overall interpretation.
Table 4
Median dimeric inhibin a levels
Median 2.5% Lower limit 97.5% Upper limit

Population n (pg/mL) (pg/mL) (pg/mL)
Normal cycling females
(days from LH surge)
Early follicular phase ( 14 to 10) 211 6.4 1.8 17.3
Mid follicular phase ( 9 to 4) 264 11.7 3.5 31.7
Late follicular phase ( 3 to 1) 121 29.0 9.8 90.3
Mid-cycle (day 0, LH surge) 96 41.8 16.9 91.8
Early luteal phase (1 to 3) 122 43.7 16.1 97.5
Mid luteal phase (4 to 11) 344 38.3 3.9 87.7
Late luteal phase (12 to 14) 138 12.5 2.7 47.1
Postmenopausal females 58 1.1 <1.0 2.1
Males 67 1.1 <1.0 2.0
1.1 Assay Principle To measure AFP, β-hCG, and dimeric inhibin A levels, the Beckman
Access2 instrument utilizes chemiluminescent noncompetitive
two-site immunoassays. Briefly, the patient samples are incubated
with two types of analyte-specific antibodies: (1) monoclonal anti-
bodies immobilized on paramagnetic particles and (2) monoclonal
antibodies conjugated to alkaline phosphatase (AP).
In the AFP immunoassay the sample is incubated with both the
paramagnetic particle-immobilized antibodies and the
AP-conjugated antibodies in one step. The reaction vessel is then
washed in the presence of a magnetic field to remove the unbound
reaction components, including any unbound AP-conjugated anti-
bodies. Importantly, the paramagnetic particle-immobilized anti-
bodies, the AFP, and the-AFP-bound AP-conjugated antibodies
remain in the reaction vessel.
The β-hCG and inhibin A immunoassays use the same principle
but are performed in two steps. The samples are first incubated with
the analyte-specific paramagnetic particle-immobilized antibodies
alone and the reaction vessel is then washed in the presence of a
magnetic field to remove unbound reaction components. In the
second step, the analyte-specific AP-conjugated antibodies are
added and the wash step is repeated.
Following antibody incubation and washing, the reaction vessel
is incubated with a chemiluminescent AP substrate (Lumi-Phos*
530). The light generated from the AP-substrate reaction, which is
proportional to the analyte concentration, is detected using a
luminometer. The serum levels of AFP, β-hCG, and inhibin A are
determined from standard calibration curves.
To measure unconjugated estriol levels, the Beckman Access2

instrument uses a heterogenous competitive binding immunoassay.
Briefly, the patient serum is added to a reaction vessel that contains
AP-conjugated estriol, anti-estriol rabbit antibodies, and paramag-
netic particles coated with goat anti-rabbit antibodies. The estriol
from the sample competes with the AP-conjugated estriol for the
binding sites of the anti-estriol rabbit antibodies. The resulting
antigen-antibody complex is in turn bound by the anti-rabbit anti-
bodies coating the magnetic particles. After incubation, the reac-
tion vessel is washed in the presence of a magnetic field such that
the substances not bound to the paramagnetic particles are washed
away. As described above, Lumi-Phos* 530 is then added to the
reaction mixture and the light generated by the reaction, which is
inversely proportional to the estriol concentration, is detected
using a luminometer. The serum level of the estriol is determined
from a standard calibration curve.
1.2 Specimen The recommended sample is maternal serum collected between

15 and 20 weeks of gestation (see Note 2). The samples must be
accompanied by a requisition form that contains the complete
patient demographics. This includes date of birth (not chronologi-
cal age), expected day of delivery (based on measurement of fetal
biparietal diameter or last menstrual period), specimen collection
date, gestational age (number of completed weeks and additional
days, see Note 4), weight at the time of sample collection, race,
insulin dependent diabetes status prior to the pregnancy, whether it
is a repeat sample, number of pregnancies and deliveries (para-
gravida), and prior history of neural tube defects.
2 Materials
2.1 Common 1. Benetech PRA—software package used to calculate the risk for
Material and Reagents a particular disorder based on MoM values (Benetech, Tor-
onto, ON, Canada).
2. Beckman Access 2 automated analyzer and associated consum-
ables, see below items 3–10 (Beckman Coulter Inc., Brea, CA).
3. Reaction vessels.
4. Fiber-free applicator sticks.
5. Deionized water.
6. Citranox (cleaning solution).
7. Contrad 70 cleaning solution.
8. 2.0 mL sample cups.
9. Calibrated pipettes.
10. Pipette tips.
2.2 Reagents for AFP Beckman Access AFP reagent and calibrator kit; this includes items
Measurement 1–6 listed below (Beckman Coulter Inc., Brea, CA).
1. R1: Access AFP reagent pack.
(a) R1a—reagent pack containing mouse monoclonal anti-
AFP antibody conjugated with paramagnetic particles.
(b) R2a—reagent pack containing mouse monoclonal anti-
AFP antibody conjugated with alkaline phosphatase.
2. AFP calibrators (S0 to S6)—calibrator material that corre-
sponds to 0, 2.5, 5, 25, 100, 500, and 3000 ng/mL AFP.
AFP assay requires calibration every 28 days. Calibrators
should be run in duplicate to prepare calibration curve.
3. Calibration Card
4. Beckman access substrate: Lumi-phos*530.
5. Beckman access wash buffer II (R3 wash buffer II).
6. Beckman access AFP sample diluents—for dilution of samples
with values outside of the analytical measurement range.
7. Level 1, 2, 3 controls, Lyphocheck maternal serum controls
(BioRad, Hercules, CA, USA).
8. Control for amniotic fluid AFP (this is a pooled patient sample,
and is prepared in our laboratory).
2.3 Reagents Beckman Access β-hCG(fifth IS) reagent and calibrator kit, which
for Total β-hCG(Fifth includes the items listed below (Beckman Coulter Inc., Brea, CA).
IS) Quantitative 1. R1: Access Total β-hCG(fifth IS) reagent pack.
Determination
(a) R1a—reagent containing paramagnetic coated goat anti-
mouse IgG: monoclonal mouse anti-βhCG.
(b) R1b—protein (goat, murine, and recombinant).
(c) R1c—reagent containing alkaline phosphatase conjugated
rabbit anti-β-hCG.
2. Total β-hCG(fifth IS) calibrators.
(a) S0: buffered bovine serum albumin (BSA) matrix with
surfactant, <0.1% sodium azide, and 0.5% ProClin**300.
Contains 0.0 IU/L hCG.
(b) S1-S5: hCG at levels of approximate
6,35,195,620,1350 IU/L, respectively, in buffered BSA
matrix with surfactant, <0.1% sodium azide, and 0.5%
ProClin 300.
(c) Calibration Card.
3. Beckman access substrate—Lumi-phos* 530.
4. Beckman access wash buffer II.
5. Level 1, 2, 3 controls, Liquicheck maternal serum (BioRad,
Hercules, CA, USA).
6. The analytical range of the assay is 0.5–1000 IU/L. If β-hCG

value is greater than 1000 IU/L, manual dilution can be
performed by diluting 1 part of patient sample to 199 of
wash buffer II. Result from manual dilution should then be
multiplied by 200 to obtain sample result.
2.4 Reagents Beckman Access inhibin A reagent and calibrator kit, which
for Dimeric Inhibin A includes the following items (Beckman Coulter Inc., Brea, CA):
Quantitative
1. R1: Access Inhibin A reagent pack.
Determination
(a) R1a—paramagnetic particles coated with mouse mono-
clonal anti-inhibin A.
(b) R1b—mouse monoclonal anti-inhibin A alkaline phos-
phatase (bovine) conjugate.
(c) R1c—Tris buffered saline, BSA, proteins (bovine,
murine).
(d) R1d—phosphate buffer, oxidizer.
(e) R1e—Tris buffer, detergent.
2. Inhibin A calibrators.
surfactant, <0.1% sodium azide, and 0.5% ProClin300.
Contains 0.0 IU/L inhibin A.
(b) S1–S6: Recombinant human inhibin A at levels of approx-
imate 10, 50, 100, 500, 1000, 1500 pg/mL, respectively,
in buffered BSA matrix with surfactant, <0.1% sodium
azide, and 0.5% ProClin 300.
(c) Calibration Card.
3. Beckman access substrate.
5. Level 1, 2, 3 controls, Beckman Access inhibin A QC.
6. Beckman Access diluent buffer II.
7. The linearity for this assay is between 1 and 1500 pg/mL.
2.5 Reagents Beckman Access unconjugated Estriol (uE3) reagent and calibrator
for Unconjugated kit, which includes the following items (Beckman Coulter Inc.,
Estriol (uE3) Brea, CA).
Measurement 1. R1: Access Total unconjugated Estriol reagent pack.
(a) R1a—paramagnetic particle coated with goat anti-
rabbit IgG.
(b) R1b—Rabbit anti-estriol.
(c) R1c—Estriol-alkaline phosphatase conjugate
2. Unconjugated Estriol calibrators.

surfactant, <0.1% sodium azide, and 0.5% ProClin300.
Contains 0.0 IU/L unconjugated Estriol (uE3).
(b) S1–S6: Recombinant human unconjugated Estriol (uE3)
at levels of approximate 0.07, 0.17, 0.34, 0.86, 3.4,
6.9 ng/mL, respectively, in buffered human serum matrix
containing <0.1% NaN3 and 0.025% Cosmocil CQ.
3. Calibration Card.
4. Beckman access substrate Lumi-Phos 530.
6. Level 1, 2, 3 controls, BioRad Liquicheck maternal serum
(BioRad, Hercules, CA, USA).
7. For samples with uE3 concentration greater than 6.9 ng/mL,
dilute sample with the zero calibrator and re-run the test. To
obtain sample result, multiply by the appropriate dilution
factor.
3 Methods
1. Mix the contents of unpunctured reagent packs by gently

inverting several times until homogenous mix is observed.
2. Load reagents into the Beckman Access 2 instrument.
3. Calibrate the instrument. Follow the instruction provided by
the vendor.
4. Confirm if the instrument calibration was successful.
5. Run controls and verify that the controls are within the accept-
able limits.
6. Place patient samples on a rack and then load the rack in the
instrument.
7. Select AFP, hCG5, inhibin or estriol free (for unconjugated
estriol-3 assay) and run.
8. For samples with AFP concentration greater than 2550 ng/
mL, order dilution (d-AFP).
9. Enter the result from this analysis into Benetech PRA, which
will subsequently calculate MoM values and the risk of having
open spina bifida, Down syndrome, and Trisomy 18. See also
Notes 4–5.
3.1 Establishing 1. Run QC in replicates, and on different days. Each clinical

and Implementing QC laboratory should have a defined standard operating procedure
Range for this; we suggest a minimum of 20 replicates run over
20 days to capture all sources of variability.
2. Calculate mean, SD, and %CV. This should be reviewed and

signed by clinical laboratory director, or appropriate designee.
3. Enter this QC range into the laboratory information system
(LIS).
4. Run QC at the beginning of every shift and make sure that the
controls are within acceptable limits, for each analyte.
4 Notes
1. Positive amniotic fluid-AFP results will trigger a reflex for

additional testing (cholinesterase and fetal hemoglobin); we
do not perform this testing in-house, it is sent to an outside
reference laboratory.
2. Specimen for the measurement of AFP from maternal serum
should be collected prior to amniocentesis.
3. Bloody amniotic fluid can have falsely elevated AFP level. In
such circumstance, the amniotic fluid must be tested for the
level of fetal hemoglobin using Kleihauer-Betke test, electro-
phoresis, or other appropriate tests.
4. Inaccurate gestational age is one of the most frequent causes of
inaccurate screening test results. Results can be recalculated
once accurate gestational age is obtained. For second trimester
testing, recalculation should only be done when the difference
in calculated and revised gestational age is >7 days.
5. Diagnosis of ONTD should not be made based on elevated
AFP values only. Elevated AFP values have been found in other
fetal abnormalities such as congenital nephrosis, omphalocele,
Turner syndrome, or fetal demise [19, 20].
6. Amniotic fluids contain very high levels of AFP and hence when
assaying for AFP from amniotic fluids, the sample should be
pre-diluted. Dilute 1 part of amniotic fluid to 10 parts of
Beckman Access sample diluents. This result should then be
multiplied by 11 to obtain the final result.
References
1. Benn PA (2002) Advances in prenatal screen- with the quadruple test. Lancet 361
ing for Down syndrome: I. general principles (9360):835–836
and second trimester testing. Clin Chim Acta 4. Macri JN, Baker DA, Baim RS (1981) Diagno-
323(1–2):1–16 sis of neural tube defects by evaluation of amni-
2. Benn PA et al (2003) Incorporation of inhibin- otic fluid. Clin Obstet Gynecol 24
A in second-trimester screening for Down syn- (4):1089–1102
drome. Obstet Gynecol 101(3):451–454 5. Rose NC, Mennuti MT (1993) Maternal
3. Wald NJ, Huttly WJ, Hackshaw AK (2003) serum screening for neural tube defects and
Antenatal screening for Down’s syndrome fetal chromosome abnormalities. West J Med
159(3):312–317
6. Combining maternal serum alpha-fetoprotein measurement. A controlled study. Lancet 1

measurements and age to screen for Down (7861):765–767
syndrome in pregnant women under age 13. Penney LL, Klenke WJ (1980) Variability in
35 (1989) New England regional genetics unconjugated and total estriol in serum during
group prenatal collaborative study of down normal third trimester pregnancy. Clin Chem
syndrome screening. Am J Obstet Gynecol 26(13):1800–1803
160(3):575–581 14. Bogart MH, Pandian MR, Jones OW (1987)
7. Merkatz IR et al (1984) An association Abnormal maternal serum chorionic gonado-
between low maternal serum alpha-fetoprotein tropin levels in pregnancies with fetal chromo-
and fetal chromosomal abnormalities. Am J some abnormalities. Prenat Diagn 7
Obstet Gynecol 148(7):886–894 (9):623–630
8. Cuckle HS, Wald NJ, Lindenbaum RH (1984) 15. Cole LA (2010) Biological functions of hCG
Maternal serum alpha-fetoprotein measure- and hCG-related molecules. Reprod Biol
ment: a screening test for Down syndrome. Endocrinol 8:102
Lancet 1(8383):926–929 16. Sturgeon CM, McAllister EJ (1998) Analysis of
9. Palomaki GE et al (1995) Risk-based prenatal hCG: clinical applications and assay require-
screening for trisomy 18 using alpha- ments. Ann Clin Biochem 35(Pt 4):460–491
fetoprotein, unconjugated oestriol and human 17. van Zonneveld P et al (2003) Do cycle distur-
chorionic gonadotropin. Prenat Diagn 15 bances explain the age-related decline of female
(8):713–723 fertility? Cycle characteristics of women aged
10. Kjessler B, Johansson SG (1977) Monitoring over 40 years compared with a reference popu-
of the development of early pregnancy by lation of young women. Hum Reprod 18
determination of alpha-fetoprotein in maternal (3):495–501
serum and amniotic fluid samples. Acta Obstet 18. Wald NJ et al (1997) Antenatal screening for
Gynecol Scand Suppl 69:5–14 Down’s syndrome. J Med Screen 4
11. Brock DJ, Sutcliffe RG (1972) Alpha- (4):181–246
fetoprotein in the antenatal diagnosis of anen- 19. Crandall BF (1981) Alpha-fetoprotein: the
cephaly and spina bifida. Lancet 2 diagnosis of neural-tube defects. Pediatr Ann
(7770):197–199 10(2):38–48
12. Wald NJ, Brock DJ, Bonnar J (1974) Prenatal 20. Macri JN, Weiss RR (1977) The utilization of
diagnosis of spina bifida and anencephaly by alpha-fetoprotein in prenatal diagnosis. Birth
maternal serum-alpha-fetoprotein Defects Orig Artic Ser 13(3D):191–199
Chapter 21
Isolation of Cell-Free DNA from Maternal Plasma

James Stray and Bernhard Zimmermann
Abstract
Noninvasive prenatal genetic tests analyzing the cell-free fetal DNA in the circulation of expectant mothers
are now performed routinely in clinical diagnostic laboratories. Leveraging the power of next generation
sequencing (NGS), these tests can detect variation in chromosomal copy number or microdeletions early in
gestation. All methods begin with blood collection followed by transport to the diagnostic lab, plasma
separation, and purification of ccfDNA from the plasma to prepare it for molecular analysis. Preservation of
ccfDNA in blood samples and highly efficient purification from plasma are paramount since the quality and
quantity of target nucleic acids determine the sensitivity and therefore success of these assays. Maximizing
quality and quantity and minimizing variation in extraction yield pose significant challenges for diagnostic
labs, many of which use manual isolation methods for plasma volumes greater than 5 mL. One way to
reduce variability is to automate the extraction processes and, to the extent possible, minimize hand-on
operations. This chapter details two procedures for isolating ccfDNA from 10 mL plasma by manual and
automated means using the QIAamp Circulating Nucleic Acid Kit and the QIAsymphony Circulating DNA
Kit. The ccfDNA recovered is suitable for downstream processing in noninvasive prenatal tests for aneu-
ploidy detection.
Key words Circulating cell-free DNA, Automation, Fetal, Noninvasive prenatal testing, QIAsymph-
ony, QIAamp, Blood sample stabilization, Circulating DNA extraction, SNP-based aneuploidy
detection
1 Introduction
Extracellular DNA detectable in the cell-free fraction of blood

(plasma or serum; see Note 1) is defined as circulating cell-free
DNA (ccfDNA). The half-life of naked DNA in blood and plasma
is extremely short, on the order of minutes [1], kept so by active
plasma nucleases including DNAse I, DNAse II, and phosphodies-
terase I [2]. Circulating cell-free DNA (ccfDNA) survives slightly
longer in the blood and avoids rapid degradation by remaining in
complex with the histone octamer (2 (H2A, H2B, H3, H4)) as
mono- and oligo-nucleosomes, with or without linker histones
(chromatosomes), which act to protect 140 to 160 base pairs
(bp) of DNA per histone core. Nucleosomes/chromatosomes and
309
310 James Stray and Bernhard Zimmermann
poly-nucleosomes/poly-chromatosomes are shed into the blood-

stream as part of normal physiology. The concentration of circulat-
ing cell-free DNA (ccfDNA) varies widely [3], in the ng/mL range,
and reflects the rate at which chromatin fragments enter the blood-
stream, less the rate of destruction by nucleases, and clearance by
hepatic uptake and immune-depletion. Healthy individuals main-
tain an average steady state ccfDNA concentration of ~13 ng/mL,
ranging between 0 and 100 ng/mL plasma [4, 5] (see Note 2). No
one cell type or mechanism accounts for the accumulation of
nuclear DNA in the blood, though the primary source is erythro-
poietic apoptosis, with variable contributions from general cell
turnover, post engulfment macrophage secretion, and traumatic
or necrotic cell death [6–8]. In addition to plasma ccfDNA in
tight association with histones, cellular DNA and various RNAs
survive plasma destruction protected within membrane microvesi-
cles, exosomes, shed continually from many cell types through the
normal process of exocytosis or blebbing [9].
Since the discovery that fetal ccfDNA is detectable and
increases in concentration in the circulation of women throughout
gestation [10], ccfDNA has become a key analyte for noninvasive
prenatal diagnostics. Fetal DNA of placental origin is shed into
maternal blood [11]. The properties of ccfDNA found in maternal
plasma pose technical challenges: first among them is the extremely
low total ccfDNA concentrations and the fact that the fetal ccfDNA
is present on average at only 10%, but frequently less than 5%, in
plasma samples [12]. In particular, the percentage of fetal DNA in
the total ccfDNA is lowest during the first trimester of pregnancy.
This issue is addressed by processing high sample volumes between
1 mL and 10 mL, depending on information content to be
obtained from the sample. The second challenge is the high degree
of ccfDNA fragmentation, which requires an efficient method for
recovery of small DNA fragments. The length distribution of frag-
mented ccfDNA shows a predominant peak at approximately
165–175 bp, consistent with a majority of DNA-histone-protein
complexes of a single nucleosome/chromatosome in size. The fetal
fraction of ccfDNA is demonstrably shorter and only monosomal in
length, with an average of ~145–150 bp protected [13–15]. If a
sufficient volume of plasma is processed to give an increased con-
centration of ccfDNA in eluates, then secondary peaks migrating
later relative to the predominant peak can be detected, consistent
with oligonucleosomes/oligochromatosomes of 2 to 3 multi-
nucleosomes in length (Fig. 1). Primarily, plasma input volumes
of 2–4 mL are used for prenatal diagnostics [16]. But for a detailed
analysis of NIPT samples (such as quantitative measurements of
fetal chromosome dosage in, e.g., aneuploidy detection), even
higher sample input volumes, up to 10 mL plasma, are preferred
to ensure reliable detection in cases having low ccfDNA content
and/or a low fraction of fetal ccfDNA.
Scalable ccfDNA Isolation 311
Fig. 1 Size distribution of extracted ccfDNA from 4 mL and 10 mL plasma. ccfDNA extracted from 4 mL plasma
(blue) and 10 mL plasma (red) using the QIAsymphony Circulating DNA Kit. 1 μL eluate subjected to Agilent
High Sensitivity DNA Chip. FU ¼ fluorescent units, marker peak at 35 bp and 10,380 bp
In this chapter we will discuss the extraction of ccfDNA from

large volumes of plasma (up to 10 mL), using the manual QIAamp
Circulating Nucleic Acid Kit, used by Natera for plasma ccfDNA
purification since offering the noninvasive prenatal test (Pano-
rama™), and the newly developed QIAsymphony Circulating
DNA Kit, an automated scalable method that operates on the
QIAsymphony SP robotic platform.
In the last few years the rapid and widespread adoption of
reliable NIPT screening has increased demand for an automated
solution for large volume ccfDNA extraction methods. To meet
demand and address this issue, a new chemistry dedicated for
automated ccfDNA extraction with modified, scalable protocols
for processing up to 10 mL was developed and compared to a
manual ccfDNA extraction method also modified for a plasma
input volume of 10 mL. Eluates from such extraction methods
are used for prenatal diagnostics using Natera’s Panorama™ V2
prenatal test (Fig. 2), detailed in [17, 18].
Developing an automated process for handling such high input
volumes in parallel to drive extraction of 96 samples in one com-
plete run in <2.5 h per batch of 24 samples is challenging, and
required a new chemistry concept. The new chemistry is based on
Anion-Exchange beads which require only a low volume of binding
buffer added to samples which results in reduced overall processing
volumes, and where sample volumes can be scaled with linear
extraction efficiencies (Fig. 3a), allowing for parallel processing
and higher throughput compared to chaotropic salt/silica-based
Genotype
0% FF 10% FF 15% FF 25% FF Fetal Mat.

1.0 AA
0.9 AA
AB
0.8
Proportion of A-Allele SNPs
0.7
AA
0.6
0.5 AB/AB AB
0.4
BB
0.3
0.2
0.1
AB
BB
0 BB
13 18 21 13 18 21 13 18 21 13 18 21
Chromosome Chromosome Chromosome Chromosome
Maternal
Genotype AA AB BB
Fig. 2 Graphic representation of sequencing data. The Panorama SNP data are presented in a simplified
fashion as the ratio of the two most likely alleles, labeled arbitrarily as A or B. This is not how the algorithm
makes ploidy calls in practice, but is one way to visualize this very large data set. Y-axis: (A allele reads)/(Total
allele reads). X-axis: Linear SNP position along each chromosome. Fetal and maternal genotypes are indicated
to the right, Red: the homozygous maternal (AA) allele cluster, Green: the heterozygous maternal (AB) allele
cluster, and Blue: the homozygous maternal (BB) allele cluster. Each spot represents the sum of maternal and
fetal A allele read proportions from analysis of the ccfDNA, and depicts patterns typical for euploid autosomes
at varying % fetal fraction (FF) of 0%, 10%, 15%, and 25%. The center triple green cluster, most evident at
25% fetal fraction, and the presence of blue and red peripheral clusters indicate the presence of two copies of
chromosomes 13, 18, and 21 in these euploid examples
automated nucleic acid extraction methods with a comparable

extraction efficiency (Fig. 3b).
The reason for failed laboratory diagnostic tests can often be
attributed to the preanalytical sample processing [19] which
emphasizes the importance of standardized workflows for ccfDNA
applications: The ccfDNA extraction is embedded within this pre-
analytical process but the workflow starts at the sample collection,
which includes blood collection, time until processing and stabili-
zation. Since the transport of whole blood from the site of collec-
tion to the laboratory for processing can take up to several days,
which determines the time span between blood collection and
plasma separation, the choice of the blood collection tube (BCT)
must not be undervalued. Next to the well-established anticoagu-
lant preservatives EDTA and Citrate BCTs, several suppliers like
Streck (Cell-free DNA BCT), PreAnalytiX (PAXgene Blood
ccfDNA Tube) now offer tubes designed to prevent clotting, stabi-
lize cells, and preserve ccfDNA using different stabilizing agents.
Usage of these new BCTs has to be considered carefully because
they can affect the quality and integrity of extracted ccfDNA and
downstream assay compatibility (see Note 3).
Fig. 3 Linearity of ccfDNA extraction from 1 to 10 mL plasma input for an automated ccfDNA extraction
method (a) and performance comparison to a manual ccfDNA extraction method (b). (a) Pooled plasma
generated from Cell-Free DNA BCT (Streck) was used. ccfDNA was extracted from 1 mL, 2 mL, 4 mL, and
10 mL plasma input on the QIAsymphony instrument using the QIAsymphony Circulating DNA Kit in combina-
tion with different protocols. ccfDNA yield was determined using an in-house real-time PCR assay for the 18S
coding sequence (66 bp amplicon). Results were calculated as target copies (haploid genome copies) per μL
eluate. (b) Blood draw from 48 single donors was performed in Cell-Free DNA BCT (Streck). ccfDNA was
extracted from 4 mL plasma input using the QIAsymphony Circulating DNA Kit and the QIAamp Circulating
Nucleic Acid Kit in comparison. ccfDNA yield was determined using an in-house real-time PCR assay for the
18S coding sequence (66 bp amplicon; ref. [20]). Results were calculated as target copies (haploid genome
copies) per mL plasma input
2 Materials
2.1 Plasma Venous maternal blood is collected into Streck Cell-free DNA
Separation from Whole BCT™ collection tubes and stored or shipped at ambient tempera-
Blood ture prior to plasma isolation. The plasma separation is performed
in the diagnostic laboratory as described in Subheading 3.1.
2.2 QIAamp The QIAamp Circulating Nucleic Acid Kit provides reagents and
Circulating Nucleic components for manual extraction of ccfDNA. The extraction
Acid Kit (QIAGEN, chemistry is based on using chaotropic salt, detergent, and protein-
Germantown, MD, ase K to release nucleic acids from membrane vesicles and bound
USA) proteins (such as nucleosome complexes) and to make these nucleic
acid molecules available for binding to a silica membrane surface.
The binding of these fragmented nucleic acids to the silica surface is
then achieved by a combination of chaotropic salt and isopropanol,
which make the binding of DNA and RNA to the hydrophilic
surface thermodynamically favored. The kit contains the following
reagents:
1. Lysis buffer ACL: contains guanidine thiocyanate, optional
addition of 1 μg carrier RNA per sample.
2. Binding buffer ACB: contains guanidine thiocyanate, final con-
centration of isopropanol is 40%.
3. Wash buffer ACW1: contains guanidine hydrochloride, final
concentration of ethanol is 56%.
4. Wash buffer ACW2: final concentration of ethanol is 70%.
5. Elution buffer: Buffer AVE (low ionic-strength solution).
6. Proteinase K, recombinant.
In addition, the following material is required for fast and
efficient vacuum processing of up to 24 spin columns in parallel
(samples and wash solutions are drawn through the column mem-
branes by vacuum instead of centrifugation, providing greater
speed and reduced hands-on time in purification procedures):
(A) QIAvac 24 Plus vacuum manifold (QIAGEN, Germantown,
MD, USA), (B) QIAvac Connecting System (QIAGEN, German-
town, MD, USA) and (C) Vacuum Pump capable of producing a
vacuum of 800 to 900 mbar.
2.3 QIAsymphony The QIAsymphony Circulating DNA Kit provides reagents and
Circulating DNA Kit components for automated extraction of ccfDNA using the QIA-
(QIAGEN, Germantown, symphony SP instrument (QIAGEN, Germantown, MD, USA).
MD, USA) The extraction chemistry is based on using proteinase K to release
nucleic acids from bound proteins (such as nucleosome complexes)
and anion-exchange beads to bind the negatively charged ccfDNA
to the positively charged bead surface under slightly acidic condi-
tions. During three wash steps impurities (mainly proteins) are
removed using a combination of increased pH of wash buffers and

salt concentration. Elution takes place under slightly alkaline con-
ditions to achieve a quantitative elution of bound ccfDNA while
making sure that the eluted DNA is intact (e.g., double stranded).
In addition to the QIAsymphony SP instrument and QIA-
symphony Circulating DNA Kit, the following QIAsymphony SP
disposable plasticware is required:
1. Sample Prep Cartridges, 8-well cartridges (QIAGEN, German-
town, MD, USA).
2. 8-Rod Covers (QIAGEN, Germantown, MD, USA).
2.4 Panorama The Panorama prenatal test is a next generation genetic test that
Prenatal Test (Version uses plasma ccfDNA from pregnant women early in gestation as an
2) input to screen for genetic aneuploidies and microdeletions that
affect child development. The assay begins by preserving ccfDNA
in a library through a series of highly efficient matched molecular
reactions designed to repair and amplify fragments in preparation
for downstream processing. An aliquot of the library is used as
input for a massively multiplex-specific target amplification reaction
which reduces the complexity of the sequencing query space and
greatly increases the sensitivity and specificity of the assay. Amplified
targets are barcoded and readied for NGS sequencing. Sequence
data is processed and an allele designation is assigned for each SNP
to the mother and child. The plot in Fig. 2 depicts allele ratios
obtained for SNPs arrayed linearly along a chromosome, and gen-
erated with input ccfDNA deriving from maternal blood having
different estimated fetal fractions (FF). The higher the fetal frac-
tion, the greater the proportion of fetal SNPs to the total SNPs
(fetal + maternal) detected. This translates into a greater divergence
between the fetal genotypes from the underlying maternal SNP
genotypes (Figs. 2, 0% FF) where maternal AA, AB, and BB clusters
are shown as red, green, and blue dots, respectively. The algorithm
does not graphically interpret the results and can discern differences
that are not obvious by visual inspection. However, this represen-
tation illustrates how the assay responds to fetal fraction and that
signal to noise of Panorama, and similar assays, can benefit from
efficient ccfDNA isolation methods that recover low abundant,
short DNA fragments from large plasma volumes.
The Panorama V2 Sample Preparation Reagents were config-
ured and optimized to generate a population of ccfDNA fragments
in eight separate steps:
1. DNA End Repair, which converts 30 or 50 overhangs of ccfDNA
fragments into blunt ends using a polymerase.
2. A-Tailing, to add a single 30 A nucleotide to blunt-ended
ccfDNA fragments to prepare ends for annealing and ligation
to adapters.
3. Ligation, the covalent coupling of synthetic adapter sequences

to both ends of A-tailed ccfDNA fragments.
4. Library Amplification, a PCR amplification of adapter-ligated
ccfDNA fragments with forward and reverse primers comple-
mentary to adapters.
5. Library purification.
6. Specific target amplification, a massively multiplexed PCR
reaction whose input is amplified ccfDNA library created in
steps 1–4. The reaction amplifies a select set of 13,392 target
loci across a subset of chromosomes, the information from
which can be used to determine the presence or absence of
certain aneuploidies.
7. Barcoding, final PCR reaction that appends a unique identifier
to the end of each target amplicon and readies fragments for
Illumina sequencing.
8. Pooling, quantification, and loading of flow cells for Illumina
Sequencing. Individually barcoded PCR products are pooled
with those originating from other cases and the concentration
of the pool is determined and the pool diluted to a concentra-
tion that achieves the optimum cluster density for sequencing a
given depth of read.
3 Methods
An overview of the complete ccfDNA workflow from blood collec-

tion to downstream analysis is depicted in a processing scheme
(Fig. 4).
Fig. 4 Processing scheme for manual and automated ccfDNA extraction. After blood collection and plasma
isolation two ccfDNA extraction methods are compared to each other: a manual extraction method based on
denaturating agents in combination with a silica surface to bind DNA and a automated extraction method
based on non-denaturating conditions using anion-exchange beads to bind DNA. Both methods result in
different usable elution volumes before purified ccfDNA is subjected to downstream anlysis
3.1 Plasma 1. Obtain the blood samples to be processed. Visually check the
Separation from Whole sample IDs to ensure that there are two blood collection tubes
Blood (BCTs) per case.
2. For each pair of BCTs, label one 15 mL conical tube and one
50 mL conical tube with the corresponding sample ID.
3. Centrifuge the BCTs at 2000 rcf with maximum acceleration/
deceleration for 20 min at 22 C.
4. Use a 10 mL serological pipette to transfer the plasma from
each pair of BCTs to the corresponding 15 mL conical tube (see
Note 4).
5. Centrifuge the 15 mL conical tubes at 3220 rcf with maximum
acceleration/deceleration for 30 min at 22 C (see Note 5).
6. Transfer the plasma from the 15 mL conical tube to the
corresponding 50 mL conical tube (see Note 4).
7. For each plasma sample, record the total plasma volumes and
hemolysis grade:
l Plasma Color (Hemolysis Grade): Yellow (None), Pink/
Orange (Moderate), Red/Dark Red (Severe).
l For none or moderately hemolyzed samples: continue to
process for severely hemolyzed samples OR samples with a
total volume of <6 mL: re-draw for a new sample.
8. For plasma samples with a volume 6 mL–9.9 mL that will be
extracted immediately, add PBS to bring the total volume to
10 mL.
9. Discard the 15 mL conical tubes in a biohazard waste bin.
10. Store plasma samples that won’t be extracted immediately at
4–8 C or proceed to ccfDNA Extraction.
3.2 Manual DNA 1. Add 200 mL 100% isopropanol to 300 mL Buffer ACB con-
Isolation from Plasma centrate to obtain 500 mL Buffer ACB. Mix well by inverting
Using a Modified 10 times after adding isopropanol.
QIAamp Circulating 2. Add 25 mL 96–100% ethanol to 19 mL Buffer ACW1 concen-
Nucleic Acid Kit trate to obtain 44 mL Buffer ACW1. Mix well by inverting
10 times after adding ethanol.
3.2.1 ccfDNA Preparation
3. Add 30 mL 96–100% ethanol to 13 mL Buffer ACW2 concen-
trate to obtain 43 mL Buffer ACW2. Mix well by inverting
10 times after adding ethanol.
3.2.2 ccfDNA Extraction 1. For each 50 mL conical, label a LoBind 1.5 mL microcentri-
fuge tube with the corresponding sample ID.
2. For plasma samples with a volume of <10 mL that were stored
at 4–8 C, add PBS to bring the sample up to 10 mL.
3. Dispense 900 μL of Proteinase K into each plasma sample. Cap
and invert each sample tube 5 times to mix the contents.
4. Dispense 8.8 mL of ACL to each plasma sample. Vortex each

sample tube for 5 s.
5. Incubate the 50 mL plasma sample tubes in a 60 C water bath
for 45 min (see Note 6).
6. Dispense 18 mL of ACB to each plasma sample. Cap and invert
each sample tube 10 times to mix the contents.
7. Incubate the plasma sample tubes in a wet ice bath for 10 min.
8. Label a QIAamp Mini spin column and column extender
corresponding to the sample ID indicated on the 50 mL coni-
cal tubes.
9. Insert the column extenders into the corresponding spin
columns.
10. Insert spin column/extender assemblies into VacConnectors
on the QIAvac 24 Plus (see Note 7).
11. Connect the vacuum tubing to the QIAvac 24 Plus vacuum
port.
12. Carefully pour each plasma sample lysate into the
corresponding column extender. Take care not to overfill the
extenders.
13. Engage the vacuum by turning on the compressor and adjust-
ing the pressure to 800 mbar (see Note 8).
14. Refill the tube extenders as the plasma sample lysate volumes
are drawn down.
15. Discard the 50 mL conical tubes and column extenders in a
biohazard waste bin.
16. Turn on heat block and set to 56 C (see Note 9).
17. Once the entire volume of the plasma sample lysates have been
drawn through, dispense 600 μL of Buffer ACW1 into each
column. Leave the lids open and allow to drain, with the
vacuum compressor still running (see Note 10).
18. Dispense 750 μL of Buffer ACW2 into each column. Leave the
lids open and allow to drain with the vacuum compressor still
running.
19. Disengage the vacuum by lowering the pressure to 0 mbar and
turning the power off.
20. Dispense 750 μL of 100% ethanol to each spin column, leaving
lids open.
21. Engage the vacuum by turning on the compressor and adjust-
ing the pressure to 800 mbar and allow columns to drain.
22. Turn off the vacuum by lowering the pressure to 0 and turning
the power off.
23. Close the caps and remove the columns from the vacuum
manifold. Place the columns into 2.0 mL collection tubes.
24. Discard the VacConnectors in a biohazard waste bin.
25. Place a pre-aliquoted tube of DNA Suspension Buffer in a heat
block at 56 C for at least 10 min.
26. Centrifuge the column/collection tube assemblies in a micro-
fuge at 16,800 g for 3 min.
27. Once the 3 min spin has finished, transfer the columns into
new 2.0 mL collection tubes, open the lids and place in a heat
block at 56 C for 10 min.
28. Assemble the labeled 1.5 mL LoBind microcentrifuge tubes
into a 96-well tube rack.
29. Dispense 50 μL of the preheated DNA Suspension Buffer into
the center of each filter column (see Notes 11 and 12).
30. Transfer the filter columns from the 2.0 mL collection tubes to
the corresponding 1.5 mL LoBind microcentrifuge tubes.
31. Close the column lids and incubate at room temperature
(15 C to 25 C) for 7 min.
32. Centrifuge the spin column/microcentrifuge tube assemblies
in a microfuge at 16,800 g for 1 min. Check that the volumes
appear consistent, and that no residual buffer is in the filter (see
Note 13).
33. Discard the filter columns; cap the microcentrifuge tubes that
now contain the DNA eluate and store at 30 C to 10 C in
a pre-PCR room.
34. Pour the vacuum waste into a labeled chemical waste container.
35. Rinse the vacuum line twice with water.
3.3 Automated DNA 1. Turn on the instrument and log on after initialization.
Isolation from Plasma 2. Load the required elution rack into the “Eluate” drawer in
Using a Modified “Elution slot 1.”
Protocol for the
3. Make sure the “Waste” drawer is properly prepared and per-
QIAsymphony form an inventory scan of the “Waste” drawer, including the tip
Circulating DNA Kit chute and liquid waste.
4. Load the required reagent cartridge(s) and consumables into
the “Reagents and Consumables” drawer. Vortex the bead
trough containing the magnetic particles vigorously for at
least 3 min before first use. Make sure that the piercing lid is
placed on the reagent cartridge and the lid of the magnetic-
particle trough has been removed or, if using a partially used
reagent cartridge, make sure the Reuse Seal Strips have been
removed. Perform an inventory scan of the “Reagents and
Consumables” drawer.
5. Aliquot the required plasma volume (see Note 14) into a

sample tube and place the samples into the appropriate sample
carrier, and load them into the “Sample” drawer. Plasma
volumes from 1 mL to 10 mL can be used as sample input
volume.
6. Using the touchscreen, enter the required information for each
batch of samples and for Proteinase K to be processed which
includes: sample information (depending on sample racks
used), protocol to be run (1–10 mL) and elution volume and
output position.
7. Place the Proteinase K into the appropriate sample carrier on
positions 1 and 2, and load them into slot A of the “Sample”
drawer. 11. Define the Proteinase K by pressing the IC button.
8. Press the Run button to start the purification procedure. All
processing steps are fully automated. At the end of the protocol
run, the status of the batch changes from RUNNING to
COMPLETED.
9. Retrieve the elution rack containing the purified nucleic acids
from the “Eluate” drawer (see Note 15). The DNA is ready to
use or can be stored at 2–8 C, 20 C, or 80 C.
4 Notes
1. CcfDNA is detectable in serum but due to the agglutination

process when blood draw takes place in serum BCTs ccfDNA
fraction in supernatant is strongly contaminated with fragmen-
ted genomic DNA. Therefore usage of serum BCT is not
recommended for ccfDNA isolation.
2. Detection of ccfDNA in eluates: Due to the very low concen-
trations of ccfDNA in sample materials, measurement of DNA
concentration by using a spectrophotometer is not recom-
mended. For determination of concentration of circulating
cell-free DNA, a sensitive and accurate fluorescence-based
quantitation assay such as Quant-iT or PicoGreen-based assays
(Thermo Fisher Scientific, Waltham, MA, USA), or a real-time
PCR assay targeting single or multi-copy genes such as 18S
RNA gene [20], or hRNaseP and hTERT, or hAlu and hrDNA
repeats should be used. Because carrier RNA interferes with
fluorescence-based assays, a real-time PCR assay is preferred for
DNA quantification.
3. Blood Collection tubes offered for ccfDNA applications target
two aims: Firstly, to stabilize the extracellular DNA (ccfDNA)
in the plasma fraction of whole blood and secondly, the pre-
servatives in such BCT tubes stabilize nucleated blood cells for
extended times, thereby inhibiting the release of cellular
(nuclear) DNA, so that additional maternal genomic DNA is

prevented from further diluting the fetal fraction [16]. Hence,
these stabilizing BCTs allow for delaying the plasma separation
up to several days after blood draw which makes it possible to
ship the primary blood tube to a testing laboratory where the
plasma is then separated. It is technically challenging to achieve
such a stabilization effect for several days without affecting the
integrity and purity of the ccfDNA fraction while ensuring, in
parallel, the quantitative bioavailability of the ccfDNA for sen-
sitive downstream applications after extraction from plasma.
Therefore, the need for sample stabilization and the need for
obtaining highly pure ccfDNA should be considered carefully
because both aspects may have an impact on the reliability of
results, making it necessary to compare the different tubes
offered. The recently available PAXgene Blood ccfDNA Tube
uses a non-crosslinking chemistry to stabilize whole blood
while maintaining the quality and yield of ccfDNA.
4. Do not disrupt the blood cells.
5. After the first centrifugation step to separate the blood cell
fraction from the plasma fraction and subsequent transfer of
the plasma to a secondary tube, a second centrifugation step
may be required due to the following considerations: (A) to
ensure that remains of the buffy coat layer which are acciden-
tally transferred after first centrifugation are completely
removed from plasma to prevent genomic DNA contamination
of ccfDNA fraction (low to medium speed centrifugation)
and/or (B) to remove microvesicles (mainly exosomes) which
also contain cell-free but membrane enclosed RNA and DNA
which may also be extracted and measured in downstream
analysis in parallel to the ccfDNA fraction (high speed centrifu-
gation required).
6. Ensure that the isopropanol has been added to the Buffer ACB.
7. To avoid air leaks, make sure all connectors are firmly secure
and close unused slots with plugs, or close the inserted
VacValve.
8. Do not readjust the pressure, once the vacuum process has
started.
9. Ensure that ethanol was added to the Buffer ACW1.
10. Ensure that ethanol was added to the Buffer ACW2.
11. Do not touch the filter membrane with the pipette tip.
12. DNA Suspension Buffer used alternatively to elution buffer
AVE for elution of ccfDNA.
13. If residual buffer is present on the lip where the filter joins the
column, or in the cap, pipet the buffer to the center of the filter
membrane and centrifuge in a microfuge at 16,800 g for

1 min.
14. Automated DNA isolation: Make sure that correct sample
volume including void volume is used to avoid sample flagging
when using liquid-level detection mode (e.g., for 10 mL pro-
tocol 10.5 mL volume is required). If not sufficient sample is
available fill up sample with PBS to obtain the required sample
volume before loading onto the instrument. Alternatively, FIX
labware has been designed to minimize dead volumes (e.g., for
10 mL protocol 10.1 mL volume is required). But it must be
considered that FIX labware does not support liquid-level
detection or clot detection. As further alternative the “Enable
less sample” mode has been designed to use all available liquid
in combination with liquid-level detection and clot detection
in case less volume than required has been detected during
sample transfer (e.g., for 10 mL protocol at least 9.4 mL
volume is required). “Enable less sample” mode results in
unclear flagged samples.
Bubbles/foam in sample input tube may result in false
liquid level detection and subsequent incomplete sample trans-
fer. Remove bubbles in sample tube and avoid pipetting of
sample on the inner wall of the vessel during sample loading.
15. Automated DNA isolation: It is recommended to remove the
eluate plate from the “Eluate” drawer immediately after the
run has finished. Depending on temperature and humidity,
elution plates left in the QIAsymphony SP after the run is
completed may experience condensation or evaporation.
In general, magnetic particles are not carried over into eluates.
If carryover does occur, magnetic particles in eluates will not
affect most downstream applications. If magnetic particles
need to be removed before performing downstream applica-
tions, tubes or plates containing eluates should first be placed
in a suitable magnet and the eluates transferred to a clean tube.
Disclaimer
The scalable ccfDNA isolation methods presented here are for

performance evaluations only.
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978-90-481-9382-0_35
Chapter 22
Noninvasive Detection of Fetal Aneuploidy Using Next

Generation Sequencing
Kirsten J. Curnow, Rebecca K. Sanderson, and Sue Beruti
Abstract
Cell-free DNA (cfDNA)-based noninvasive prenatal testing (NIPT) utilizing next generation sequencing
(NGS) is a highly sensitive and specific approach designed to screen for fetal aneuploidy. NIPT was first
introduced in 2011 and has been rapidly adopted in a clinical setting because of the improved performance
afforded compared with traditional prenatal serum screening options. We describe a PCR-free, paired-end
sequencing-based NIPT, the VeriSeq NIPT Solution. This NIPT screens for fetal aneuploidy of chromo-
somes 21, 18, 13, X, and Y. Using the validated approach detailed here, users can achieve high sensitivities
and specificities for trisomies 21, 18, and 13 as well as sex chromosome aneuploidies with low failure rates.
The automated workflow can be completed in 1 day, with only 2 h of hands-on time from a single
technician.
Key words Next generation sequencing, Paired-end sequencing, Noninvasive prenatal test, Cell-free
DNA, PCR-free, Prenatal screening, Aneuploidy, Trisomy, Down syndrome
1 Introduction
Prenatal screening is currently used for the detection of common

fetal aneuploidies. Traditional screening options include measure-
ment of first and/or second trimester serum biomarkers and ultra-
sound examinations; these tests are primarily designed to screen for
trisomy 21 (Down syndrome), with some tests also reporting a risk
for trisomy 18 and trisomy 13. Although these tests have a sensitiv-
ity of around 80–90% for trisomy 21 [1], they have a false positive
rate of around 5%. As a result, these tests have low positive predic-
tive values (~5%), which means that only around 1 in 20 patients
with a positive test have an affected pregnancy. Thus, there was a
need for a more accurate approach with lower false positive rates
thereby reducing the number of women undergoing unnecessary
invasive procedures.
The discovery of cell-free DNA (cfDNA) in blood and deter-
mination that in pregnant women it is a mixture of maternal cfDNA
325
326 Kirsten J. Curnow et al.
and fetal cfDNA (primarily placental origin) drove development of

noninvasive prenatal testing (NIPT). Molecular analysis of cfDNA
isolated from a maternal peripheral whole blood sample has proven
to be a much more accurate and comprehensive method of screen-
ing for fetal aneuploidy [2]. With sensitivities above 90% and
specificities of around 99.9%, NIPT has positive predictive values
above 50%, which is around tenfold higher than biomarker-based
tests [2]. A range of molecular technologies are used for NIPT:
microarray, targeted sequencing, and whole-genome sequencing
(WGS). Although test sensitivities and specificities are similar
between the different methodologies, there are marked differences
in the test failure rates. Whole-genome sequencing has several
advantages: lower technical failure rates than targeted sequencing
and microarray-based approaches [3–5]; data from non-test chro-
mosomes can be used as a reference during analysis; comprehensive
coverage information that can be useful in determining sample
quality and fetal fraction estimation; relative ease of expanding the
test menu, such as screening for other autosomal aneuploidies or
select microdeletion syndromes.
Herein, we describe a validated WGS-based NIPT used to
screen pregnant women of at least 10 weeks’ gestation for fetal
aneuploidy on chromosomes 21, 18, 13, X, and Y; the VeriSeq™
NIPT Solution [3] is available as an IVD in select countries and
regions. This technology is also the basis for a commercially avail-
able NIPT offered in the CLIA-certified Illumina laboratory (Fos-
ter City, CA). The principles of the method are as follows: sample
collection, plasma isolation, cfDNA extraction, library preparation,
quantification, library pooling and sequencing, and analysis. Sam-
ple collection requires a minimum of 7 mL of maternal peripheral
whole blood collected in a single Streck cell-free blood collection
tube, which prevents cell lysis and genomic contamination and
stabilizes the sample at room temperature. Subsequent sample
preparation steps are part of an automated workflow utilizing a
liquid handling system, and can be completed in just over a day
with around 2 h of hands on time from a single technician. Plasma is
isolated using standard centrifugation techniques. Cell-free DNA is
extracted by adsorption onto a binding plate, washes, and then
elution. The library preparation process is PCR-free, and involves
end repair to convert 50 and 30 overhangs to blunt ends, addition of
a single deoxyadenosine nucleotide at the 30 end, then ligation of
indexed adaptors to the cfDNA fragments. The adaptors serve two
purposes: sample identification during sequencing; library capture
on the solid surface of the sequencing flow cell for cluster genera-
tion and sequencing. After quantitation of library yield, 48-sample
batch libraries are pooled; the amount of each library is adjusted to
minimize variation in coverage. The library pool then undergoes
paired-end sequencing on a next generation sequencer. Once
sequencing is complete, the data is analyzed with nucleotide base
cfDNA-Based Noninvasive Prenatal Aneuploidy Screening 327
calls made from the signal intensity measurements taken during

sequencing. Paired-end read information is used to assess coverage
and individual fragment size. As the size profiles of maternal and
fetal cfDNA differ, fragment size information is useful in making
fetal fraction estimates. Further, analysis is refined by binning frag-
ment size, with a relative increase in the fetal signal achieved by
focusing on shorter fragments. Next, secondary data analysis is
performed: Reads are demultiplexed using the index sequences;
sequence reads are mapped to the human genome; the number of
unique reads is calculated within each 100 kb genomic bin; cover-
age is normalized on a subchromosomal level. The analysis software
employs an algorithm based on multiple data inputs, including
sequencing coverage, sequence read quality, and estimated fetal
fraction, to determine under or over representation of chromo-
somes 21, 18, 13, X, and Y. The assay uses a dynamic threshold
metric to determine whether the sample can be reported, obviating
the need for an arbitrary fetal fraction cutoff. The NIPT protocol
and reagents described herein result in the lowest failure rate
among in-lab NIPT offerings with comparably high clinical
performance.
2 Materials
2.1 Equipment 1. VeriSeq Onsite Server with preinstalled VeriSeq NIPT Assay
Software (Illumina, Inc., San Diego, CA, USA).
2. VeriSeq NIPT Microlab STAR (Hamilton Company, Reno,
NV; Hamilton Company, Bonaduz, Switzerland) with prein-
stalled VeriSeq NIPT Workflow Manager.
3. Next generation sequencing (NGS) system with the following
capabilities: 2 36 bp paired-end sequencing; compatible with
VeriSeq NIPT Sample Prep dual index adapters; automatic
production of BCL files; two dye-based chemistry; 400 million
paired-end reads per run; compatible with VeriSeq NIPT Assay
Software.
4. Centrifuge and rotor assemblies for blood collection tubes with
the following specifications: Refrigerated centrifuge capable of
1600 g with no-brake option; swinging bucket rotor with
buckets; bucket inserts, 48 or 96 tube capacity, 76 mm mini-
mum depth; insert adapters to support 16 100 mm blood
collection tubes. Recommended: Allegra 6 Series Centrifuge,
1600 g (Beckman Coulter, Indianapolis, IN); Allegra Cen-
trifuge GH-3.8 Rotor with buckets (Beckman Coulter,
Indianapolis, IN); Allegra Centrifuge Bucket Covers, set of
two (Beckman Coulter, Indianapolis, IN); Allegra Centrifuge
Adapter Assembly, 16 mm, set of four (Beckman Coulter,
Indianapolis, IN).
5. Centrifuge and rotor assemblies for microplates with the fol-

lowing specifications: Centrifuge capable of 5600 g; swing-
ing plate rotor with 96-well plate carriers, 76.5 mm minimum
depth; support base for microplates. Recommended: Sorvall
Legend XTR Centrifuge (Thermo Fisher Scientific, Waltham,
MA); HIGHPlate 6000 Microplate Rotor (Thermo Fisher
Scientific, Waltham, MA); either MicroAmp 96-Well Support
Base (Thermo Fisher Scientific, Waltham, MA) or 96-Well
PCR Plate Carrier (Thermo Fisher Scientific, Waltham, MA).
6. Gemini XPS or SpectraMax M2 Microplate reader (Molecular
Devices, San Jose, CA).
7. SpectraMax High-Speed USB, Serial Adapter (Molecular
Devices, San Jose, CA).
8. Thermal cycler with the following specifications: heated lid;
4 C to 98 C temperature range; 2 C temperature accuracy;
2 C per second minimum ramp rate; compatible with Twin.tec
PCR Plate 96-well, full skirt.
9. 1000 μL Conductive Non-Sterile Filter Tips.
12. Deep-Well Reservoir (Corning Axygen, Tewksbury, MA).
13. MagNA Pure LC Medium Reagent Tubs 20 (Roche, Basel,
Switzerland).
14. Deep-Well Plate 96 (Eppendorf, Hauppauge, NY).
15. Low Volume 384 Well Black Flat Bottom Polystyrene Micro-
plate (Corning, Tewksbury, MA).
16. Twin.tec PCR Plate 96-well with a full-skirt (Eppendorf,
Hauppauge, NY).
17. Microseal ‘F’ Foil (Biorad, Hercules, CA) or Foil seals (Beck-
man Coulter, Indianapolis, IN).
18. Cell-Free DNA Blood Collection Tube (BCT) CE (Streck, La
Vista, NE).
19. Push Caps (Sarstedt, Numbrecht, Germany).
2.2 Reagents 1. VeriSeq NIPT Sample Prep Kit (Illumina, Inc., San Diego, CA,
USA). The kit contains an Extraction Box, Library Prep Box,
Accessory Box, and Workflow Tubes and Labels.
2. 100% Ethanol for molecular biology.
3. DNase/RNAse-free water.
4. Optional: Dulbecco’s Phosphate-Buffered Saline (DPBS) for
no template control (NTC).
2.3 Cell-Free DNA For steps 1 to 3 below, reagents to be prepared are supplied in the
Extraction Extraction and Accessory Boxes of the VeriSeq NIPT Sample Prep
Kit. Label the reservoir tubs and deep-well reservoirs with the name
of the reagents.
1. Final Plasma deep-well plate. If previously refrigerated, let it
stand for 30 min to bring to room temperature. Unseal plate
before use.
2. Proteinase K: Prepare by slowly adding 3.75 mL Proteinase
Buffer to each reagent vial, cap the vial and vortex until resus-
pended. For 48 samples, use 3 reagent vials. For 96 samples,
use 4 reagent vials. Pool prepared reagent from all vials into a
reagent tub.
3. Wash Buffer II: Add 100 mL 100% EtOH to each reagent
bottle, inverting to mix. For 48 samples, prepare 1 bottle.
For 96 samples, prepare 2 bottles. Mark checkbox on bottle/s.
4. 70% EtOH cleaning solution for cleaning vacuum system: 70%
Ethanol, 30% DNase/RNAse-free water.
2.4 Library For steps 1 to 9 below, reagents to be prepared are supplied in the
Preparation Library Prep and Accessory Boxes of the VeriSeq NIPT Sample
Prep Kit. Label the reservoir tubs and deep-well reservoirs with the
reagent names.
1. End Repair Mix: Thaw at room temperature and then vortex
to mix.
2. A-Tailing Mix: Thaw at room temperature, vortex to mix, and
then briefly centrifuge.
3. Ligation Mix: Thaw at room temperature, vortex to mix, and
then briefly centrifuge.
4. Resuspension Buffer: Vortex to mix. Return to storage
after use.
5. Hybridization Buffer: Thaw at room temperature and then
vortex to mix. Return to storage after use.
6. VeriSeq NIPT DNA Adapter Plate: Thaw at room temperature,
vortex to mix, and then centrifuge at 1000 g for 20 s. Apply a
plate barcode.
7. Sample Purification Beads: Let it stand for 30 min to bring to
room temperature before mixing by vortexing or inversion
until all beads are in suspension and the mixture is homoge-
nous. Vortex vigorously before each use.
8. Combine A-Tailing Mix and Resuspension Buffer in a screw
cap tube, vortex to mix, and then briefly centrifuge. For 48 sam-
ples: 900 μL A-Tailing Mix, 1200 μL Resuspension Buffer. For
96 samples: 1800 μL A-Tailing Mix, 2400 μL Resuspension
Buffer.
9. Combine Ligation Mix and Resuspension Buffer in a screw cap

tube, vortex to mix, and then briefly centrifuge. For 48 samples:
230 μL Ligation Mix, 1713 μL Resuspension Buffer. For
96 samples: 440 μL Ligation Mix, 3278 μL Resuspension
Buffer.
10. cfDNA Elution Plate: If previously stored, thaw at room tem-
perature, vortex for 1 min, and then centrifuge at 1000 g
for 20 s.
11. 80% Ethanol: 40 mL 100% Ethanol, 10 mL DNase/RNAse-
free water.
2.5 Library For steps 1 to 3 below, reagents to be prepared are supplied in the
Quantification Accessory Box of the VeriSeq NIPT Sample Prep Kit.
1. DNA Quantification Reagent: Thaw at room temperature.
Vortex to mix, and then briefly centrifuge. Protect from light.
2. DNA Quantification Standard: Vortex to mix, and then briefly
centrifuge.
3. Resuspension Buffer: Vortex to mix.
4. Libraries Plate: If previously stored, thaw at room temperature,
vortex to mix, and centrifuge at 1000 g for 20 s.
2.6 Library Pooling For step 1 below, the reagent to be prepared is supplied in the
Library Prep Box of the VeriSeq NIPT Sample Prep Kit.
1. Hybridization Buffer: Thaw at room temperature, and then
vortex to mix. Return to storage after use.
2. Libraries Plate: If previously stored, thaw at room temperature,
vortex at 1500 rpm for 1 min, then centrifuge at 1000 g for
20 s.
2.7 Preparing 1. Pool tubes: If previously stored, thaw at room temperature.

for Sequencing Vortex briefly, then centrifuge briefly.
3 Methods
3.1 Process Blood 1. Centrifuge barcoded blood samples at 1600 g for 10 min at
Samples 4 C (see Note 1). Begin plasma isolation (see Subheading 3.2)
within 15 min of completing centrifugation. Repeat centrifu-
gation if more than 15 min has elapsed.
2. Remove samples tubes from the centrifuge and visually inspect
each tube to confirm that it contains at least 1.5 mL plasma
above the buffy coat (see Note 2).
3. Uncap tubes and load all samples and any plasma controls into
tube carriers (see Note 3).
The VeriSeq NIPT Solution utilizes a liquid handling system

(VeriSeq NIPT Microlab STAR [ML STAR]) to minimize required
hands-on time. Plasma Isolation (see Subheading 3.2), cell-free
DNA extraction (see Subheading 3.3), and library preparation (see
Subheading 3.4) steps are largely performed by the ML STAR (see
Notes 4, 5).
3.2 Plasma Isolation 1. Label 1 deep-well plate Intermediate Plasma and apply a
barcode.
3.2.1 Preparation
2. Label 1 deep-well plate Final Plasma and apply a barcode.
3.2.2 Procedure 1. On the ML STAR, open the AppLauncher, and then click
VeriSeq NIPT Method.
2. Enter the Batch ID and username, and then click OK.
3. Click New Batch, and after initiation, click OK to begin plasma
isolation.
4. Perform one of the following: To load an existing sample sheet,
select the sample sheet associated with the batch, and then click
OK; to proceed without selecting a sample sheet, click No
Sample Sheet. The sample sheet is a comma-delimited file (Sam-
pleSheet.csv) that stores the information to set up and analyze a
sequencing experiment. The file includes a list of samples and
their index sequences, as well as the workflow to be employed.
5. Select the batch size, and then click OK. Select the number of
no template controls (NTCs), and then click OK.
6. Confirm that all barcodes are affixed, and load the samples, tips,
and plates onto the carrier (see Table 1). Click OK after each
load prompt.
7. Make sure the carriers, labware, and reagents are loaded cor-
rectly, and then click OK in the Pre-Spin Deck Verification
screen.
8. When alerted by the Workflow Manager, make sure that the
ML STAR loading deck is free of any obstructions to allow the
ML STAR to unload the carriers, and then click Unload.
9. Remove the Intermediate Plasma deep-well plate. Visually
inspect the plate for consistent volumes in each well, the
expected volume is 1000 μL. Make note of any inconsistencies
and record at the end of the Plasma Isolation procedure. Seal
the plate and centrifuge at 5600 g for 10 min.
10. Click Yes to proceed to final Plasma Preparation.
11. Remove the plate seal and reload the plate as detailed in
Table 1.
12. Select the Intermediate Plasma plate has been spun checkbox,
and then click OK.
Table 1
ML STAR loading details for the plasma isolation procedure
Procedural Sample batch Carrier Site

step size type Track Item position
6 48 Tip 7–121000 μL tips 5
Tube 15 Prepared blood samples tubes 1–24 1–24
Multiflex 19–24
Empty deep-well plate, Final Plasma - 4
barcoded
Multiflex 19–24 Empty deep-well plate, Intermediate 5
Plasma – barcoded
Reagent 47 [Optional] DPBS for no template control 5
96 Tip 7–12 1000 μL tips 4, 5
Multiflex 19–24 Empty deep-well plate, Final Plasma - 4
barcoded
Multiflex 19–24 Empty deep-well plate, Intermediate 5
Plasma – barcoded
Reagent 47 [Optional] DPBS for no template control 5
11 48, 96 Multiflex 19–24 Intermediate Plasma deep-well plate 5
DPBS Dulbecco’s phosphate-buffered saline
ML STAR to unload the carriers, and then click Unload.
14. When prompted by the Workflow Manager, empty the carriers
and deck.
15. Remove the Final Plasma deep-well plate. Visually inspect the
plate for consistent volumes in each well (the expected volume
is 900 μL), visible cell pellets, and excessive hemolysis. If you
observe a visible cell pellet or excessive hemolysis, invalidate the
affected sample at the end of the Plasma Isolation method or
use Batch Manager.
16. When prompted by the Workflow Manager, click OK.
17. Enter comments about affected wells, and then click OK.
18. Perform 1 of the following: To continue to cfDNA Extraction,
click Yes; To stop, click Exit (see Note 6).
3.3 Cell-Free DNA 1. Label 1 new full-skirt plate Intermediate, and apply a plate
Extraction barcode.
3.3.1 Preparation 2. Label 1 new full-skirt plate cfDNA Elution, and apply a plate
barcode.
3. Label 1 new deep-well plate Extraction Intermediate, and apply
a deep-well plate barcode.
4. Apply a plate barcode to the DNA Binding plate.
5. Prepare the vacuum system: Remove the vacuum manifold and
clean with 70% EtOH; empty the vacuum waste; ensure the ML
STAR vacuum system is on.
3.3.2 Procedure 1. Click OK to start cfDNA Extraction. If the VeriSeq NIPT

Method is not already open: Open the AppLauncher, and
click VeriSeq NIPT Method; enter the Batch ID and username,
and then click OK.
2. Load tips onto the tip carriers as detailed in Table 2, and then
click OK.
3. Load counted tips onto the tip carriers as detailed in Table 2.
4. Enter the location of the first and last tips for each tip rack, and
then click OK.
5. Scan Extraction Box barcodes. Enter the user name or reagent
preparer initials, and then click OK.
6. Scan Accessory Box barcodes. Enter the user name or reagent
7. Confirm that barcodes are affixed, unseal the Final Plasma
deep-well plate if necessary, and load plates onto the plate
carrier as detailed in Table 2, and then click OK.
8. Confirm that the DNA Binding plate is barcoded, then
click OK.
9. For a 48-sample batch size, cut a seal in half width-wise and
apply over unused columns 7–12 of the plate before loading
onto the vacuum manifold.
10. Load the DNA Binding plate onto the vacuum manifold with
the barcode facing right, and then click OK.
11. Load reagent tubs onto the reagent carrier as detailed in
Table 2, and then click OK.
12. Transfer the specified reagents into the deep-well reservoirs,
and then load the reservoirs onto the deep-well carriers as
detailed in Table 2, and then click OK.
13. Wait for the automated reagent volume check to complete.
14. Confirm that vacuum waste is not more than half full (empty
recommended), and then click OK.
Table 2
ML STAR loading details for the cfDNA extraction procedure

2 48 Tip 1–6 1000 μL tips 1, 2
7–12 300 μL tips 1
96 Tip 1–6 1000 μL tips 1, 2, 3, 4
7–12 300 μL tips 1
3 48, 96 Tip 49–54 1000 μL tips 1
300 μL tips 2
50 μL tips 3
7 48, 96 Multiflex 19–24 New full-skirt plate, Intermediate - 1
barcoded
New full-skirt plate, cfDNA Elution - 2
barcoded
New deep-well plate, Extraction 4
Intermediate - barcoded
Final Plasma deep-well plate - barcoded 5
11 48 Reagent 47 16 mL Elution Buffer 1
11 mL Proteinase K 2
96 Reagent 47 16 mL Elution Buffer 1
15 mL Proteinase K 2
12 48 Deep- 39–44 125 mL Wash Buffer II 1
well 125 mL Wash Buffer I 2
60 mL 100% EtOH 3
100 mL Lysis Buffer 4
60 mL DNase/RNase-free water 5
96 Deep- 39–44 200 mL Wash Buffer II 1
well 125 mL Wash Buffer I 2
100 mL 100% EtOH 3
100 mL Lysis Buffer 4
100 mL DNase/RNase-free water 5
15. Confirm the placement of all carriers, labware, and reagents,

and then click OK in the Extraction Deck Verification screen.
16. After the final vacuum step, centrifuge the DNA Binding plate,
and then click OK: Remove the DNA Binding plate and clean
the bottom surface with 70% EtOH; seal any uncovered wells
on the DNA Binding plate and place it on the empty Final
Plasma deep-well plate; centrifuge the DNA Binding plate/
Final Plasma plate assembly at 5600 g for 10 min with the
brake on.
17. During DNA Binding plate centrifugation, complete the vac-
uum cleaning: Wait for the automated waste disposal to com-
plete; clean the vacuum manifold and inside the vacuum system
with 70% EtOH, and then replace the vacuum manifold; select
the Manifold is on Vacuum checkbox, and then click OK.
18. Remove vacuum manifold, and then click OK.

19. After centrifugation, unseal the wells containing samples on
the DNA Binding plate and place it on the cfDNA Elution
plate. The cfDNA Elution plate is on the vacuum manifold.
Load the DNA Binding plate with the barcode to the right, and
then click OK.
20. After the incubation step, select the Plates are assembled as
indicated checkbox, confirming that the DNA Binding/
cfDNA Elution plate assembly is on a support base
(if required by centrifuge).
21. Seal the uncovered wells on the DNA Binding plate and cen-
trifuge at 5600 g for 2 min with the brake on, and then
click OK.
22. Visually inspect the cfDNA Elution plate for consistent
volumes in each well. Expected volume is approximately 55 μL.
23. Seal and retain the cfDNA Elution plate for library preparation.
ML STAR to unload the carriers, and then click Unload to
unload the deck.
25. Unload all carriers and clean the ML STAR deck, and then
click OK.
27. Perform one of the following: To continue to Prepare
Libraries, click Yes; to stop. Click Exit (see Note 6).
3.4 Library 1. Label 1 new full-skirt plate Libraries and apply a plate barcode.
Preparation 2. Make sure that the ML STAR thermal control is on.
3.4.1 Preparation
3.4.2 Procedure 1. Click OK to start Library Preparation. If the VeriSeq NIPT

Method is not already open: Open the AppLauncher, and click
VeriSeq NIPT Method; enter the Batch ID and username, and
then click OK.
2. Confirm that the following are prepared as indicated in the
Reagent Preparation screen: A-Tailing Mix, Ligation Mix, and
80% EtOH; Sample Purification Beads, End Repair Mix, and
VeriSeq NIPT DNA Adapter Plate.
3. Select the checkboxes, and then click OK.
4. Scan Library Prep Box barcodes. Enter the user name or
reagent preparer initials, and then click OK.
Table 3
ML STAR loading details for the library preparation procedure

6 48 Tip 1–6 50 μL tips 1, 2
7–12 300 μL tips 1, 2, 3, 5
96 Tip 1–6 50 μL tips 1, 2, 3, 4
7–12 300 μL tips 1, 2, 3,
4, 5
7 48, 96 Tip 49–54 1000 μL tips 1
300 μL tips 2
50 μL tips 3
9 48, 96 Multiflex 19–24 cfDNA Elution plate - barcoded 1
DNA Adapter plate – barcoded 2
New 96-well full-skirt plate, libraries - 3
barcoded
New 96-well full-skirt plates 4, 5
10 48, 96 Deep- 39–44 50 mL 80% EtOH in a deep-well 1
well reservoir
New 96-well full-skirt plates 2, 3, 4, 5
11 48, 96 Reagent 47 2.5 mL End Repair Mix 1
Prepared A-Tailing Mix 2
Prepared Ligation Mix 3
10 mL Sample Purification Beads 4
12 mL Hybridization Buffer 5
6. Load tips onto the tip carriers as detailed in Table 3, and then
click OK.
7. If you stopped the protocol after the cfDNA Extraction proce-
dure, load counted tips onto the tip carriers as detailed in
Table 3.
8. Enter the location of the first tip for each tip rack, and then
click OK.
9. Confirm that barcodes are affixed, and load plates (barcode
facing right) onto the plate carrier as detailed in Table 3, and
then click OK.
10. Load the deep-well carrier as detailed in Table 3, and then
click OK.
12. Make sure that the carriers, labware, and reagents are loaded as
indicated, and then click OK in the Library Deck Verification
screen.

unload the deck.
15. Visually inspect the Libraries plate for consistent volumes in
each well.
16. Seal and retain the Libraries plate.
17. Unload the carriers, clean the deck, and then click OK.
19. Perform one of the following: To continue to Quantify
Libraries, click Yes; to stop, click Exit (see Note 6).
3.5 Library 1. Turn on the fluorometer 10 min before use.

Quantification 2. Apply a plate barcode to a new 384-well plate.
3.5.1 Preparation 3. Apply a plate barcode to a new full-skirt plate.
3.5.2 Procedure 1. Click OK to start quantification. If the VeriSeq NIPT Method

is not already open: Open the AppLauncher, and click VeriSeq
NIPT Method; enter the Batch ID and username, and then
click OK.
3. Load tips onto the tip carrier as detailed in Table 4, and then
click OK.
4. Confirm that barcodes are affixed, unseal the Libraries plate,
and load plates (barcode facing right) onto the Multiflex carrier
as detailed in Table 4, and then click OK.
5. Load reagent tubes without caps into the tube carrier as
detailed in Table 4, and then click OK.
7. If you stopped the protocol after the Library Preparation pro-
cedure, load counted tips onto the tip carriers as detailed in
Table 4.
8. Enter the location of the first tip for each tip rack, and then
click OK.
indicated, and then click OK in the Quant Deck Verification
screen.
Table 4
ML STAR loading details for the library quantification procedure

3 48 Tip 1–6 300 μL tips rack 1
50 μL tips rack 2
96 Tip 1–6 300 μL tips rack 1
50 μL tips rack 2, 3
4 48, 96 Multiflex 19–24 New full-skirt plates - 1
barcoded
New 384-well plate - barcoded 2
Libraries plate – barcoded 3
New 96-well full-skirt plates 4, 5
5 48, 96 Tube 46 DNA Quantification Standard 1
DNA Quantification Reagent 2
6 48, 96 Reagent 47 New reagent tubs 1
16 mL Resuspension Buffer 2
7 48, 96 Tip 49–54 1000 μL tips 1
300 μL tips 2
50 μL tips 3
unload the deck.
12. Unload the Libraries plate. Visually inspect the plate for con-
sistent volumes in each well then seal the Libraries plate and
store at room temperature until the fluorometric data analysis
is complete.
13. Unload the remaining 96-well plates and visually inspect for
consistent volumes in each well. Gross errors in volume can
indicate an issue with pipetting steps.
14. Unload the 384-well plate and visually inspect for liquid in the
appropriate wells. Seal the plate with a foil seal; centrifuge at
1000 g for 20 s; then incubate at room temperature for
10 min, protected from light.
15. Unload all carriers and clean the ML STAR deck, and then
click OK.
16. After incubation, remove the foil seal and load the 384-well
plate onto the microplate reader. Make sure that A1 is in the
top left corner, and click Read.
17. Export the data as an. XML file as follows: Right-click Barcode,
select rename, scan the barcode of the Qualification plate, and
then click OK; at the upper-left corner, click the plate icon, and
then select Export from the menu; select the Expt1 checkbox,
set the output format to XML, and click OK; Set the output file
path and file name, and then click Save.
3.5.3 Analysis 1. On the ML STAR, in the Scanner Information screen, enter the
fluorometer ID.
2. Enter comments about the fluorometer run, and then
click OK.
3. Navigate to the. XML quantification file that contains the
fluorometric data, and then click OK.
4. Review the standards curve and sample concentration analysis
results, and then click OK.
5. If you need to rerun the plate, click Rescan. As samples are time
and light sensitive, perform the Rescan immediately (when
necessary).
7. Assess the results and proceed as follows: If the results pass
specification, proceed to Pool Libraries; If the results fail speci-
fication, the system aborts the method and you must repeat the
quantification procedures (see Subheading 3.5.2).
8. Perform one of the following: To continue to Pool Libraries,
click Yes; to stop, click Exit (see Note 6).
3.6 Library Pooling 1. Label an empty pooling tube Pool A. For 96 samples, label a
second empty pooling tube Pool B.
3.6.1 Preparation
2. Save the following denature program on a thermal cycler with a
heated lid: Choose the preheated lid option and set to 102 C;
set the reaction volume to 50 μL; set the ramp rate to 4 C per
second; incubate at 96 C for 10 min, and then 0 C for 5 s;
hold at 4 C.
3.6.2 Procedure 1. Place the Libraries plate on the preprogrammed thermal cycler
and run the denature program.
2. Click OK to start pool libraries. If the VeriSeq NIPT Method is
not already open: Open the AppLauncher, and click VeriSeq
NIPT Method; enter the Batch ID and username, and then
click OK.
3. Select the pool concentration, and then click OK.
4. The target cluster density is 220–260 k/mm2. If necessary,
adjust the pooling concentration to achieve the target cluster
density.
5. When prompted by the Workflow Manager, perform one of the
following: To load a sample sheet, select the sample sheet
Table 5
ML STAR loading details for the library pooling procedure
Procedural step Sample batch size Carrier type Track Item Site position
7 48, 96 Tip 7–12 50 μL filter tips 1
8 48, 96 Multiflex 19–24 Denatured Library plate 1
9 48 Tube 46 New 2 mL tube, Pool A 1
96 Tube 46 New 2 mL tube, Pool A 1
New 2 mL tube, Pool B 2
10 48 Reagent 47 3 mL Hybridization Buffer 1
96 Reagent 47 3 mL Hybridization Buffer 1
11 48, 96 Tip 49–54 1000 μL tips 1
300 μL tips 2
50 μL tips 3
associated with the batch, and then click Load; to use system
default values for remaining sample types or sex reporting, click
Use Default for each setting.
6. Click Start to begin timer for denaturing plate.
7. Load tips onto the tip carriers as detailed in Table 5.
8. Load the Denatured Library plate (barcode facing right) onto
the Multiflex carrier as detailed in Table 5, and then click OK.
9. Load pooling tubes onto the tube carrier as detailed in Table 5,
and then click OK.
11. Load tips onto the tip carriers as detailed in Table 5. Enter the
location of the first and last tips for each tip rack, and then
click OK.
indicated, and then click OK in the Pooling Deck Verification
screen.
unload the deck.
14. Unload the tube carrier. Cap each pooling tube, vortex, and
then centrifuge briefly.
15. Sequence libraries as soon as possible after pooling. Store the
Libraries plate at 25 C to 15 C for up to 7 days to enable
repooling, if necessary. The Libraries plate is stable for up to
7 days cumulative storage at 25 C to 15 C.
16. Click OK.

18. Click OK at Pooling Complete screen (see Note 6).
3.7 Preparation 1. Prepare the next-generation sequencing system with the fol-
for Sequencing lowing settings: Paired-end run with 36 x 36 cycle reads; dual
indexing with 8-cycle index reads; run name the same as
3.7.1 Preparation
Pool Name.
3.7.2 Procedure 1. Add buffer and library pool directly to the sequencer sample
cartridge as follows: 900 μL Hybridization Buffer; 450 μL Pool
A; pipette to mix.
2. Proceed with sequencing using a next-generation sequencing
system according to the manufacturer’s instructions.
3. Confirm correct run configuration when prompted.
4. Repeat procedure for Pool B, if necessary.
3.8 Sequencing Data 1. After sequencing is complete, sequencing data is automatically

Analysis and Result sent to the VeriSeq NIPT Assay Software for analysis and
Interpretation sequencing report generation (please note, this is not a clinical
report). The report includes classifications for each sample in
the batch as well as an assessment of all run QC metrics. The
analysis process takes approximately 4 h for a 48-sample batch.
2. The VeriSeq NIPT Solution employs an algorithm based on
multiple data inputs, including sequencing coverage, sequence
read quality, and estimated fetal fraction, to determine fetal
chromosomal representation.
3. For samples that pass QC, the VeriSeq NIPT Assay Software
automatically generates a result of ANEUPLOIDY
DETECTED or NO ANEUPLOIDY DETECTED for chro-
mosomes 21, 18, and 13 for each patient sample. A result of
ANEUPLOIDY DETECTED indicates that the sample has
screened positive for trisomy of the given chromosome.
4. Results on the fetal sex chromosome status are automatically
generated and optionally reported. When no sex chromosomal
aneuploidy is detected the report will state NO ANEU-
PLOIDY DETECTED appended with the sex classification:
XX (female fetal sample) or XY (male fetal sample). Sex chro-
mosome aneuploidies are reported as ANEUPLOIDY
DETECTED appended with the particular aneuploidy
detected: XXX, XXY, XYY, or XO (monosomy X). In rare
cases, the sex chromosome values fall outside the reportable
range and the system generates a result of SEX CHROMO-
SOMES NOT REPORTABLE; results for autosomal aneu-
ploidy may still be reported for these samples.
5. The VeriSeq NIPT Assay Software uses statistics generated

during sequencing to provide a fetal fraction estimation
(FFE) for each sample. The FFE is the estimated fetal cfDNA
component which is recovered by the assay and reported as a
rounded percentage for each sample. The average standard
deviation of this estimate across all samples is 1.3%. The FFE
is not to be used in isolation to exclude samples when reporting
results.
6. To make chromosomal representation calls, the VeriSeq NIPT
Assay Software utilizes the individualized Fetal Aneuploidy
Confidence Test (iFACT), a dynamic threshold metric that
indicates whether the system has generated sufficient sequenc-
ing coverage, given the fetal fraction estimate for each sample.
The system makes chromosomal representation calls only if a
sample meets the iFACT threshold. If a sample fails to achieve
this threshold, the QC assessment displays FAILED iFACT and
the system does not generate a result. The iFACT assessment is
applied to all samples.
7. In addition to iFACT, the VeriSeq NIPT Assay Software
assesses several other QC metrics during analysis. The addi-
tional metrics include assessments of coverage uniformity on
reference genomic regions and the distribution of cfDNA frag-
ment lengths. The QC assessment displays either a QC flag or a
QC failure for any metrics outside of the acceptable range (see
Table 6). In the case of QC failure, the system does not gener-
ate a result for the sample. If a sample fails QC, a second plasma
aliquot can be processed provided there is sufficient plasma
volume in the blood collection tube.
Table 6
ML STAR loading details for the library pooling procedure
Recommended
Failure mode Possible result Interpretation action Comments
Insufficient Sample QC Insufficient plasma Redraw Based on visual
input plasma failure volume inspection of
plasma volume.
Blood tube No separation of Sample was not Make sure that the
failure blood into centrifuged centrifuge started
layers and the tube was
spun at the correct
force. Redraw
sample.
Improper sample Redraw Frozen samples will
storage or not separate.
transport
(continued)
Table 6
(continued)
Recommended
Failure mode Possible result Interpretation action Comments
Sample clog Plasma Individual samples Inspect sample, if
OR slow contamination may clog the remaining plasma
flow binding plate if in tube is red or
there is significant milky, cancel
contamination in sample and request
the plasma sample redraw. If sample
appears normal,
retest sample.
Hardware Inadequate digestion Retest sample.
malfunction of material during
extraction
Individual Sequencing QC Insufficient genetic Check Sample Indicates a bad
Sample failure input OR Annotation. Check sample input OR a
Analysis QC Mistransfer during for similar mistransfer on the
failure sample handling performance on ML STAR.
previous samples in Insufficient
relative plate genetic material
position. Retest can come from
sample. insufficient
cfDNA in the
plasma or cell-
based DNA
causing over-
dilution of the
sample for
sequencing.
Low FF or NES Insufficient data Retest from plasma.
count generated to make
accurate reporting
Quantification Failed Insufficient process Repeat quantification. Passing standards
QC failure quantification yield curve metrics
run—Batch indicates issue
median below with library
minimum preparation.
Failed Standard curve failure Repeat quantification.
quantification due to bad
run quantification
Pooling Failure to Pooling Analysis is Reevaluate target
Failure complete unable to calculate pool concentration,
sample proper pool rerun pooling
pooling volumes analysis.
4 Notes
1. Blood samples (7—10 mL whole blood) must be collected in

Streck Cell-Free DNA BCT and must not be frozen. Blood
collection tubes containing samples need to be stored at 4 C
within 5 days of collection, with plasma isolation completed
within 10 days of sample collection.
2. Excessively hemolyzed blood samples should be cancelled. The
ML STAR checks that wells are empty after vacuum steps, so
blood samples that are too viscous will remain in the well;
samples determined not to have been transferred are excluded
from further analysis.
3. The Pluggo Decapper System (LGP Consulting) is useful for
uncapping large numbers of BCTs.
4. The ML STAR should be observed during all automated steps.
5. Improper cleaning and maintenance of the ML STAR can lead
to cross-contamination and poor assay performance.
6. There are several safe-stopping points: (1) After completion of
plasma isolation, the Final Plasma plate can be sealed and
stored at 2 C to 8 C for up to 7 days; (2) After completion
of cfDNA extraction, the cfDNA Elution plate can be sealed
and stored at 25 C to 15 C for up to 7 days; (3) After both
the library preparation procedure and the library quantification
procedure, the Libraries plate can be sealed and then stored at
25 C to 15 C for up to 7 days of cumulative storage;
(4) After library pooling, the pooling tubes can be capped and
stored at 25 C to 15 C for up to 7 days.
Acknowledgments
We thank the technical staff at the CLIA-certified Illumina labora-

tory (Foster City, CA) who developed these protocols for use in a
clinical laboratory.
References
1. Malone FD, Canick JA, Ball RH, Nyberg DA, 2. Bianchi DW, Parker RL, Wentworth J,
Comstock CH, Bukowski R, Berkowitz RL, Madankumar R, Saffer C, Das AF, Craig JA,
Gross SJ, Dugoff L, Craigo SD, Timor-Tritsch Chudova DI, Devers PL, Jones KW, Oliver K,
IE, Carr SR, Wolfe HM, Dukes K, Bianchi DW, Rava RP, Sehnert AJ (2014) DNA sequencing
Rudnicka AR, Hackshaw AK, Lambert- versus standard prenatal aneuploidy screening.
Messerlian G, Wald NJ, D’Alton ME, First, N Engl J Med 370(9):799–808. https://doi.
Second-Trimester Evaluation of Risk Research org/10.1056/NEJMoa1311037
C (2005) First-trimester or second-trimester 3. Illumina Inc (2017) VeriSeq NIPT Solution
screening, or both, for Down’s syndrome. N Package Insert. https://support.illumina.com/
Engl J Med 353(19):2001–2011. https://doi. content/dam/illumina-support/documents/
org/10.1056/NEJMoa043693
documentation/chemistry_documentation/ detection of fetal aneuploidies. Fetal Diagn

dx/veriseq-nipt-solution/veriseq-nipt-solution- Ther 40(3):219–223. https://doi.org/10.
package-insert-1000000001856-02.pdf. 1159/000442931
Accessed 5 Feb 2018 5. Jones KJ, Wang E, Bogard P, White K,
4. Ryan A, Hunkapiller N, Banjevic M, Schmid M, Stokowski R, Nicolaides KH
Vankayalapati N, Fong N, Jinnett KN, (2018) Targeted cell-free DNA analysis with
Demko Z, Zimmermann B, Sigurjonsson S, microarray quantitation for assessment of fetal
Gross SJ, Hill M (2016) Validation of an sex and sex chromosome aneuploidy risk. Ultra-
enhanced version of a single-nucleotide poly- sound Obstet Gynecol 51(2):275–276. https://
morphism-based noninvasive prenatal test for doi.org/10.1002/uog.18968
Chapter 23
Noninvasive Antenatal Screening for Fetal RHD in RhD

Negative Women to Guide Targeted Anti-D Prophylaxis
Frederik Banch Clausen, Klaus Rieneck, Grethe Risum Krog,
Birgitte Suhr Bundgaard, and Morten Hanefeld Dziegiel
Abstract
RhD negative pregnant women who carry an RhD positive fetus are at risk of immunization against the D
antigen, which may result in hemolytic disease of the fetus and the newborn. Predicting the fetal RhD status
by noninvasive antenatal screening for the fetal RhD gene (RHD) can guide targeted use of antenatal anti-
D prophylaxis.
Cell-free fetal DNA is extracted from maternal plasma from RhD negative pregnant women at a
gestational age of 25 weeks. A real-time PCR-based detection of two RHD exons enables reliable
prediction of the fetal RhD status to determine the administration of antenatal prophylaxis, as well as
postnatal prophylaxis.
Key words Noninvasive RHD typing, Cell-free fetal DNA, Anti-D, Antenatal prophylaxis, Hemolytic
disease of the fetus and newborn, Real-time PCR
1 Introduction
1.1 Noninvasive Immunization against the D antigen from the Rh blood group
Antenatal Screening system is the major cause of hemolytic disease of the fetus and
for Fetal RHD newborn (HDFN) and can cause fetal or neonatal death [1]. The
introduction of postnatal immune prophylaxis in the 1960s drasti-
cally reduced the immunization incidents in pregnant, RhD nega-
tive women [2]. In several countries, antenatal prophylaxis is
combined with postnatal prophylaxis to further minimize the
immunization risk [1].
Noninvasive antenatal screening for the fetal RhD gene (RHD)
enables antenatal prophylaxis to be targeted to only those women
who carry an RhD positive fetus [3, 4]. A targeted antenatal pro-
phylaxis thus avoids unnecessary treatment of pregnant women who
carry an RhD negative fetus and are at no risk of immunization
[5, 6]. In the European population, approximately 40% of the RhD
347
348 Frederik Banch Clausen et al.
negative women are of no risk of immunization and should not be

treated [5, 6].
Based on an analysis of cell-free fetal DNA from the plasma of
pregnant women, a real-time PCR-based detection of the RHD
predicts the fetal RhD status with high accuracy [3–6]. To target
the antenatal prophylaxis by noninvasive antenatal RHD screening,
it is essential that the assay has a high diagnostic sensitivity, thus
avoiding false negative results that may result in immunizations
and, consequently, HDFN. As a screening assay, the sensitivity is
>99.3% from a gestational age of approximately 10–11 weeks and
99.6–100% from around 25 weeks [4].
1.2 Strategies Due to the extensive polymorphism of the Rh blood group system
for Gene Targeting and high degree of homology between the RhD and the RhCE
gene, it is important to consider the most appropriate strategy to
target the RHD gene in order to obtain an accurate and reliable
prediction of the fetal RhD status in RhD negative pregnant
women. Specifically, the choice and combination of exon targets is
important. Genetic variants, such as silent genes, hybrid genes, or
point mutations causing weak D types may complicate the predic-
tion of the fetal RhD status [4, 7, 8]. As different RHD variants
have different frequencies in different populations, the choice of
gene targets may be based on the genetic composition of the
population to be tested. Different combinations of RHD exons
5, 7, and 10 are usually applied [4, 6]. However, as false positive
results are of minor clinical relevance, a simple setup will suffice for a
screening assay for the large majority of populations. It is essential
to obtain a high sensitivity as a false negative test result may lead to
omission of anti-D administration, thus putting the woman in risk
of immunization against D; the specificity is of secondary concern.
1.3 The Copenhagen According to the recommendations of the Danish Health and
Setup Medicines Authority, and as part of the RhD prophylaxis regime
implemented in Denmark in January 2010, all non-immunized
RhD negative pregnant women should be offered an antenatal
screening for the fetal RHD at gestational age of 25 weeks
[9]. The result of this test will guide antenatal anti-D prophylaxis
administered at a gestational age of 29 weeks [9].
The Copenhagen setup is based on automated DNA extraction
followed by a real-time PCR analysis of RHD exons 7 and 10 that
are duplexed into one assay using the same reporting dye to
increase sensitivity [10].
For the first 2 years of routine analysis, we have obtained a
sensitivity of 99.97% [6]. During the last 4 years, we have encoun-
tered only one false negative result out of approximately 10,000
tested samples. Consequently, we have terminated the serological
postnatal cord blood typing. Presently, both the antenatal and the
postnatal anti-D prophylaxis are administered based on the recom-
mendations from the antenatal RHD screening program in
Denmark.
Antenatal RHD Screening 349
2 Materials
2.1 Blood Sample 1. A standard blood sample centrifuge.

Processing 2. Sample racks for the QIAsymphony SP DNA extraction instru-
ment (Qiagen Inc., Basel, Switzerland).
2.2 DNA Extraction 1. The QIAsymphony SP (Qiagen Inc., Basel, Switzerland).

2. The QIAsymphony DSP Virus/Pathogen Midi Kit using car-
rier RNA (Qiagen Inc., Basel, Switzerland) (see Note 1).
3. Sarstedt tubes without skirts (Sarstedt, Nümbrecht, Germany).
4. Reagent box, provided by the kit.
5. Plastic utensils for the QIAsymphony SP (Qiagen Inc., Basel,
Switzerland), including pipetting tips.
6. AVE buffer, provided by the kit.
7. Sterile DNA free, RNase and DNase-free pipetting tips with
filters for manual pipetting (MμltiGuard™ Barrier Tips, Sor-
ensen BioScience, Inc., Salt Lake City, UT, USA).
8. Prepare the carrier RNA stock.
(a) Add the provided 2 mL AVE buffer into the provided
1350 μg carrier RNA tube.
(b) Vortex 3 and centrifuge briefly to avoid liquid in the lid.
(c) Make eight aliquots of 165 μL each (designated cRNA)
and store at 4 C.
2.3 Real-Time PCR 1. Instrument for automated PCR setup: QIAsymphony AS (Qia-
gen Inc., Basel, Switzerland).
2. Instrument for PCR: The real-time PCR ABI 7500 detection
system (Applied BioSystems, Foster City, USA).
3. Software for PCR interpretation: 7500 System SDS software
Version 1.4 (Applied BioSystems, Foster City, USA).
4. PCR plates: MicroAmp® Optical 96-Well Reaction Plates
(Applied BioSystems, Foster City, USA).
5. PCR plate cover: MicroAmp™ Optical Adhesive Film (Applied
BioSystems, Foster City, USA).
6. Sterile H2O (Mini Plasco®, Braun Melsungen, Melsungen,
Germany), store at 4 C.
7. Sarstedt tubes with skirts for reagents to be inserted into the
QIAsymphony AS, and Sarstedt tubes without skirts for stor-
age or other purposes not involving the QIAsymphony AS.
8. Plastic utensils for the QIAsymphony AS, including
pipetting tips.
Table 1
Primers and probes for real-time PCR detection of DNA in maternal plasma
Name Sequence Specificity

0 0
RHDex7s 5 -CAGCTCCATCATGGGCTACAA-3 RHD
RHDex7a 50 -CCGGCTCCGACGGTATC-30 RHD
RHDex7p 50 -FAM-CAGCACAATGTAGATGATCTCTCCAAGCAG-TAMRA-30 RHD
0 0
RHDex10s 5 -CCTCTCACTGTTGCCTGCATT-3 RHD/RHCE
0 0
RHDex10a 5 -AGTGCCTGCGCGAACATT-3 RHD
RHDex10p 50 -FAM-TACGTGAGAAACGCTCATGACAGCAAAGTC-TAMRA-30 RHD/RHCE
GAPDHs 50 -CCCCACACACATGCACTTACC-30 GAPDH
0 0
GAPDHa 5 -CCTAGTCCCAGGGCTTTGATT-3 GAPDH
0 0
GAPDHp 5 -FAM-AAAGAGCTAGGAAGGACAGGCAACTTGGC-TAMRA-3 GAPDH
Ex exon, s sense, a antisense, p probe
9. Sterile DNA free, RNase and DNase-free pipetting tips with

filters for manual pipetting (Sorensen BioScience, Inc., Salt
Lake City, UT, USA)).
10. Primers and probes are listed in Table 1.
(a) Reconstitute primers in sterile H2O to a concentration of
90 μM and store at 20 C.
(b) Reconstitute the probes in sterile H2O to a concentration
of 40 μM and store at 20 C (see Note 2).
(c) Transfer 310 μL of each primer to a Sarstedt tube per
primer to make up working solutions, store at 30 C.
(d) Transfer 100 μL of each probe to a Sarstedt tube per
probe and add 300 μL sterile H2O to each tube to make
up 10 μM working solutions, store at 20 C.
(e) For each assay, transfer 100 μL each from the working
solutions of sense primer, antisense primer, and the probe
into one Sarstedt tube, named SAP, for example
RHDex7SAP (see Note 3).
11. TaqMan Universal PCR Master Mix 2 (Applied BioSystems)
including uracil-N-glycosylase (UNG). Store at 4 C.
12. Control DNA (prepared in-house from whole blood). DNA
from an RHD negative woman is used as RHD negative con-
trol, and DNA from an RHD positive (RHD hemizygous)
individual is used as RHD positive control.
(a) Extract the DNA from whole blood using the QIAamp
DNA Blood Mini Kit (Qiagen) (see Note 4).
(b) Quantify eluted DNA using a spectrophotometer, and
dilute with sterile H2O to approximately 5 ng/μL, and
store at 20 C in 8 μL aliquots (see Note 5).
(c) The first time for each new donor, use the PCR assay for
RHD to ensure that the DNA from an RhD negative
donor is truly RHD negative, and that the RhD positive
donor DNA is truly RHD positive, before using the DNA
as control (see Note 6).
3 Methods
We receive a blood sample from the general practitioner (GP).

Sample registration, processing, and DNA extraction is done in
Area 1. Area 1 is a clean area, which is meticulously and constantly
kept free of PCR products in order to preclude any risk of carry-
over. Preparation of reagents and mastermix is done in a dedicated
flow bench, also in Area 1.
Gloves are used all the time and changed frequently. Real-time
PCR is done in Area 2. Areas 1 and 2 must be physically separated,
and to avoid carry-over of PCR products, no transport is allowed
from Area 2 to Area 1.
We test for glyceraldehyde 3-phosphate dehydrogenase
(GAPDH) as a marker for total DNA. We do not use a control
for fetal DNA; this is not necessary for a screening at a gestational
age of 25 weeks.
3.1 Blood Sample A blood sample taken in 6 mL EDTA blood tubes from a pregnant
Validation RhD negative woman is received from the woman’s GP. The blood
sample must meet the following criteria to be included for further
analysis:
1. The blood sample must be from a pregnant RhD negative
woman.
2. The blood sample must be from a non-immunized pregnant
RhD negative woman, thus without detectable anti-D (see
Note 7).
3. Transportation time in days from venipuncture to centrifuga-
tion must not exceed 7 days (see Note 8).
4. The time of venipuncture must be at a gestational week of
minimum 22 weeks (see Note 9).
5. The blood sample should appear undamaged.
6. The blood sample must not have been opened before arriving
at the laboratory for analysis.
7. If one of the criteria from 1 or 3–6 is not met, a new sample is
requested from the GP.
3.2 Blood Sample 1. After arriving at the laboratory, store samples at room temper-
Processing ature until further analysis the same day. If a sample is stored
overnight, place the sample at 4 C.
2. Centrifuge the 6 mL EDTA blood tube at 1700 g for 10 min
(see Note 10).
3. From here, all handling of the sample is done using gloves.
4. Inspect the sample visually for appropriate approximate plasma
volume (minimum 1.2 mL) and for hemolysis. If there is
insufficient plasma material, or if the blood sample displays
severe hemolysis, request a new blood sample from the general
practitioner.
5. The centrifuged blood tube is then placed directly into a sam-
ple rack that can be inserted into the QIAsymphony SP.
3.3 DNA Extraction 1. Prepare the QIAsymphony SP instrument for DNA extraction
(see Note 11).
2. Prepare and insert the reagent box.
3. Insert plastic utensils (see Note 12).
4. Insert the plate for DNA elution (see Note 13).
5. Prepare the carrier RNA.
(a) Add 805 μL AVE buffer to each of two Sarstedt tubes
without skirt.
(b) Add 70 μL cRNA (from 2.2.6c) to each tube.
(c) Vortex 3 and centrifuge briefly.
6. Insert the carrier RNA, both tubes.
7. Insert the sample rack, containing the blood tube, into the
QIAsymphony SP. A barcode reader in the QIAsymphony
scans the tube for sample ID when inserting the sample rack.
8. Select the integrated run, which includes the programming of
the QIAsymphony AS (see Note 11).
9. Select the “Cellfree 1000_V6_DSP default IC”-assay for DNA
extraction.
10. Run the program.
3.4 Real-Time PCR 1. For the preparation of the RHD mastermix, we use a total PCR
reaction volume of 25 μL consisting of 15 μL mastermix and
10 μL template-DNA. For the preparation of the GAPDH mas-
termix, we use a total PCR reaction volume of 25 μL consisting of
20 μL mastermix and 5 μL template-DNA (see Note 14).
2. Prepare the RHD mastermix.
(a) Calculate the required volume of mastermix, according to
number of samples tested (see Note 15).
(b) Mark a Sarstedt tube with skirt “RHD mastermix.”

(c) Add 1 μL sterile H2O per PCR well.
(d) Add 12.5 μL Universal Master Mix (2) per PCR well.
(e) Add 0.75 μL RHDex7SAP per PCR well.
(f) Add 0.75 μL RHDex10SAP per PCR well.
(g) Vortex 3–5 times and centrifuge briefly.
3. Prepare the GAPDH mastermix.
(a) Calculate the required volume of mastermix.
(b) Mark a Sarstedt tube with skirt “GAPDH mastermix.”
(c) Add 6.75 μL sterile H2O per PCR well.
(d) Add 12.5 μL Universal Master Mix (2) per PCR well.
(e) Add 0.75 μL GAPDH SAP per PCR well.
(f) Vortex 3-5 times and centrifuge briefly.
4. Prepare the control DNA.
(a) Thaw the RHD negative control DNA aliquot (5 ng/μL),
vortex, and centrifuge briefly.
(b) Add 54 μL sterile H2O each into three Sarstedt tubes with
skirts.
(c) Add 6 μL of the aliquot into the first tube, vortex, and
centrifuge briefly.
(d) Add 6 μL from the first tube into the second as well as into
the third to obtain two tubes with 50 pg/μL each, vortex,
and centrifuge briefly. These are the final RHD negative
controls (see Note 16).
(e) Thaw the RHD positive control DNA aliquot (5 ng/μL),
vortex, and centrifuge briefly.
(f) Add 54 μL sterile H2O each to four Sarstedt tubes with
skirts.
(g) Add 6 μL of the aliquot into the first tube, vortex, and
centrifuge briefly.
(h) Add 6 μL from the first tube into the second tube, vortex,
and centrifuge briefly.
(i) Add 6 μL from the second tube into the third and fourth
tubes to obtain two tubes with 5 pg/μL each, vortex, and
centrifuge briefly. These are the final RHD positive con-
trols (see Note 17).
5. Load the PCR reagents into the QIAsymphony AS, following a
software protocol designed specifically as described above with
triplicate analysis of RHD and a single analysis of GAPDH for
each sample, non-template control samples, and RHD negative
and RHD positive control samples.
6. Start the automated transfer of the elution plate from the

QIAsymphony SP to the QIAsymphony AS.
7. Start the automated PCR pipetting setup.
8. When the PCR setup is completed, remove the plate and seal it
with a cover (MicroAmp™ Optical Adhesive Film).
9. Insert the plate into the 7500 ABi PCR instrument; apply a
template setup or follow the wizard instructions for setup (see
Note 18).
10. Use the following thermal PCR profile: 2 min at 50 C and
10 min at 95 C, followed by 45 cycles of 95 C for 15 s and
60 C for 1 min.
11. To avoid carry-over of PCR products, it is essential to ensure a
proper discarding of the PCR plates as well as to avoid return-
ing to Area 1.
3.5 Interpretation 1. For software analysis of the PCR results, use automated back-
of Results ground and a fixed threshold of 2.0.
2. Consider a PCR reaction positive, if the cycle threshold
(Ct) value is <42 (see Note 19).
3. General rules for the controls:
(a) The non-template control can be positive in a maximum
of two out of three (2/3) RHD PCR reactions (with a
Ct-value <40).
(b) The RHD positive control must be positive in minimum
2/3 PCR reactions (Ct 38).
(c) The RHD negative control can be positive in a maximum
1/3 PCR reactions (Ct < 40).
(d) The non-template control must be negative for GAPDH
(Ct > 38).
(e) The RHD positive and RHD negative controls must be
positive with GAPDH.
4. Interpretation rules for samples:
(a) Consider a sample RHD positive, if all triplicate PCR
reactions are positive (see Note 20).
(b) Consider a sample RHD negative, if one or none PCR
reactions are positive (Ct > 40), and if the GAPDH
Ct-value is <35.
(c) Consider a sample Inconclusive, if two PCR reactions are
positive.
(d) Consider a sample Inconclusive, if the RHD Ct-values
are less than or equal to the Ct-values of GAPDH,
which may indicate a possible RHD positive woman (see
Note 21).
5. We recommend that all women with an RHD positive or

Inconclusive sample result are offered antenatal anti-D
prophylaxis.
6. If the criteria from 3a-e are not met, all the samples from the
PCR run are retested using backup plasma, or a new blood
sample is requested. If the GAPDH Ct-value is 35 for a
seemingly RHD negative result, a new blood sample is
requested, as a high Ct-value may indicate a poor DNA extrac-
tion with the risk of a false negative result. Overall, a result is
only concluded and reported, when all the listed criteria are
met (see Note 22).
4 Notes
1. We have not tested whether including the carrier RNA makes

any difference as regards to the cffDNA yield, and it is possible
that the maternal DNA fulfills the role of the carrier RNA.
However, we have chosen to follow the manufacturer’s
recommendations.
2. We use our primers and probes quickly. If testing few samples, a
higher concentration may be considered for the primer and
probe stock solutions. We use HPLC purified primers. Primers
can be purchased from, e.g., Eurofins MWG Operon, Edels-
berg, Germany or TAG Copenhagen, Copenhagen, Denmark.
3. The SAP is named for Sense primer, Antisense primer, and
Probe. We use this SAP solution containing all oligos of an
assay simply to have fewer pipetting steps in the downstream
analysis, thereby saving time and decreasing the risk of pipet-
ting mistakes. In our experience, the SAP solution works well
for about a year, and perhaps longer. However, as a conse-
quence of the routine testing, we make a fresh SAP approxi-
mately every second week.
4. Other DNA extraction systems can be used.
5. We quantify the elution, then dilute to 10 ng/μL and quantify
again, and then dilute to 5 ng/μL and re-quantify and note the
estimated concentration. This procedure with two quantifica-
tion steps ensures a more accurate final DNA concentration
compared to a procedure with only a single quantification step.
In principle, the RHD positive control DNA can be used as a
reference DNA for a relative quantification of the cell-free fetal
DNA, using the delta Ct-method. However, we find that using
the control DNA for the delta Ct-method results in an inaccu-
rate quantification, and we recommend using a universal stan-
dard curve, which does not need to be included on each PCR
plate [11].
6. We use DNA from an RHD hemizygous individual to simulate

the presumable hemizygous DNA from an RHD positive fetus.
If necessary, the zygosity of the RHD positive control DNA
can be assessed or confirmed by a zygosity test [12].
7. If the pregnant woman has a known anti-D, she is treated as an
immunized woman. The sample is analyzed but with a different
algorithm, where the minimum acceptable gestational week is
10 weeks.
8. Cell-free fetal DNA can remain stable even during prolonged
transportation in EDTA-tubes. We have shown that the detec-
tion of cell-free fetal DNA, using the duplex RHD assay
described in this method, is unchanged despite a transportation
time of up to 9 days and despite an increase in the level of
maternal DNA [11]. This finding is corroborated by others
[13]. Consequently, we allow a sample to be 7 days old. How-
ever, we still require from the general practitioners that the
anticipated transportation time must not exceed a maximum
of 4 days, so that the transportation time does not exceed
7 days when samples are in transit over weekends or public
holidays.
9. On a few occasions, samples from early pregnancy have been
sent by mistake, such as gestational weeks 8, and this has been
shown to result in false negative results [6]. We chose not to
analyze samples before 22 gestational weeks, mainly because
our validation was done with samples from 20 gestational
weeks and onward, but also to help the general practitioners
to comply with the request that the samples should be taken in
gestational weeks 25. The overall system, however, is robust
enough to analyze samples from earlier than gestational weeks
22.
10. We do not use a second centrifugation step to eliminate excess
maternal DNA background. We have shown that this is not
necessary for the detection of fetal RHD at gestational weeks
25 [11]. Avoiding a second centrifugation step also avoids
further handling that can result in sample mixup or increased
risk of contamination.
11. Preparing and running the QIAsymphony involves several
steps in the QIAsymphony software. These have been left out
here, as these steps will be incorporated into an analysis
through a training program provided by Qiagen personnel.
When the program is selected, one can select an integrated
program that allows for a simultaneous setup of the QIA-
symphony SP and AS. In this way, the program is initiated for
both the DNA extraction as well as for the downstream PCR
pipetting. However, the system allows you to prepare the
QIAsymphony AS with utensils and reagents after starting the
program.
12. It is important that the plastic utensils are inserted correctly;

otherwise an instrument failure may occur.
13. It is also possible to use tubes for DNA elution.
14. When we extract DNA from 1 mL plasma, elute into 60 μL,
and apply 10 μL for PCR testing, we are really testing an
equivalent of 1/6 of 1 mL plasma (1000 μL 10 μL/
60 μL ¼ 167 μL). This is also called the plasma-equivalent
per PCR. A high plasma-equivalent per PCR is essential for a
high sensitivity. Theoretically, if the plasma-equivalent is dou-
bled, then the sensitivity is equally doubled; however, using a
lower elution volume or a higher template volume may also
increase the amount of PCR inhibitors in the PCR reaction. We
use a PCR setup where we test three reactions for RHD. Using
multiple reactions will increase the sensitivity, if one reaction is
allowed to be negative.
15. The required mastermix volumes depend on the number of
samples tested plus the controls, and some additional volume.
We test the RHD positive and negative controls, as well as the
non-template control as samples, thus accounting for three
extra samples. In addition, a substantial dead volume is
required. The QIAsymphony software will inform you of the
number of reactions and estimate the required mastermix
volume.
16. The reason for making two control tubes of each control is that
the PCR setup program requires a set of reagents for each assay,
RHD as well as GAPDH.
17. The difference of concentration in the two controls relates to
their function. The RHD negative control serves as control to
show that the RHD assay does not produce an unspecific PCR
signal, which is shown at a higher concentration than the
concentration to be expected from the maternal plasma. The
RHD positive control with approximately 50 pg per PCR
serves as control for a reliable detection of low-level DNA,
yet kept at a concentration that results in positive amplifications
in all triplicate reactions each time.
18. A txt.-file describing the sample IDs and the PCR setup can be
exported from the QIAsymphony and imported to the PCR
software, thus ensuring automated setup in the PCR instru-
ment and sample ID tracking; the txt.-file carries the barcode of
the PCR plate inserted into the QIAsymphony. The barcode of
the plate can then be read upon inserting the plate into the
PCR instrument, thus ensuring that the PCR setup and the
sample IDs are connected to the correct plate.
19. If an amplification curve deviates from an expected curve
shape, consult the raw data (e.g., “compliment” in the
software) to make a visual evaluation; use the data from the

positive and negative controls to guide you.
20. An estimated 95%-limit of detection (LoD) for the RHD
positive conclusion based on 3/3 positive PCR reactions is
9 copies per mL plasma. For the inconclusive results with
2/3 positive PCR reactions, the LoD is 6 copies per mL
plasma.
21. Women that we suspect to be RHD positive are recommended
anti-D prophylaxis. In addition, although without any clinical
implications, we test the maternal blood (left from the DNA
extraction) serologically for the RhD status using anti-D IgG
LOR17 (reacting with most D variants including DVI) and
GAN4B.5 (reacting with most D variants but not with DVI) as
well as for RhC and RhE [6]. The results are kept to distinguish
the outcomes: maternal weak D, DVI, and silent, nonfunc-
tional RHD genes. During the first two years of routine test-
ing, we also included further individual DNA testing of the
maternal DNA for RHD exon 5, 7, and 10.
22. Additional comments that might be useful: In our laboratory
we run approximately 80 samples per week, with a maximum of
20 samples per DNA extraction and PCR test. The full proce-
dure can be done within a day and by one person. The DNA
extraction step takes about 2 h and the PCR is about 1.5 h;
registration, processing, and reporting take about 2 h. The
current price, which covers reagents and personnel, is about
100 EUR per sample.
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GR, Nielsen LK, Tabor A, Dziegiel MH pd.2889
INDEX
A Comparative genomic hybridization (CGH) .........12–14,

73–84, 86, 252
Amniocentesis .......................4, 6–9, 26, 45, 46, 50, 188, Congenital bilateral absence or atrophy of the vas deferens
225, 237, 287, 288, 298, 306 (CBAVD) .................................................. 221, 224
Amniotic fluid ...................................... 3, 4, 6, 10–12, 45,
Copy-number variation (CNV) .................. 37, 101, 187,
106–109, 112, 113, 115, 117–119, 129, 132, 188, 251, 252
133, 136, 139, 147, 152, 153, 164, 165, 188, Cystic fibrosis (CF) ................6, 11, 24, 29, 61, 221–230
203, 236, 237, 242–244, 288, 290, 292, 293,
297, 298, 303, 306 D
Aneuploidy .............................. v, 5, 6, 11, 24, 26, 35–38,
46–49, 51–53, 61, 63, 73–85, 119, 121, 129, DNA extraction........................... 63–66, 76, 77, 92, 105,
139, 164, 168, 310, 325–344 123, 124, 153, 156, 165, 172–177, 203, 215,
Antenatal prophylaxis........................................... 347, 348 225, 230, 239, 249, 329, 331, 333–335,
Anti-D......................................................... v, 50, 347–358 348–350, 352, 355, 356, 358
Array CGH ...............................28, 36–38, 140, 153, 156 DNA extraction kit ..................................... 107, 172, 270
Automation ................................................................... 309 DNA prep .................................. 149, 151–153, 155, 156,
227–230, 245–247
B Down syndrome...................v, 4–6, 45–47, 53, 297–300,
305, 325
Biopsy ...................... 7, 8, 24, 26–29, 31, 36, 37, 62, 63,
69, 74, 76, 83, 86, 91 F
Blastomere ................................24, 26, 27, 29, 36, 37, 62
Blood sample stabilization............................................ 309 Fetal anomalies ................................................... 4, 45, 268
Fetal cells ............6, 45, 48, 49, 106, 129, 147, 149, 237
C Fetal defect markers ...................................................... 297
Fluorescence in situ hybridization (FISH) ............ 11, 12,
Cell culture ..................................... 4, 107, 111, 129, 242 24, 28, 73, 86, 129, 140, 252, 268
Cell free amniotic fluid ............................... 108, 112, 242
Chorionic villi.............................. 7, 8, 12, 106–108, 110, G
117–119, 131, 149, 152, 188, 211, 215, 236
Chorionic villus sampling (CVS) ..................... 4, 7–9, 11, Gene dosage .................................................................. 161
45, 46, 50, 106, 107, 110, 139, 148, 149, 156, Genotyping.............. 61, 64, 66–70, 117, 125, 207, 208,
172, 188, 225, 237, 242, 243 210, 214, 215, 223, 224, 228–230, 237,
Chromosomal abnormalities ............................ 24–25, 28, 246–248, 283
53, 164, 251, 253, 288
H
Chromosomal microarray analysis (CMA) ............... v, 13,
130, 187, 188, 268 Hemolytic disease of the fetus and newborn
Chromosomal rearrangement .......................... 26, 34–35, (HDFN)................................................ v, 347, 348
74, 81, 164, 251–263 HEXA gene ................................................. 234, 235, 245
Chromosomal SNP microarray ..........187, 194, 197–200 HEXA sequencing......................................................... 245
Chromosome aneuploidies ..................... 53, 81, 139, 340 Hexosaminidase A (Hex A) deficiency .......234–236, 243
Chronic pulmonary disease .......................................... 221 History of prenatal diagnosis ...................................45–54
Circulating cell-free DNA (ccfDNA)...........................v, 6,
49–52, 106, 107, 113, 309–322, 325–331, I
333–336, 342–344, 348, 355, 356
In vitro fertilization (IVF) ..........v, 24, 29, 73, 83, 85, 91
Circulating DNA extraction ....................... 311, 313, 314
Inorganic liquid phase extraction................................. 107
Interphase FISH................................................... 141, 149
Brynn Levy (ed.), Prenatal Diagnosis, Methods in Molecular Biology, vol. 1885, https://doi.org/10.1007/978-1-4939-8889-1,
361
PRENATAL DIAGNOSIS
362 Index
J Polymerase chain reaction (PCR) ....................10, 11, 24,
28, 51, 62, 64, 67–70, 73, 75–78, 87, 88, 90–93,
Jumping libraries ........................................................... 252 95, 97, 99, 121–124, 126, 140, 153–157, 162,
163, 165–167, 170, 188–190, 192–196, 198,
M
204, 207, 215, 216, 224, 227–230, 239,
Maternal cell contamination (MCC) ..................... 14, 49, 245–248, 252–259, 269, 270, 274–276, 316,
107, 115, 117, 136, 141, 147, 176, 225, 237, 328, 348, 350, 352–355, 357, 358
238, 241, 244, 248, 249, 270 Precipitation ....................... 66, 106–108, 110, 111, 151,
Maternal screening....................................................46, 48 174, 176, 192, 247
4-Methylbelliferil-N-acetyl-β-glucosamine Preimplantation Genetic Diagnosis (PGD)............23–38,
(4-MUG) ........................ 234–236, 238, 242, 243 61–70, 74, 210, 224
4-Methylumbelliferyl-6-sulfo-2-acetamido-2-deoxy-β-D- Preimplantation genetic screening (PGS)..................... 26,
glucopyranoside (4-MUGS) ................... 234–236, 63, 73, 74, 78, 82, 83
238, 239, 242, 244 Preimplantation genetic testing (PGT) ................... vi, 25,
Microarray ................................v, 12–14, 62, 73–84, 107, 28–31, 34
115, 118, 172, 173, 180, 181, 183, 187, 188, Preimplantation genetic testing for aneuploidy
190, 269, 326 (PGT-A) .........................................................35, 38
Microsatellite markers .........................140, 158, 237, 241 Prenatal diagnosis (PND)................................. v, vi, 3–15,
Molecular cytogenetics ................................................... 12 45–54, 105, 107, 117, 129, 156, 161, 187, 194,
Mosaicism ......................... 7, 9, 12, 25, 37, 38, 130, 148, 197–200, 207, 208, 210, 211, 221–230,
164, 168, 222 233–249, 251, 267–283
Multiplex ligation-dependent probe amplification Prenatal screening .........4, 45, 46, 52, 53, 297–306, 325
(MLPA)..........................................vi, 11, 130, 161
Multiplex polymerase chain reaction ................. 151, 153, Q
161, 163, 228, 246 QIAamp ........................63–65, 108, 110, 111, 113, 165,
Mutation........................... 10, 13, 29, 30, 32, 51, 62, 66, 311, 313, 314, 317–319, 350
67, 69, 121, 125, 130, 150, 163, 207–215, QIAsymphony ........................... 230, 249, 311, 313–315,
222–225, 228–230, 234, 235, 237, 245–248, 318, 320, 322, 349, 352–354, 356, 357
251, 267, 348 Quad screen.......................................................... 297–306
Quantitative fluorescence-polymerase chain reaction
N
(QF-PCR) ............................vi, 11, 117, 130, 140,
Next generation sequencing (NGS) .................... v, vi, 13, 141, 148–150, 153–155, 158
28, 37, 51, 86, 224, 251–263, 268, 270, 315, Quantitative polymerase chain reaction technology
325–344 (qPCR).....................................50, 62, 73, 86, 259
Noninvasive prenatal diagnosis (NIPD) ......... 45–52, 310
Non-invasive prenatal testing (NIPT) ....................v, vi, 6, R
46, 49, 51–53, 310, 311, 326–331, 333, 334, Rapid prenatal test ........................................................ 131
336, 338, 340, 342 Real-time PCR .............................. 69, 70, 208, 210, 211,
Non-invasive RHD typing................................... 347–358 214–217, 313, 320, 348–352
Nuchal translucency (NT) ............................... v, 5, 47, 48 Recessive ....................................6, 24, 27, 30, 33, 51, 67,
221, 234, 283
P
Paired-end sequencing......................................... 326, 327 S
PCR-free ........................................................................ 326 Sample identity................................ 65, 66, 68, 158, 317,
Percentage Hex A activity (Hex A%) ..........................236, 318, 352, 357
243, 244 Serum screening ................................................... 6, 46, 47
PGT-aneuploidy (PGT-A) ......................... 27, 28, 30, 33, Short tandem repeats (STR) ................11, 117, 122, 237
35–38, 86, 91, 100, 101 Single gene disorder (SGD) .......................24–25, 29–34,
PGT-human leukocyte antigen (PGT-HLA) ..........33–34 51, 61–70, 172
PGT-monogenic (single gene) (PGT-M) ................29–34 SNP-based aneuploidy detection ................................. 309
PGT-structural (chromosomal) rearrangement Specific Hex A activity ................................ 234, 236, 244
(PGT-SR).......................................................34–35 Structural variation (SV).....................251, 252, 261, 263
Pitfalls ............................................................................ 130
PRENATAL DIAGNOSIS
Index 363
T U
Targeted mutation ............................. 223–225, 228–230, Ultrasound............................... 4–7, 9, 12, 13, 45–48, 50,
237, 246–248 53, 164, 188, 225, 267, 272, 274–276, 278, 281,
Tay-Sachs disease (TSD).................................. 8, 233–249 287
β-Thalassemia and sickle cell syndromes.....................207, Ultrasound abnormalities ...................164, 267–283, 287
209, 210, 212, 213, 216
Trisomy .......................................... 5, 6, 8, 11, 38, 46, 47, W
51–53, 121, 129–131, 136, 141, 142, 148–150,
Whole exome sequencing (WES)..................13, 267–283
154, 157, 163, 168, 169, 297–299, 305, 325, 340 Whole genome amplification (WGA) ...... 75, 86, 91, 172
Trophectoderm .......................................... 26–28, 37, 62, Whole genome sequencing (WGS)...........v, 13, 252, 326
73, 74, 76, 83, 91

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Methods in

Molecular Biology 1885

Brynn Levy Editor

For further volumes:

ISSN 1064-3745 ISSN 1940-6029 (electronic)

© Springer Science+Business Media, LLC, part of Springer Nature 2019

New York, NY, USA Brynn Levy

PART I HISTORICAL INTRODUCTION

PART II PREIMPLANTATION GENETIC TESTING

4 Molecular Testing for Preimplantation Genetic Diagnosis

PART III TRADITIONAL PRENATAL DIAGNOSIS

11 Multiplex Ligation-Dependent Probe Amplification (MLPA)

PART IV NON-INVASIVE PRENATAL TESTING

20 Quad Screen Test, A Multiplexed Biomarker Assay for Prenatal Screening

Traditional Prenatal Diagnosis: Past to Present

including Down syndrome, Turner syndrome, Klinefelter syn-

2.2 Biochemical In 1977, Wald et al. published a large multi-center collaborative

additional screening in the second trimester [45]. With the avail-

3.1 Amniocentesis Amniocentesis is the technique for withdrawing amniotic fluid

standard of care to recommend an amniocentesis during the second

3.2 Fetoscopy Fetoscopy is an endoscopic procedure that provides access (for

later, in 1973, Sandahl and Kullander biopsied the placenta in

3.4 Percutaneous Percutaneous Umbilical Blood Sampling (PUBS) involves obtain-

4 Techniques for Assessing Fetal Genetic Information

4.1 Karyotype The ability to diagnose chromosome abnormalities from prenatal

abnormalities in pregnancy. As such, advanced maternal age, gen-

together, dramatically increased the speed and sensitivity of the

4.3 Fluorescence In The combination of traditional cytogenetic and molecular techni-

4.4 Comparative The development of chromosomal comparative genomic hybridi-

could scan the entire genome without having to preselect specific

number of probes used on a given array. Today, most clinical

Amidst the progression in both procedural sampling and analysis

need for additional geneticists and genetic counselors to interpret

acid sequences by multiplex ligation- in situ hybridization. Am J Hum Genet 49

hybridisation in fetuses with multiple malfor- Genet 19(12):1230–1237. https://doi.org/

Overview of Preimplantation Genetic Diagnosis (PGD):

Preimplantation genetic diagnosis (PGD)1 was first accomplished

2 Pioneers in Detection of Single Gene Disorders and Chromosomal Abnormalities

PGD can be said to have begun in 1968 when Gardner and

observed mosaicism in cleavage stage embryo; Munné also recog-

3 Professional and Scientific Organizations

Scientific presentations and clinical application of preimplantation

ASRM members. Presidents (currently Chairmen) serve 2 year

PGD requires access to DNA from gametes or embryos within

primary oocyte should have the normal allele. Oocytes deduced to

of 22.4% and 37.4% after removal of one versus two cells,

4.3 Blastocyst Biopsy of trophectoderm in the 5- to 6-day approximately 120-cell

higher pregnancy rates than “same-cycle” transfer. The presump-

5 Single Gene Disorders (PGT-M)

5.1 Unique As already discussed, only PGT allows information to be obtained

embryo biopsy, if necessary. So despite the complexity of PGT-M

relative were affected. PGT-M to exclude transmission of any adult

5.1.4 HLA-Compatible Among sibs, one in four is HLA compatible (identical).

By the present time, the list of disorders for which PGT-HLA

6 Chromosomal Rearrangements (PGT-SR)

Chromosomal rearrangements (translocation or inversion) are well

that this was achieved with an average of 1.24 cycles is unexpected

7 PGT-Aneuploidy (PGT-A) to Improve Pregnancy Rates

PGT aneuploidy testing is increasingly performed solely to improve

l Hold: 50 C for 2 min, 95 C for 10 min.