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Methods. 2007 February ; 41(2): 143150.

Analysis of Cell Cycle Phases and Progression in Cultured

Mammalian Cells

Christoph Schorl and John M. Sedivy*

Department of Molecular Biology, Cell Biology and Biochemistry, and Center for Genomics and
Proteomics, Brown University, 70 Ship Street, Providence, RI 02903, USA.

Abstract
Fluorescence Activated Cell Sorting (FACS) analysis has become a standard tool to analyze cell
cycle distributions in populations of cells. These methods require relatively large numbers of cells,
and do not provide optimal resolution of the transitions between cell cycle phases. In this report we
describe in detail complementary methods that utilize the incorporation of nucleotide analogs
combined with microscopic examination. While often more time consuming, these protocols
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typically require far fewer cells, and allow accurate kinetic assessment of cell cycle progression. We
also describe the use of a technique for the synchronization of adherent cells in mitosis by simple
mechanical agitation (mitotic shake-off) that eliminates physiological perturbation associated with
drug treatments.

1. Introduction
Among the crucial events required for the development of malignancies are the loss of
responsiveness to negative regulators of cell cycle progression and/or the acquirement of
independence from mitogenic signals (Hanahan and Weinberg, 2000). Not surprisingly,
expression profiles of genes involved in governing cell cycle progression can be used as
molecular markers to predict responsiveness to therapeutic intervention and patient survival
in various human neoplasias (reviewed in Singhal et al., 2005, Yasui et al., 2005, Quinn et al.,
2005). Over the last 30 years, beginning with the revolutionary discoveries of the genes
involved in cell cycle control by Hunt, Nurse and Hartwell (reviewed in Nurse, 2000), intensive
research efforts have led to significant progress in identifying the molecular machinery
involved in cell cycle progression. Today this information is widely used for the development
of highly specific therapeutic interventions in cancer treatment.
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While FACS is a useful technique that has become a standard tool to analyze the DNA content
of cells, it provides only a snapshot of the cell cycle distribution at any given point in time.
FACS also requires relatively large numbers of cells to achieve adequate statistical
significance, suffers from a variety of sample preparation artifacts, and does not distinguish
accurately between closely spaced events, for example, late G1 phase from early S phase. In
contrast, incorporation of nucleotide analogs, such as Bromo deoxyuridine (BrdU), even for
periods as short as a few minutes, can very clearly and reproducibly mark cells in S phase when
combined with sensitive immunological detection methods and microscopic observation. In
conjunction with physiological methods for cell synchronization that avoid the use of drugs,

*Corresponding author: John M. Sedivy, phone: 401-863-7631, fax: 401-863-9653, e-mail: john_sedivy@brown.edu.
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 Schorl and Sedivy Page 2

these approaches allow the accurate determination of dynamic cell cycle phase progression in
living cells.
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The methods presented in this article have been developed using the Rat-1 cell line (Prouty et
al., 1993). This is an established, immortalized fibroblast cell line derived from a mid-gestation
rat embryo. It shows good contact inhibition and does not display any significant transformed
phenotypes such as anchorage independent growth or tumor formation in immuno-
compromised mice. In most aspects it is very similar to the several murine fibroblast cell lines
established by the 3T3 protocol: NIH-3T3, Balb/c-3T3, Swiss-3T3, etc. The techniques
described here can also be readily adapted to primary fibroblast cultures, such as mouse embryo
fibroblasts (MEF) or normal human diploid fibroblasts (HDF) from a variety of sources. Other
cell types may require significantly different culture conditions, and transformed cells typically
cannot be adequately synchronized in the G0 cell cycle phase by serum deprivation and/or
contact inhibition; however, the methods for labeling and sample processing in exponential
phase should be readily adaptable.

2. BrdU labeling
BrdU and uridine (Sigma, St. Louis, MO, cat. no. B5005, U3003) are made up as 1000 x and
100 x stock solutions, respectively, in distilled water (dH20), filter sterilized, and stored
protected from light at 4 C. To avoid unequal distribution and locally high concentrations,
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both solutions should be pre-added to the medium, rather than added to plated cells. Uridine
is added to prevent incorporation of BrdU into RNA. The final concentration in medium for
BrdU is 1 g/ml and for uridine is 1 mg/ml. While BrdU can have cytotoxic effects, at the
concentrations used in our studies we did not detect any deleterious effects. It is however
important to stress that once BrdU has been added all subsequent handling of the cells should
be done under safe light conditions (orange or red illumination), as even brief exposure to
standard overhead fluorescent lights can elicit toxicity. Incorporation of BrdU is terminated
by addition of L-ascorbic acid (Sigma, cat. no. A4544) directly to the culture medium to a final
concentration of 0.067 M (Moscovitis et al., 1980). This has the effect of rapidly killing the
cells without perturbing their morphology or causing detachment. The particular advantage of
this method is that L-ascorbic acid can be rapidly pipetted into a single well of a multi-well
plate, which can then be returned to the incubator for continued culture of cells in adjacent
wells. This is very useful in time course experiments that may take 24 h or longer to complete.
Addition of ascorbic acid will change the color of the medium to bright yellow. The stock
solution of L-ascorbic acid is made as 0.4 M in dH2O, filter sterilized, and stored in the dark
at 4 C. It can be used until the color of the stock solution changes from pale opaque to yellow.
After termination of the experiment cells can be kept under the medium/ascorbic acid mixture
for up to 48 h at 4C without detrimental effects on subsequent staining.
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3. Experiments to analyze cell cycle progression

3.1. Culture conditions
Both the quality of the serum used to supplement the medium as well as the culture conditions
can dramatically affect the proliferation rates of cells. It is highly recommended to carefully
test different batches of serum for their effects on the parameters under study. We test our Rat-1
fibroblasts using two criteria: 1) maximum rate of proliferation under exponential growth
conditions; 2) minimum apoptosis during a 48 h serum-deprivation period (0.25% serum) under
100% confluent conditions. Other assays may be applicable in other systems. We have
observed variations as large as 2530% in exponential growth rates between individual serum
batches. Once the desired batch of serum has been identified sufficient amounts should be
purchased for all planned experiments.

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In terms of culture conditions that may affect cell cycle progression, frequent replenishment
of the medium and keeping the cells in a well dispersed and subconfluent state are the most
important. In our hands, for cells with population doubling times of < 24 h, replacing the
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medium every 2 days and not allowing the cells to exceed 50% confluency results in
homogenously growing cultures and highly reproducible doubling times. For slower growing
cells less frequent media changes may be used, but confluency should not be allowed to exceed
50%. We define confluency as the % of the total culture vessel surface that is occupied by cells.
This can be estimated from phase contrast micrographs taken at low magnification. Although
maximum desirable cell density may vary between cell types, the main objective is to avoid
cell-cell contact, which can lead to contact inhibition and hence a departure from exponential
growth kinetics. Transformed cell lines typically do not show contact inhibition, so this
criterion may perhaps be relaxed; however, such cells tend to metabolize rapidly so that close
attention should be paid to medium changes.

We have also found that subculture of cells at ratios between 1 : 4 and 1 : 6 is optimal, because
it tends to minimize growth in patches where only the outer cells are free from significant
contact inhibition. While this may be more labor intensive because it necessitates a frequent
subculture regimen, it also promotes maximum exponential phase cycling of the cultures.
Finally, to ensure asynchronous and homogeneous cycling, any experiment should start with
a minimum of two passages under the above conditions before any measurements are made.
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3.2. Synchronization of cultures

For kinetic analyses of cell cycle progression it is highly desirable to synchronize the cells.
One frequently used method is to arrest the cells in the G0 phase, which can be achieved by
growing the culture to confluency followed by serum deprivation (0.1% to 0.25% serum) for
48 h. At the start of the serum deprivation period it is important to thoroughly rinse the culture
dishes several times with PBS to remove residual serum. The degree of growth arrest is best
determined by FACS of ethanol fixed and propidium iodide stained cells (0.05 g/ml final
concentration; Shichiri et al., 1993). Good growth arrest should result in 95% or greater cells
with a G1/G0 DNA content.

Some cell types, for example MEF or Balb/c-3T3 can be adequately synchronized by contact
inhibition alone. In these cases the cultures are simply allowed to reach 100% confluency and
then incubated without medium change for a further 4872 h. Release into the cell cycle is best
achieved by subculture; while re-feeding with medium containing fresh serum without
subculture should induce some cell cycle entry, this may not recruit all cells and also may not
produce adequate synchrony of progression.

Cells that have been synchronized by the combination of contact inhibition and serum
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deprivation can typically be recruited into the cell cycle very efficiently (>95% entry in Rat-1
cells) by simple re-feeding with medium containing fresh serum. While both methods are
believed to induce minimal physiological perturbations, contact inhibition alone would be
considered to be the gentler of the two. Transformed cell lines typically cannot be synchronized
by these methods. Serum deprivation of subconfluent cultures is usually avoided because it
commonly results in apoptosis.

The second commonly used synchronization point is mitosis (M). A method that produces
good yields of cells is incubation with the microtubule inhibitor nocodazole (Sigma, cat. no.
M1404) at 40 -200 ng/ml final concentration (depending on cell type), diluted from a stock
solution of 2 mg/ml in DMSO. Ideally, incubation time should be for a period of at least one
cell cycle (to allow all cells to accumulate in M), but this may produce toxic effects, and is
almost certain to perturb subsequent cell cycle progression as the cells recover from the drug
treatment. Shorter periods of incubation will result in lower yields of mitotic cells, which then

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need to be separated from the non-synchronized cells. Fortunately, since cells in mitosis
become rounded and lose most of their attachment to the substratum, they can be dislodged
using relatively gentle mechanical agitation.
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The optimal procedure to minimize physiological perturbations, commonly called "mitotic

shake-off", is to eliminate the drug treatment altogether. This technique is rapid, very gentle,
and produces highly synchronized cultures; however, the yields can be very low. Since mitosis
lasts approximately 30 min, in a culture with a 24 h doubling time only ~5% of the cells will
be in M phase, and not all can be harvested by the shake-off. Thus, a considerable number of
dishes as well as some teamwork may be needed to reduce the processing time as much as
possible, since otherwise the synchronization of the culture is compromised. 30 to 40 10-cm
plates of exponentially growing cells will yield enough mitotic cells for 12 wells of a 24 well
microtiter plate. To harvest the mitotic cells it is easiest to gently tap 3 stacked plates at a time
against a hard surface for about one minute while turning the dishes in 90 steps. In order to
avoid contamination it is important to avoid spillage of the medium onto the lid, and hence this
technique requires some practice. The medium from all plates is pooled and cells are harvested
by centrifugation. At least two people should do the tapping of the plates while a third person
handles the pooling and centrifugation. The dishes can be recycled by adding back fresh
medium, returning to the incubator, and harvesting additional mitotic cells at a later time (e.g.,
the next day).
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3.3 Pulse labeling

A straightforward experiment to determine the fraction of cells in S phase is to pulse label an
asynchronous culture with BrdU for 15 to 60 min and immediately harvest for analysis. Using
this highly sensitive method, it is possible to capture cells at the very beginning and end of S
phase, and thus to obtain more accurate representation of S phase content than is possible by
FACS.

The pulse labeling experiment can be augmented by using synchronized cells. For example,
after mitotic shake-off cells are seeded in 24 well microtiter plates, BrdU/uridine containing
medium is added to the wells as successive time points (every 24 h), and after 30 min
incubation with BrdU each well is quenched with ascorbic acid (Fig. 1A). Using this protocol
one should see a relatively rapid rise in BrdU-positive cells as the culture enters S phase. As
cells exit S phase the labeling should drop off, resulting in a clearly defined peak of BrdU
incorporation. If the synchrony of the culture is maintained, a second increase in BrdU labeling
will occur when the cells enter the subsequent S phase. In this type of experiment cultures with
shorter cell cycles typically maintain better synchrony than those with longer cell cycles (Fig.
1A; compare c-myc +/+ cells with c-myc / cells). Under ideal conditions it should be possible
to determine the length of one entire cell cycle (midpoint of the first rise to the midpoint of the
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second rise), the length of G1 phase (beginning of the experiment to the midpoint of the first
rise), and the length of S phase (mid point of the first rise to the midpoint of the first decline).
G2 can then be estimated indirectly by subtracting the length of G1 and S from the entire cell
cycle (under most conditions M phase can be estimated to be 3060 min).

There are two reasons why the S phase peak may not reach 100%. First, not all the cells in the
culture may be cycling; the fraction of active cells can best be estimated by continuous labeling
(below). Second, if the culture synchrony is not good, the fastest cells may exit S phase before
the slowest cells enter it. Thus, even if all cells are cycling, there may not be any one time when
all the cells are in S phase.

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3.4 Continuous labeling

In a continuous labeling protocol BrdU is added at the beginning to all the wells, and individual
wells are quenched with ascorbic acid at successive time points. This provides a very accurate
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determination of S phase entry either from M phase or G0 phase that is not compromised by
S phase exit. In a well synchronized, active culture one should see a rapid, continuous rise to
100% labeling (Fig. 1B). The half point (50% BrdU positive cells) is usually taken to represent
the G0 to S or M to S transition interval. In most normal cell types the interval from G0 to S
is significantly longer (up to 2-fold) than the interval from M to S.

In an asynchronous culture one should see all the S phase cells labeled at the first time point,
followed by a straight rise to 100% labeling (the interval from the start of the experiment to
the point of 100% labeling represents the length of G2+M+G1 phases). In either case failure
to reach 100% labeling indicates that not all the cells are cycling. Significant departures from
a linear rise in labeling indicate the presence of subpopulations with different cell cycle kinetics.

If cells are seeded at the start of the experiment (for example, after mitotic shake-off) the volume
can be reduced by up to 50%; the dishes should then be kept absolutely still in the incubator
to promote attachment. We have routinely taken the first time point at 2 h at which point the
cells, although not completely spread out, are sufficiently attached to allow processing. With
many fibroblast cell lines it should be possible to push back the first time point to 1 h. If
attachment is a problem (or if earlier time points are desired), gentle centrifugation (500 x g,
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room temperature, 5 min) in a swinging bucket rotor equipped to accommodate microtiter

plates is a good option. Another option is to coat the wells with adhesion-promoting substrates
(e.g., collagen, fibronectin, matrigel, etc.).

3.6 Determination of the restriction point

The restriction point separates G1 into 2 sub-phases referred to as G1pm (G1 post mitosis) and
G1ps (G1 pre synthesis) (Zetterberg, et al. 1995). G1pm is necessary for several signaling
processes that are required for cell cycle commitment, whereas G1ps is characterized by rapid
macromolecular synthesis in preparation for S phase. The exact molecular landmarks defining
the restriction point are still debated, but there is a general agreement that the key events are
the phosphorylation and successive inactivation of the retinoblastoma protein. Thus once a cell
has passed the restriction point it becomes largely independent of mitogenic signaling, and is
committed to complete one cell cycle even if exogenous stimuli are withdrawn (Fig. 2A). Cells
synchronized either in M or G0 can be used in these experiments. Typically, the G1pm interval
in exponentially cycling cells (M synchronization) is very short (Fig. 2B,C), whereas during
the cell cycle recruitment of resting cells (G0 synchronization) it is much longer. In contrast,
the duration of G1ps is similar under both conditions (Schorl and Sedivy, 2003).
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Based on the property of mitogen-independence, we modified earlier assays to allow the facile
determination of the restriction point (Fig. 2AC). Synchronized cells are seeded in 24 well
clusters in the presence of BrdU/uridine-containing complete medium and allowed to attach.
At successive time points one well is washed 3 times with PBS, serum-free medium containing
BrdU/uridine is added back, and incubation is continued for a time previously determined by
continuous labeling to give complete S phase entry (high labeling index). In this experimental
setup, the key (and only) variable is the time at which serum is withdrawn, since all cells are
incubated for the same total time and BrdU is present throughout. The halfway point in the rise
of BrdU-positive cells thus demarks the time at which S phase entry becomes non-responsive
to serum withdrawal.

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3.6 Labeled mitoses

This assay allows a direct kinetic determination of G2 length in exponentially cycling
asynchronous cultures. The assay is based on the ability to distinguish mitotic cells
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microscopically by their characteristic dumbbell shape (mitotic figures). Asynchronously

growing cells are pulse labeled with BrdU for 30 min, washed several times with PBS,
incubation is continued in normal medium in the absence of BrdU, and successive time points
are taken (Fig. 3A). We use 6 cm dishes and score the entire dish for each time point. The brief
pulse labels all S phase cells, and the initial appearance of BrdU-labeled mitotic figures thus
denotes the time needed for cells labeled in late S phase to traverse into mitosis. Typically, all
mitotic figures become labeled rapidly, and the G2 interval is taken as the half point of this
rise. The number of BrdU-positive cells should drop sharply again as the cells labeled in early
S phase complete the traverse, and the steepness of the down curve is a good indicator of
synchrony. In such cases, the width of the peak at the half way point is another independent
means of calculating the length of S phase (Fig. 3A).

Several possible complications need to be taken into account. First, because of their rounded
shape, it may not be easy to distinguish BrdU-positive and BrdU-negative cells under phase
illumination, and observations should thus be verified by examination under bright field
illumination. Second, since mitosis is the shortest phase of the cell cycle, large numbers of total
cells need to be scanned to find a statistically significant number of mitotic cells. Finally,
mitotic cells are relatively loosely attached to the plates and can thus be easily washed off
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during the antibody staining and processing steps.

3.7 Double labeling

One parameter that is not easily measured in unperturbed cultures is the time for one complete
cell cycle. Since even gentle synchronization such as mitotic shake-off can affect cell cycle
progression, the ideal experiment would simply employ exponential, asynchronously growing
cultures. Such an approach is made possible by the existence of antibodies specific for two
different nucleotide analogs, Chloro deoxyuridine (CldU) and Iodo deoxyuridine (IdU). Cells
are marked in S phase using the first analog, and entry of these cells into the next S phase is
then monitored with the second analog. In practice, exponentially growing cells are pulse
labeled with CldU for 30 min, washed twice with PBS, and cultured with excess thymidine for
1 h to dilute the intracellular pool of CldU. The thymidine-containing medium is then removed,
and incubation is continued with normal medium. At subsequent time points a second 30 min
pulse using IdU is delivered, the samples are quenched, fixed, and stained for
immunofluorescent observation using the CldU- and IdU-specific antibodies. A representative
experiment is shown in Fig. 3B.
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4 Processing and analysis

4.1 Fixation
While ascorbic acid is a superb quenching reagent, and cells can be kept in its presence in
culture medium for up to 48 h, we prefer to proceed to the fixation steps as soon as the
experimental regimen allows. Prior to fixation, the plates should be washed quickly but very
thoroughly 23 times with PBS. During all steps it is important to avoid pipetting directly on
top of the cells, as this can dislodge them. Likewise, it is important to prevent the cells from
drying out at any point. We therefore do not stain more than 8 to 10 plates at one time, and
remove solutions by simply decanting the medium into a large beaker, as opposed to using an
aspirator, which takes much longer. After the PBS washes to remove the medium and ascorbic
acid, cells are fixed with 100 % ice-cold methanol for 10 min at 4 C, followed by 3 washes
with PBS. At this point the plates can be stored under PBS sealed with parafilm (to avoid
desiccation) at 4 C for several weeks.

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4.2. Denaturation
In order to provide accessibility to the antibodies, the DNA needs to be rendered single-stranded
to expose the incorporated nucleotide analogs. The classical method is to denature the DNA
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with acid (1.5 M HCl final concentration, 1 h at room temperature, gentle rocking). Immediately
afterwards the plates are washed 3 times (34 min each wash) with 0.1 M borate buffer (boric
acid, BioRad, Hercules, CA, cat. no. 161-0751, pH adjusted to 8.5 with NaOH), followed by
3 similar washes with PBS. At this point it is possible to store the plates under PBS at 4C as
described before.

Another way to expose the incorporated nucleotide analogs is by partial degradation with
benzonase, a genetically engineered nuclease which degrades both single and double stranded
DNA (VWR International, West Chester, PA, cat. no. 80108-808). In this protocol the exposure
of single-stranded regions and binding of the antibody are accomplished simultaneously. Cells
are incubated in a humidified chamber at 37C for 2 h with anti-BrdU monoclonal antibody
(Becton Dickinson, San Jose, CA, cat. no. 555627), diluted 1 : 200 in PBS containing 1 mM
Mg2+ and 125 U/ml benzonase. This is followed by the desired secondary antibody (Herbig et
al., 2004).

4.3 Staining
Non specific staining is prevented by first blocking with 0.1% PBS-BSA (PBSA) solution
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(0.1% w/v BSA Fraction V, Fisher Scientific, cat. no. BP1600-100) dissolved in PBS in a
humidified chamber at 37C for 1 h without shaking. Primary anti-BrdU monoclonal antibody
(Becton Dickinson, cat. no. 555627, 0.5 mg/ ml stock concentration) is used at 1 : 200 dilution
in PBSA for 1 h, also at 37 C in the humidified chamber. The diluted antibody can be re-used
4- to 5-times without loss of signal intensity and should be stored at 20 C. All subsequent
steps are carried out at room temperature and utilize the Vectastain ABC Elite Mouse IgG Kit
from Vector Laboratories (Burlingame, CA, cat. no. PK-6102), combined with the Vector
Laboratories NovaRED Substrate Kit for peroxidase (cat. no. SK 4800).

Plates are first rinsed briefly several times with PBS, and then agitated gently on a rotating
platform for 10 min under PBS before adding the biotinylated secondary antibody. We have
adopted the protocol suggested by the manufacturer in order to avoid over-staining. The
secondary antibody solution is comprised of 10 ml PBS, 115 l normal horse serum, and 23.5
l biotinylated anti-mouse secondary antibody. 7501000 l of this solution are added per well
of a 24 well culture dish, and incubated for 30 min.

During this time Vectastain solution AB is prepared (10 ml PBS, 4 drops solution A, 4 drops
solution B, mixed thoroughly after adding each solution). Solution AB is pre-incubated for 30
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min at room temperature before addition to the cells. The time required for a PBS rinse followed
by a 10 min PBS wash (above) after the secondary antibody should be taken into account while
preparing solution AB. 7501000 l of solution AB is used per well, incubated for 30 min, and
again followed by a PBS rinse and a 10 min PBS wash (above). Finally, a brief rinse with
dH2O is used to remove salts, which is critical for even staining. The cells can are kept in
dH20 while the NovaRED staining solution is prepared by mixing 10 ml dH2O with 3 drops
each of reagents 1 through 3 (immediate mixing after adding each component is important).
Finally, 2 drops of hydrogen peroxide solution are added, mixed in, and the staining solution
is added to the cells. The staining intensity is checked immediately and frequently in a
microscope at 20 x magnification. After good BrdU incorporation nuclear staining is typically
easily detectable after 30 to 40 sec (but can be strong enough as early as 10 sec), at which time
the reaction can be stopped by performing several dH2O washes. Prolonged exposure of the
cells to the staining solution is unnecessary and will result in non-specific staining. Stained

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plates can be stored at 4C under dH2O (sealed with parafilm) for several weeks without loss
of signal intensity.
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4.4 Two color immunofluorescent detection of CldU and IdU incorporation

This procedure is adapted from the one published by Aten et al. (1992). CldU and IdU can be
purchased from Sigma (cat. no. C6891 and I7125, respectively). While histochemical detection
of BrdU incorporation can be performed directly on cells grown in plastic culture plates, for
immunofluorescent detection cells should be seeded on glass coverslips which can then be
cultured in appropriately sized culture vessels. Exponentially growing cells are incubated
initially for 30 min with 2.6 g/ml CldU (10 M final, stock solution is 1000 x or 10 mM
prepared in the same way as for BrdU), followed by 2 washes with PBS and incubation for 1
h in medium containing 200 mM thymidine. Cells are again washed twice with PBS and
incubation is continued using normal medium. At subsequent times the medium is changed for
a 30 min labeling period to medium containing 3.5 g/ml IdU (10 M final, stock solution is
1000 x or 10 mM prepared in the same way as for BrdU). If needed, the labeling is quenched
with ascorbic acid as indicated above.

To process the samples the cells are first washed three times in 0.05 % Tween-PBS (PBS-T).
All antibody incubations are carried out in a humidified chamber at room temperature. Rat
monoclonal antibody clone BU1/75 (ICR1; Harlan, Indianapolis, IN, cat. no. MAS 250) diluted
1 : 520 in PBSA-T (PBS-T containing 0.1% BSA) was used to detect CldU. For detection of
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IdU we used mouse monoclonal antibody clone B44 (BD Biosciences, San Jose, CA, cat. no.
347-580) diluted to a final concentration of 2.5 g/ml in PBSA-T. The secondary antibodies
were a Cy3-conjugated donkey anti-rat antibody (Jackson Immunoresearch, West Grove, PA,
cat. no. 712-165-153), and Alexa 488-conjugated goat anti-mouse antibody (Molecular Probes,
Eugene, OR, cat. no. A-11029), both used at a final concentration of 3.0 g/ml.

The antibodies were used sequentially, with the anti-CldU antibody being used first, followed
by the donkey anti-rat antibody, then the anti-IdU antibody, and finally the goat anti-mouse
antibody. A high salt wash (28 mM Tris, pH 8.0, 500 mM NaCl, 0.5% Tween 20) for 10 min
followed by a 10 min wash in PBSA-T was used after each antibody incubation to reduce non-
specific signals. Finally, cells were counterstained with 0.1 g/ml 4,6-diamidino-2-
phenylindole (DAPI) for 15 min to visualize the nuclei, followed by a final PBS wash.
Microscopic analysis was performed at 200 x magnification. Randomly selected fields were
photographed with a Spot-II digital camera (Diagnostics Products, Los Angeles, CA) and the
images were scored. Exposure to the excitation beam as well as to overhead fluorescent lighting
should be minimized to avoid photo bleaching.

5.1 Analysis of proliferative capacity using CFSE staining

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This procedure is very useful to assess the longer term proliferative capacity of cultures,
specifically, whether all cells are cycling uniformly and continuously, and to address the
possible existence of slower or faster growing sub-populations ( Lyons, 1999, Lyons et al.,
2001). Under ideal conditions it allows the monitoring of cultures for multiple cell cycles, to
a degree that cannot be achieved using nucleotide incorporation methods. 5-(and 6-)
carboxyfluorescein diacetate succinimidyl ester (CFSE; Invitrogen, Molecular Probes, cat. no.
C-1157) diffuses into cells where it attaches to amine groups of cytoplasmic proteins and is
metabolized by cellular esterases to a fluorescent dye. CFSE has very similar spectral properties
to fluorescein and hence can be detected in the FL-1 channel of a flow cytometer.

CFSE is prepared as a 5 mM stock solution in DMSO, and stored at 20 C protected from

light (do not freeze or thaw more than 3 times). 13 x107 cells are trypsinized, collected by
low speed centrifugation, resuspended in 1 ml complete medium containing 10 M CFSE, and

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incubated for 10 min at 37C. 14 ml of ice-cold complete medium is added and the incubation
is continued on ice for another 5 min. Cells are again collected by centrifugation and washed
once in CFSE-free medium. A small aliquot is removed and immediately analyzed by FACS
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as intact, live cells (without fixation).

The remaining cells are returned to culture in 10 cm dishes, making sure to seed them at a low
enough density to allow multiple cell cycles. One dish is analyzed by FACS each subsequent
day (the time intervals should be matched roughly to the doubling time). This method relies
on the fact that after an initial rapid turnover period (manifested as a large decrease in
fluorescence during the first 1224 h), subsequent turnover is minimal and the dye-modified
proteins are distributed equally to the daughter cells. Thus, most dye dilution results from cell
growth and division, such that daughter cells are roughly half as bright as mother cells. These
successive generations can be visualized as discrete peaks by FACS, shifted to the left in the
FL-1 channel compared with the peak of the initial cell population. The reduction in
fluorescence intensity can be followed over several cell cycles, and shoulders or small peaks
remaining to the right of the main peak indicate the existence of sub-populations of non-
dividing cells.

Conclusion
In this review we have provided detailed protocols for several methods that allow kinetic
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analyses of cell cycle progression in living cells. These protocols can be easily modified to the
experimental needs of the investigator. Undoubtedly the near future will see great progress in
the understanding of the cell cycle and cell cycle progression, and the knowledge gained from
studying the cell cycle will result in improved therapies for numerous diseases. Indeed, small
inhibitor molecules targeting the dysregulated activities of key cell cycle regulators are
currently undergoing clinical trials, and it is hoped that they will significantly improve cancer
therapy.

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Figure 1.
Pulse labeling (A) and continuous labeling (B) of c-myc+/+ and c-myc/ cells synchronized
by mitotic shake off (reproduced from Schorl and Sedivy (2003) Mol. Biol. Cell 14, 823835).
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Figure 2.
(A) Schematic representation of the method to determine the restriction point. Phases of the
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cell cycle: M, mitosis; G0, quiescence (resting cells); G1pm, G1 phase post mitosis; G1ps, G1
phase pre DNA synthesis; S, S phase; G2, G2 phase. R designates the restriction point. Filled
nuclei depict cells that have incorporated BrdU. For further details see text. (B) and (C)
Restriction point determination in cycling cells using mitotic shake off for synchronization
(reproduced from Schorl and Sedivy (2003) Mol. Biol. Cell 14, 823835). (B) c-myc+/+ cells.
(C) c-myc/ cells.

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Figure 3.
(A) Labeled mitoses experiment performed with exponentially cycling c-myc +/+ cells. (B)
Double labeling with CldU and IdU to determine entry into the second S phase in exponentially
cycling c-myc+/+ and c-myc/ cells. The data (labeling indices) are presented as the percentage
of CldU/IdU double-labeled cells in the total pool of CldU-positive cells. For further details
see text (reproduced from Schorl and Sedivy (2003) Mol. Biol. Cell 14, 823835).

Methods. Author manuscript; available in PMC 2007 March 20.