Am J Physiol Cell Physiol 304: C927–C938, 2013.
First published February 7, 2013; doi:10.1152/ajpcell.00006.2013.
Review
METHODS IN CELL PHYSIOLOGY
Classic “broken cell” techniques and newer live cell methods for cell cycle
assessment
Lindsay Henderson,1 Dante S. Bortone,1 Curtis Lim,1 and Alexander C. Zambon2,3
1
Department of Biology, University of California San Diego, La Jolla, California; 2Department of Medicine, University of
California San Diego, La Jolla, California; and 3Department of Pharmacology, University of California San Diego,
La Jolla, California
Submitted 8 January 2013; accepted in final form 1 February 2013
cell cycle pathway; live cells; sensors; analysis methods; review
THE CELL DIVISION CYCLE, referred to here as the cell cycle, is a
highly conserved pathway that plays a central role in tissue
development and cellular homeostasis. The cell cycle consists
of three “gap” phases, G0, G1, and G2, that are interspersed
between two other phases, i.e., the S (DNA synthesis) phase,
where DNA synthesis results in chromosome duplication, and
the M (mitosis) phase, where the process of karyokinesis is
often coupled to cytokinesis to complete cell division (Fig. 1).
Cellular dormancy is when normal cells exit the cell cycle and
enter G0. Cells in G0 often display a decrease in overall
metabolic activity, reduced rRNA synthesis, decreased translation, and decreased cell size, rendering them resistant to a
variety of stresses (2, 116). However, distinguishing between
cells in G0 and cells that are arrested in G1 can be technically
challenging, and thus many adult nondividing somatic cell
types (e.g., cardiomyocytes, neurons) are commonly classified
as being in G0/G1 arrest (28, 87, 95, 124).
Address for reprint requests and other correspondence: A. C. Zambon, 9500
Gilman Dr., La Jolla, CA 92093-0636 (e-mail: [email protected]).
http://www.ajpcell.org
Many of the core genes that control the cell cycle were first
discovered in yeast, and this model organism continues to shed
light on new pharmacological agents that perturb this important
pathway (72). Dysregulation of the cell cycle can lead to the
development and progression of a malignant phenotype. It is
thus not surprising that the majority of currently available
cancer therapeutics target core aspects of the cell cycle pathway (119).
The cell cycle is controlled by an orchestrated network of
genes and their encoded proteins that are temporally regulated by transcription, dynamic protein-protein interactions,
and posttranslational modifications (e.g., phosphorylation,
ubiquitination). Many mammalian cells require 12–24 h to
complete cell division, with differences in cycle time goverened primarily by time spent in the various gap phases
(i.e., G0, G1, and G2) (105). This “circadian” nature of cell
cycle kinetics is likely influenced by the regulation of core
cell cycle genes (e.g., P21, Ccnd1, Ccnb2, Ccna1) by
circadian clock genes or their encoded proteins (9). However, in some circumstances, e.g., neural stem cells in vivo,
cell cycling times can be much shorter (12 h) or longer (ⱕ15
0363-6143/13 Copyright © 2013 the American Physiological Society
C927
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Henderson L, Bortone DS, Lim C, Zambon AC. Classic “broken cell”
techniques and newer live cell methods for cell cycle assessment. Am J Physiol
Cell Physiol 304: C927–C938, 2013. First published February 7, 2013;
doi:10.1152/ajpcell.00006.2013.—Many common, important diseases are either
caused or exacerbated by hyperactivation (e.g., cancer) or inactivation (e.g., heart
failure) of the cell division cycle. A better understanding of the cell cycle is critical
for interpreting numerous types of physiological changes in cells. Moreover, new
insights into how to control it will facilitate new therapeutics for a variety of
diseases and new avenues in regenerative medicine. The progression of cells
through the four main phases of their division cycle [G0/G1, S (DNA synthesis), G2,
and M (mitosis)] is a highly conserved process orchestrated by several pathways
(e.g., transcription, phosphorylation, nuclear import/export, and protein ubiquitination) that coordinate a core cell cycle pathway. This core pathway can also receive
inputs that are cell type and cell niche dependent. “Broken cell” methods (e.g., use
of labeled nucleotide analogs) to assess for cell cycle activity have revealed
important insights regarding the cell cycle but lack the ability to assess living cells
in real time (longitudinal studies) and with single-cell resolution. Moreover, such
methods often require cell synchronization, which can perturb the pathway under
study. Live cell cycle sensors can be used at single-cell resolution in living cells,
intact tissue, and whole animals. Use of these more recently available sensors has
the potential to reveal physiologically relevant insights regarding the normal and
perturbed cell division cycle.
Review
C928
APPROACHES FOR CELL CYCLE ASSESSMENT
Mitosis
G2
G0
G1
S-phase
Fig. 1. Main phases of the cell cycle.
The Core Signaling Pathway that Controls the Cell Cycle
The transition into G1 prepares the cells for DNA synthesis
and is controlled by interactions of cyclin-dependent kinase
(CDK)4 and CDK6 with their heterodimeric partners: the
cyclin D CCND1–3 family members (the choice of partner is
poorly understood) (12, 107). The induction of CDK4/6 and
CCND proteins facilitates their translocation to the nucleus,
where they become further activated by phosphorylation from
the CDK-activating kinase (CAK) complex (comprised of
CDK1, CDK2, and cyclin A or B). Subsequently, CDK4/
CCND1 phosphorylates retinoblastoma (RB) protein family
members catalyzing RB exchange with transcription factor
Dp1 (Tfdp1) family members for binding to and the subsequent
activation of the E2F family of transcription factors. Seven
E2F (1–7), two Tfdp (1, 2), and three RB (p-RB, p-107, and
p-130) genes have been identified in humans and mice (24, 25).
Activated E2F/TFDP heterodimers bind to DNA consensus
elements in the promoters of genes that carry out DNA synthesis during the S phase (Fig. 2). CDK2/CCNE heterodimers
complete the transition into the S phase by phosphorylation of
the initiators of DNA replication among others (74).
A variety of cell cycle checkpoint genes control progression
through the cell division cycle. These CDK inibitors (CKIs)
inhibit cell cycle progression by binding to and sequestering/
inhibiting CDK activity (18). CKIs are regulated by both
transcriptional and posttranscriptional mechanisms (e.g., subcellular sequestration, ubiquitination). They fall into two classes: the INK4 family (p14, p15, p16, p18, and p19) and the
KIP/CIP family (p21, p27, and p57). These two families are
structurally unrelated and have differing specificities for regulating cell cycle progression (Fig. 2) (11).
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days or more) depending on cell type and anatomical location (71).
Classic methodology to assess for cell cycle progression,
such as quantification of DNA content, DNA synthesis, and
immunohistochemical analysis of proliferative marker genes,
has enabled many seminal insights into signaling pathways and
pharmacological agents that control it. Many of these classic
experimental approaches relay on cell fixation and/or synchronization, and thus aspects of cell proliferation cannot be examined by such “broken cell” methods. For example, classic
approaches cannot be employed for longitudinal studies examining heterogeneity of cycling kinetics in cultured cells, tissues,
or intact animals.
The development of new methods that enable the assessment
of cell cycle progression in living cells, tissues, and intact
animals has set the stage for unprecedented studies in the area
of cell cycle regulation. These tools are revealing new insights
into the dynamic temporal and heterogeneous kinetics of the
cell cycle. For example, it is now possible to label subpopulations of living cells in each of the main phases of the cell cycle
with genetically encoded fluorescent proteins, thereby providing a molecular beacon to pinpoint single cells as they traverse
through the cell cycle. This approach enables one to quantify
cells in specific phases of the cell cycle over time without
perturbing cell or tissue structure, thereby maintaining cell or
tissue integrity for downstream applications. For example,
using imaging one can simultaneously track and quantify cells
in specific phases of the cell cycle while quantifying the
duration of each cell in specific phases. Alternatively, these
live cell sensors facilitate high-throughput screening of compounds or cDNAs that can induce or inhibit the cell cycle with
standard fluorescent plate readers.
Live cell sensors enable in vivo studies that were not
previously possible. They can be used to visualize heterogeneity of tumor cell proliferation in vivo as a function of tissue
microenvironment and/or drug treatments. These live cell cycle
sensors can also be used to isolate and purify cell populations
in distinct phases of the cell cycle for genomic studies such as
transcriptional profiling or proteomics. Such studies could be
used to probe complex gene regulatory networks and/or quantify changes in cellular proteins during specific phases of the
cell cycle. Other, more detailed specific examples of the
application of these novel reporters are provided and referenced below.
Another application of these novel live cell cycle sensors is
in the exciting and burgeoning area of single-cell analysis. The
goal of examining cellular heterogeneity over many days or
even weeks at the single-cell level was once just a hope. This
goal is now attainable with the advent of live cell cycle
reporters and rapidly developing technologies, which include
single-cell PCR, advances in live cell and whole animal in vivo
imaging, and laser capture microdissection. Integration of these
approaches has enabled simultaneous multiparameter tracking
and quantification of the cell cycle within single cells in vitro
and in vivo.
In this review, we first provide a general overview of the
core signaling pathway that controls the cell cycle. The fundamental molecular processes at the heart of this pathway, such
as DNA synthesis and the expression of proteins during specific phases of the cell cycle, serve as the basis for the
development of methods to assess cell cycle progression. We
then discuss classic methods that generally rely on broken cell or
fixed tissue techniques. Finally, we discuss newer approaches that
use cell-permeable or genetically encoded live cell cycle sensors
that have enabled studies of the cell proliferation at single-cell
resolution in intact cells, tissues, and whole animals.
Several reviews and books on cell cycle biology and techniques have been published (e.g., see Refs. 24, 43, 77, 88, and
119). The purpose of this review is to integrate newer findings
and methods in the context of classic ones and their inherent
limitations. We anticipate that organizing this review in such a
way will provide an intellectual platform for the creative
integration of old and new techniques to ask important new
questions in this area.
Review
APPROACHES FOR CELL CYCLE ASSESSMENT
KIP/
CIP
Cdc25c
p21
p27
p57
CAK
Wee1
Myt1
Cdk7
Ccnh
P
Ccnb
Cdk1
Ccna
Cdk1
M
Late G2
Mitosis
P
G0
INK4
p14
p15
p16
p18
p19
Ccnd
Cdk4
Ccnd
Cdk6
P
G0-G1
Ccnb
Cdk1
C929
Rb
E2f
G1
G2
P
P
Rb
G2
S-phase
Ccne
Cdk2
Ccna
Cdk2
Late
G1-S
S-G2
P
KIP/
CIP
p21
p27
p57
Dp1
E2f
Transcription of
cell cycle machinery
for S phase
Initiators of DNA
Replication
P21 and P27(KIP1) negatively regulate CDK activity and
also help assemble CCND/CDK4 and CDK6 complexes in G1,
thereby both inhibiting and to a smaller extent facilitating cell
cycle progression (99). This balance between pro- and antiproliferative signaling is mediated by a complex balance of stoichiometric interactions and changes in cellular compartmentalization and by mechanisms that have yet to be fully elucidated. P21 and P27 promote CCND/CDK4 and CDK6
interactions by stablizing the complexes and by directing the
complexes to the nucleus (99). Low concentrations of P21
promote assembly and kinase activity, whereas higher concentrations of P21 inhibit kinase activity (58). The inhibition of
CDK4 by P27 depends on the absence or presence of P27
tyrosine phosphorylation, which modifies P27 from a bound
inhibitor to a bound noninhibitor (91). During the G1-to-S
transition, P27 is ubiquitinated by S phase kinase-associated
protein 2 (SKP2) and then targeted for degradation (99). The
degradation of P27 is necessary for the transition from G1 into
S; this is associated with an increase in E2F-dependent transcriptional activity (64). As discussed below, some of these
changes in protein stability and cellular compartmentation have
been exploited to generate sensors for assessment of cell cycle
regulation.
During the progression through the S phase, CCNA/CDK2
complexes form and phosphorylate several proteins, including
cell division cycle 6 protein (CDC6; Fig. 2). CDC6 is involved
in the formation of the initiation complex and origin liscensing
(40, 46). Mechanisms that prevent DNA rereplication in eukaryotic cells inhibit origin licensing. Origin licensing occurs
by the binding of the origin recognition complex, a multisubunit ATPase, to DNA at the replication origins and the recruitment of CDC6 and the chromatin licensing and DNA replica-
tion factor 1 (CDT1) protein. These events are required prior to
the recruitment and binding of minichromosome maintenance
proteins 2–7 onto chromatin (69). The minichromosome maintenance complex, once bound, is the DNA helicase that opens
the helix at the replication origin and unwinds the two strands
as replication forks travel along the DNA (17).
In G2, the fidelity of chromosome replication is assessed by
the DNA damage response pathway as the cell prepares to
enter the M phase. A primary function for G2 is to prevent
chromosome segregation in the presence of unreplicated or
damaged DNA (102). CCNB/CDK1 complexes drive early G2
transition, and their activity is regulated by phosphorylation
and ubiquitination (77). If DNA damage is detected, signaling
pathways (e.g., activity of ATM/ATR kinases and their downstream substrates CHK1 and CHK2 kinases) lead to the phosphorylation of the phosphatase CDC25C sequestering it in the
cytoplasm and preventing it from dephosphorylating and activating nuclear CCNB/CDK1 triggering G2 progression (Fig. 2)
(54). Subsequently, the cell enters G2 arrest (91). When DNA
replication is complete, CCNA/CDK1 complexes regulate the
transistion into the M phase.
During the entry into the M phase, the activity of CDC25C
with CDK1 is greater than the activities of opposing kinases
WEE1 and MYT1. CDC25C and WEE1 work in positive and
negative feedback loops, respectively, to fully activate CCNB/
CDK1. The activated CCNA/CDK1 and CCNB/CDK1 complexes phosphorylate substrates that are important for nuclear
envelope breakdown and centrosome separation (77). During
G2, centrosomes prepare for duplication so that they can
organize the spindle poles during mitosis. In mammalian cells,
centrosomes are comprised of two centrioles surrounded by
pericentriolar material. The single centrosome duplicates prior
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Dp1
Fig. 2. Simplified pathway representation of the major regulators of cell cycle progression. CDK, cyclin-dependent kinase;
CAK, CDK-activating kinase; RB, retinoblastoma; P, phosphorylation; G, gap phase; INK4, inhibitor of cyclin-dependent
kinase 4; KIP, kinase inhibitor protein; CIP, cyclin inhibitor
protein.
Review
C930
APPROACHES FOR CELL CYCLE ASSESSMENT
Classic Methodology to Assess for Cell Cycle Progression
Cell counting and incorporation of labeled nucleoside
analogs. The most direct approach for assessing an end point
of cell cycle activity, i.e., cytokinesis, is to simply count cells
over time. In light of the fact that some cells become polynucleated and/or polyploid in vitro and in vivo and under stress
[e.g., cardiomyocytes (59, 65, 104, 118)], determining total cell
number is necessary when assessing cell proliferation by other
methods (e.g., increased DNA synthesis, as discussed below).
Cell counting is often conducted in the presence of a dye (e.g.,
trypan blue), which is used to identify cells with compromised
cellular membranes. This enables one to exclude apoptotic or
necrotic cell populations. These types of studies are usually
conducted with a hemocytometer that employs a grid system
that bounds a defined volume, thereby enabling cell density
determination.
Coulter counters are also able to assess for cell number while
simultaneously recording distributions of cell size. These machines work on the principle that cells are nonconducting
particles that disrupt a current when passing through a channel.
The size of current disruption is proportional to cell size. In
recent years, a number of smaller and cheaper instruments that
are based on the coulter principle (e.g., Scepter; EMD-Millipore) or image quantification Countess (Invitrogen) have become available as alternatives to coulter counters.
Although counting cell numbers as a function of time is a
straightforward method for assessing cell proliferation, in some
situations it may not be sensitive enough to detect small but
significant changes in proliferation rates. Moreover, it cannot
be used to assess cell proliferation in vivo. Cell-permeable
nucleoside analogs {e.g., [3H]thymidine, bromodeoxyuridine
(BrdU)} are a sensitive means to assess for DNA synthesis as
an indication of cell proliferation (63, 79). These analogs are
incorporated into newly synthesized DNA during S phase
progression. Importantly, they can be used in vitro and in vivo
in pulse chase experiments to assess cell proliferation during a
defined window of time (79, 104). The use of BrdU has largely
replaced that of [3H]thymidine since the latter requires radioactive handling procedures that can pose hurdles in some
laboratory settings. Moreover, the use of BrdU facilitates
multiparameter assessment (e.g., immunohistochemical staining for cellular antigens as well as changes in DNA synthesis).
Other “BrdU-like” nucleoside analogs with modified halogen moieties such as chlorodeoxyuridine or iododeoxyuridine
have been developed to enable sequential S phase cell labeling
in vitro and in vivo. This approach can be useful for identifying
cells that have undergone two rounds of DNA synthesis (115,
117). These analogs label cells that are traversing S phase
with equal efficiency when administered in equimolar concentrations, thereby enabling both qualitative and quantitative studies.
BrdU and possibly other halogenated thymidine analogs
have limitations that include adverse effects on cell proliferation and viability (63). In addition, in some cellular settings
(e.g., neurons), incorporation of nucleoside analogs occurs in
cells undergoing DNA repair, turnover, and/or apoptosis (5,
57). It is postulated that a DNA damage response is triggered
in terminally differentiated neurons subjected to death-inducing stimuli in vitro or in vivo as well as in Alzheimer’s disease
(33). A variety of death-inducing stimuli can upregulate cyclins, CDKs, and DNA synthesis, thereby increasing BrdU
incorporation in postmitotic neurons undergoing apoptosis (5,
33). Staining for BrdU, chlorodeoxyuridine, and iododeoxyuridine with anti-sera also requires strong denaturing conditions
such as the use of concentrated HCl or mixtures of methanol
and acetic acid and high heat, which can degrade the structure
of the specimen. These detection conditions can disrupt
epitopes in colabeling experiments.
In light of these limitations, 5-ethynyl-2=-deoxyuridine
(EdU) was developed as an alternative to BrdU (94). EdU can
be detected with fluorescent azides in a Cu(I)-catalyzed “click
chemistry” reaction that is highly sensitive and much faster
than BrdU detection. EdU does not require an antibody, and
thus the label has a higher diffusion rate that penetrates tissue
much more effectively. Moreover, denaturation of the specimen is not required, thus maintaining cell epitope integrity for
colabeling experiments. However, a variety of fluorophores
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to cell entry into mitosis. The resulting daughter centrosomes
create the poles of the spindle once the nuclear envelope is
broken down in mitosis (41).
Mitosis occurs in five phases: prophase, prometaphase,
metaphase, anaphase, and telophase. During mitosis the
CCNB/CDK1 complex phosphorylates CDC25C, and this creates a positive feedback loop to induce activation of CCNB/
CDK1 at the G2/M transition (Fig. 2) (45, 83). The CCNB/
CDK1 complex helps initiate various events in mitosis. In
prophase, the chromosomes condense and centrosomes promote nuclear envelope breakdown (4). Here, the CCNB/CDK1
complex phosphorylates the centrosome-associated motor protein Eg5 (8), which leads to centrosome separation. In nuclear
lamina breakdown, the lamins disperse, the metaphase mitotic
spindle is created, and the lamins complete their dispersal
before the sister chromatids separate in anaphase (81). The
CCNB/CDK1 complex helps with breaking down the nuclear
lamina and cell rounding that occurs by the disassembly and
reassembly of microfilaments (121). The chromosome condensation that occurs in prophase is controlled in part by activity
of mitotic histone H3 kinases. Aurora kinase family members
and other kinases can phosphorylate the NH2 terminus of H3 at
Ser10 and Ser28 (39) to initiate chromosome condensation. As
discussed below, both Aurora kinases and phospho-H3 (Ser10)
are commonly used immunohistochemical markers for cell
proliferation. CCNA and CCNB must be degraded to exit
mitosis; this degradation is accomplished by a ubiquitin-mediated pathway that is regulated by the anaphase-promoting
complex/cyclosome (APC/C) ubiquitin ligase (31, 76). This
cell cycle phase-dependent change in protein stability by
APC/C has also been harnessed to generate live cell cycle
sensors, as discussed below.
Cytokinesis, the physical separation of a single cell into two
daughter cells, is regulated by the Rho proteins, which are part
of the Ras family of GTPases. The Rho proteins are involved
with positioning the cell division plane as well as the spatial
and temporal regulation of the contractile ring (49). The RhoA
protein is concentrated at the division plane before cell division
occurs and is activated in the equatorial plane. The contractile
ring is formed from actin filaments and myosin (82). RhoA
promotes actin assembly as well as myosin II motor activation,
and then the actin filaments elongate, as is required for the
creation of the contractile ring (49).
Review
C931
APPROACHES FOR CELL CYCLE ASSESSMENT
Table 1. DNA-labeling dyes and their characteristics
Name of Stain
Propidium iodide
7-AAD
DAPI
Draq5
Hoescht 33258
Hoescht 33342
Hoescht 34580
Vybrant DyeCycle
Vybrant DyeCycle
Vybrant DyeCycle
Vybrant DyeCycle
Violet
Green
Orange
Ruby
Table 2. Immunohistochemical markers of cell proliferation
Cell
Permeable
Excitation
Maximum, nm
Emission
maximum, nm
No
No
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
536
488
350
488–647
352
350
392
405
488
488, 532
488, 633/5
617
655
470
665
461
461
440
437
534
563
686
7-AAD, 7-amino-actinomycin D; DAPI, 4=,6=-diamidino-2-phenylindole.
G0/G1
# Cells
1500
1000
Sub-G1
(apoptotic cells)
G2/M
500
S
0
0
200
400
600
800
1000
Pacific Blue-A: Violet DNA Pacific Blue-A
Fig. 3. Cytometry-based DNA content assessment-HT1080 cells were incubated with DyeCycle Violet DNA stain (111) for 5 min and analyzed by flow
cytometry. Populations in the G0/G1, S, and G2/M phases are shaded in pink,
yellow, and green, respectively.
Ki67
Proliferating cell nuclear
antigen
Aurora kinase A
Aurora kinase B
Phospho H3 (Ser10) histone
Phospho-RB (Ser807–811)
Cellular
Compartment
Labeled Phase
MKI67
PCNA
Nucleus
Nucleus
G1, S, G2, M
S
AURKA
AURKB
H3
RB
Nucleus
Nucleus
Nucleus
Nucleus, cytosol
G2, M
G2, M
M
G1-S transition
RB, retinoblastoma.
DNA content (88). Since cells typically cycle heterogeneously,
use of these dyes to study specific transitions within the cell
cycle often requires cell synchronization prior to assessment
for DNA content (38).
Immunohistochemical markers of cell proliferation. A number of cell cycle pathway-associated proteins are used as immunohistochemical markers of cellular proliferation (Table 2). The
most commonly used markers are Ki67, PCNA (29), and
phospho-histone H3 (Ser10). Ki67 is a 360-kDa nuclear protein
that can be used to detect and quantify proliferating cells (89).
It is induced when quiescent, arrested cells enter the late G1-S
transition (36) and continues to be expressed through the G2
and M phases. Its expression is undetectable in cells in G0 (26).
Ki67 expression is elevated in a variety of human tumor tissues
and is a diagnostic marker that inversely correlates with survival rates in a variety of cancers (15, 48, 89). PCNA is a
␦-DNA polymerase cofactor involved in DNA replication and
DNA repair (13, 36). A variety of commercially available
phosphospecific antibodies for cell cycle pathway members
can be used to indicate whether these proteins contain
activating or inactivating phosphorylation states (e.g., phospho-Ser RB).
Biochemical assays for cell cycle assessment. CDK activity
can be measured after immunoprecipitation of specific CDKs
from cellular or tissue lysates or with recombinant purified
kinases. CDK kinase activity is quantified by transfer of
radiolabeled phosphate from [␥-32P]ATP or by immunoblotting using phosphospecific antibodies for CDK substrates (e.g.,
RB, vimentin, nuclear protein in the AT region (NPAT)) (10).
Direct CDK inhibitors such as flavopiridol UCN-01, paullones,
hymenialdisine, and roscovitine can be incorporated into such
assays to validate kinase specificity (98). Cells can be synchronized in a specific phase of the cell cycle by inducing a block
at a specific checkpoint and then releasing the block and
subsequently assaying as a function of time (38). Serum
starvation is one way to arrest and synchronize cells in culture.
In addition, a variety of compounds can be used to synchronize
cells in a specific phase of the cell cycle (e.g., double-thymidine block to arrest cells in G1 and nocodazole to arrest cells in
G2) (38). G1 CDKs (e.g., CDK4/6) and G2/M kinases can
efficiently phosphorylate RB and histone H1 in vitro, respectively. However, cross-reactivity at lower efficiency between
G1 and G2 kinases and the aforementioned substrates can be
observed (10, 12).
A second biochemical approach to assess for cell cycle
activity exploits the functional requirement of specific transcription factor families during cell cycle progression (e.g.,
E2F family members) (7). Electric mobility shift assay is a
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(e.g., GFP, R-phycoerythrin) are diminished by Cu(I)-catalyzed detection of EdU; alternative EdU detection strategies
should be employed when codetection of these molecules is
desired (6).
Cell-impermeable DNA stains. Fluorescent molecules such
as propidium iodide (PI) and 7-amino-actinomycin D (7-AAD)
were found to provide suitable chemical characteristics for
univariate analysis of cellular DNA content by flow cytometry
(Table 1) (88). PI and 7-AAD exhibit a high affinity for nucleic
acids and red and far-red fluorescence, respectively. For use in
cell cycle analysis, cells are fixed, permeabilized, and then
treated with RNase prior to staining. This approach reveals
distribution of cells in three clustered phases of the cycle
(G0/G1, S, and G2/M) and makes it possible to detect apoptotic
cells by fractional DNA content (sub-G1 populations) (88).
Cells in G0/G1 exhibit roughly one-half the fluorescence as
cells in G2/M, and cells in the S phase exhibit a range of
fluorescence, as they synthesize DNA (Fig. 3).
PI or 7-AAD can also be used in conjunction with BrdU or
EdU pulsing to determine the rates of cell cycle progression
(29). Pulse-chase labeling experiments are used to provide
information about the kinetics of cell cycle progression during
distinct phases. Cells are first pulsed with a labeled nucleotide
analog for a duration of time, and then cells are fixed and
assayed for nucleotide incorporation (new DNA synthesis) and
Gene
Name
Protein Name
Review
C932
APPROACHES FOR CELL CYCLE ASSESSMENT
Methods for Cell Cycle Assessment in Living Cells and Tissue
Cell-permeable dyes for live cell labeling of DNA content
and cell proliferation. A variety of membrane-permeable dyes
have been developed to determine DNA content in living cells
(Table 1) (70, 101, 111). Optimal dye characteristics for live
cell cycle assessment include cell and tissue permeability, low
cytotoxicity and phototoxicity, photostability, and stochiometric specificity for DNA with minimal affinity for RNA. Also, if
possible, spectral characteristics that render it compatible with
other fluorescent proteins (e.g., GFP, RFP) facilitate multiparameter assessment. DNA content staining is useful for directly
quantifying the number of cells in distinct phases in the same
way that cell-impermeable DNA stains are used, as described
above. However, they have the added advantage that intact live
cells can loaded with the dye. This is important if one wants to
quantify changes in DNA content in vivo after a perturbation
within a defined window of time (e.g., drug dose, injury).
Alternatively, these dyes can be used to purify or quantify cells
in distinct phases of the cell cycle or containing different
amounts of DNA ploidy (e.g., G0/G1, S, 2N, 4N, etc.) while
maintaining cell integrity for downstream applications (e.g.,
quantifying specific transcripts or proteins by methods that
would be sensitive to cell permeabilization, e.g., high-content
sequencing analysis).
Of the commercially available cell-permeable DNA dyes, 4=,6=diamidino-2-phenylindole (DAPI), Draq5 {1,5-bis[2-(methylamino)
ethyl]amino-4,8-dihydroxyanthracene-9,10-dione}, Hoescht
(bis-benzimides), and Vybrant DyeCycle dyes have characteristics that include sufficient affinity and permeability to facilitate supravital staining and cytometry-based assessment of cell
cycle phase distribution. DAPI (21) and the Hoescht dyes were
the first dyes used in living cells because of their cell permeability characteristics and their specificity for DNA. DAPI and
Hoescht dyes are excited in the UV range (350 –390 nm) and
emit fluorescence in the blue range. They bind strongly to the
minor groove of DNA, have fairly broad emission spectra,
increase fluorescence after binding to DNA, and favor AT-rich
sequences. Two Hoescht stains (33258 and 33342) are used for
DNA labeling and cell cycle analysis (61, 62). Hoescht stains
are also substrates for the ATP-binding cassette transporter that
is highly expressed on stem and progenitor cells, a property
that is exploited for the use of these dyes to identify and purify
stem cell populations from adult tissues (111). In general,
chronic exposure to DNA-intercalating dyes can have a variety
of negative effects on normal cell function, including adverse
effects on cellular proliferation (100).
The spectral characteristics of DAPI and the Hoechst stains
require that they be excited by UV light, thus limiting their use
in long-term live cell imaging studies. Prolonged exposure of
living tissue to UV light causes photobleaching and DNA
damage responses that alter cell cycle progression and induce
apoptosis. Moreover, establishing stable levels of nuclear dye
retention for long-term imaging can be difficult and often
requires repetitive loading. Continuous imaging of Hoechststained living cells can result in rapid cell death (70). Moreover, although UV light sources are common on microscopes,
this is not the case for flow cytometers, and retrofitting a flow
cytometer with UV light and detectors can be expensive. In
light of these limitations, the DyeCycle Violet stain was first
developed and was based on Hoechst 33342 structural characteristics. It exhibits a right-shifted excitation and a broader
emission spectra than does Hoechst 33342 (Table 1) (111).
Similarly to the Hoechst stain, DyeCycle Violet significantly
increases fluorescence after binding to DNA. Other DyeCycle
stains that emit in the green, orange, and red wavelengths that
are excitable with 488-nm lasers have been developed. These
AJP-Cell Physiol • doi:10.1152/ajpcell.00006.2013 • www.ajpcell.org
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sensitive method for detecting transcription factor DNA interactions. Cell lysates, either whole cell or nuclear, are incubated
with double-stranded labeled oligonucleotides (e.g., 32P or
biotin end-labeled) that contain DNA binding sequences for
specific transcription factors. Samples are then resolved by
nondenaturing polyacrylamide gel electrophoresis. Transcription factor activity is reflected in a higher-molecular-weight
shift in the migration of labeled DNA oligonucleotides. Specificity of binding is confirmed by several controls, including
competition with unlabeled nucleotides, and, if possible, “supershifting” by preincubation with antibodies raised against
specific transcription factors (7). Electric mobility shift assays
have been used to detect E2F family activity during the G1-S
transition (68) and for Foxo family members that are transiently activated during G2/M transition (e.g., FoxM1) (56, 84).
Luciferase reporters to assess for cell cycle regulation.
Luciferase assays have gained popularity due to their relative
simplicity and applicability to high-throughput assays. Such
assays are conducted by fusing a cell cycle-responsive promoter upstream of a destabilized firefly luciferase gene (Photinus pyralis) in an expression vector. Cells are transfected and
synchronized at a specific phase of the cell cycle and then
released, treated, lysed, and assessed for luminescence using a
luminometer. Signals must then be normalized. A variety of
methods for doing so include the use of untransfected cells,
cells transfected with an empty luciferase vector, and cells
transfected with a constitutive promoter not regulated by the
cell cycle [e.g., EF1␣ (122)].
The dual-luciferase normalization method consists of
cotransfecting cells with a control plasmid containing a ubiquitous promoter (e.g., phosphoglycerate kinase 1) upstream of
Renilla luciferase (Renilla reniformis). The firefly luciferase
signal is measured and quenched, and Renilla activity is then
assessed in the same lysates. This is the most efficient method
to achieve signal normalization; efficient application of this
method is facilitated by use of a luminometer with built-in
injectors. This promoter-based system can also be adapted to
assess for cell cycle activity in living cells by switching the
reporter gene to a fluorescent gene (e.g., GFP, as discussed
below) or by using cell-permeable luciferase gene product
substrates (32). Luciferase reporter studies are possible in live
cells while controlled tissue culture growth conditions are
maintained, but they require specialized laboratory equipment
(e.g., LumiCycle from Actimetrix) and culture media.
Cell cycle-responsive promoters have been successfully utilized with the luciferase system to assess for cell cycle progression. The ⬃100-bp (⫺89/⫹11) proximal promoter of the
human cyclin A gene is repressed in G1 phase and induced
upon S phase entry [as shown in NIH 3T3 cells (97)]. The
1.5-kb proximal promoter of the E2f1 gene is repressed in G0
and early G1 and is induced during the G1-S transition in
REF-52 cells (47) or U2OS cells (32). Six forkhead transcription factor binding sites can drive luciferase expression during
G2/M in U20S cells (32, 60).
Review
C933
APPROACHES FOR CELL CYCLE ASSESSMENT
Live Cell Sensor
Design
Ki67p-GFP
1.5kb Ki67p
activity is attenuated, and thus it could be used to indirectly
quantify the number of cells that have exited the cell cycle
specifically by determining the number of Ki67p-GFP cells
that are GFP⫺.
The labeling of proliferating cells with Ki67p-GFP may aid
in the study of the cell proliferation in the nervous system.
Recent evidence suggests that neurons generated in the same
time window preferentially form connections with one another
(22). To assess the ability of Ki67p-GFP to label proliferating
subpopulations of highly interconnected neurons in vivo, we
electroporated the developing cerebral cortex of an embryonic
14.5-day embryo in utero with Ki67p-GFP and a constitutively
active chicken ␤-actin promoter (90) driving RFP (CAGGpRFP) reporter constructs. Fluorescence was assessed after 24 h
(Fig. 5). Ki67p-GFP labeled fewer cells than did the CAGGpRFP construct, suggesting that low numbers of cells are actively proliferating. Moreover, in the subventricular zone, cell
bodies labeled positive for Ki67p-GFP colocalized as pairs,
suggesting that they are in the late stages of the M phase or
have recently undergone cytokinesis (Fig. 5).
In rapidly cycling immortalized cells that are constantly
reentering the G1-S transition, a limitation of using GFP as a
reporter protein for cell cycle studies is its long half-life (⬃26
h) (19). Thus, we attempted to use the Ki67p to drive expression of a destabilized GFP (66), but this resulted in very dim
fluorescence that was inadequate for labeling proliferating cells
(unpublished observations). We anticipate that using Ki67p to
drive expression of the cell cycle-destabilized proteins (discussed below) will increase the utility of the Ki67p in immortalized cell types and in vivo.
Reporters based on cell cycle-dependent changes in protein
localization and/or stability. GFP fusions with core cell cycle
proteins such as polo-like kinase 1 (3), Ccnb1 (42), yeast cell
cycle proteins (110), P27 (123), and histone 2B (50) helped
shed new light on the dynamic macromolecular changes that
occur during cell cycle progression. However, overexpression
of cell cycle-related proteins that retain enzyme activity can
perturb the stoichiometric balances that are required for proper
cell cycle regulation and are thus not optimal for serving as live
cell cycle sensors (16).
Reference
GFP
Labeled Phase
Labeled Cellular
Compartment
G1, S, G2, M
G1-c; S-c; G2-c; M-all
[105,112]
All
G0-n; G1-n; S-n,c; G2-c; M-all
late S, G2, M
S-c; G2-n; M-all
S, G2, M
S-n; G2-n; M-all
[122]
Stealth G1/S
UbCp
Stealth G2/M
Ccnb1p
Ccnb1 (1/170)
GFP
[105,114]
CCNB1-GFP
Pgkp
Ccnb1 (1/170)
GFP
[35,55]
G0/G1 Fucci
CCAGp
mKO2
hCdt1 (30/120)
[93]
G0,G1, early S
G0-n; G1-n; early S-n
G2/M Fucci
CCAGp
mAG
hGem (1/110)
[93]
S, G2, M
S-n; G2-n; M-all
G0/G1 Fucci2
G2/M Fucci2
R26p
R26p
GFP 131aa Helb PSLD
PNpA
mCherry hCdt1 (30/120)
[1,92]
G0,G1, early S
G0-n; G1-n; early S-n
PNpA
mVenus hGem (1/110)
[1,92]
S, G2, M
S-n; G2-n; M-all
Fig. 4. Genetic reporters for cell cycle assessment in live cells and tissue; nucleus (n), cytoplasm (c), promoter (p), chicken ␤-actin promoter (CCAG), endogenous
rosa 26 promoter (R26), and floxed PGK-neomycin resistance gene-poly A cassette (PNpA), green fluorescent protein (GFP).
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dyes have a proprietary structure (Invitrogen) but are reported
to be useful for sorting cells by flow cytometry such that the
sorted cells can be cultured after incubation with the dye.
DRAQ5 is a deep red fluorescing bisalkylaminoanthraquinone (Ex 646, Em 681) capable of reporting three-dimensional
nuclear structure and location in live cells, even in the presence
of green fluorescent protein (GFP) (70, 101). Anthraquinones
are synthetic DNA-binding agents that are structurally related
to DNA-intercalating anthracycline antibiotics. The far-red
emission spectrum of DRAQ5 facilitates its use in conjunction
with commonly used green- and red-emitting fluorophores and
fluorescent proteins. It stains cellular nuclei and penetrates into
tissue on the second-to-minute timescale. It is cytotoxic, with
10% survival of cells exposed at 10 nM for 24 h, and thus it is
not recommended for live cell sorting, where long-term survival is desired (70, 101).
Carboxyfluorescein diacetate succinimidyl ester can be used
to estimate the number of times that cells have divided (67).
Carboxyfluorescein diacetate succinimidyl ester passively diffuses into cells and is cleaved by intracellular esterases to
produce a highly fluorescent carboxyfluorescein succinimidyl
ester. The succinimidyl ester reacts with intracellular amines,
forming fluorescent conjugates that are retained in daughter
cells after cellular division. Each daughter cell exhibits approximately one-half of the fluorescent signal as the parent cell.
Cell cycle-responsive promoters for sensing cell cycle entry
in live cells. As mentioned above, cell cycle-dependent promoters have enabled the application of the luciferase system to
assess cell cycle regulation. These promoters can also be used
to drive expression of fluorescent genes to assess regulation of
the cell cycle in living cells. Our laboratory generated, cloned,
and characterized the 1.5-kb proximal promoter of the human
Ki67 gene and used it to drive enhanced GFP expression
(Ki67p-GFP; Fig. 4) (122), hypothesizing that it would be able
to distinguish between quiescent and cycling cells.
The Ki67p-GFP reporter has been used to distinguish between subpopulations of cells that are arrested in G0 from those
that are actively transitioning through it (27). We reported that
Ki67p-GFP colocalizes with endogenous Ki67 and can be used
to label proliferating cells in three-dimensional tissue cultures
(122). It has the added advantage that in quiescent cells its
Review
C934
APPROACHES FOR CELL CYCLE ASSESSMENT
Seminal work from Stubbs and Thomas (105) and Thomas
(112) exploited cell cycle-dependent changes in protein localization to develop two genetically encoded “stealth” live cell
cycle sensors that label cells during the G1-S and G2-M
transitions. As noted above, changes in protein localization
(e.g., cytoplasmic/nuclear transport) play a major role in cell
cycle regulation (86). Stubbs and Thomas (105) developed a
stealth G1/S reporter construct by using the ubiquitin C constitutive promoter (90) to drive expression of a cDNA encoding
a GFP fused to the 131-amino acid phosphorylation-dependent
subcellular localization control domain of the helicase B gene
(Helb) (Fig. 4). The HLB gene is a DNA-dependent ATPase
that catalyzes the unwinding of DNA during DNA replication
and repair during S phase entry (34, 109). The G1/S reporter
labels nuclei of cells in G0/G1 and the early S phase. During S
phase transition, the fusion protein is shuttled out of the
nucleus and is only in the cytoplasm during G2/M (Fig. 4).
The stealth G2/M sensor harnesses the 949-bp proximal
promoter of Ccnb1 (44) to drive expression of a cDNA encoding Ccnb1 amino acids 1–170 fused to the NH2 terminus of
enhanced GFP (Fig. 4). This Ccnb1 proximal promoter is
active in late S phase and drives expression of the Ccnb1enhanced GFP fusion construct. The expressed reporter translocates from the cytoplasm to the nucleus at prophase and is
degraded by the encoded Ccnb1 destruction box during mitosis
(16, 105, 113). Because the Ccnb1 destruction box contains an
APC/C E3 ubiquitination target domain of CCNB1, the reporter protein is stable during the S phase until metaphase. The
reporter is expressed and degraded in concert with endogenous
Ccnb1 but does not compete for binding to CDK since it lacks
the COOH-terminal sequences that comprise the Ccnb1/
CDK interaction domains. Extensive clonal selection and
characterization was carried out to identify a clonal U2OS
line that exhibits the appropriate signal intensity and normal
cell cycle kinetics (114).
Robust application of the G1/S reporter that relies on changes in
cellular localization requires an environmentally controlled
confocal microscope and image-processing software to automate quantification of changes in reporter localization. Moreover, it is not known whether these constructs can be used in
other commonly used model organisms (e.g., Drosophila or
zebrafish). However, because advanced imaging is used to
assess cell cycle reporters that are dependent on changes in
protein localization, one could envision quantifying changes in
other cellular structures (e.g., nuclear envelop breakdown) or
labeled proteins in the context of cells in specific phases of the
cell cycle.
Recently, Klochendler et al. (55) generated a transgenic
mouse line containing a Ccnb1-GFP fusion construct described
above (16) under control of the mouse phosphoglycerate kinase
1 promoter (Fig. 4). The expression construct had been developed and characterized previously in cultured cells (35). The
transgenic mouse line was used to isolate and profile the
expression of dividing and nondividing adult and juvenile liver
cells. Interestingly ⬃10% of CCNB1/GFP-positive cells do not
traverse the S phase (as measured by BrdU incorporation),
suggesting that a subpopulation of cells have altered APC/C
activity (55, 76).
A similar approach of fusing cell cycle protein destabilization domains to fluorescent reporters was employed by SakaueSawano et al. (93) to develop the fluorescent ubiquitinationbased cell cycle indicator (Fucci). This live cell sensor consists
of a duel transgenic system that labels nuclei of cells in the
G0/G1 and early S phases with the fast-folding monomeric
Kusabira Orange (mKO2) fluorescent protein (51) and cells in
late S/G2 and early M phases with a monomeric version of the
green fluorescent protein Azami Green (mAg) (52). Cells in the
S phase are labeled by both proteins (Fig. 4).
The mKO2 reporter is a fusion of amino acids 30 –120 of the
human Cdt1 protein to the carboxyl terminus of mKO2
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Fig. 5. Ki67p-GFP expression in the developing nervous system. The developing cerebral cortex of an embryonic mouse was electroporated in utero with a
plasmid containing the 1.5-kb proximal promoter of the human Ki67 gene upstream of GFP (Ki67p-GFP) (122) and a ubiquitously expressed reporter,
CAGGp-RFP (90), at embryonic day 14.5 and fixed after 24 h. The ventricular (VZ) and subventricular zones (SVZ) contain neural precursors. These precursors
produce postmitotic neurons that will migrate through the intermediate zone (IZ) to the cortical plate (CP) via the radial glial fibers projecting to the pial surface.
Arrows indicate presumptive postmitotic neurons (red arrows) and mitotically active precursors (yellow arrows).
Review
APPROACHES FOR CELL CYCLE ASSESSMENT
difficult-to-transfect and nondividing cell types. Generation of
replication-deficient lenti-viral particles is straightforward
when a suitable packaging cell line (e.g., HEK-293FT) and
packaging vector plasmids are available. Moreover, isolation
of Fucci- and Fucci2-expressing cells from Fucci transgenic
mice circumvents the need for gene transfer into primary
isolated cell types.
Summary and Future Perspectives
Cell cycle progression is a fundamental process in all living
organisms. Classically, this process has been studied using
immortalized cell lines and/or approaches that require tissue
fixation, thereby limiting our understanding of important features of cell cycle progression, including its heterogenous
nature. In vivo, the cell cycle is dynamically and tightly
regulated by multiple mechanisms and signaling pathways that
receive inputs that vary depending on cell type and cellular
niche.
This review highlights a number of classic and emerging
techniques for cell cycle assessment. These emerging techniques will likely shed light on new genes and proteins that can
be targeted to block or induce cell cycle progression in specific
cell populations for experimental studies and therapeutic purposes. For example, the cell cycle pathway is a primary target
in cancer therapy. However, tumor cell heterogeneity contributes to cancer dormancy, a process that is poorly understood.
Cancer dormancy occurs when residual disease is present but
the patient remains asymptomatic. This process is common in
many cancers [e.g., 20 – 45% of patients with breast or prostate
cancer will relapse years or decades later (23, 53, 85)].
The transition of a cancer cell into cellular dormancy (cell
cycle arrest or cellular quiescence) is one of several proposed
mechanisms that cause cancer dormancy (2). Live cell cycle
sensors that enable single-cell resolution of cell cycle kinetics
in vivo over extended times (perhaps even on the order of
months to years) are likely to increase our understanding of this
important but poorly understood process and will hopefully aid
in the discovery of new therapeutics in this area.
Conversely, the process of inducing cell cycle progression in
other therapeutic settings (e.g., during cell therapy for regenerative medicine) is also a promising concept, assuming this
process can be tightly controlled. For example, cardiac disorders are a leading cause of death and disability worldwide, in
part as a consequence of the limited capacity for adult cardiac
myocytes (CMs) to reenter the cell cycle and proliferate.
Cardiac ischemia, in particular as a consequence of myocardial
infarction, can destroy CMs and, in addition, other cell types
essential for a normal functioning myocardium (e.g., endothelial and smooth muscle cells in the vasculature).
Although significant progress has been made in promoting
the differentiation of embryonic and adult stem cells into
terminally committed lineages (e.g., CMs), far less emphasis
has been placed on targeting pathways to increase the proliferative expansion of progenitor cells for therapeutic applications. Conceptually, such cells could be of greater therapeutic
benefit if they could be transplanted in sufficient quantities, by
controlling their proliferative expansion in vitro or in vivo, and
then guided to differentiate into multiple cell types in the target
organ. We anticipate that these and other newly developed cell
cycle reporter systems will enable studies in these areas.
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[mKO2-hCdt1(30 –120)]. Amino acids 30 –120 contain a Cy
motif that is targeted by SKP2 E3 ligase, a ligase that ubiquitinates a variety of cell cycle proteins during S/G2, targeting
them for proteolysis (76, 108). As mentioned above, Cdt1 is a
ubiquitin ligase that is involved in origin licensing and the
formation of the prereplication complex during DNA replication (17, 120).
The late S and G2/M Fucci reporter fuses residues 1–110 of
the human geminin (Gem) protein to the carboxyl terminus of
mAG [mAG-hGem(1–110)]. Gem inhibits DNA replication
during the late S and G2 phases by binding to and inhibiting
Cdt1 activity to prevent incorporation of minichromosome
maintenance proteins into the prereplication complex (120). It
is degraded during the metaphase-anaphase transition (73). The
Gem domain encoded within residues 1–110 is ubiquitinated
by APC/C during late M/G1 phase, targeting it for degradation
in a manner similar to that described above for CCNB1. This
activity results in nuclear accumulation of mAG-hGem(1–110)
during the late S and G2/M phases (Fig. 4) (93). Domains from
the zebrafish orthologs of Gem and Cdt1 were used to generate
a zebrafish Fucci system that has been used to generate exceptionally detailed movies of cell cycle progression in developing
zebrafish embryos (106). The system has also been adapted to
Drosophila (75) and Ciona (80).
CAG promoter-driven Fucci cassettes (78) were used to
develop transgenic mice (93). Double-Fucci transgenic mice
were generated in which every somatic cell nucleus in the
developing embryo exhibits either red or green fluorescence.
Fucci2 mice have been developed recently using mCherryhCdt(30/120) and mVenus-hGem(1/110) fluorescent chimeras that provide better color contrast compared with the
first-generation Fucci reporters (Fig. 4) (1). Fucci2 transgenes have also been targeted to the Rosa26 locus, thereby
reducing in vivo variability caused by the CAG promoter
and transgenic insertion events. Moreover, the targeting
construct was developed to enable cell type-specific Fucci2
expression by Cre-mediated loxP recombination (1).
The Fucci system has greatly facilitated studies of spatial
and temporal cell cycle regulation in vitro and in vivo. Fucciexpressing cells have been used to examine the complex
relationships between cell cycle kinetics and fundamental cellular processes such as signaling (93), differentiation (14), cell
size (103), protein compartmentation (96), and heterogeneity
of tumor cell responses to anti-cancer agents (92). In the latter
study, individual cells were tracked with the reporter system
within a population of unsynchronized cells, and heterogeneous effects of drug doses were quantified at the single-cell
level.
The Fucci system has proven to be an invaluable tool for in
vivo applications such as tumor and xenograft models, where
its activity as a readout of tumor cell proliferation can be
tracked in intact animals using whole animal imaging (20, 93).
It has also been used for advanced imaging of cell proliferation
in the neural system in clarified intact brain tissue (37) and the
proliferation of astrocyte populations in postnatal brains (30),
among others.
As with any genetically encoded sensors, limitations of gene
transfer can be significant. Many cell types, both primary and
immortalized, can be difficult to transfect. In light of this, both
the Ki67p-GFP and Fucci reporter constructs have been cloned
into lenti-viral vectors, thereby facilitating their delivery into
C935
Review
C936
APPROACHES FOR CELL CYCLE ASSESSMENT
ACKNOWLEDGMENTS
We thank the University of California San Diego Neuroscience Microscopy
Facility (P30 NS047101) for the use of their imaging equipment.
GRANTS
D. S. Bortone was supported by an National Institutes of Health (NIH)
Research Service Award Grant (National Institute of Neurological Disorders
and Stroke: 1-F32-NS-076185-01A1). A. C. Zambon was supported by an
American Heart Association Grant (10SDG2630130) and NIH Grants 1-U54HL-108460, 8-UL1-TR-000100, and P01-HL-098053. L. Henderson was supported by National Institute of General Medical Sciences Grant 2-R25-GM083275.
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the authors.
L.H., C.L., and A.C.Z. contributed to conception and design of the research;
L.H., D.S.B., and A.C.Z. analyzed the data; L.H., D.S.B., C.L., and A.C.Z.
prepared the figures; L.H. and A.C.Z. drafted the manuscript; L.H., D.S.B.,
C.L., and A.C.Z. edited and revised the manuscript; L.H., D.S.B., C.L., and
A.C.Z. approved the final version of the manuscript; D.S.B. and A.C.Z.
performed the experiments; D.S.B. and A.C.Z. interpreted the results of the
experiments.
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