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2.4: The Cell Cycle and Changes in DNA Content - Biology

2.4: The Cell Cycle and Changes in DNA Content - Biology


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Four stages of a typical cell cycle

The life cycle of eukaryotic cells can generally be divided into four stages and a typical cell cycle is shown in Figure (PageIndex{13}). This lag period is called Gap 1 (G1), and ends with the onset of the DNA synthesis (S) phase, during which each chromosome is replicated. Following replication, there may be another lag, called Gap 2 (G2), before mitosis (M). Cells undergoing meiosis do not usually have a G2 phase. Interphase is as term used to include those phases of the cell cycle excluding mitosis and meiosis. Some cells never leave G1 phase, and are said to enter a permanent, non-dividing stage called G0. Understanding the control of the cell cycle is an active area of research, particularly because of the relationship between cell division and cancer.

Measures of DNA content and chromosome content

The amount of DNA within a cell changes following each of the following events: fertilization, DNA synthesis, mitosis, and meiosis (Fig 2.14). We use “c” to represent the DNA content in a cell, and “n” to represent the number of complete sets of chromosomes. In a gamete (i.e. sperm or egg), the amount of DNA is 1c, and the number of chromosomes is 1n. Upon fertilization, both the DNA content and the number of chromosomes doubles to 2c and 2n, respectively. Following DNA replication, the DNA content doubles again to 4c, but each pair of sister chromatids is still counted as a single chromosome (a replicated chromosome), so the number of chromosomes remains unchanged at 2n. If the cell undergoes mitosis, each daughter cell will return to 2c and 2n, because it will receive half of the DNA, and one of each pair of sister chromatids. In contrast, the 4 cells that come from meiosis of a 2n, 4c cell are each 1c and 1n, since each pair of sister chromatids, and each pair of homologous chromosomes, divides during meiosis.


2.4: The Cell Cycle and Changes in DNA Content - Biology

A comprehensive review of assays for the cell cycle, and the results from a Labome survey of formal publications.

The cell cycle is the process by which eukaryotic cells duplicate and divide. The cell cycle consists of two specific and distinct phases: interphase, consisting of G1 (Gap 1), S (synthesis), and G2 (Gap 2), and the mitotic phase M (mitosis) (Figure 1). During interphase, the cell grows (G1), accumulates the energy necessary for duplication, replicates cellular DNA (S), and prepares to divide (G2) [2]. At this point, the cell enters the M phase, which is divided into two tightly regulated stages: mitosis and cytokinesis. During mitosis, a parent cell's chromosomes are divided between two sister cells. In cytokinesis, the division of the cytoplasm occurs, leading to the formation of two distinct daughter cells. Each phase of the cell cycle is tightly regulated, and checkpoints exist to detect potential DNA damage and allow it to be repaired before a cell divides. If the damage cannot be repaired, a cell becomes targeted for apoptosis. Cells can also reversibly stop dividing and temporarily enter a quiescent or senescent state G0. The first checkpoint is at the end of G1, making the decision if a cell should enter S phase and divide, delay division, or enter G0. The second checkpoint, at the end of G2, triggers mitosis if a cell has all the necessary components.

Several methods to assess the cell cycle are discussed below. However, it is important to remember that these methods are not mutually exclusive, and for the best and most reliable data multiple dyes and/or analytes can be combined in a single experiment or multiple assays used.

The most common method for assessing the cell cycle is to use flow cytometry to measure cellular DNA content. During this process, a fluorescent dye that binds to DNA is incubated with a single cell suspension of permeabilized or fixed cells. Since the dye binds to DNA stoichiometrically, the amount of fluorescent signal is directly proportional to the amount of DNA. Because of the alterations that occur during the cell cycle, analysis of DNA content allows discrimination between G1, S, G2 and M phases. The simple protocol for cellular analysis is outlined in Figure 2. Briefly, cells are fixed and permeabilized to allow the dye(s) to enter the cell and to prevent them from being exported out. Staining with the DNA binding dye then occurs after cells have been treated with RNase to ensure only DNA is being measured. Several datasets, including forward scatter vs. side scatter, pulse area vs. pulse width, and cell count vs. propidium iodide, are collected to ensure only single cells are measured. Examples of these traces are shown in Figure 3.

There are several different dyes that can be used in these assays, including propidium iodide (PI) [3, 4], 7-amino actinomycin-D (7-AAD), Hoechst 33342 and 33258, and 4’6’-diamidino-2-phenylindole (DAPI). For example, Chopra S et al labeled mouse bone marrow–derived dendritic cells and paw single cell suspensions with 0.5 μg/ml DAPI from Thermo Fisher during flow cytometry on a BD LSR II instrument and cell sorting with a BD Aria II SORP cell sorter [5]. Zhang H et al combined mitosis-specific anti-pMPM2 antibody ( 05-368 from MilliporeSigma) staining with DAPI to obtain prometaphase cells through FACS [6]. However, most FACS machines commonly used contain only single argon-ion lasers, and as such dyes requiring UV activation such as DAPI and Hoescht 33342 are less frequently used. A derivative of Hoechst dye, SiR-Hoechst, has excitation at 640 nm, and thus may find widespread use [7]. Hoechst 33258 has also been used to image polycomb bodies [8]. Rhodes JDP et al estimated the proportion of mitotic cells through antibody staining of serine 10 phosphorylated histone H3 in FACS [8]. Phosphorylated histone H3 staining has also been used in immunohistochemistry to identify mitotic cells in mouse [9, 10] and killifish [11].

When carrying out these analyses, it is important to recognize that simple single stained FACS analysis using 7-AAD or PI is unable to distinguish between cells in G1 or G2 from those in very early or very late S phase, and similarly those in G2 or M. It is therefore sometimes necessary to combine these dyes with a proliferative marker such as BrdU [3]. For example, Calvanese V et al combined 7AAD and BrdU-PE staining in flow cytometry to assess the cell cycle stages of cultured hematopoietic stem or progenitor cells [12]. This requires additional steps at the beginning of the study, where live cells still in culture are incubated with BrdU for a period of approximately 30 minutes, before incubation with anti-BrdU and fluorescence-conjugated secondary antibodies. Cells are then assayed as described above.

There are a number of important considerations when carrying out analysis of cell cycle FACS data. The forward scatter/side scatter plots are an integral part of the analysis and should not be overlooked, since this is how single cells are identified. If doublets (when the DNA content of two cells in G1 are recorded as a single G2/M event) are allowed in the analysis, it can lead to over-representation of G2/M. Cellular aggregates and flow rates below 1000 cells/second should also be avoided to allow a low sample pressure differential to be used, which leads to an optimal coefficient of variance (CV). Finally, reference samples containing normal diploid DNA should be included as an additional control.

The Nicoletti assay [13] is a modified form of cell cycle FACS analysis that concurrently allows apoptosis to be assessed by measuring cells with low intact DNA content, and high fragmented DNA content [14] (the pre-G1 peak). The Nicoletti method is very similar to that described above, with the exception that a hypotonic buffer (such as HFS buffer containing sodium citrate and Triton X-100, or a hypotonic fluorochrome solution) is used to permeabilize the cells. Apoptotic cells stain weaker in these assays due to the activation of cellular nucleases and the diffusion of low molecular weight DNA out of the cell. Fixing and permeabilizing cells stimulates the release of oligo- and mononucleosomes. The use of a hypotonic buffer facilitates the loss of fragmented DNA, resulting in a shift of the pre-G1 peak.

The images in Figure 4 demonstrate healthy cells (top), cells in which a sub-population is beginning to undergo apoptosis (middle), and a population of cells with extensive apoptosis (bottom). However when using the Nicoletti assay, care must be taken to discriminate apoptotic nuclei from cell debris, and to ensure that DNA shearing does not occur during the fixing and staining processes.

The cyclins are key regulatory components of the cell cycle machinery. The cyclin family comprises the classical cyclins, cyclin-dependent kinases [15, 16] (CDKs) and Cdk inhibitors (CKIs). Although there is much redundancy between the individual cyclins and CDKs [17], the activity and expression of the individual proteins fluctuate during each distinct phase of the cell cycle, playing an important regulatory role. Although this is a complex and highly regulated process, in general cyclins can be divided into sub-groups governed by the phase of the cell cycle they regulate, summarized in Figure 5. For example, Cyclin D1 is required for the passage of cells from G0 to G1. Once expressed, it forms a complex with Cdk4, which activates retinoblastoma protein, leading to the upregulation of Cyclin E. Cyclin E, in combination with cyclin A, then interacts with Cdk2 to promote G1/S transition. In contrast, cyclins B1 and B2 are expressed during M phase where they interact with Cdk1 to form part of the MPF (M phase/maturation promoting factor), an assembly that regulates a cascade of processes leading to mitotic spindle assembly and ultimately cell division. The expression of each human cyclin and their interaction with Cdks are summarized in table 1.

Cyclin Peak phase expressed Cdk binding partners Top three suppliers
DG1Cdk4, Ckd6CCND1:Invitrogen MA1-39546 (335), Cell Signaling Technology 2978 (93), Santa Cruz Biotechnology sc-20044 (38)
EG1/SCdk2CCNE1:Santa Cruz Biotechnology sc-247 (41), Cell Signaling Technology 4129 (36), Invitrogen MA5-14336 (22)
AS/G2Cdk1, Cdk2CCNA1:Cell Signaling Technology 4656 (23), Santa Cruz Biotechnology sc-271682 (5), R&D Systems MAB7046 (2)
BMCdk1CCNB1:Santa Cruz Biotechnology sc-245 (87), Cell Signaling Technology 4135 (32), Invitrogen MA5-14319 (23)

These distinct expression patterns can therefore be exploited during cell cycle analysis. The total levels and/or phosphorylation status of individual cyclins can be easily and rapidly measured using specific antibodies by immunoblotting [3]. In addition, specific ELISA kits are available for individual cyclin family components, allowing for a more quantitative assessment of expression. Finally, fluorescently conjugated antibodies can be used in immunohisto- or immunocyto-chemical approaches, or in flow cytometry. Combining cyclin staining with FACS methods examining DNA content provides a powerful and quantitative tool to analyze the cell cycle accurately [3].

Tetraploid cells are associated with the formation of malignancy and often possess the stem-cell characteristics. Thus, with relevance to cancer biology [18, 19] and regenerative tissue homeostasis [20], it is conceivable that the analysis of tetraploid cells would be of importance. The tetraploid G1 cells and diploid G2/M cells are difficult to detect as they possess the same ploidy that is 4C DNA content.

FUCCI (Fluorescence ubiquitination-based cell cycle indicator) system is a technology that utilizes the cell cycle phase-specific expression of proteins and their degradation by the ubiquitin-proteasome degradation system [21, 22]. The technology analyzes the living cells in a spatio-temporal manner using dual-color protein-fluorescent chimeras. Moreover, it enables to overcome the problem of isolating the cells in different phases, which is otherwise difficult to differentiate only with the DNA-based stains such Hoechst. It is composed of two proteins - Cdt1 (Cdc10 dependent transcript 1) and Geminin. Both proteins are used in their truncated forms (hCdt1 and hGeminin) and are conjugated to two different fluorescent proteins. They express alternately in the two different cell cycle phases. Cdt1 is a conserved replication factor required for licensing the chromosome for DNA synthesis. Cdt1 is expressed throughout the G1 phase and is ubiquitinated by the ubiquitin ligase complex SCFSkp2 during S and G2/M phases followed by its degradation by the proteasome. In contrast, geminin inhibits the licensing activity of Cdt1 by interfering with the binding of licensing factors to the replication origin during the S phase. It is present during S/G2/M phases. At the end of M phase and throughout the G1 phase, geminin is ubiquitinated by the E3 ligase complex APCcdh1 and degraded by the proteasome [21].

Supplier Kit References
Caltag MedsystemsFUCCI
MBL Life SciencesFUCCI [23]
Takara Pharmaceuticals FUCCI vectors
ThermoFisher Scientific Premo™ FUCCI Cell Cycle Sensor (BacMam 2.0) [24, 25]

Figure 6 depicts the scatterplot representing the live cells expressing different fluorochromes implying their diverse cell cycle phases. Depending on the probe selection, the two chimeras emit different fluorescence. Several probes are available commercially. One of the examples is stated below along with a diagram. Fucci-G1 Red is a fusion protein of a fragment of human Cdt1 (amino acids 30-120) with the red fluorescent mCherry-RFP, that detects the cells in G1 phase. Fucci-S/G2/M Green is a fusion protein of a fragment of human geminin (amino acids 1-110) with the green fluorescent protein mAG1 (monomeric Azami-Green1) that visualizes S, G2 and M phases. Thus, the G1 tetraploids emit red fluorescence and G2/M tetraploids emit green fluorescence. By employing this technology, it is also possible to distinguish between the mononucleated diploid cells from the binucleated cells.

Table 2 lists some of the commercial suppliers of FUCCI kits. The FUCCI sensors from Thermo Fischer Scientific, for example, were used to analyze G1 phase in mouse stem cells and evaluate mechanisms of UV-mediated damage [24] and investigate the awakening and proliferation of dormant metastatic cells by neutrophil extracellular networks [25]. M Barnat et al obtained pCAG-Geminin-GFP and pCAGCdt1-mKO2 from A. Miyawaki, RIKEN Brain Science Institute, Japan [10].

One of the limitation of the FUCCI system is that these systems requires the expression of multiple reporter constructs intracellularly and reduces the chance to image other targets spectrally. This problem has been overcome by modification of this system. Zerjatke et al developed fluorescently tagged endogenous proliferating cell nuclear antigen (PCNA) as an all-in-one cell cycle reporter. This reporter with PCNA-mRuby alters in brightness and localization in different phases. Consequently, it provides a readout of the cell cycle phase including quiescence and quantitative dynamics of individual fate determinants of cell cycle regulation [26].

Another limitation is that FUCCI system and its variants allow visualizing whether cells are within one of the proliferative phases (S, G2, or M) of the cell cycle, they do not report simultaneous visualization of the three phases cells in real time. Bajar et al developed a robust method that enables simultaneous imaging of the all four phases. They established an intensiometric reporter for the S/G2 transition and further engineered a far-red fluorescent protein, mMaroon1, to track the process of mitotic chromatin condensation. They designed a new version called Fucci4 by combining the new reporters with the FUCCI system and incorporating four orthogonal fluorescent indicators that enable to capture all cell cycle phases in the living cells [27]. Fucci4 has diverse applications in development, physiology, and cancer. Fucci4 allows 1) how diverse changes at the molecular, genetic and extracellular signaling level alter the cell cycle, 2) molecular mechanisms regulating specific phase transitions and 3) screening for drugs that affect a particular cell cycle phase or cell cycle distribution.

There are several applications of FUCCI system in various branches of biology and medicine. For studying development biology, Sugiyama et al generated transgenic Zebrafish lines expressing the non-mammalian FUCCI counterparts [28]. They were employed to study the spatio-temporal regulation of cell-cycle progression during major morphogenetic events/processes (gastrulation, metamorphosis, involution, invagination and branching) [21, 29]. Zielke et al engineered Drosophila-specific FUCCI system (Fly-FUCCI) that involves tissue-specific expression of the FUCCI probes [30]. This allows one to distinguish G1, S, and G2 phases of interphase. This serves as a valuable tool for visualizing cell-cycle activity during development, tissue homeostasis, and neoplastic growth.

The FUCCI system can be used in tumors for in vivo cell cycle profiling by stably transfecting cell lines with FUCCI reporters and development of the xenograft tumors [31]. Nico Battich et al correlated the synthesis and degradation rates of mRNAs along the cell cycle indicated by the FUCCI system [32].

Sawano et al re-engineered the Cdt1-based sensor from the original Fucci system to respond to S phase-specific CUL4Ddb1-mediated ubiquitination alone or in combination with SCFSkp2-mediated ubiquitylation. This system is known as Fucci(CA) and it demarcates interphase with boundaries between G1, S, and G2. The applications of Fucci(CA) included tracking the transient G1 phase of rapidly dividing mouse embryonic stem cells and identifying UV-irradiation damage in S phase [24].

Several new reagents for cell cycle analysis, such as chromobodies and Cycletest reagent, have recently been developed. Chromobodies are fusion proteins, which contain fluoresceins linked to the antigen binding domain of heavy chain antibodies. These reagents are used to detect the expression of various intracellular proteins within the cellular compartments and dynamic changes of their distribution during different phases of the cell cycle. Furthermore, chromobodies can be applied for the detection of both cytoskeletal and nuclear proteins. With regard to cytoskeletal target proteins, changes in vimentin expression have been analyzed by specific chromobodies in a study that generated vimentin knock-out cell line [33]. As to the analysis of nuclear proteins by chromobodies, Proliferating Cell Nuclear Antigen (PCNA), which plays a crucial part in the replication process in the nucleus, was detected by the red fluorescent protein-bound chromobody [34]. Also, an advanced modification of PCNA detection by chromobodies, which was based on 4D quantitative analysis of the PCNA expression and distribution during the replication phase, has recently been reported [35].

Moreover, chromobody assays can be combined with other techniques. For example, a combination of chromobody-based analysis of the cell cycle with the Chto Tox-Glo cytotoxicity method has been described. In that study, the visualization of PCNA expression in subcellular compartments by chromobodies was followed by the evaluation of protease activity in vitro [36]. With regard to the applications of chromobodies for in vivo research, the chromobody-based method has been originally applied to studies in zebrafish [37]. In addition, Wegner et al have analyzed the expression of actin in the mouse brain using anti-actin chromobodies labelled with fluorescent protein mNeptune2. [38].

suppliersreagents/kits methods sample references
Becton Dickinson7-amino-actinomycin D [39]
BioLegendCytoPhase violetflow cytometry [40]
MilliporeSigma7-amino-actinomycin D [41]
MilliporeSigmaHoechst 33258 [42]
MilliporeSigmaHoechst 33342 [39]
MilliporeSigmapropidium iodide
Thermo Fisher7-amino-actinomycin D [43]
Thermo FisherHoechst 33258 [44]
Thermo FisherHoechst 33342
Thermo Fisherpropidium iodide

In addition, Cycletest PLUS reagent kit produced by BD Biosciences is recommended for cell cycle analysis. This method includes the elimination of the cell membrane and cytoskeleton with a detergent and trypsin, respectively, followed by the digestion of RNA and stabilization of the chromatin. The kit contains propidium iodide (PI), which binds to the extracted nuclei. The stained nuclei are analyzed by flow cytometry to measure the binding of PI to DNA. This reagent was successfully used in several recent studies. For instance, Cycletest has been applied to evaluate the anti-tumor effects of thymoquinone in human breast tumor MCF-7 cells [45] and the effects of α-solanine in colorectal tumor cells [46].

This section is provided by Labome to help guide researchers to identify most suited cell cycle analysis assay kits. Labome surveys formal publications. Table 3 lists the major suppliers for reagents/kits used in the cell-based assays and their numbers of publications in the Labome survey. The review article on cell proliferation lists the survey results on BrDU.

This article is derived from an earlier version of an article authored by Dr. Laura Cobb "Cell-Based Assays: the Cell Cycle, Cell Proliferation and Cell Death", written in February 2013. Dr. Samayita Das contributed to the section on the FUCCI system in September 2019.


Bliss Biology

The cell cycle refers to the life cycle of a Eukaryotic cell. At any given time body cells are in one of two different phases of their lives:

Interphase: makes up 90% of the life of a cell. This is the phase where the cell is growing, carrying out normal functions, and duplicating its genetic material. There are three stages of interphase:

G1 Phase – The cell is growing.

S Phase – Synthesis phase. Here, the DNA is replicating

G2 Phase – In this phase, the cell is double-checking to make sure there were no errors made when copying the DNA

Mitosis: A very short part of the cell cycle. In mitosis, the cell is splitting in two to create two brand-new, identical daughter cells. There are several stages that make up mitosis.

Prophase – the cell PREPARES to divide. The chromatin DNA condenses into “X”-shaped chromosomes. The spindle of the centrioles begins to form.

Metaphase – Chromosomes line up in the MIDDLE of the cell cytoplasm. The spindle of the centrioles attaches to the centromere of the chromosomes.

Anaphase – The chromosomes pull APART, with each sister chromatid pulling to either side of the cell.

Telophase – With the chromosomes now located on the far opposite side of the cell, the nuclear membrane begins to reform around them. The spindle fibers of the centrioles disappear and the cell membrane begins to pinch together to become TWO separate cells.

Cytokinesis – This stage of the cell cycle occurs after mitosis ends. In this phase, the cell membrane cleaves, or CUTS in half and two new daughter cells are formed. These daughter cells are identical to the parent cell. In plant cells, a new section of cell wall called the Cell Plate is built to separate the two new daughter cells.


Release from cell cycle arrest with Cdk4/6 inhibitors generates highly synchronized cell cycle progression in human cell culture

Each approach used to synchronize cell cycle progression of human cell lines presents a unique set of challenges. Induction synchrony with agents that transiently block progression through key cell cycle stages are popular, but change stoichiometries of cell cycle regulators, invoke compensatory changes in growth rate and, for DNA replication inhibitors, damage DNA. The production, replacement or manipulation of a target molecule must be exceptionally rapid if the interpretation of phenotypes in the cycle under study is to remain independent of impacts upon progression through the preceding cycle. We show how these challenges are avoided by exploiting the ability of the Cdk4/6 inhibitors, palbociclib, ribociclib and abemaciclib to arrest cell cycle progression at the natural control point for cell cycle commitment: the restriction point. After previous work found no change in the coupling of growth and division during recovery from CDK4/6 inhibition, we find high degrees of synchrony in cell cycle progression. Although we validate CDK4/6 induction synchronization with hTERT-RPE-1, A549, THP1 and H1299, it is effective in other lines and avoids the DNA damage that accompanies synchronization by thymidine block/release. Competence to return to cycle after 72 h arrest enables out of cycle target induction/manipulation, without impacting upon preceding cycles.

1. Background

Synchronized progression through the cell division cycle throughout a population supports the ability to extrapolate the biochemical and functional attributes of the synchronized bulk population back to infer behaviour in an individual cell [1,2]. Many approaches are popular. Bulk levels of DNA or cell cycle markers support fractionation of live, or fixed, cell populations into pools enriched for discrete cell cycle stages [3,4]. Although yields are low, selection synchronization based upon size, or mitotic shake off, are highly effective approaches to isolate cells in one cycle phase from a large population of asynchronous cells with minimal impact upon the proteome [5–9]. However, the ease of induction synchrony makes it the most widely applied approach.

Induction synchrony exploits the ability of transient exposure to a particular context to accumulate cells at a discrete cell cycle stage, before removal of the context simultaneously releases all cells in the population, to progress synchronously through subsequent phases of the cell division cycle [1]. In yeasts, transient ablation of cell cycle regulators through reversible conditional mutations and the addition of mating pheromones predominate [10,11]. Although the advent of analogue-sensitive versions of cell cycle kinases has introduced analogous chemical genetic approaches into human tissue culture studies [12–15], induction synchrony via serum starvation [16], contact inhibition [17] or activation of either the DNA replication, or spindle assembly, checkpoints [7,18] remain the most widely used. However, it is important to note that each form of arrest superimposes specific changes in the transcriptome and proteome upon the core cell cycle arrest signature [19–21]. These specific routes to cell cycle exit are reflected in different routes of return to the synchronized cycles [22]. Thus, the apparent reflection of normal cell cycle progression achieved with selection synchronization is yet to be matched by current approaches to induction synchronization [4,9,21].

The discovery that transient treatment with thymidine synchronized mitotic progression [18] led to protocols that sharpened the degree of synchrony by imposing a second thymidine block before releasing cells into the cycle of study [23–26]. This ‘double thymidine block’ remains one of the most popular choices however, the power of this approach, its reliance upon the DNA replication checkpoint to arrest S phase progression, with stalled DNA replication forks, comes at a cost. Although the cell cycle arrest is robust in many lines, the stalled forks are prone to collapse over the extended arrest and subsequent attempts at repair introduce damage and chromosomal rearrangements [27–31]. There are also reports of understandable impacts upon RNA biology and proteome during the extended S phase arrest [21,32,33]. Thus, this popular approach can be of limited utility in the study of DNA replication and some transcriptional and chromatin-associated events.

When early cell cycle events are to be analysed, induction synchrony via release from a mitotic arrest in the previous cell cycle provides an attractive alternative. However, like other forms of induction synchrony that arrest within the cycle, prolonged cell cycle arrest will generate an imbalance in the many regulators, whose levels fluctuate with cell cycle progression, as a consequence of stage-dependent transcription and/or destruction [21,34,35]. Consequently, the next cycle may well be altered by excessive regulatory activities, or substrates, inherited from the preceding, arrested, cycle. Incisive studies by Ginzberg et al. [36] revealed how counter-measures to accommodate some imbalances promote adjustments in growth rates at two points in the cycle. Furthermore, prolonged mitotic arrest can initiate the atypical mitotic exit termed mitotic slippage [35,37], trigger apoptotic pathways [38] and/or leave a memory of the mitotic arrest that modifies cell cycle progression in the next cycle and beyond [39–42].

Thus, while highly informative for some questions, data obtained through traditional induction synchrony approaches, that rely upon arrest within the cycle, have to be interpreted with caution. They must be consolidated with complementary data from alternative approaches to reveal the commonalities that exclude the artefacts incurred in each distinct approach to synchronization.

A further challenge in synchronizing cell cycle progression throughout a population arises when there is a need to assess the impact of protein depletion, induction or replacement. It is critical to ensure that the destruction, induction or activation of a mutant variant starts after the synchronizing procedure is complete. If not, then the phenotype can be a legacy arising from perturbation of progression through the previous cycle, rather than a direct impact upon the cycle being studied. Advances in degron and PROTAC (PRoteolysis TArgeting Chimera) technologies may overcome many of these challenges [43–45]. However, even with many induction synchronization approaches, the switch from one version of a protein to another must be exceptionally rapid and complete if perturbation of the preceding cycle is to be avoided.

Inspired by the power of pheromone induction synchronization at G1 phase of yeast cell cycles [11,46], we explored the utility of induction synchrony with CDK4/6 inhibitors palbociclib, ribociclib and abemaciclib. These inhibitors arrest cell cycle progression of mammalian tissue culture cells at the restriction point in G1 phase, prior to commitment to the cell cycle [47,48]. Synchronization by induction from the natural pause point in the cycle has a number of appealing attributes. Firstly, the cell cycle programme is yet to be set in motion. Secondly, extended arrest via Cdk4/6 inhibition does not invoke compensatory changes in cell cycle or growth controls rather it simply adjusts cell size control [36]. Finally, palbociclib-imposed cell cycle arrest has less impact upon the transcriptome than the cell type-specific changes seen upon synchronization via contact inhibition and serum deprivation [19,20].

Cdk4 and Cdk6 kinases determine commitment to the cell cycle of many cells [48]. They partner Cyclin D and the Kip family members p21 and p27 to generate active trimeric kinase complexes that phosphorylate the C terminus of the retinoblastoma (Rb) family protein [49–54]. This mono-phosphorylation supports further phosphorylation of Rb by Cdk1/Cdk2–Cyclin E and Cdk1/Cdk2–Cyclin A complexes [13]. Hypo-phosphorylated Rb binds tightly to the transcription factors of the E2F family, to block the transcription of genes required for cell cycle commitment. Rb dilution and hyper-phosphorylation relieves this inhibition, to promote transcription of cell cycle genes, including Emi1, Cyclin E and Cyclin A [55]. Induction of these Cyclins rapidly boosts Rb phosphorylation by Cyclin E and Cyclin A Cdk complexes [56]. Emi1, Cyclin E and Cyclin A complexes then seal commitment to the cycle by inhibiting the anaphase-promoting complex/cyclosome (APC/C) activating component Cdh1, thereby stabilizing APC/C Cdh1 targets, including Cyclin A [57–61].

The key role played by Cdk4–Cyclin D and Cdk6–Cyclin D in presiding over cellular proliferation, and the contrast between the ability of mice to survive genetic ablation of Cdk4, Cdk6 and Cyclin D and the addiction of cancer lines to these kinases, prompted the development of clinically successful Cdk4/6 inhibitors [62–66]. These inhibitors bind to the inactive Cdk4–Cyclin D and Cdk6–Cyclin D dimers, rather than the active trimeric complexes, yet they impose a very efficient cell cycle arrest [54]. This counterintuitive impact has been proposed to stem from the sequestration of the inactive dimers, or monomeric kinases by the drugs. In this model, this sequestration releases p21 and p27 to elevate concentrations of these Cdk2 inhibitors, to a level where they block the ability of the Cdk2–Cyclin A and Cdk2–Cyclin E complexes to promote the feedback loops and S phase events that drive commitment to the cycle [48,54,67]. Each drug has a specific ‘off target’ profile. Ribociclib's impressive specificity is almost matched by palbociclib, while abemaciclib shows significant off-target impacts, with notable activities towards Cdk1, Cdk2, Cdk7 and Cdk9 complexes, that can induce an arrest in G2 alongside G1 [68]. Paradoxically, this off-target impact may account for abemaciclib's greater efficacy in some clinical settings [68,69].

We describe how Cdk4/6 induction synchrony generates highly synchronous progression through the cell cycles of a number of lines, without the marked appearance of a marker of DNA damage, foci of staining with antibodies that recognize serine 139 of the histone γ-H2AX when it is phosphorylated (γ-H2AX), that accompanies thymidine induction synchronization. The ability to return to cycle after 72 h arrest in G1 with palbociclib will support a broad range of manipulations in the arrested, non-cycling state [70]. Thus, any impacts upon progression through the cycle of study will not be a secondary consequence of perturbation of the preceding cell cycle.

2. Methods

2.1. Cell lines and cell culture

The cell lines used in this study are listed in electronic supplementary material, table S1. Upon receipt, lines were expanded and frozen into aliquots of 1 × 10 6 cells that were expanded by at least two cycles of continual passage prior to each experiment. Unless otherwise stated, hTERT-RPE-1, A549, GP2d, LoVo, U2OS, DLD-1, HCT 116, HeLa, A172, HMBC, HEK293, HEK293T and H157 were maintained in high glucose Dulbecco's modified eagles medium (DMEM: D6546, Sigma Aldrich) supplemented with 10% FBS (Hyclone, South American Origin, SV30160.03, GE Healthcare) 2 mM GlutaMAX (Gibco, 35050061) and 1000 U ml −1 Penicillin/Streptomycin (Gibco, 15140122). THP1, H1299, H1975, H1437, H2052, H2452, H520, H1915, NB-19, BEAS2B and H1395 were maintained in Roswell Park Memorial Institute 1640 medium (RPMI) supplemented with 2 mM GlutaMAX (61870044, Gibco), 10% FBS (Hyclone, South American Origin, SV30160.03, GE Healthcare) and 1000 U ml −1 Penicillin/Streptomycin (Gibco, 15140122). MCF 10A was maintained in Dulbecco's modified eagles medium/Nutrient Mixture F-12 Ham (D6421, Sigma Aldrich), supplemented with 10% Horse Serum (16050130, ThermoFisher), 10 µg ml −1 Insulin (I9278, Sigma Aldrich), 0.5 µg ml −1 Hydrocortisone (H0888, Sigma Aldrich), 20 ng ml –1 human epidermal growth factor (hEGF: E9644, Sigma Aldrich) and 1000 U ml −1 Penicillin/Streptomycin (Gibco, 15140122). A stock solution of 1 mg ml −1 hydrocortisone was prepared in ethanol and a 100 µg ml −1 stock solution of hEGF was prepared in water. Both were stored in aliquots at −20°C. Cells were maintained at a confluency of less than 70% and populations were not passaged more than 10 times before any experiment. In figure 2e, where hTERT-RPE-1 cells are grown in RPMI, cells were originally grown from the frozen stock in DMEM, passaged twice in RPMI, then plated in RPMI at the start of the experiment.

2.2. Drug treatment

Palbociclib (PD-0332991), abemaciclib (LY2835219) and ribociclib (LEE011) were purchased from Selleck (catalogue numbers S1116, S5716 and S7440, respectively), while thymidine (T1895) and nocodazole (M1404) were from Sigma-Aldrich. Palbociclib, abemaciclib, ribociclib and nocodazole were dissolved in DMSO to generate stock solutions of 10 mM in each case. Thymidine was dissolved in water to make a stock solution of 100 mM. All stocks were stored in aliquots at −20°C.

For adherent cell lines, cells were released from the substrate by treatment with trypsin (15400054, Gibco), pelleted by centrifugation at 300g for 5 min before resuspension in growth media. Cells were counted using a TC20 automated cell counter (BioRad) and 2.5 × 10 5 cells were plated in a 10 cm dish (353003, Falcon) with 10 ml media (plating density = 4.4 × 10 3 per cm 2 ). For the THP1 suspension line, cells were pelleted by centrifugation at 300g for 5 min, resuspended in growth media and counted, before 1 × 10 6 cells were seeded in 10 ml media in a 10 cm dish. Cells were then incubated for 6 or 12 h (see figure legends) before drug was added.

2.3. Flow cytometry

For cell cycle analysis, cells were washed once with phosphate-buffered saline (PBS) before releasing from the substrate with trypsin, and washed in PBS before fixation in ice-cold 70% Ethanol and freezing at −20°C for at least 18 h up to a maximum of two weeks. Fixed cells were pelleted, washed three times in PBS at room temperature before 50 µl 100 µg ml −1 RNase (NB-03-0161, Generon) was added to the final pellet, followed by 500 µl 50 µg ml −1 propidium iodide (P4170, Sigma-Aldrich) dissolved in PBS. Samples were analysed within a window of between 30 min and 6 h after addition of propidium iodide. Data were acquired on a BD LSR II flow cytometer (BD Biosciences) using FACSDiva™ software (BD Biosciences) and analysed with FlowJo software (BD Biosciences). A total of 1 × 10 4 cells were counted for each sample.

S phase status was monitored by measuring 5-ethynyl-2'-deoxyuridine (EdU) incorporation with the Click-iT™ Plus EdU Alexa Fluor 488 Flow Cytometry Assay Kit (ThermoFisher, C10632) according to the manufacturer's instructions. To monitor cumulative DNA content between the time of release and the point of fixation in figure 9c, 1 µM EdU was added at the time of release from palbociclib. Cells were trypsinized and washed with 3 ml of 1% BSA in PBS, then incubated with 100 µl of Click-iT™ fixative for 15 min, pelleted, washed with 3 ml of 1% BSA in PBS and left at 4°C overnight. Cells were resuspended in 100 µl of 1× Click-iT™ permeabilization and wash reagent and incubated with 500 µl of Click-iT™ Plus reaction cocktail for 30 min, before washing once with 3 ml of 1× Click-iT™ permeabilization and wash reagent and resuspended in 500 µl of 1× Click-iT™ permeabilization and wash reagent. FxCycle Violet stain (F10347, Invitrogen) was added to a final concentration of 1 µg ml −1 . Samples were analysed on a BD LSR II flow cytometer (BD Biosciences) using FACSDiva software (BD Biosciences) and analysed with FlowJo software (BD Biosciences). A total of 10 4 cells were counted per sample. For quantification, the peak of 2 N cells was used to gate an assessment of 2 N DNA content, as the overlap between S phase and G2/M made it challenging to categorically assign cell cycle status on the basis of 4 N DNA content alone. Thus, throughout the manuscript, we monitor cell cycle progression as a reduction in 2 N DNA content.

2.4. Immunofluorescence

Cells were grown on 13 mm coverslips (No. 1.5, VWR, 631-0150P) in 10 cm dishes (353003, Falcon) using the appropriate growth conditions. For EdU staining, the Click-iT™ EdU Cell Proliferation Kit for Imaging, Alexa Fluor™ 594 dye (Invitrogen, C10339) was used. One hour prior to harvesting cells, 10 µM EdU was added to the growth media. Cells were washed in PBS before fixing for 20 min in 2% paraformaldehyde in PBS, before three washes in PBS + 0.1% Tween and storage in PBS + 0.1% Tween at 4°C overnight. Permeabilization in PBS + 0.5% Triton-X-100 was followed by three washes in PBS + 0.1% Tween. The Click-iT™ reaction for EdU detection was performed according to the manufacturer's instructions. Cells were washed three times in 5% Bovine Serum Albumin (BSA ThermoFisher Scientific, 11423164) in PBS then incubated with primary antibody to H2AX (Ser139), clone JBW301 (Millipore Cat no. 05-636, RRID:AB_309864) at a dilution of 1 in 500 for 1 h before three washes in PBS + 5% BSA and incubation with a 1 in 500 dilution of goat anti-mouse IgG Alexafluor 488 antibody (Thermo Fisher Scientific Cat no. A32723, RRID:AB_2633275), and 2 mg ml −1 4′,6-diamidino-2-phenylindole (DAPI, ThermoFisher Scientific, 11916621) for 1 h. After a further three washes in PBS + 0.1% Tween, coverslips were mounted by inversion onto 2 µl of Vectashield Antifade Mounting Medium (Vector Labs H1000). Slides were analysed using an Axioskop2 (Zeiss, Inc.) microscope.

2.5. Protein analysis

To collect hTERT-RPE1 protein samples, culture medium was removed, by aspiration, from a 10 cm plate, before two washes in PBS and addition of 400 µl TruPAGE LDS sample buffer (Sigma-Aldrich, PCG-3009) containing complete protease inhibitor cocktail (Roche, 11697498001), PhoSTOP (Sigma-Aldrich, 4906837001) and DTT Sampler Reducer (Sigma-Aldrich, PGC-3005) to the plate. Cells were scraped from plate for transfer into a 1.5 ml microfuge tube and snap frozen in liquid nitrogen for storage at −80°C. For THP1, cells were centrifuged at 300g for 3 min and the pellet resuspended in sample buffer as above, frozen in liquid nitrogen and stored at −80°C. Samples were heated at 70°C for 10 min before loading on a 10 cm, 10% precast TruPAGE gel (Sigma-Aldrich PCG2009) with TruPAGE SDS running buffer (Sigma Aldrich PCG3001) and transferred to PVDF membrane (BioRad, 1 620 177) by wet transfer with TruPAGE transfer buffer (Sigma-Aldrich PCG3011). Membranes were blocked in 5% milk and incubated in 2% BSA with primary antibodies to GAPDH (Cell Signaling Technology Cat no. 13084, RRID:AB_2713924), Eg5 (Sigma Aldrich Cat no. SAB4501650, RRID:AB_10747045), Wee1 (Cell Signaling Technologies Cat no. 13084, RRID:AB_2713924) or pHH3 ser10 (custom antibody to peptide ARTKQTARKS*TGGKAPRKQLASK: Eurogentec) overnight at 4°C. After washing, membranes were incubated for 1 h at room temperature, with the appropriate secondary antibody (Anti-mouse IgG phosphatase-conjugated antibody (Sigma-Aldrich Cat no. A3688, RRID:AB_258106), or Anti-rabbit IgG phosphatase-conjugated antibody (Sigma-Aldrich Cat# A3687, RRID:AB_258103)), washed and developed with chromogenic 5-Bromo-4-chloro-3-indolyl phosphate (BCIP, Sigma-Aldrich, B6149).

3. Results

3.1. Efficient synchronization of hTERT-RPE1 cell cycle progression with transient palbociclib treatment

The telomerase immortalized hTERT-RPE1 cell line is widely used for cell cycle and mitotic studies, yet is refractory to double thymidine block induction synchronization. We, therefore, chose this popular line to assess the efficiency of G1 arrest and release by transient exposure to palbociclib.

Cells were grown to 1.5 × 10 4 cells cm −2 in serum supplemented (10%) Dulbecco's modified eagles medium (DMEM), before release from the substrate with trypsin and plating at a density of 4.4 × 10 3 cells per cm 2 . Six hours later, palbociclib was added in a range of concentrations from 50 nM to 1 µM to two identical populations. Twenty-four hours later, one population was fixed, while the palbociclib containing medium for the other was replaced with pre-warmed medium containing 330 nM nocodazole, before this sample was fixed a further 24 h later. DNA content assessment by fluorescence-activated cell sorting (FACS) analysis of propidium iodide (PI)-stained samples revealed a tight arrest at the 24 h time point with 2 N DNA content at all palbociclib concentrations of 100 nM and higher (figure 1a,b). Cells readily released into a 4 N arrest after being arrested in G1 for 24 h with 100 nM and 200 nM palbociclib however, release was less efficient when the arrest was imposed by 500 nM and 1 µM palbociclib (figure 1b) as the 2 N DNA content remained high 24 h after nocodazole addition.

Figure 1. Palbociclib induction synchrony of hTERT-RPE1 cells. (a) hTERT-RPE1 cells were grown to 1.5 × 10 4 cells cm 2 in DMEM (+10% serum), trypsinized and plated into 10 cm dishes at 4.4 × 10 3 cm −2 . Six hours later, 150 nM palbociclib was added to the culture. After 24 h, cells were washed twice with pre-warmed medium before the addition of pre-warmed medium that contained 330 nM nocodazole before incubation for a further 24 h. Samples (one 10 cm dish per data point) were stained for propidium iodide FACS analysis at the following time points: just before palbociclib addition (U, untreated), at the switch from palcociclib to nocodazole medium (P) and 24 h after this switch to nocodazole (P + N). The strength of the nocodazole-induced spindle checkpoint arrest was revealed by the addition of nocodazole to an asynchronous population (U + N) for 24 h. The bimodal peak in the upper panel (U) shows 2 N (G1, left) DNA and 4 N (G2/M, right) DNA content of an asynchronous, untreated population. This experiment was repeated six times. (b) Cell populations were treated in the same way as (a) with the palbociclib concentration changed to the indicated value and one sample being left untreated. The average frequency of 2 N cells from at least three biological repeats is plotted for the palbociclib (grey bars) and nocodazole (blue bars) arrest points. Error bars show the limits of 1 s.d.. (c,d) hTERT-RPE1 cells were grown to around 1.5 × 10 4 cells cm −2 in DMEM (+10% serum), trypsinized and plated into 10 cm dishes at 4.4 × 10 3 cm −2 . Twelve hours later, 150 nM palbociclib was added to the culture, before three washes in pre-warmed DMEM and sampling one dish every hour to generate the propidium iodide FACS profiles in (c) from which the plots of 2 N content shown in (d) were derived. The numbers next to the plots in c indicate time (hours) since release with U indicating an untreated control population. The 13–25 h plots (red (c): open squares (d)) were taken simultaneously alongside the 0–12 h population (grey (c): filled circles (d)). The synchronization shown in (c,d) has been performed six times and always reveals comparable results however, variations in the precise synchronization profiles in each experiment (as can be seen in subsequent figures in the manuscript) mean that it is not appropriate to merge them into a single dataset.

Monitoring DNA content at hourly intervals after release from 150 nM palbociclib arrest revealed a synchronous progression from G1 arrest with 2 N DNA, through S phase into G2/M phases with 4 N DNA content before a return to 2 N DNA at 20 h (figure 1c,d). As with all experiments presented herein, 24, or 48, h periods were covered by sampling parallel populations indicated by black and red lines/shading in the FACS plots and circles and squares in the graphs. For example, in the 0–13 h samples of a 24 h experiment, palbociclib was removed at the start of sampling, while the release had been done 13 h earlier for samples that were simultaneously collected for the 13–24 h samples. Consequently, many graphs we present have 13 h time points from each set.

As the second cycle after release is less likely to be influenced by the physiological challenges of induction synchrony, it is often desirable to monitor this second cycle, rather than the first cycle after release [1]. We, therefore, extended our assessments to monitor total DNA content over 48 h after release from 24 h treatment with 150 nM palbociclib. A notable degree of synchrony persisted in the second cycle with 2 N cells declining to constitute 46% of the population 30 h after release (figure 2a electronic supplementary material, figure S1). Thus, it will be possible to monitor some trends in biochemical behaviour associated with cell cycle progression in the second cycle after release.

Figure 2. Context and perdurance for palbociclib induction synchronization. For (a), hTERT-RPE1 cells were grown to around 1.5 × 10 4 cells cm −2 in DMEM (+10% serum), trypsinized and plated in 10 cm dishes at 4.4 × 10 3 cm −2 . Twelve hours later, 150 nM palbociclib was added. Twenty-four hours after palbociclib addition, cells were washed twice with medium that did not contain any palbociclib before incubation in pre-warmed DMEM (+10% serum) with sampling one dish every 2 h for propidium iodide FACS staining to generate the profiles in 12 h batches covering a 48 h release period of the same population of cells. Samples for the 0–12 (filled circles), 14–24 (open squares), 26–36 (filled circles) and 36–48 (open squares) were taken in parallel from subpopulations to which the palbociclib had been added at staggered intervals. This experiment was performed twice, with similar results in each iteration. (be) Plots derived from the same population of cells. For (b), hTERT-RPE1 cells were grown to 1.5 × 10 4 cells cm −2 in DMEM (+10% serum), trypsinized and plated in 10 cm dishes at 4.4 × 10 3 cm −2 . Twelve hours later, 150 nM palbociclib was added. Twenty-four hours after drug addition, the cells were washed twice with medium before incubation in pre-warmed DMEM (+10% serum) that lacked palbociclib and two batches of the same population of hTERT-RPE1 cells were followed, sampling one 10 cm dish for each data point. In one batch (open symbols), 150 nM palbociclib was re-added to the population 12 h after the initial release from palbociclib, while the other was left to transit the restriction point into the second cycle (red). For (c), the density at which cells from the same population used for the initial plating in (b and d) were plated at a fourfold higher density of 1.76 × 10 4 cells cm −2 in 10 cm dishes. (d) The expansion of the same starting population used in (b) and (c) lacked one cycle of splitting so that the starting population that was synchronized in (d) had been grown to confluence (early contact inhibition) before plating 12 h before palbociclib addition. (e) Cells from the same vials used to seed the populations used in bd were grown in RPMI for two passages, alongside the cells used in bd, before the entire synchronization outlined in figure 1c was conducted in RPMI. For (be), samples were simultaneously taken from two batches of the same population: palbociclib was added to one at the start of a 12 h sampling period (circles), while it had been added to the other 12 h earlier (squares). For the FACS plots from which these 2 N DNA contents were derived, see electronic supplementary material, figures S1 and S2.

When studying events at the end of the cycle, the fastest cycling cells in the population will progress into the next cell cycle to initiate a second round of cell cycle events before the events in the first cycle are completely finished in some members of the population. This natural and subtle variation in the timing of cell cycle progression generates a merged dataset, in which information from the second cycles of some cells is superimposed upon that from the cells that are still approaching the end of their first cycle. In this way, the finer points of the kinetics of change in the first cycle can be obscured by overlapping events in the subsequent cycle. One simple solution to overcome this confusion would be to ensure that exit from the first cycle was blocked. This can be easily achieved by re-addition of palbociclib mid-way through the first cell cycle. We, therefore, asked whether such re-addition of palbociclib, after release from the first arrest, would compromise the progression through the observed cycle. Encouragingly, a second application of Cdk4/6 inhibitor 12 h after release had no impact upon progression through the cycle under study (figure 2b), compare the red (no re-addition) and open black symbols (palbociclib added back at 12 h: electronic supplementary material, figure S2a,b). Thus, palbociclib re-addition can be used to insulate observations from consequences of entrance into the next cycle, thereby giving greater insight into the kinetics of cell cycle events.

3.2. Growth conditions are key for optimal synchronization

The impact of metabolism, growth control and quiescence upon the control of commitment to the cell cycle prompted us to assess the impact of context upon the efficiency of palbociclib induction synchronization of hTERT-RPE1 cells. There was a notable reduction in the efficiency of synchronization of hTERT-RPE1 cells when the density of the population seeded onto plastic 6 h prior to palbociclib addition was increased fourfold to 1.76 × 10 4 cells cm −2 . At this density, the proportion of the population with a 2 N DNA content only declined to 38% rather than the dip to 16% in the identical population plated at 4.4 × 10 3 cm −2 (figure 2c electronic supplementary material, figure S2c). The efficiency of both arrest and release was also compromised when the identical population used in figure 2b,c had been grown to confluence to induce ‘contact inhibition’ before splitting to generate the populations that were arrested for the synchronization (declined to only 23% 2 N, figure 2d electronic supplementary material, figure S2d). Finally, in simultaneous studies of cells from the same initial population that had been passaged twice in Roswell Park Memorial Institute 1640 Medium (RPMI) before synchronization in this medium, fewer cells arrested cell cycle progression after 24 h in Palbociclib (81% versus 95%) and the proportion of 2 N cells only dipped to 37% of the population in RPMI, rather than the decline to 16% in DMEM (figure 2b,e electronic supplementary material, figure S2a,e). Thus, culture conditions alter the efficiency of synchronization and, once initial studies indicate that a line is competent for synchronization, a variety of conditions should be assessed and care should be taken to ensure that cells remain within active proliferation in the expansion in the lead up to synchronization.

3.3. Oscillations in established cell cycle markers accompany progression through synchronized cycles

To assess the utility of the approach for biochemical assays, extracts from a palbociclib synchronized culture were probed with antibodies to monitor markers whose levels fluctuate as cells transit cycles synchronized by other means. Anticipated fluctuations in the levels of the kinesin 5 Eg5 and phosphorylation on serine 10 of histone H3 highlight the utility of this approach to monitoring biochemical changes throughout the population (figure 3).

Figure 3. Cdk4/6 induction synchrony can reveal transient cell cycle events. hTERT-RPE1 cells were grown to around 1.5 × 10 4 cells cm −2 in DMEM (+10% serum), trypsinized and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. Twelve hours later, 150 nM palbociclib was added. Twenty-four hours after palbociclib addition, the cells were washed twice in growth medium before the medium was replaced with pre-warmed DMEM (+10% serum) that did not contain any palbociclib. One 10 cm dish was taken for each sample every hour to generate the propidium iodide FACS profiles in 13 h batches (a) to gauge the fluctuations in 2 N DNA content in the population (b), while sampling to monitor the indicated markers by western blot every 2 h (c). The numbers next to the plots in (a) indicate hours since release with U indicating an untreated control population. Both the mitotic kinesin 5 motor protein Eg5 and phosphorylation of the serine of histone H3 at position 10 peak as cells return from the 4 N state to the 2 N state (mitosis and cell division). This plot shows one of the three repeat experiments, which revealed similar fluctuations in the same cell cycle markers.

3.4. Synchronization of multiple lines with palbociclib, ribociclib and abemaciclib

We next asked whether palbociclib induction synchronization would be similarly effective in other lines by assessing the efficiency of arrest and subsequent release into nocodazole of a further 24 cell lines following addition of a range of palbociclib concentrations for 24 h followed by replacement with 330 nM nocodazole medium and incubation for a further 24 h. These experiments were exploratory in nature and so were not done to the same degree of rigour as the studies of hTERT-RPE1 or the detailed A549, H1299 and THP-1 analyses described elsewhere in this manuscript. Specifically, for some lines, fewer than 10 000 cells were counted and only two biological repeats (each with two technical repeats) were conducted. These exploratory investigations revealed no appreciable impact of palbociblib upon cell cycle progression in 7 of the 24 lines (figure 4), partial responses in 9, with exploitable arrest and release profiles observed in 8: MCF10A, LoVo, H1975, NB19, DLD1 (figure 5), THP-1, H1299 and A549. Robust arrest release in the H1299, A549 and THP-1 was confirmed in more rigorous testing of at least three biological repeats, over 10 000 cell counts (figure 6).

Figure 4. Palbociclib induction synchronization screening: refractory cell lines. The indicated cell lines were grown to around 1.5 × 10 4 cells cm −2 in in the media specified in the methods, trypsinized and plated into 10 cm dishes at 4.4 × 10 3 cm −2 . Six hours later, the cells were treated with the indicated concentration of palbociclib or left untreated. Twenty-four hours after this, samples were stained for propidium iodide FACS analysis to gauge the proportion of the population that had a 2 N DNA content. One 10 cm dish was used for each dataset. Each plot represents the average of a minimum of four datasets (at least two biological repeats, each of which had at least two technical repeats). The error bars represent 1 × s.d.

Figure 5. Palbociclib induction synchronization screening: responsive cell lines. Each line was grown in the media specified in the methods to around 1.5 × 10 4 cells cm −2 trypsinized, and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. Six hours later, 0.2 or 1 µM palbociclib was added to one-third of the dishes for each cell population. A control sample was left untreated. Twenty-four hours after this, cells were either fixed for staining (grey bars) or washed with fresh medium twice before incubation in 330 nM nocodazole for 24 h after which these samples were fixed and processed for propidium iodide FACs analysis alongside the samples that had been fixed 24 h earlier (blue bars). The proportion of the population that had a 2 N DNA content was calculated from the FACs profiles and plotted in the panels. The samples from the first 24 h incubation are shown in grey, while the subsequent nocodazole-treated samples are shown in blue. Each plot represents the average of at least four datasets (at least two biological repeats, each of which had at least two technical repeats). The error bars represent 1 × s.d. These experimental tests were purely exploratory in nature with the goal of finding lines for more detailed analysis in the rest of the study. Thus, these data should not be used to rule out the amenability of Cdk4/6i synchronization in the cell lines that show a partial response. We have not tested different inhibitors, media, longer incubations (in case, the cell cycles of certain lines emulate that of THP-1 of exceeding 24 h), or whether the addition of an MAP kinase inhibitor, or Cdk6 PROTAC may sharpen the responses. See Discussion for more details.

Figure 6. Spectrum of responses of four different lines to three Cdk4/6 inhibitors. THP1 and H1299 were grown in RPMI (+10% serum) and hTERT-RPE1 and A549 were grown in DMEM (+10% serum). Each adherent line was grown to around 1.5 × 10 4 cells cm −2 before they were trypsinized and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. The suspension THP1 cells were grown to around 4 × 10 5 ml −1 , centrifuged at 300g for 3 min before plating at 1 × 10 5 ml −1 in 10 cm dishes. One 10 cm dish was plated for each condition. Six hours later, the indicated concentration of the indicated Cdk4/6 inhibitor was added to each population. Twenty-four hours after this, samples (an entire 10 cm dish) were either fixed or washed with fresh medium twice before incubation in media containing 330 nM nocodazole for 24 h after which these samples were fixed and processed for propidium iodide FACs analysis alongside the samples that had been fixed 24 h earlier (blue bars). The proportion of the population that had a 2 N DNA content was calculated from the FACs profiles and plotted in the panels. The samples from the first 24 h incubation are shown in grey while the subsequent nocodazole arrest profiles in blue. Each plot represents the average of at least three biological repeats. The error bars represent 1 × s.d.

As the different Cdk4/6 inhibitors exhibit distinct pharmacological responses [68], we compared induction synchronization profiles in four different lines to gauge the spectrum of responses and whether a reliance upon palbociclib alone as a means to judge competence for CDK4/6 inhibition induction synchrony was a valid approach, or whether other inhibitors may show even greater efficacy. A549, H1299, THP1 and hTERT-RPE1 populations were exposed to a range of palbociclib, ribociclib and abembaciclib concentrations, before the CDK4/6 drug containing medium was swapped for medium containing nocodazole 24 h into the arrest. These nocodazole-containing cohorts were then harvested 24 h after the swap into nocodazole (figure 6).

In keeping with the efficacy of palbociclib, these selected lines showed strong Cdk4/6 inhibitor induction synchrony with the two other inhibitors (figure 6). Palbociclib and ribociclib gave excellent and comparable arrest/release profiles across a broad range of concentrations. By contrast, the window of competence was much narrower for abemaciclib (figure 6). When the efficiency of G1 arrest is used to identify concentrations of drug that have comparable impact upon restriction point passage, the ability to exit the G1 arrest into the G2 block was markedly lower when the inhibition had been imposed by abemaciclib. For example, for hTERT-RPE1 cells, 200 nM abemaciclib is required to attain the same block in G1 as 100 nM palbociclib, yet there was great variation in the ability to release and an average of 57% of the population remained arrested with 2N, rather than the 15% that persists in the palbociclib-treated population (figures 1b and 6d). This inefficiency is a recurrent theme in all lines (figure 6a–c). Comparisons between palbociclib and ribociclib reveal subtle distinctions to suggest that, when extensive work is to be performed with a specific line, there would be merit in testing both inhibitors when fine tuning the dose–response profile.

The acute myeloid leukaemia-derived line THP-1 is an attractive line for cell cycle studies because cell retrieval via pelleting avoids the challenges of harvesting from adherence to a matrix. We, therefore, assessed both the robustness of synchrony in this line (figure 7a,c, d electronic supplementary material, figure S3) and the behaviour of signature cell cycle markers as a population transited a synchronized culture (figure 7a,c). As noted for hTERT-RPE1 cells in figure 3, markers oscillated with their characteristic periodicities, all be it at a slower rate, in reflection of the longer, 30 h first cell cycle of THP1 cells (figure 7b compare this profile with the 22 h first cycle of hTERT-RPE1 cells, figure 2a).

Figure 7. Utility of THP1 suspension cell line for cell cycle analysis. THP1 cells were grown to around 4 × 10 5 cells ml −1 in RPMI (+10% serum), isolated by mild centrifugation at 300g for 3 min, before resuspension in RPMI at a concentration of 1 × 10 5 cells ml −1 in 10 cm dishes. Twelve hours later, 150 nM palbociclib was added for 24 h before cells were isolated by centrifugation at 300g for 3 min and once more resuspended in RPMI. Samples (one 10 cm dish per sample) were taken every 2 h to generate the propidium iodide FACS profiles in 12 h batches (a) to gauge the fluctuations in 2 N DNA content in the population (c), while sampling to monitor the indicated markers by western blot every 2 h (b). The numbers next to the plots in (a) indicate hours since release with U indicating an untreated control population. This analysis of cell cycle samples by western blotting shown in ac was done three times. (d) Cells grown as in (a) with the exception that sampling of 12 h batches was extended over a 48 h release period of the same population of cells. For this batch of cells, samples were only processed for PI FACs analysis of DNA content to generate the plot of the frequency of 2 N cells show in the panel. Samples for the 0–12 (filled circles), 14–24 (open squares), 24–36 (filled circles) and 36–48 (open squares) were taken in parallel from subpopulations to which the palbociclib had been added at staggered intervals. The propidium iodide FACs plots from which the data in (d) are derived are in electronic supplementary material, figure S3. This analysis of a 48 h progression of THP1 was done once. Note that the time taken for THP1 cells to transit the first cell cycle, under these conditions, is 32 h.

3.5. Inhibitor cocktails match the efficacy of single-agent synchronization

Off-target inhibition is a concern for interpretation of phenotypes arising from chemical perturbations. With the goal of minimizing the off-target impacts of each individual inhibitor [68], we asked whether a cocktail of all three inhibitors would be as effective as single agent? We used a mix of one-third of the most effective concentration for each individual drug for each individual drug for each cell line (as indicated in the figure legend) to test the level of cell cycle arrest and release. Encouragingly, all lines gave robust arrest and efficient release (figure 8).

Figure 8. Cocktails support efficient arrest and release in four responsive lines. THP1 and H1299 were grown in RPMI (+10% serum). hTERT-RPE1 and A549 were grown in in DMEM (+10% serum). Adherent cell lines were grown to around 1.5 × 10 4 cells cm −2 before they were trypsinized and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. The suspension line THP1 was grown to around 4 × 10 5 ml −1 and centrifuged at 300g for 3 min before plating at 1 × 10 5 ml −1 in 10 cm dishes. Four 10 cm dishes were seeded for each cell line. Six hours later, two dishes for each line were left as the untreated control, while the indicated cocktail comprising 33 nM abemaciclib + 33 nM palbociclib + 300 nM ribociclib for THP1 and H1299, 33 nM abemaciclib + 33 nM palbociclib + 67 nM ribociclib for A549 and 33 nM abemaciclib + 33 nM palbociclib + 167 nM ribociclib for hTERT-RPE1 was added to the other. Twenty-four h later, one dish for each condition was fixed for staining, while the medium in the remaining dish was replaced with medium containing 330 nM nocodazole. Media exchange for A549, H1299 and hTERT-RPE1 was achieved by aspiration while the media switch for the THP1 was achieved by mild centrifugation at 300g for 3 min followed by resuspension in the nocodazole-containing medium. Twenty-four hours later, these three nocodazole samples were fixed and processed for propidium iodide FACs analysis alongside the samples that had been fixed 24 h earlier. The proportion of the population that had a 2 N DNA content was calculated from the propidium iodide FACs profiles and plotted in the panels. The inhibitor cocktail arrested samples from the first 24 h incubation are shown in grey while the subsequent nocodazole arrest profiles in blue. Samples treated with the inhibitor cocktail are indicated by C, while the untreated population by U. Each plot represents the average of at least three biological repeats. The error bars represent 1 × s.d.

3.6. γ-H2AX staining reveals minimal DNA damage in palbociclib induction synchrony

A major limitation of the widely used ‘double thymidine block’ approach is the accumulation of DNA damage during the early S phase arrest [28]. This damage likely arises from collapse of, and attempts to repair, the DNA replication forks that stalled because nucleotide provision was compromised [27]. The damage accrued is likely to account for the abnormal anaphase profiles in the divisions after release from thymidine block [29].

We, therefore, monitored the accumulation of a marker of DNA double-strand breaks, foci of phosphorylation of γ -H2AX at serine 139, to assess the level of damage during palbociclib arrest and the ensuing cycle after release. As these foci form naturally during S phase, when replication forks generate double-strand breaks, we counterstained cells to identify those undergoing DNA replication in the hour before sampling by adding 10 µM of the nucleotide 5-ethynyl-2'-deoxyuridine (EdU) 1 h before processing each sample. All cells that are stained with this 10 µM EdU pulse will have been actively replicating DNA in the hour before fixation. This enabled us to distinguish EdU-positive cells with γ-H2AX foci, in which we assume the foci are a consequence of DNA replication, from those with no EdU staining, in which we assume the foci are indicative of sites of repair of DNA damage. We counted a cell as positive for γ-H2AX foci when immunofluorescence staining revealed more than two foci in a nucleus. To consolidate the insight into the timing of S phase from the assessment of total DNA content throughout the population (figure 9a,b), we monitored progression through S phase by quantifying the cumulative incorporation of EdU into the DNA after the addition of the lower concentration of 1 µM EdU at the time of release to persist throughout the experiment and label of all DNA synthesized after the release. S phase was largely complete by 16 h after release from 150 nM palbocilib (figure 9c).

Figure 9. γ-H2AX foci associated with DNA replication in hTERT-RPE1 cells. hTERT-RPE1 cells were grown to around 1.5 × 10 4 cells cm −2 in DMEM (+10% serum), trypsinized and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. Three 10 cm dishes were plated for each timepoint, one for propidium iodide staining (a,b), one for cumulatative EdU incorporation and one with coverslips for immunofluorescence. Twelve hours later, 150 nM palbociclib was added. Twenty-four hours later, cells were washed twice before incubation in pre-warmed medium that did not contain any palbociclib. At this point, one-third of dishes were left without EdU addition, 1 µM EdU was added to one-third of dishes at the time of release from palbociclib arrest, to monitor the cumulative accumulation of this marker of DNA replication (i.e. the extent of S phase progression at any one given point). In the dishes containing coverslips, 10 µM EdU was added to each dish just 1 h before these samples were fixed, in order to identify cells that were highly likely to have been actively replicating at the time of fixation. The cells with no EdU treatment were stained with propidium iodide, to monitor DNA content (a) that was used to calculate the value for the proportion of the population with 2 N DNA content (b), alongside the proportion of the population that had incorporated the EdU at any one time point (c). The samples from the population that had been subjected to the 1 h 10 µM EdU pulses were counterstained with γ-H2AX antibodies (d, e). The numbers next to the plots in (a) indicate hours since release, with U indicating an untreated control population. (d) Plots show the frequency of cells with staining of more than two γ-H2AX foci (light fill), EdU (intermediate fill) or those cells positive for both markers (black). (e) Focuses on the γ-H2AX-positive cells. The plots show the following at the indicated time points after palbociclib removal: the proportion of the whole population that stain positive for γ-H2AX alone (black, i.e. damaged cells, unlikely to be in S phase at the time of fixation), or for both γ-H2AX and EdU (grey, cells in S phase at the time of fixation). For each time point, at least 200 cells were counted to score each characteristic as a proportion of the total counted. The two populations sampled in parallel after staggered palbociclib addition are differentiated by grey and red in (a), closed circles and open squares in (b,c). This experiment was repeated three times with the similar outcomes each time however, the finer kinetics of the synchrony plots differed and so it was not appropriate to combine the datasets.

Figure 9d shows plots from a population of hTERT-RPE1 cells synchronized by palbociclib arrest/release. The frequency of cells incorporating the EdU pulse (dark grey shading), those showing more than 2 γ-H2AX foci (light grey shading) and those showing staining with both (black) are indictated. For the plots in figure 9e, we only scored cells that contained γ-H2AX foci. The portion of cells in the population with foci, yet no EdU are shaded black. By contrast, cells S phase cells that stained positive for EdU incorporation are represented by grey shading (figure 9e).

These plots in figure 9e show that the vast majority of γ-H2AX-positive cells are those that are in the process of replicating their DNA. Very few cells in the population had the DNA damage marker, but no EdU staining. Furthermore, cells that have only just initiated S phase may not have incorporated detectable levels of the EdU nucleotide pulse, even though they will have γ-H2AX foci associated with replication forks. Consequently, the assessment by scoring a positive incorporation of the EdU pulse label will give a modest underestimate of S phase cells. Taking this into consideration, and the fact that many cells that incorporated the EdU pulse did not have any γ -H2AX foci (figure 9d), it would appear that minimal DNA damage accompanies palbociclib induction synchronization (figure 9d,e).

As our unpublished experiments confirmed the widely shared view that hTERT-RPE1 cells are refractory to synchronization with thymidine (data not shown), we used H1299 to directly compare the levels of DNA damage arising during palbociclib induction synchrony and that accumulating at the cell cycle arrest and release following transient addition of 2 mM thymidine to the same starting population (figure 10 electronic supplementary material, figure S4). Although the degree of synchrony achieved by palbociclib arrest release is not as high in H1299 as in hTERT-RPE1, the results were clear.

Figure 10. DNA damage at arrest and throughout release in thymidine but not palbociclib synchronized H1299 cells. H1299 cells were grown to 1.5 × 10 4 cells cm −2 in RPMI (+10% serum), trypsinized and plated at 4.4 × 10 3 cm −2 in 10 cm dishes. Twelve hours later, either 150 nM palbociclib (ac) or 2 mM thymidine (df) was added, as indicated. Two 10 cm dishes were used for each condition and each timepoint, one for FACS analysis and one containing coverslips for immunofluourescence. Twenty-four hours after inhibitor addition, cells were washed twice with pre-warmed medium before the addition of pre-warmed medium that did not contain any inhibitor. Samples (the contents of one entire 10 cm dish per sample) were removed for fixation at two hourly intervals. Ten micromolar EdU was added to each dish containing coverslips 1 h before fixation, alongside processing for combined propidium iodide staining to monitor DNA content by FACs (electronic supplementary material, figure S4) from which we derived the plots of 2 N DNA content shown in (a). Processing the EdU staining revealed the cells that were likely to be in S phase at the time of fixation, while the γ-H2AX staining revealed cells with double-strand breaks in their nuclear DNA that could arise as a consequence of DNA damage or active replication. (b,e) Plots of the frequency of cells with staining of more than two γ-H2AX foci (light fill), EdU (intermediate fill) or those cells positive for both markers (black). (c,f) focus only on the population of γ-H2AX-positive cells. Two categories are scored: those that show only a γ-H2AX signal and no EdU signal (black) and those positive for both γ-H2AX and EdU and so are highly likely to be in S phase at the time of fixation (light). For each time point, at least 200 cells were counted to score each characteristic as a proportion of the total counted. The two populations sampled in parallel after staggered palbociclib addition are differentiated by closed circles and open squares in (a,d). This experiment was repeated three times with the similar outcomes each time however, the finer kinetics of the synchrony plots differed and so it was not appropriate to combine the datasets.

Consistent with previous reports [28], and in stark contrast to the minimal levels of DNA damage with palbociclib induction synchronization (figure 10ac), synchronization by exposure to a single dose of thymidine led to persistent γ-H2AX-positive scores for many cells after release from the thymidine arrest point in early S phase (figure 10df). Importantly, there were foci in many cells that had not incorporated the 1 h EdU pulse label at the time of fixation (the hallmark of actively replicating cells) in cells a long time after the block had been released (figure 10e,f). These data suggest that some damage that accumulated at the arrest point persisted throughout the subsequent release, beyond the period of replication. The persistence of damage is consistent with previous reports of chromosomal aberrations during divisions synchronized by thymidine induction synchrony [29].

Although we cannot exclude forms of DNA damage that do not generate γ-H2AX foci, palbociclib induction synchronization of cell cycle progression does not appear to compromise genome integrity to the same degree as thymidine based synchronization.

3.7. Competence to release maintained over 72 h

Removal, induction or replacement of a molecule of interest gives great insight into its function. When the manipulation is done in synchronized cultures, it is imperative that the manipulation does not occur in the preceding cycle, otherwise some of the phenotype observed could be an indirect consequence of irrelevant damage accrued in the preceding cycle as cells approach the block point. While impossible to avoid in selection synchronization, it remains a major challenge in many forms of induction synchronization that rely upon arrest within the cycle. One major appeal of halting cell cycle progression at a point when one cycle is complete and the next is yet to start is that the impact of any molecular manipulation of the arrested population will generate a phenotype that is a direct consequence of this manipulation upon progression through the ensuing cycle. None of the consequences will be attributable to problems in completing the cycle leading up to the block from which the cells are released.

We, therefore, compared the competence to return to cycle after Cdk4/6 inhibition for 24, 48 and 72 h. Pilot experiments established that the ability to hold the arrest, or release after arrest, varied between cell lines and so we selected different palbociclib concentrations to rigorously quantitate the competence to release following protracted arrest (figure 11). The efficiency of synchronization was assessed by the addition of the indicated concentration of palbociclib to cells 6 h after they were split from a subconfluent population and incubation for the indicated times before sampling one half of the population for FACs analysis of DNA content (grey). The palbociclib containing growth media for the other half of the population was replaced with medium containing 330 nM nocodazole before this population was sampled for FACs analysis a further 24 h later (blue). The nocodazole incubation trapped the cells released from the G1 block in the next mitosis as a consequence of activation of the spindle assembly checkpoint (SAC) [71]. While the ability to maintain the G1 arrest declined to varying degrees in the different lines, with hTERT-RPE1 being the most proficient at maintaining arrest, all lines exited the arrest to reduce the number of 2 N cells after 24 h in nocodazole (figure 11).

Figure 11. Substantial recovery from extended palbociclib-imposed cell cycle arrest. THP1 and H1299 were grown in RPMI (+10% serum), and hTERT RPE1 and A549 were grown in DMEM (+10% serum). Adherent lines were grown to around 1.5 × 10 4 cells cm −2 , isolated by trypsin digestion and plated at 4.4 × 10 3 cells cm −2 in 10 cm dishes. The suspension cell line THP1 was grown to around 4 × 10 5 cells ml −1 , isolated by centrifugation at 300g for 3 min and plated at 1 × 10 5 ml −1 in 10 cm dishes. Eighteen 10 cm dishes were plated for each cell line. Six hours later, dishes were either treated with the indicated concentration of palbociclib or left untreated. At the indicated time intervals, one plate for each condition was processed for propidium iodide FACs analysis, while another was incubated in palbociclib-free medium containing 330 nM nocodazole for a further 24 h before it too was processed for propidium iodide FACs analysis. The proportion of the population that had a 2 N DNA content was calculated from the FACs profiles and plotted in the panels. The samples after the incubation for the time shown under the plot are shown in grey, while the paired sample of cells of this population that had been released from the palbociclib arrest by medium replacement with nocodazole medium before fixation 24 h later are shown in blue. Each plot represents the average of at least three biological repeats. The error bars represent 1 × s.d.

4. Discussion

Our examination of the potential of Cdk4/6 inhibition as a novel approach to induction synchrony reveals a highly effective approach to synchronization of cell cycle progression throughout a population of either adherent, or suspension, human cell lines. We believe that a number of attributes make it a valuable addition to the broad portfolio of cell cycle synchronization technologies.

The release from palbociclib-imposed cell cycle arrest at the natural decision point for cells, the restriction point, is not associated with changes in growth rates or rates of progression through the cycle into which they are released [36,72]. Rather Cdk4/6-Cyclin D activity appears to set size control [36,72]. Conceptually, this resonates with the original definition of the restriction point, as a rate-limiting gateway upon which multiple regulatory systems converge to regulate passage through the gateway into commitment to division [47].

A second appeal of this approach lies in the apparently limited impact upon genome integrity. Although the double thymidine block approach has predominated for over 50 years, the DNA damage that accompanies the arrest persists through the release (figure 10) [28,29]. This cumulative damage limits the utility of this approach for the study in several fields, including DNA replication, DNA repair and chromatin. While assessing genome integrity through the acquisition of γ-H2AX foci is not an exhaustive assessment of damage, our data suggest that Cdk4/6i induction synchrony is not accompanied by the damage that arise following release from thymidine [29]. Extensive analyses of mitotic progression following palbociclib induction synchronization revealed no sign of any of the chromosomal abnormalities that accompany thymidine induction synchronization [73] (Jon Pines and Mark Jackman 2020, personal communication). Cdk4/6i induction synchronization, therefore, has the potential to open up a number of previously intractable questions to study in synchronous cultures.

Perhaps the most important appeal of this approach is that cells can remain arrested outside of the cell cycle in full serum for very protracted timescales while retaining the competence to return to a synchronized cycle. This stasis means that a molecule of interest can be completely depleted, while cells are out of the cycle, without having any impact upon the progression through the previous cell division cycle. If a mutant version of this target was simultaneously induced, it would support highly directed questions about protein function when the cells are released to synchronously transit the ensuing cycle. The study of regulation of centriole biogenesis by Viol et al. [70] provides a compelling illustration of the power of protein induction in a palbociclib arrest prior to release.

While the benefits of this approach are particularly attractive, as with all approaches to cycle synchronization, disadvantages will inevitably emerge as these protocols are more widely adopted. Although it seems that cell mass accumulation at the arrest point does not impact upon cell cycle kinetics after release [36], studies in model organisms suggest that there is likely to be adaptation to modified size control at some point after commitment [74]. Recent studies suggest that adaptation is unlikely to occur after the second cycle after release [72]. The approach is also not effective in many lines when the level of control exerted by Cdk4/6 versus Cdk2 is tipped more heavily in favour of Cdk2 control [13,20,22,48,54,61–63,72,75–79].

When a particular line is refractory to Cdk4/6 induction synchronization, several approaches may switch the line to confer sensitivity to Cdk4/6 inhibition. For lines in which Cdk4/6 inhibition alone has little impact, reducing flux through to Cdk4/6 Cyclin D by reducing serum, or inhibiting the downstream MAP kinase pathways can compromise translation to reduce Cyclin D levels and tip the balance to impose cell cycle arrest by palbociclib, or Cdk2 inhibition [13,59,72,80]. The cell cycle arrest in HCT116 when Trametinib complements palbociclib is a good example of this synergy [81]. The recent revelation that the Cdk4/6 inhibitors are targeting the inactive, rather than active Cdk4 and Cdk6, complexes suggests another option for refractory lines when resistance arises from greater reliance upon Cdk6, rather than Cdk4 [54]. Cdk4/6 inhibitors impose the arrest because they sequester Cdk4 and Cdk4–CyclinD away from the Hsp90 chaperone system to reduce the number of molecules that can form an active complex with p27 [54,67,82]. The lower affinity of Cdk6 for the Hsp90 chaperone complex enables it to more readily assemble into active trimers that have no affinity for the inhibitors than Cdk4 does. This has prompted the suggestion that the predominance of Cdk6–Cyclin D-p27 trimers depletes p27 from the pools of Cdk2 to elevate Cdk2 activities and confer palbociclib resistance in cancers in which Cdk6 expression is elevated [67,83,84]. Thus, cell lines that rely upon Cdk6 rather than Cdk4 to drive passage through the gateway into the cycle will have reduced sensitivity to Cdk4/6 inhibitors. In such lines, a Cdk6-specific PROTAC may reduce reliance upon Cdk6 to impose sensitivity to Cdk4/6 inhibition and so support synchronization with the inhibitors [85]. Finally, because many cell lines will bypass the requirement for Cdk4/6 by exploiting Cdk2 to drive cells into cycle, partial inhibition of Cdk2 in these lines [13,86–88] should sensitize cells to Cdk4 inhibition. One challenge with this approach is the accompanying risk that strong Cdk2 inhibition will impact not only upon DNA replication in the preceding cycle, but upon the regulation of mitotic commitment because Cdk2–Cyclin A has been tied directly to regulation of Wee1 and the G2/M transition [13,89–91].

Given the variety of means by which resistance to Cdk4/6 inhibitors arises [48], it is perhaps easiest to empirically test whether a line will be amenable to Cdk4/6i induction synchrony. One of two simple assessments will identify a line as being Cdk4/6i induction synchrony compliant. The arrest/release into nocodazole that we show in figures 1 and 4–6 will indicate competence of most lines to synchronize by Cdk4/6i induction synchrony: a line will synchronize if a population accumulates 2 N DNA content one doubling time after inhibitor addition, before switching to 4 N DNA content a further doubling time after release into nocodazole. However, this assay relies upon the strength of the SAC [71] that can be so weak in some lines that it fails to impose a long cell cycle arrest with 4 N DNA content [92]. The second approach is independent of SAC integrity. Cells are released from palbociclib into 1 µM EdU before entrance into the next cycle is blocked by re-addition of palbociclib 12 h after release. If EdU is incorporated into most genomes throughout a population that has arrested cell cycle progression with a 2 N DNA content in response to the second dose of palbociclib, then the line will be competent for synchronization.

Our manipulations of cell density and culture history highlight the importance of growth state and culture history when customizing the synchronization protocol to gain maximum synchrony in the chosen line. The plasticity of the G1 control of commitment to the cycle is well documented. Distinct transcriptional and proteomic changes accompany cell cycle exit when a switch from the cycle is triggered by contact inhibition, serum starvation or inhibition of DNA replication, such that a cell's history alters the configuration of signalling in actively cycling cells and those returning from quiescence [19–22,59,72,80]. Thus, a recent history of contact inhibition reduces the efficiency of synchronization upon release from palbociclib (compare the profile in figure 2b of cells that have been kept in subconfluent culture prior to manipulation, with that if figure 2d which shows the same starting population of cells that had experienced a brief spell of contact inhibition). Such memory of contact inhibition is likely to have broad ranging impacts, as illustrated by the way in which it changes the loading of replication factors at origins in hTERT-RPE1 cells [93]. Consequently, care should be exercised to ensure that a population's path to synchronization has been freely dividing and as ‘healthy’ as it can be.

While we outline the utility of palbociclib induction synchrony in the study of cell cycle control and execution, pausing the cycle at the restriction point could have considerable benefit in other fields, such as synchronizing the formation of the primary cilium by serum depletion from Cdk4/6i arrest, or in studying differentiation in systems, such as haematopoesis, where a pause in G1 can be crucial for differentiation.

In conclusion, the successful quest to generate drugs to control proliferation in the clinic [65] has generated tools that will be of great utility in studies of the functional and biochemical changes that accompany and drive cell cycle progression.

5. Conclusion

Cdk4/6 induction synchrony is a simple, reproducible approach that will support biochemical interrogation, induction, depletion and replacement of molecules in any scale of cell line culture. Simple assays can determine competence of this approach to synchronize cell cycle progression in a novel line. We anticipate that a number of approaches may convert a recalcitrant line into a Cdk4/6i synchronization compliant line.


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Huberman, J. A., Tsai, A., and Deich, R. A., Nature, 241, 32 (1973).


Hanaoka, F., and Yamada, M., Biochem. Biophys. Res. Commun., 42, 647 (1971).

Tremblay, G. Y., Daniels, M. J., and Schaechter, M., J. Mol. Biol., 40, 65 (1969).

Ben-Porat, T., Stere, A., and Kaplan, A. S., Biochim. Biophys. Acta, 61, 150 (1962).

Levis, A. G., Krsmanovic, V., Miller-Faures, A., and Errera, M., Europ. J. Biochem., 3, 57 (1967).

Friedman, D. L., and Mueller, G. C., Biochim. Biophys. Acta, 174, 253 (1969).

Mizuno, N. S., Stoops, C. E., and Sinha, A. A., Nature New Biology, 229, 22 (1971).

Mizuno, N. S., Stoops, C. E., and Peiffer, jun., R. L., J. Mol. Biol., 59, 517 (1971).

Pearson, G. D., and Hanawalt, P. C., J. Mol. Biol., 62, 65 (1971).

Probst, H., Ulrich, A., and Krauss, G., Biochim. Biophys. Acta, 254, 15 (1971).

Hatfield, J. M. R., Exp. Cell Res., 72, 591 (1972).

O'Brien, R. L., Sanyal, A. B., and Stanton, R. H., Exp. Cell Res., 70, 106 (1972).

Comings, D. E., and Kakefuda, T., J. Mol. Biol., 33, 225 (1968).

Huberman, J. A., and Riggs, A. D., J. Mol. Biol., 32, 327 (1968).

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Comings, D. E., and Okada, T. A., Exp. Cell Res., 62, 293 (1970).

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Kikuchi, T., and Sandberg, A. A., J. Nat. Cancer Inst., 32, 1109 (1964).

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The model represents the change in the DNA content of a cell during the cell cycle.

The life cycle of eukaryotic cells can generally be divided into four stages and a typical cell cycle is shown in Figure 2.13. When a cell is produced through fertilization or cell division, there is usually a lag before it undergoes DNA synthesis (replication). This lag period is called Gap 1 (G1), and ends with the onset of the DNA synthesis (S) phase, during which each chromosome is replicated. Following replication, there may be another lag, called Gap 2 (G2), before mitosis (M). Cells undergoing meiosis do not usually have a G2 phase. Interphase is as term used to include those phases of the cell cycle excluding mitosis and meiosis. Many variants of this generalized cell cycle also exist. Some cells never leave G1 phase, and are said to enter a permanent, non-dividing stage called G0. On the other hand, some cells undergo many rounds of DNA synthesis (S) without any mitosis or cell division, leading to endoreduplication. Understanding the control of the cell cycle is an active area of research, particularly because of the relationship between cell division and cancer.

The amount of DNA within a cell changes following each of the following events: fertilization, DNA synthesis, mitosis, and meiosis (Fig 2.14). We use “c” to represent the DNA content in a cell, and “n” to represent the number of complete sets of chromosomes. In a gamete (i.e. sperm or egg), the amount of DNA is 1c, and the number of chromosomes is 1n. Upon fertilization, both the DNA content and the number of chromosomes doubles to 2c and 2n, respectively. Following DNA replication, the DNA content doubles again to 4c, but each pair of sister chromatids is still counted as a single chromosome (a replicated chromosome), so the number of chromosomes remains unchanged at 2n. If the cell undergoes mitosis, each daughter cell will return to 2c and 2n, because it will receive half of the DNA, and one of each pair of sister chromatids. In contrast, the 4 cells that come from meiosis of a 2n, 4c cell are each 1c and 1n, since each pair of sister chromatids, and each pair of homologous chromosomes, divides during meiosis.

water is a polar molecule. a chemical bond is a force of attraction between atoms or ions. bonds form when atoms share or transfer valence electrons. atoms form chemical bonds to achieve a full outer energy level, which is the most stable arrangement of electrons. so, false

in h2o molecule, two water molecules are bonded by a hydrogen bond but the bond between two h - o bonds within a water molecule are covalent. the greater the electronegativity difference, the more ionic the bond is. bonds that are partly ionic are called polar covalent bonds. nonpolar covalent bonds, with equal sharing of the bond electrons, arise when the electronegativities of the two atoms are equal.

the cells uses some of the atoms of the glucose molecules to atp molecules. the atoms of the glucose molecules will then be converted into energy which will be eventually stored in atp. the energy produced which are stored in atp are coming from the glucose’s chemical bonds.


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MATERIALS AND METHODS

Yeast Strains and Plasmids

The wild-type (PSY580) and NTF2 deletion strains (ACY114, ACY115) used in this study have been described (Corbett and Silver, 1996). The MAD2 deletion strain, KH141, (MATa, MAD2::URA3,leu2,3–112, trp1–63, ade2,his3–11,15) was the generous gift of Andrew Murray. The yeast plasmids, pPS882 (CEN, LEU2, NTF2), pPS883 (CEN, URA3, NTF2), pPS919 (CEN, LEU2, ntf2–1), and pPS920 (CEN, LEU2, ntf2–2) used in this study have been described (Corbett and Silver, 1996). The bacterial expression plasmid for Ntf2p (pPS982) has been described (Wong et al., 1997). The NLS-green fluorescent protein (GFP) plasmid (pGADGFP) used for the NLS-GFP import assay has been described (Shulga et al., 1996). The plasmid for the expression of C-terminal fusion to the GFP has been described (Kahana et al., 1995). Fusion constructs to scRan (Gsp1p) were constructed by engineering an in-frameXhoI site at the termination site of the GSP1coding region and a PstI 710 bp upstream of the 5′ start site of the GSP1 coding region (pAC410). The bacterial expression plasmids for Ntf2–1p (pAC41) and Ntf2–2p (pAC43) were generated by cloning an ∼480-bp PCR fragment containing a 5′-BamHI restriction enzyme site and a 3′-HindIII restriction enzyme site introduced by PCR amplification of pPS991 (ntf2–1ts) or pPS920 (ntf2–2ts) with the primers AC21 (5′CGGGATCCATGTCTCTCG ACTTTAACAC3′) and AC88 (5′CCCGAAGCTTCGCTATCGCCTTATACATCG3′). The PCR product was cloned into the T7-based expression vector pMW172 (Way et al., 1990). All procedures including yeast transformations, culture manipulations, and extract preparations were performed by standard methods (Adamset al., 1997).

Ntf2p Purification and Immobilization

Yeast Ntf2p was purified from Escherichia coli as previously described for rat Ntf2p (Clarkson et al., 1996). Expression plasmids for Ntf2p (pPS982), Ntf2–1p (pAC41), or Ntf2–2p (pAC43) were transformed into E. coli BL21 (DE3). Transformants were inoculated into 2X tryptone-yeast extract medium containing 100 μg of ampicillin/ml and grown overnight at 30°C. It was not necessary to induce expression as the basal level of expression of the T7 polymerase yielded a large amount of Ntf2p. Bacteria were harvested by centrifugation and were stored at −80°C until required.

Ntf2p was isolated by thawing the cell pellet and resuspending it in 25% sucrose, 50 mM Tris-HCl (pH 8.0), 5 mM MgCl2, 1 mM EDTA, 0.1 mM phenylmethylsulfonyl fluoride (PMSF). Cells were lysed by French press and treated with DNAse at 25°C for 30 min. The soluble fraction was isolated by centrifugation at 40,000 × g for 20 min and dialyzed overnight against 20 mM Tris-HCl (pH 8.0), 2 mM MgCl2, 1 mM dithiothreitol (DTT), 0.1 mM PMSF (NTF2 buffer A). The lysate was clarified at 40,000 ×g for 30 min at 4°C, was applied to DE52 ion exchange column (10 × 3 cm), and was washed with NTF2 buffer A. Ntf2p was eluted from the column with a gradient of 0 to 400 mM NaCl. Fractions containing Ntf2p were pooled, concentrated using a Centriprep-10 (Amicon, Charlotte, NC) concentrator, and applied to a column of Sephacryl SR100 preequilibrated in 20 mM Tris-HCl (pH 7.4), 50 mM NaCl, 2 mM MgCl2, 1 mM DTT, 0.1 mM PMSF (NTF2 buffer B). Fractions containing Ntf2p were collected and pooled.

Purified Ntf2p was cross-linked to cyanogen bromide (CNBr)-sepharose beads as previously described (Clarkson et al., 1996). Briefly, CNBr-sepharose beads (Pharmacia, Uppsala, Sweden) were swollen and washed in 1 mM HCl. Beads were transferred to coupling buffer (100 mM NaHCO3 (pH 8.3), 500 mM NaCl) and were added to 2–5 mg of Ntf2p in coupling buffer. Coupling was carried out at 4°C overnight. Residual active groups were blocked with 1 M Tris-HCl (pH 8.0) for 2 h at room temperature. Beads then were washed successively and extensively four times in coupling buffer and acid wash buffer (0.1 M sodium acetate (pH 4.0), 500 mM NaCl).

Binding Assays

Yeast cell extracts were prepared from cultures grown overnight at 30°C or at room temperature for ts strains in yeast extract-peptone-dextrose (YEPD) medium to confluency. Cells were harvested by centrifugation and washed once with water. Cells then were resuspended in one volume of PBSMT (1 × PBS, 2.5 mM MgCl2, 0.5% Triton X-100) supplemented with protease inhibitors (0.5 mM PMSF and 3 μg each of aprotinin, leupeptin, chymostatin, and pepstatin per milliliter). One volume of glass beads was added, and cells were lysed with 10–15 60-s pulses in a beadbeater (lysis was monitored by light microscopy to >70% lysis). The resulting lysate was clarified by centrifugation and assayed for protein concentration by using the Bio-Rad (Cambridge, MA) Protein Assay Kit.

Two milligrams of yeast lysate was incubated with 50 μl of Ntf2p-sepharose beads. Binding was carried out in PBSM (total volume, 500 μl) at 4°C for 1 h. Beads then were washed two times for 10 min in PBSM and one time for 10 min in PBSMT. Bound proteins were eluted with 100 μl of sample buffer and were resolved by SDS-polyacrylamide gel electrophoresis and transferred to nitrocellulose for immunoblotting.

Immunoblot Analysis

Immunoblot analysis was performed essentially as described (Towbin et al., 1979) with the following modifications: following transfer, the nitrocellulose filter was blocked in TBST buffer (10 mM Tris, 140 mM NaCl, 0.05% Tween-20) plus 5% milk for 15 min. Filters to be used for the detection of scRan were blocked with TBS1/2T (TBS buffer containing 0.025% Tween-20) for 15 min. scRan-GFP was detected by incubation with a 1:5000 dilution (in TBST plus 5% milk) of an anti-GFP rabbit polyclonal antibody (the generous gift of P. Silver and J. Kahana, Harvard Medical School, Boston, MA). scRan was detected by incubation with a 1:1000 dilution (in TBS1/2T plus 5% milk) of an antiscRan rabbit polyclonal antibody (the generous gift of D. H. Wong and P. Silver, Harvard Medical School). Following several washes with TBST (or TBS1/2T for scRan), the filter next was incubated in a 1:5000 dilution of horseradish peroxidase-linked goat antirabbit polyclonal antiserum (Promega, Madison, WI) for 1 h at room temperature. The filter was again washed in TBST (or TBS1/2T for scRan) and detection was carried out with a 1:1 mixture of (chemiluminescent) ECL reagents (Amersham, Little Chalfont, UK) as recommended by the manufacturer.

Localization of scRan-GFP and Nuf2-GFP

The scRan-GFP and Nuf2-GFP (Kahana et al., 1995) fusion proteins were localized by directly viewing the GFP signal in living cells through a GFP-optimized filter (Chroma Technology, Brattleboro, VT) using an Olympus (Tokyo, Japan) BX60 epifluorescence microscope equipped with a Photometrics (Tucson, AZ) Quantix digital camera.

Indirect Immunofluorescence Microscopy

Ten-milliliter cultures were grown to log phase in YEPD overnight, then the cultures were split, and half was maintained for 3 h at 25°C and half was shifted to 37°C for 3 h. Cultures then were fixed with 600 μl of 37% formaldehyde for 30 min. Following fixation, cells were harvested by centrifugation and were washed once with 0.1 M potassium phosphate buffer (pH 6.5), once with P solution (1.2 M sorbitol, 0.1 M potassium phosphate buffer, pH 6.5), and were resuspended in 1 ml of P solution. DTT was added to 25 mM, and the cells were incubated at 30°C for 10 min with gentle agitation followed by the addition of 300 μg of zymolyase (United States Biological, Swampscott, MA). The digestion of the cell wall was monitored by microscopy. Digested cells were collected by centrifugation, were washed once with P solution, and were resuspended in 1 ml of P solution. Cells then were applied to Teflon-faced microscope slides precoated with 0.3% polylysine. After cells were adhered to slides, they were fixed with methanol at −20°C for 6 min and dried in cold acetone for 30 s. Cells were blocked by incubating for 15 min with PBS plus 0.5% bovine serum albumin (BSA). Antitubulin was diluted 1:100, and anti-Npl3p was diluted 1:1000 in PBS plus 0.5% BSA and incubated with the cells overnight at room temperature. Cells were washed several times with PBS plus 0.5% BSA and were incubated for 2 h with a 1:1000 dilution of either fluoroscein isothiocyanate (FITC)-labeled antimouse antibodies or Texas Red-labeled antimouse antibodies (Jackson ImmunoResearch, West Grove, PA) for antitubulin or with FITC-labeled antirabbit for Npl3p as indicated and with 4′,6-diamido-2-phenylindole (DAPI antibodies) (1 μg/ml).

Growth and Viability

NTF2, ntf2–1, and ntf2–2 cells were grown in YEPD overnight at 25°C, diluted to 2 × 10 6 cells/ml and shifted to 37°C. Growth was monitored by counting cells every 2 h using a hemacytometer. Viability was monitored every 2 h by plating 200 cells/plate onto YEPD plates. Plates were incubated at 25°C for 4 days, and colonies were counted.

Determination of Yeast Cell DNA Content

Cells were prepared for the FACS by staining with propidium iodide (Epstein and Cross, 1992). Briefly, cells were ethanol fixed overnight at 4°C, washed, and resuspended in 1 ml of 50 mM sodium citrate, pH 7.0. Cells then were treated with 0.08 mg/ml Rnase A for 1 h at 50°C followed by 0.25 mg/ml proteinase K before incubation in 8 μg/ml propidium iodide. Each sample was analyzed with a FACS Caliber cytometer from Becton Dickinson (Franklin Lakes, NJ).

Construction and Analyses of MAD2Δ Strains

To construct the ntf2ts MAD2Δ, double mutants, KH141 (MAD2Δ) was mated with ACY115 (NTF2Δ) containing either pPS919 (ntf2–1ts) or pPS920 (ntf2–2ts). The resulting diploids were sporulated, and tetrads were dissected to isolate the double-mutant strains. Single and double mutants as well as a wild-type controls were grown overnight in YEPD at 25°C. Cells were diluted to 0.2 × 10 6 cells/ml in 50 ml of YEPD, with half grown at 25°C and half shifted to 37°C. Growth was monitored by measuring the OD600. To test the benomyl sensitivity of the yeast strains indicated, cells were grown overnight at 25°C, and 100,000, 10,000, 1,000, 100, and 10 cells were spotted on 10-μg/ml benomyl plates and incubated at 25°C. To test the nocodazole sensitivity of the yeast strains indicated, cells were grown overnight at 25°C and diluted to 0.2 × 10 6 cells/ml into YEPD containing 15 μg/ml nocodazole, and growth was monitored by OD600.

NLS-GFP Import Assay

The NLS-GFP import assay was performed as previously described (Shulga et al., 1996). Briefly, cells were grown to early-midlog phase in synthetic media containing 2% glucose at 25°C, were pelleted, were resuspended in 1 ml of 10 mM sodium azide and 10 mM 2-deoxy- d -glucose in glucose-free synthetic medium, and were incubated at 25°C for 45 min. The cells then were pelleted, were washed with 1 ml of ice-cold ddH20, were repelleted, were resuspended in 100 μl of glucose-containing synthetic medium prewarmed to 37°C and were incubated at 37°C. For scoring, 2-μl samples were removed every 2.5 min, and cells observed and counted through a GFP optimized filter (Chroma Technology) using an Olympus BX60 epifluorescence microscope. Cells were scored as “nuclear” if the nucleus was both brighter than the surrounding cytoplasm and a nuclear-cytoplasmic boundary was visible. At least 50 cells were counted at each time point.


Cell Cycle Control

The ‘life cycle’ of a dividing eukaryotic non-embryonic cell starts with the cell triggered to enter the cell cycle and ends with the equal partitioning of the genetic material and cleavage of the cell during cytokinesis. The whole process is called the cell cycle and consists of four main phases.

Entry to the cycle is made in Gap 1 (G1) phase and this is followed in sequence by a DNA synthesis (S) phase, Gap 2 (G2) phase, and Mitosis (M). After mitosis (M) some cells enter the G1 phase of a new cell cycle whilst others may diverge at the start of G1 into a phase called Gap O (zero). Phases G1, S and G2 are often grouped and called ‘interphase’.

Cells in G-0 (zero) are quiescent and not dividing (hence zero), this may be permanent or temporary.
Mitosis (M phase) had been observed and described in some detail by the start of the 20th century, but it was not until about 50 years later that it was discovered that DNA synthesis took place as a separate process ahead of mitosis. Between mitosis (M) in a previous cell cycle and DNA synthesis (S) there was a time gap. This was designated Gap 1 (G1). The time gap between DNA synthesis (S) and mitosis (M) was designated Gap 2 (G2).

After cell division the daughter cells follow ONE of several pathways:

  • cells that can divide again immediately enter the G1 phase of a new cell cycle.
  • other cells enter G-0 phase. Some of them are quiescent for a time but then re-enter G1 phase.
  • some specialist cells, for example nerve cells, do not divide again.
  • other cells can be triggered by activities such as wound repair to enter G1 phase of the cell cycle to divide as required.

Mitosis is visually very dramatic but it only occupies about 5% of the total cell cycle time. Within the ‘big picture’ of the life cycle of a dividing cell, interphase (phases G1, S, and G2) account for the other 95% of cell cycle time. Research evidence shows interphase (once called resting phase!) operates in a beautifully ordered systematic way. It has also been shown to be more complicated than previously thought.

The time taken for a eukaryotic cell to divide varies widely with cell type and environment. Yeast cells take from 1.5 to 3 hours, intestinal epithelial cells about 12 hours and cells in culture about 22 hours. In different organisms and at different developmental times the details of the cell cycle vary. Embryonic cells in many organisms run a cycle that is shorter than similar cells in the adult. Cells of yeast and mammal show differences in cycle detail but the general mechanism of the cell cycle has been highly conserved over the years.

During the cell cycle cytoplasmic chemistry influences to a large extent the activities of the whole cell. At all other times we think in terms of the cell nucleus determining cytoplasmic activity.
KEY CONCEPTS:

  • The cell cycle is a high quality cell production facility.
  • An efficient production facility must have a controlled, co-ordinated and sequential system for producing products. Also needed is a system to monitor incoming raw materials, product processing, the processing environment and the facility to anticipate and deal with production problems.
  • The eukaryotic cell is a ‘good manufacturer’ – most of the time! During the cell division cycle it has the biological equivalent of the following systems in place: Quality Assurance (QA), Quality Control (QC) and as part of QC, Internal Security (IS).
  • At certain points in the cycle critical ‘reviews and decisions’ are made by QA and QC. These events are called ‘checkpoints’.

The cell cycle system of a eukaryotic organism includes:

  1. a biomolecular surveillance capability that integrates extracellular and intracellular signals and informs the cell’s QA and QC about: the need for a new cell the availability of raw materials including a considerable amount of chemical energy whether the microenvironment is suitable for cell division and, the integrity of the genome and the fidelity of its replication (Mainly QA).
  2. a system that controls the cycling of the cycle and ensures the cell only enters the next phase when passage through the previous one is complete also that the phases of the cell cycle are entered in the correct order. Satisfactory passage through all four phases and usually the various checkpoints of the cell cycle are critical for cell division (QC).
  3. a control system that initiates and terminates chemical reactions in the cell cycle and, with the use of a form of ‘licensing’ ensures that DNA is replicated once and once only during S phase (QC and IS).
  4. cell cycle arrest and delay functions (QC).
  5. DNA damage detection and repair facilities and a system that detects unreplicated DNA and, in mitosis, a system to ensure adequate spindle assembly and chromosome attachment (QA/QC), – otherwise known as “checkpoints”, and
  6. the capability and facility to trigger programmed cell death (apoptosis) (QC).

A cell is ‘cycled’ through each phase and from phase to phase by the action of proteins including specific cyclins and cyclin dependent kinases (cdks). Different cyclins and cdks rise and fall in activity during the cell cycle.

Sometimes faults go undetected (as in industry). Quality control and assurance systems can also fail. Quality control system failure is associated with cell and organ disease and probably as many as 50% of cancers (but this does not mean one single system failure alone will cause cancer).
KEY CONCEPTS: ENTRY LEVEL SUPPORTING INFORMATIONCells of eukaryotes normally only divide when instructed to do so. Chemicals called mitogens signal cells to start dividing.Cells competent to divide join the cell cycle in G1 phase and remain in this phase for a bit less than half of the total cell cycle time. This is the longest phase and microenvironmental conditions and signals received from other cells can shorten or lengthen G1.Early G1 phase
Recent work suggests that a licensing system for ‘one DNA copy only’ is set up early in G1 phase and expires only after DNA replication has started in S phase. During G1 phase only one set of the genome is present.

Late G1 phase
The main drivers of progression through the cell cycle are called protein kinases. There are several of these and each is a combination of a cyclin and a catalytic enzyme called a cyclin dependent kinase (cdk).
The different combinations operate in specific parts of the cell cycle and rise and fall in activity during the cycle. In so doing they contribute to the mechanism of phase entry and exit. The raised level of activity of the various cyclin and cdk combinations is terminated by proteolysis of cyclins after polyubiquitination. (Ubiquitins are a group of proteins that are covalently linked to proteins targeted for degradation. After molecular binding the target proteins, in this case cyclins, are degraded by proteolysis – protein loosening from Greek lusis, ‘loosening’)

The first cyclin combination (cyclin D and cdk 4 and 6) trips in about three quarters of the way through G1 phase to be joined later by the cyclin E and cdk 2 combination, both cyclins driving the cell into S phase

At about this time the cell passes through the ‘Restriction Point’ (in yeasts called START). This is a point of ‘no return’, the cell cycle equivalent of ‘Caesar crossing the Rubicon’. Once the cell passes this point it is restricted, there is no going back the cell is committed to replicate in ‘S’ phase.

Checkpoints

As the cell progresses through the cycle, checkpoints are encountered. Although not absolutely essential for cell division they are well-established features of most cells.
Checkpoints in the cell cycle are not unlike those found at some border crossings where passports, papers and merchandise are checked. At the molecular level a cell cycle checkpoint consists of (1) sensor/detector (2) a signal sender and (3) a receiver/effector.

G1 checkpoint
The first of the ‘surveillance checkpoints’ is found towards the end of Gap1 (G1) phase and is the G1 DNA damage checkpoint. At this checkpoint and just ahead of it the DNA of the cell selected to divide is subjected to biomolecular surveillance for integrity. In humans DNA damage is ‘self-signalled’ mainly by the damaged DNA itself raising the level and activity of the protein products of certain genes, especially those of the tumour suppressor gene p53 from chromosome 17. Gene p53 is called the ‘guardian of the genome’ and is largely responsible for determining whether the cell should be admitted to the next synthesis (S) phase. If DNA is not too damaged a repair may be possible and the cell cycle arrested and slowed down until repairs are effected. Cell cycle ‘quality control’ may determine that the damage cannot be repaired. In this case ‘p53’ will trigger the programmed cell death (apoptosis) facility.

The division cycle of a cell with badly damaged DNA may end at this checkpoint.

Unfortunately, quality control in cells is not perfect. Very occasionally DNA damage is missed by QC and slightly changed DNA slips through the system. Sometimes this is because the damaged DNA does not trigger the checkpoint system. The quality control system may also be damaged itself. DNA of the p53 gene can be damaged by sunlight (u.v. radiation) and mutagenic chemicals including those from cigarette smoke and aflatoxin from, for example, mouldy peanuts.

At the G1 checkpoint in yeast cell cycle, a check is made to ensure cell size is correct for division, but cell size has been found not to be so critical a factor in some cells of higher eukaryotes

S (DNA synthesis) phase

S phase occupies about a third of total cell cycle time. It is here that under the ‘licensing’ system only one new copy of the cell’s DNA is synthesized. This includes acceptable alterations but also questionable ones not detected by QC because of system errors and breakdown. The failure of the ‘guardian of the genome’ p53 gene to operate due to damage within its own DNA is an example of this.

In S phase the double-stranded DNA unwinds into two component strands that serve as templates for the synthesis of a new strand on each. Newly formed units of the bases adenine (A), thymine (T), guanine (G) and cytsosine (C) are attached to compliment bases on the unwound DNA. One set of DNA is now produced for each of the two daughter cells. The process of DNA synthesis consumes a considerable amount of energy. (For details of this process see the textbooks listed on our website for example: Pollard, T. D. & Earnshaw, W. C., ‘Cell Biology’ 1st Edit. 2002. 2nd printing, with additions 2004. Publ: W. B. Saunders).

S phase checkpoints
There appear to be three types of checkpoint in S phase. Not surprisingly all respond in some way to problems with DNA replication. These problems range from a shortage of deoxyribonucleotides for making new DNA, to the presence of enzyme inhibitory chemicals and breaks in the DNA molecule. The cell cycle can arrest here if the DNA is unreplicated or in any way incomplete and not competent to proceed to phase G2.
If everything goes smoothly by the end of S phase the cell will contain two identical sets of its genome.

The cycle is driven through S phase and into G2 phase by the cyclin A and cdk 2 combination.

G2 (Gap 2) phase
G2 phase is generally shorter than that of G1. Much of this time is spent in developing and preparing organelles for dividing and sharing during cytokinesis at the end of mitosis. Cyclins A and B coupled to cdk 1 drive the cell through the end of S phase, through G2 phase and M phase This phase contains the G2 checkpoint. After G2 the cell is committed to division.

G2 Checkpoint
This DNA structure checkpoint is encountered towards the end of the G2 phase. The G2 checkpoint is very critical in that it has the ‘responsible’ function of providing a quality assurance check before the cell enters mitosis.

  1. monitoring takes place to ensure that there is no unreplicated DNA and that TWO identical sets of the genome are now present and intact.
  2. both sets of the genome are ‘proof-read’ with a high level of surveillance. A check is made for molecular damage within the DNA and this evidence will determine whether the genome will be retained, repaired or rejected.
  3. if repair of the damaged DNA is possible the cell cycle is arrested for as long as the repair takes. Cell cycles operate in ‘real time’ so delay can be accommodated within the system by QC delaying future events.
  4. badly damaged DNA will be identified and the p53 gene products will trigger the programmed death (apoptosis) programme.
  5. ‘Quality control’ will check the overall competence of the cell to enter mitosis. On the list of competences will be such items as “is the mechanism leading to the separation of the sister chromatids in place?”<

M phase (mitosis)
Mitosis presents the drama of division. An awesome and beautiful presentation, it occupies just 5% of the total cell cycle time, or roughly an hour in higher eukaryotic cultured cells. It is the shortest phase but the final production is the culmination of work done during the rest of the cell cycle.

Cyclin A and B coupled to cdk 1 drives the cell through mitosis (the student is referred to a textbook at the desired level for detailed information about mitosis). At the end of mitosis the sister chromatids, joined as pairs by cohesive ‘glue’ since they were replicated in S phase, are separated to form two new equivalent sets of chromosomes. This event is chromosome segregation.

Some organelles in the cytoplasm are disassembled into molecular units to be divided during division of the cytoplasm (kinesis), and then new ones constructed in the daughter cells. Other cytoplasmic inclusions are shared (not always evenly) between the daughter cells.

M phase (mitosis) checkpoints

The working of one checkpoint within mitosis, the metaphase checkpoint, is well established. There may be more checkpoints but further work is needed to establish their existence.

Metaphase checkpoint

Also called ‘spindle assembly or kinetochore attachment checkpoint’ it operates during metaphase and before the cell enters anaphase. It checks for misaligned chromosomes and also that microtubules are attached to kinetochores – a very critical mechanism. The start of anaphase is delayed until all the chromosomes are aligned and appropriately attached.

When telophase is complete cytokinesis – (the division of cytoplasm) takes place. After this the two daughter cells will be directed to G-0 (zero) or G1 phase.

CONCEPT MESSAGES:

  1. Research evidence shows very clearly that cell division in eukaryotes should now be viewed as an active multiphase process called the CELL CYCLE. The process is highly conserved and covers all the phases of preparation as well as the actual cell division. Satisfactory passage through the phases and checkpoints of the cycle are critical for cell division to take place.
  2. Mitosis whilst obviously very important and dramatic should be viewed as part of the ‘Big Picture’ of cell division. Mitosis occupies just 5% of the total cell cycle time.
  3. Working within the cell cycle are various forms of molecular quality assurance and control. These controls operate to assist the cell division programme and prevent damage to the genome.

CHALLENGE YOUR CRITICAL THINKING:

  • Cancer is about cells dividing in an unregulated way. The nature of the cell cycle control system lends itself to the design of anti-cancer drugs that disrupt the cell cycle and initiate programmed cell death.
    Consider the total cell cycle. Select two or more strategic sites as possible targets for the action of anti-cancer drugs? Think through in terms of cell and molecular biology what you would want the drug to do in order to interfere with the cell cycle.
  • If DNA replication and cell division were always perfect in every case what would be the implications for evolution?

SELECTED WEBSITES:
http://nobelprize.org
Click on Cell Cycle game in the education panel.

http://www.cancerresearchuk.org
Click on ‘Science & Research’ in the text. (Then select Annual Reports especially LRI 2002 & Sc.Yearbook 02/03)

http://www.cellsalive.com
Click on ‘cell cycle’ (select ‘checkpoints for animation’).

Grateful thanks are due to Margarete Heck for her valuable contribution.


Watch the video: The Cell Cycle and cancer Updated (May 2022).