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Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, RussiaFederal Center of Brain Research and Neurotechnologies, Federal Medical Biological Agency, Moscow, Russia
Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, RussiaFederal Center of Brain Research and Neurotechnologies, Federal Medical Biological Agency, Moscow, RussiaCenter for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, Moscow, Russia
Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, RussiaCenter for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, Moscow, Russia
Detection of thymidine analogues after their incorporation into replicating DNA represents a powerful tool for the study of cellular DNA synthesis, progression through the cell cycle, cell proliferation kinetics, chronology of cell division, and cell fate determination. Recent advances in the concurrent detection of multiple such analogues offer new avenues for the investigation of unknown features of these vital cellular processes. Combined with quantitative analysis, temporal discrimination of multiple labels enables elucidation of various aspects of stem cell life cycle in situ, such as division modes, differentiation, maintenance, and elimination. Data obtained from such experiments are critically important for creating descriptive models of tissue histogenesis and renewal in embryonic development and adult life. Despite the wide use of thymidine analogues in stem cell research, there are a number of caveats to consider for obtaining valid and reliable labeling results when marking replicating DNA with nucleotide analogues. Therefore, in this review, we describe critical points regarding dosage, delivery, and detection of nucleotide analogues in the context of single and multiple labeling, outline labeling schemes based on pulse-chase, cumulative and multilabel marking of replicating DNA for revealing stem cell proliferative behaviors, and determining cell cycle parameters, and discuss preconditions and pitfalls in conducting such experiments. The information presented in our review is important for rational design of experiments on tracking dividing stem cells by marking replicating DNA with thymidine analogues.
Somatic stem cells are cells able to self-renew and to produce progeny that undergo differentiation into functional organ- or tissue-specific cells. Somatic stem cells support continuous physiological organ or tissue renewal and/or regeneration of injured organs or tissues. Disturbance of somatic stem cell functioning is thought to underlie many degenerative diseases in humans, and in situ stem cell activation and transplantation into injured organs are considered as promising therapeutic strategies to delay or resist the onset of diverse degenerative disorders.
Stem cells undergo mitotic division to realize their functions within an organism. Delivery and subsequent detection of thymidine analogues that incorporate into replicating DNA during the S-phase of the cell cycle are one of the basic approaches for tracing the fate of dividing stem cells and their progeny in diverse in vitro and in vivo systems. Several comprehensive reviews have already described the application of nucleotide analogues for marking replicating DNA (
). These reviews addressed questions regarding the technical aspects of nucleotide analogue detection using antibodies or bioorthogonal chemical reactions, approaches for double S-phase labeling, applications of modified nucleotides for stem cell research, investigation of spatiotemporal features of DNA replication, multiparametric cell cycle analysis by flow cytometry, and labeling of living cells. In this review, we will focus on (i) critical points regarding delivery, dosage, and detection of nucleotide analogues for single- and multilabel marking of replicating DNA, (ii) applications of pulse-chase and cumulative labeling schemes and their combinations for determining cell cycle parameters and for revealing specific modes of cell cycle behavior, such as re-entering and exiting the cell cycle, (iii) caveats to consider when applying labeling with modified nucleotides, and (iv) the most recent advances in detection of replicating DNA. These topics are largely absent in previous reviews.
Delivery and detection of nucleotide analogues
A brief overview on marking replicating DNA
Replication of genetic material is a key process underlying cell division. It is essential for creating multicellularity and multiplication of all organisms. A cell replicates its DNA when passing through the S-phase of the cell cycle. Tagging replicating DNA enables nuclei of dividing cells and their progenies to be marked due to ability of the tag to remain within the replicated DNA for prolonged periods.
Labeling replicating DNA with the radioactive nucleoside 3H-thymidine, which is a precursor of one of the four chemical building blocks of DNA, and its detection by autoradiography was initially introduced by Taylor et al. in 1957 (
). In their work they used autoradiographic analysis of chromosome preparations from Vicia faba seedlings treated with 3H-thymidine and revealed that only one of the two sister chromatids in each chromosome was radioactive in the cells of the roots collected after the second replication cycle. Thus, during replication, daughter chromosomes receive an original and a new strand. This observation supported the semiconservative replication model. Later, the delivery of 3H-thymidine and another radioactive nucleoside, 14C-thymidine, with subsequent autoradiographic detection revealed features and mechanisms of DNA replication in pro- and eukaryotic cells, such as unwinding of the double helix, formation of the replication fork, spatial patterning of DNA replication, and creation of the lagging DNA strand through intermittent synthesis of Okazaki fragments (reviewed in (
)). By tracing dividing cells and their progeny by autoradiographic detection, 3H-thymidine was widely employed in developmental biology, regenerative biology, and stem cell research. For instance, this approach enabled birth dating of neurons within different cortical layers during corticogenesis in mammals (
). The major disadvantages of 3H-thymidine are handling of a radioactive substance and the use of the time-consuming autoradiography method for detection. Detection of 5-bromo-2′-deoxyuridine (BrdU) (Table 1), a synthetic nucleoside analogue of thymidine, is an alternative technique for the determination of DNA replication and has overcome these disadvantages (
). BrdU incorporated into DNA is recognized by a specific polyclonal or monoclonal antibody produced against bromouridine or iododeoxyuridine complexed to a carrier protein such as bovine serum albumin. The ability to combine BrdU labeling with the detection of cell-type-specific markers via specific antibody staining or reporter gene expression has become a gold standard for studying cell division and differentiation, which are major cellular processes underlying development in multicellular organisms and tissue renewal and regeneration in adulthood.
Table 1Summary on modified nucleotides
Doses used for intraperitoneal injections in rodents
Solubility in water-based solvents (maximal concentrations that have been reported), mg/ml
Due to their nonoverlapping detection techniques, BrdU and 3H-thymidine can be combined in one sample allowing for temporal discrimination of DNA synthesis in dividing cells to reveal their progression through the cell cycle (
). However, the detection of truly double labeled nuclei in tissue sections using immunohistochemistry and autoradiography may be compromised, because anti-BrdU antibodies stain cell nuclei and the radioactive label causes formation of silver grains in the photographic emulsion covering the tissue section surface. Therefore, if the nuclei of two cells overlap when we observe a tissue section through the microscope, we are unable to judge unequivocally whether we see a truly double labeled cell or two cells that have incorporated different labels. Application of the halogenated thymidine analogues 5-chloro-2′-deoxyuridine (CldU) and 5-iodo-2′-deoxyuridine (IdU) that resemble BrdU in their ability to tag replicating DNA (Table 1) and extension of the antibody panel for their recognition have led to the development of a method for concurrent detection of two distinct labels within a sample, allowing for birth dating of various cohorts of cells (
The major drawback of detection of halogenated thymidine analogues using antibodies is the necessity of DNA denaturation, usually in a 0.5 N–4 N hydrochloric acid solution, prior to sample processing, because of the inaccessibility of the BrdU epitope within complimentary paired bases. The DNA denaturation procedure erodes cell and tissue components limiting the use of various concurrent molecular assays. In particular, acidic DNA denaturation prior to BrdU detection by antibodies leads to poor staining of samples with widely used nuclear dyes such as propidium iodide, DAPI, or Hoechst stains. A variety of approaches were proposed to overcome this drawback. These include (i) treatment with sodium hydroxide, which disrupts the DNA structure via deprotonation of the nucleobases, (ii) incubation with various nucleases (for instance, exonuclease III) or nuclease mixtures to generate single-stranded regions, in which the antibody is able to bind to BrdU, (iii) exposure to monovalent copper ions, which, in the presence of oxygen, oxidizes deoxyribose moieties, producing DNA breaks, (iv) ultraviolet light photolysis, and (v) heating (
). The method is based on the incorporation of EdU into replicating DNA and its subsequent detection by the covalent coupling of a fluorescent azide to a terminal alkyne group through a Cu(I)-catalyzed [3 + 2] cycloaddition reaction, frequently called a ‘‘click’’ reaction (see Fig. 1 in (
)). This method does not require denaturation of DNA and, therefore, lacks the major drawback related to the use of halogenated thymidine analogues. EdU labeling followed by detection using the click reaction preserves the integrity of cell and tissue components allowing for a variety of concurrent molecular assays to be performed. The click reaction is a bioorthogonal chemical reaction, a type of reaction that does not interfere with biochemical reactions occurring in live cells (
The latest advances in creating strategies for probing biological macromolecules with a synthetic tag containing a bioorthogonal functional group have extended the toolset for labeling replicating DNA. Reported novel nucleotide analogues include (2′S)-2′-deoxy-2′-fluoro-5-ethynyluridine (F-ara-EdU), 5-(azidomethyl)-2′-deoxyuridine (AmdU), and 5-vinyl-2′-deoxyuridine (VdU) (Table 1). VdU is detected by a nonoverlapping bioorthogonal chemical reaction (an alkene–tetrazine ligation reaction). Therefore, it can be combined with other nucleotide analogues that are detected via the azide–alkyne click reaction and even to halogenated nucleotide analogues to produce multilabel marking of replicating DNA (
) reported a novel strategy that has revolutionized identification of BrdU incorporated into DNA. This novel strategy utilizes the Suzuki–Miyaura reaction to detect BrdU with fluorescent boronic acid probes instead of the traditionally used anti-BrdU antibodies.
Besides simple tracing of dividing cells and their progenies, several labeling schemes that combine pulse-chase and cumulative labeling with multilabel marking of replicating DNA have been developed for revealing specific modes of stem cell proliferative behavior and for evaluating key cell cycle parameters (Table 2). Application of such labeling schemes (see below) enables creating descriptive models for tissue histogenesis and renewal and stem cell maintenance, differentiation, and elimination.
Table 2Summary on assays that can be performed using single, double, and triple S-phase labeling with modified nucleotides
Single S-phase labeling
Double S-phase labeling
Triple S-phase labeling
Pulse-chase labeling with the chase period less than the average cell cycle length (
). Larvae and adults of marine invertebrates (sea urchins, sponges, flatworms, etc.) are considered permeable for most pharmacological agents. Therefore, to mark replicating DNA in these species, nucleotide analogues are dissolved at micro- and millimolar concentrations in the ambient seawater they are maintained in (
). To label replicating DNA in fish (zebrafish (Danio rerio)) or frogs (Xenopus laevis), larvae and even adults can also be bathed in ambient water supplemented with millimolar concentrations of a nucleotide analogue (
Thymidine analogues can be delivered into most vertebrate species through intraperitoneal, intravenous, or intramuscular injections, drinking water, and osmotic minipumps. Intraperitoneal injection is the easiest way to deliver a nucleotide analogue. Nucleotide analogues resemble natural thymidine; therefore they readily absorb into the blood stream after being injected intraperitoneally, spread broadly in the body through the blood circulation system, and penetrate virtually all organs and tissues of an organism, including those separated by barriers (brain, testis, placenta). Intraperitoneal injections are usually used in small animals, such as rodents (
Treatment with a thymidine analogue (usually BrdU) dissolved in drinking water is frequently used in rodent studies and is employed when long-term labeling of dividing cells is necessary. The concentration of BrdU in drinking water is typically 0.8 to 1 mg/ml (
). To overcome aversion to the taste of BrdU and to enhance intake of BrdU containing water to elevate labeling of dividing cells, drinking water is frequently supplemented with sucrose or orange juice (
). This route of a nucleotide analogue delivery is used when distress evoked by daily intraperitoneal injections perturbs the resultant experimental data, or high animal mortality is observed. However, due to the circadian dependence of water intake, treatment with BrdU dissolved in drinking water marks different numbers of dividing cells during the light and dark phases of the day (
Independently from the route of delivery, the thymidine analogue dosage is of great importance when labeled cells are evaluated quantitatively and comparison between the experimental and control groups is necessary. We must ensure that changes in the number of labeled cells originate from alterations in proliferation, not from alterations in the nucleotide analogue uptake. A saturating dose, which is defined as the dose of a nucleotide analogue necessary to label most cells in the S-phase, satisfies this condition. The saturating dose of a nucleotide analogue depends on the species used, the organ studied, and the life stage analyzed. Theoretically, the saturating dose must be determined for each individual experimental condition. An accurate determination of saturating doses for BrdU and EdU has been reported for dividing cells in the hippocampal dentate gyrus of adult rodents (
). Interest in the hippocampal dentate gyrus in regard to determining the saturating dose of a nucleotide analogue is due to the following. The hippocampal dentate gyrus is a spatially limited structure. In the dentate gyrus, dividing cells do not extensively migrate. They do not form very dense clusters, allowing for easy discrimination and counting of individual labeled nuclei using optical microscopy. Therefore, quantitative evaluations of proliferating cells in the whole dentate gyrus of animals of the same age are in a good agreement between many research groups (
To evaluate the saturating dose practically, animals separated into several experimental groups receive single intraperitoneal injections of various doses of a nucleotide analogue. The doses tested are usually in the range of 10 to 600 mg/kg body weight. Then, numbers of labeled cells are counted, and the dependence of cell counts on thymidine analogue dose is determined. Typically, the dependence is represented by an initial raise in the number of labeled cells along with the increase in the thymidine analogue dose, followed by a plateau where dose elevation does not increase the number of labeled cells (
)). Current estimations of the saturating dose of BrdU determined by quantitation of labeled cells in the hippocampal dentate gyrus after a single intraperitoneal delivery are 150 mg/kg body weight in mice (
) determined 50 mg/kg body weight as the saturating dose of BrdU for labeling dividing cells residing in the walls of the lateral ventricles of a songbird zebra finch. Lower doses result in underestimation of the proliferating cell number, whereas higher doses provide the same number of the labeled cells as the saturating dose.
) reported determination of the BrdU saturating dose for the optimal labeling of dividing cells in the rat forestomach. The thymidine analogue was delivered continuously for 2 days through the subcutaneous implementation of flat-faced cylindrical matrices containing BrdU. A 200 mg dose (or on average 540 mg/kg body weight for 2 days) was found to label most dividing cells in the rat forestomach. The saturating dose of the rest of the halogenated nucleotide analogues remains to be determined. Although the saturating doses of BrdU have been accurately determined, most studies reported the use of the standard dose of BrdU, 50 to 100 mg/kg body weight in rodents (Table 1) (
). The doses within this range enable labeling of 60 to 90% of the proliferating cells that would be detected by a single delivery of the saturating dose. Accurate measurements have validated partial labeling by the standard doses for quantitative analysis and comparisons between experimental groups (
). This difference in the labeling level between BrdU and EdU seems to originate from the method of detection rather than from individual distinctions in labeling capacity of the thymidine analogues. BrdU was detected using immunohistochemistry combined with DAB (3,3′-Diaminobenzidine)-staining (
), whereas EdU was detected using the fluorogenic click reaction. Even suggesting equal sensitivity of both detection methods, the transmitted light microscopy required for observing DAB-stained samples provides lower contrast than fluorescent microscopy. There is a high risk of skipping slightly labeled nuclei when counting cells in DAB-stained samples. Therefore, underestimation of the proliferating cell number observed in DAB-stained samples can originate from quantification of labeled cells using transmitted light microscopy. To fill this gap, it is necessary to reevaluate the BrdU saturating dose using a fluorescently tagged antibody.
In sum, the observations mentioned above exhibit that dosage of a nucleotide analogue is a challenging problem, and preliminary tests are strongly recommended to elucidate behavior of the label under specific experimental circumstances. Importantly, dosage of a nucleotide analogue is also a compromise between labeling efficiency and adverse effects, such as overall cytotoxicity and cell cycle arrest (see below).
Detection of nucleotide analogues by antibodies and click reaction
Several excellent reviews and detailed protocols have already described staining procedures for revealing thymidine analogues in diverse biological samples (
). Here, we will delineate a workflow and briefly discuss principal stages for the detection of the most frequently used nucleotide analogues using antibodies and the click reaction.
Independently of biological sample type, the halogenated nucleotide analogues (BrdU, CldU, and IdU) necessitate DNA denaturation prior antibody detection (Table 1). The most frequently used method of DNA denaturation is the immersion of a sample into a 0.5 N–4 N hydrochloric acid solution. Solution temperature (usually 37 °C) and immersion time (usually 10–60 min) can be adjusted to obtain desirable staining quality. The drawbacks of acidic DNA denaturing and ways to bypass these drawbacks have been mentioned above (see also (
). This procedure is desirable when processing whole organisms or tissue sections because it partially reverses the wrinkling and shrinking of the sample evoked by hydrochloric acid exposure.
Revealing halogenated nucleotide analogues incorporated into DNA employs standard immunohistochemical (DAB) or immunofluorescent (a fluorescent tag) staining procedures, which include permeabilization, blocking nonspecific secondary antibody labeling, exposure to the primary and secondary antibodies, and signal amplification steps. Description of these procedures can be found elsewhere (
). To detect halogenated nucleotide analogues, numerous antibodies produced in various hosts are available from diverse commercial sources. This provides great flexibility in the concurrent detection of halogenated nucleotide analogues and various biomolecules in the same specimen using different labeling techniques. However, the level of BrdU labeling has also been found to strongly depend on the primary antibody used for BrdU detection both in terms of labeled cell numbers and in terms of fluorescence intensity (brightness of labeled cells) (
). Both reports have unambiguously demonstrated that primary anti-BrdU antibodies originating from distinct sources are not equally sensitive to nucleotide analogues. Most primary anti-BrdU antibodies have been found to cross-react to EdU, excluding mouse monoclonal anti-BrdU antibody (clone MoBU1) (
). Mouse monoclonal anti-BrdU antibody (clone B44), which is sensitive to BrdU and IdU, exhibited reduced cross-reactivity to CldU, whereas rat monoclonal anti-BrdU antibody (clone BU1/75) reacts with both BrdU and CldU and is insensitive to IdU (
). The detection of EdU is much easier than the detection of halogenated nucleotide analogues. Permeabilization is the only procedure required prior to EdU detection by the click reaction. The covalent binding of a fluorescent azide to a terminal alkyne group in the EdU residue requires the presence of monovalent copper ions. The click reaction mixture is supplemented with a reducing agent, usually ascorbic acid or its sodium salt (+)-Sodium L-ascorbate to obtain monovalent copper ions from divalent copper ions, which are usually obtained by dissolving copper-containing salts such as CuSO4 (
). To this end, live cells or acutely prepared tissue samples pretreated with EdU are administrated with a staining solution supplemented with CuSO4, ascorbic acid, and cell-membrane-permeable tetramethylrhodamine azide. However, Cu(I) ions are highly toxic, and therefore the cells do not survive staining. Although live microscopy of EdU-labeled and azide-stained cells is of limited utility, it may be useful in those cases where subsequent molecular assays are not compatible with formaldehyde fixation and/or permeabilization of cell membranes, or when they must be performed without removal of the samples from the microscope stage to overlap other experimental readouts with the EdU labeling (
Detection of both halogenated nucleotide analogues and EdU is accompanied with the exposure of a biological specimen to highly chemically active substances such as hydrochloric acid or monovalent copper ions, which can affect the biochemical properties of macromolecules and the integrity of cellular components. Therefore, if the concurrent detection of other macromolecules within the same specimen is necessary, the protocol for thymidine analogue detection must be adjusted for the specific experiment to diminish the negative effects of the staining procedures on the other components to be detected.
Critical points for the concurrent detection of several nucleotide analogues within the same sample
In the brief overview on marking replicating DNA, we have already mentioned that detection of truly double labeled nuclei may be compromised when employing the double labeling technique using BrdU and 3H-thymidine. Moreover, this double labeling technique is not compatible with current fluorescence microscopy, such as confocal or light-sheet microscopy. This limitation narrows the range of research issues that can be addressed by the double nucleotide labeling with BrdU and 3H-thymidine. Therefore, we will focus here on the current double and triple labeling techniques that exploit antibodies and bioorthogonal chemical reactions for detection of nucleotide analogues.
Double labeling using CldU and IdU was initially presented in 1992 (
). This double labeling method is based on the different sensitivity of two monoclonal antibodies against BrdU. One antibody, rat anti-BrdU antibody clone BU1/75, recognizes both BrdU and CldU, but exhibits low binding to IdU. The other antibody, mouse anti-BrdU antibody clone B44, recognizes BrdU and IdU, while displaying low binding to CldU. Procedures such as sequential application of the primary antibodies, washing in Tris-buffered saline with a high salt concentration, and determination of appropriate antibody dilutions have been proposed to remove residual cross-reactivity of rat anti-BrdU antibody clone BU1/75 to IdU and mouse anti-BrdU antibody clone B44 to CldU in both in vitro and in vivo examinations (
). Detection of a noncognate nucleotide analogue by these antibodies can also originate from nonspecific binding of the secondary antibodies that imperfectly discriminate between mouse and rat immunoglobulins. Therefore, the use of the secondary antibodies that have been additionally cross-adsorbed to the respective immunoglobulins is strongly recommended (
Detection of yet another pair, BrdU and EdU, lacks any residual cross talk, thus enabling unequivocal discrimination between truly double and single labeled cells. Initially, ten different anti-BrdU antibodies from various commercial sources were examined for their reactivity to EdU (
). Therefore, the delivery of EdU and BrdU followed by their detection via the click reaction and the application of the mouse monoclonal anti-BrdU antibody clone MoBU1 is a reliable double labeling method. Interestingly, when cultured cells labeled with EdU undergo staining with anti-BrdU antibodies, the signal of most anti-BrdU antibodies (except clone MoBU1) is not completely removed even by prolonged click reaction or the click reaction with an elevated fluorescent azide concentration (
). Among the examined nonfluorescent azides, azidomethyl phenyl sulfide was found to completely remove the signal of anti-BrdU antibodies at a low concentration of 2 mM. Therefore, the application of the click reaction with azidomethyl phenyl sulfide (the second click reaction) after the fluorogenic click reaction (the first click reaction) and prior to the anti-BrdU antibody staining is a reliable alternative approach for the concurrent detection of EdU and BrdU.
Recently, we reported triple S-phase labeling of dividing cells in vivo and the protocol for concurrent detection of CldU, IdU, and EdU (
). The reported protocol combines the detection of CldU and IdU via application of the rat anti-BrdU antibody clone BU1/75 and the mouse anti-BrdU antibody clone B44 with inhibition of antibody binding to EdU using a second click reaction with azidomethyl phenyl sulfide (Fig. 1). In our study, we validated the method for triple S-phase labeling qualitatively and quantitatively. The complete protocol for the detection of the three nucleotide analogues was applied to brain sections of mice that received a single injection of an individual nucleotide analogue. Microscopic analysis confirmed that the complete protocol specifically detects individual nucleotide analogues in these mice without any cross-reactivity. Quantitative analysis of labeled cells in the hippocampi of mice that received sequential injections of all three thymidine analogues at fixed time intervals revealed full agreement with the predicted parameters of triple labeling in a system with known cell cycle kinetics. Moreover, this quantitative analysis also confirmed equality in the labeling capacity of the examined thymidine analogues when delivered at equimolar doses. The importance of equimolar delivery was indicated for maintenance of the quantitative relationship between cell populations incorporating two labels at different time points (
Another study reported a novel approach for the concurrent detection of three other thymidine analogues, F-ara-EdU, VdU, and BrdU, using two nonoverlapped bioorthogonal chemical reactions and antibodies against BrdU (
). Applications of the nucleotide analogues that are detected via bioorthogonal chemical reactions are described below.
Labeling schemes and respective readouts for revealing stem cell proliferative behaviors and fates
“Pulse labeling” (or “a pulse dose”) can be defined as labeling dividing cells when a thymidine analogue is available for incorporation into replicating DNA within a time interval that is shorter than the average cell cycle duration of a given cell population. On average, the cell cycle of most eukaryotic cells lasts for several hours. Therefore, treating cultured cells, or bathing larvae or adult organisms with a thymidine analogue for several hours or less may be considered as pulse labeling. A single intraperitoneal or intravenous injection of a thymidine analogue may also be considered as pulse labeling because of its transient availability in the blood (the bioavailability of modified nucleotides is described below). Pulse labeling is followed by a “chase” period when the nucleotide analogue is not delivered. The chase period in this labeling scheme may be varied in a wide range, from several minutes to months and even years, depending on purposes of the study (
) (Fig. 2A). If the chase period after pulse labeling is less than the average cell cycle length of a given cell population, cells that have incorporated the label are considered to be dividing cells (cells passing the cell cycle), and quantification of these cells allows for estimation of proliferative activity (proliferation assay). Extension of the chase period enables the progeny of cells that were passing the S-phase of the cell cycle at the time point when a single nucleotide analogue was delivered to be tracked. Tracking the progeny of dividing cells combined with identifying differentiation markers allows the fates of cells generated at a certain time to be determined. Therefore, this labeling scheme is extensively used in developmental biology and stem cell research. Pulse-chase labeling with two and even three nucleotide analogue species allows for the discrimination of cells born at distinct times, thus significantly increasing the resolution of the cell fate analysis (
) (Fig. 2A). Moreover, double or triple S-phase labeling offers unmatched flexibility in experimental design and reduces the number of experimental groups, thus drastically facilitating the workflow.
“Window” labeling for tracking cells after terminal mitosis
One of the major disadvantages of the pulse-chase labeling scheme is that this type of labeling does not enable judgment about what happens to the labeled cells during the chase period. Pulse-chase labeling does not reveal whether the labeled cells undergo additional rounds of division or exit the cell cycle soon after having been labeled. Although there were attempts to discriminate between heavily labeled cells that completed their division and faintly labeled cells that underwent additional rounds of division, these data are considered doubtful. “Window” labeling overcomes this problem. It is a variation of double S-phase labeling and combines “pulse” and “cumulative” labeling (
). In this labeling scheme, the first nucleotide analogue is administrated by continuous delivery that lasts no longer than average duration of a single cell cycle in an examined cell population (Fig. 2B). Immediately after completion of the first nucleotide analogue delivery, the second nucleotide analogue is continuously delivered throughout the rest of the experiment. Cells that incorporated the first label, but not the second, underwent terminal mitosis within the time interval between the beginning of first label administration and the onset of second label exposure (Fig. 2B). Thus, this labeling scheme enables the fate of those cells that underwent terminal mitosis at a specific time window to be traced. This approach offers unmatched opportunities for tracking differentiation, migration, and survival of postmitotic stem cell progeny.
Many subsets of somatic stem cells within adult mammalian organisms are characterized by a prolonged mitotically inactive state (G0-phase). This state, frequently referred to as quiescence or dormancy, is reversible, and quiescent stem cells intermittently enter the cell cycle. Such behavior is characteristic for hematopoietic, muscular, and hair follicle stem cells (
). It has also been demonstrated that the intestine, the tissue with the most intensive self-renewal, harbors a subset of the so called “+4 stem cells,” which undergo infrequent divisions to fuel a pool of rapidly dividing stem cells in intestinal crypts (
). Such infrequently dividing somatic stem cells have been identified using the label-retaining assay. The label-retaining assay resembles pulse-chase labeling, but cells usually undergo marking over prolonged period via continuous delivery of a nucleotide analogue instead of pulse labeling (Fig. 2C). During the chase period in the absence of the label, the rapidly dividing cells dilute the incorporated label to an undetectable level by repeated cycles of the DNA replication and its distribution between daughter cells. While the rapidly dividing cells dilute the incorporated label, the infrequently dividing cells retain the label over a long period. Therefore, this population of cells is referred as label-retaining cells (LRCs). Although LRCs can be observed when pulse-chase labeling has been conducted, a prolonged labeling instead of pulse labeling is necessary to mark as many as possible infrequently dividing stem cells, which usually represent a minor population.
Labeling schemes and respective readouts for revealing key parameters of cell cycle kinetics
Preconditions for applying labeling schemes
The labeling schemes described below allow for determination of key parameters of cell cycle kinetics at the population level. Therefore, application of these labeling schemes necessitates several preliminary assumptions important for valid interpretation of the results. First, the examined proliferative cell population must be homogeneous in terms of the S-phase and entire cell cycle durations. Second, the cells in the population must be randomly distributed throughout all phases of the cell cycle, i.e., the cell population must be asynchronous. Third, the proliferative cell population must be in a steady-state growth phase, i.e., the number of cells that are currently in the cell cycle does not change significantly during the experiment. Fourth, the proliferative cell population must be spatially confined, i.e., cells should not extensively migrate outside of the site of their initial location. Fifth, cell death events are rare and can be neglected. The last two requirements are necessary for the valid quantification of labeled cells.
Cumulative labeling for determining the cell cycle and S-phase durations
Cumulative labeling is extensively applied for determining key parameters of the cell cycle and is conducted via either continuous nucleotide analogue delivery (usually for in vitro labeling) (
). Cumulative labeling is aimed to mark all cells of the proliferative population and to track the kinetics of label incorporation into cells passing the cell cycle (Fig. 3A). In this labeling scheme, a nucleotide analogue is available for incorporation into replicating DNA throughout the duration of the experiment. Subsets of specimens are periodically collected for counting the cells that have incorporated the label. Then, the dependence of cell counts from the time intervals between the beginning of label delivery and collection of the specimens for analysis is calculated. This dependence linearly increases with the lengthening of time intervals between the beginning of label delivery and collection of the specimens for analysis. The increase occurs because some cells have left S-phase, remaining labeled, while other cells have entered S-phase and therefore, have de novo incorporated the label. However, the cells that have already incorporated the label at the beginning of nucleotide analogue exposure eventually re-enter the S-phase. Therefore, all cells circulating in the cell cycle have been marked with a nucleotide analogue, and the number of labeled cells stops increasing, reaching a plateau (maximal value). An intercept between the linear regression lines of the slope and the plateau enables determination of the time interval (Δt) equal to Tcell cycle−TS-phase. If we normalize cell counts to the maximal value, an intercept between the y-axis and the continuation of the slope (y0) enables estimation of the ratio TS-phase/Tcell cycle. This enables creating an equation system:
where Δt and y0 are the parameters determined in Figure 3A, TS-phase is the S-phase duration, and Tcell cycle is the cell cycle duration. Therefore, solving the equation system (Equation 1) enables determination of the S-phase duration and the cell cycle duration.
Cumulative labeling has extended the analysis of cell proliferation beyond the routine determination of the dividing cell numbers and allowed for measurement of cell cycle kinetics in diverse experimental situations (
). It should be noted that the linear increase of the dependence indicates that the proliferative cell population satisfies the preliminary assumptions. If the increase in the dependence is not linear, the examined proliferative cell population is either not homogeneous or is not asynchronous, or both and the determination of key cell cycle parameters using cumulative labeling is impossible.
Percent labeled mitoses method for comprehensive analysis of the cell cycle kinetics
It may seem that the pulse-chase labeling method mentioned above provides very limited information regarding cell division; however, its variation, referred to as the percent labeled mitoses method, enables measurement of the duration of each cell cycle phase (Fig. 3B) (
). Pulse labeling marks a cohort of cells that are in the S-phase. Progressing through the cell cycle, this cohort enters mitosis. The percent of labeled mitotic figures starts growing, reaching a plateau at 100%. The percent of labeled mitotic figures will remain 100% until all labeled cells pass. This percentage starts decreasing when unlabeled cells reach the M-phase. The percent of labeled mitotic figures equals 0% again when all labeled cells complete mitotic division. The next rise of the percent of labeled mitotic figures occurs when labeled cells enter mitosis again. The time interval between pulse labeling and the appearance of the first labeled mitotic figure corresponds the duration of the G2-phase. The time interval when the percent of labeled mitotic figures grows from 0 to 100% is the duration of the M-phase. The time interval between time points where the percent of labeled mitotic figures initially increases to 100% and then reduces to 0% again is the duration of the S-phase. The time interval between two subsequent rises of the percent of labeled mitotic figures corresponds to the entire duration of the cell cycle.
Similarly, the distribution of cycling cells between cell cycle phases can be determined by pulse labeling combined with the detection of the proliferating cell markers Ki67 or the proliferating cell nuclear antigen (PCNA) and mitotic marker, phosphohistone H3 (
). The proportion of cells that have incorporated both labels declines progressively as the interval between the labels lengthens because the cells marked with the first label leave the S-phase, and unlabeled cells enter the S-phase and become marked with the second label only. The intercept of the declination line with the time axis (Fig. 3C) reveals the time point when all cells marked with the first label exit the S-phase, giving an estimation of the S-phase duration. When we further lengthen the time interval between the two labels, the proportion of the double labeled cells starts growing, reflecting the entry of cells that have incorporated the first label into the next S-phase. Then, the proportion declines again. The time point when the peak is reached corresponds to the time interval between two consecutive S-phases or, in other words, the cell cycle duration.
If the S-phase and cell cycle durations are known approximately for a given proliferative population, simplified double labeling can be applied for estimation of the S-phase and cell cycle lengths (
). In this labeling scheme, one group of animals receive two labels separated by a time interval shorter than the S-phase duration, and subsequent cell quantifications enable the calculation of the S-phase length. Another group of animals receive two labels separated by a time interval longer than the difference between the cell cycle length and the S-phase duration, and subsequent cell quantifications enable the calculation of the cell cycle length. This approach can be applied when we need to test whether any physiological or pathological stimulus alters the cell cycle parameters of a given population of proliferating cells (
Double and triple S-phase labeling schemes for discrimination between proliferating and quiescent subpopulations of daughter cells
Generally, there are three possible fates of daughter cells after mitosis of stem and progenitor cells. Both daughter cells remain in the cell cycle, both daughter cells exit the cell cycle, or one cell re-enters the cell cycle and the other becomes mitotically quiescent. The ratio between re-entering and exiting the cell cycle defines the kinetics of cyto- and histogenesis during normal development or physiological tissue renewal in adults. If the fraction of daughter cells that re-enter the cell cycle exceeds that of daughter cells leaving the cell cycle, the proliferative population is expanding; otherwise the proliferative population is becoming exhausted. Determination of these fractions is critical for creating kinetic models of cyto- and histogenesis (
) have suggested a double labeling scheme to trace a limited cohort of cells after mitosis in terms of their proliferative fates (Fig. 4A). In this labeling scheme, two pulse labels are delivered at a time interval (Δt1−2) shorter than the average S-phase duration in the cell population studied. The cells that incorporate the first label leave the S-phase prior to delivery of the second label, becoming a cohort of labeled cells with known position within the cell cycle. Then, one group of samples (animals or cell cultures) is collected for analysis at a time point (t) after the second label delivery (Path A in Fig. 4A) that meets the following criteria:
where TS-phase is the duration of the S-phase, Tcell cycle is the cell cycle duration, and Δt1−2 is a time interval between delivery of two labels. In parallel, another experimental group receives additional deliveries of the second label followed by analysis at the same time point (Path B in Fig. 4A). The cells that have incorporated the first label but not the second one pass mitosis and a portion of them enter the S-phase of the next cell cycle within the time interval determined by Equation 2. In the first experimental group (Path A in Fig. 4A), the population of cells that have incorporated the first label includes both the proliferative and quiescent subpopulation of daughter cells. Due to the availability of the second label during the time interval determined by Equation 2 in the second experimental group (Path B in Fig. 4A), cells that have initially incorporated the first label become double labeled when they enter the subsequent S-phase. Therefore, in this group, cells that incorporated only the first label belong to the subpopulation of mitotically quiescent progenies. In this case, the quiescent fraction (Q) is calculated as the ratio:
where is the number of cells with the first label only in the second group (Path B in Fig. 4A), and is the number of cells with the first label only in the first group (Path A in Fig. 4A). The proliferative fraction (P) is calculated as:
The suggested methodology, however, requires analysis of two experimental groups. Recently, we presented an improved protocol for the determination of cell fractions that re-enter or exit the cell cycle (Fig. 4B) (
). This protocol is based on triple labeling and does not require separate experimental groups. Similar to the previous labeling scheme, a cohort of cells with a known position in the cell cycle is highlighted by two pulse labels separated by a time interval (Δt1−2) shorter than the average S-phase duration. Then, the third label is delivered at a time point (t3) after delivery of the second label that meets the following criteria:
where TS-phase is the duration of the S-phase, Tcell cycle is the cell cycle duration, and Δt1−2 is a time interval between delivery of two labels (the first and the second). In this labeling scheme, the cohort of cells with the first label only is the quiescent fraction Q, whereas the cohort of cells that incorporated the first and the third label but not the second one is the proliferative fraction P.
Notably, both methodologies necessitate determination of the S-phase duration and the cell cycle length.
Caveats to consider when applying labeling with modified nucleotides
Remarks for application of pulse chase and cumulative labeling
Pulse-chase labeling with a short chase period is the simplest labeling scheme that is commonly used to address the question of whether an examined stimulus elicits changes in proliferative activity of a certain cell population in vivo or in vitro (proliferation assay). Comparison of labeled cell numbers between experimental (exposed to a stimulus) and control (nonexposed to a stimulus) groups is mainly interpreted in terms of increased/decreased cell proliferation. However, despite its simplicity, pulse-chase labeling can produce compromised results in certain cases. For instance, it might be assumed that, for a certain cell population, a stimulus elicits changes in duration of the individual cell cycle phases with no effect on overall number of cells circulating in the cell cycle. In this case, proliferation assay performed using pulse-chase labeling will reveal an increase/a decrease in the number of labeled cells that is proportional to a stimulus-induced increase/decrease in the ratio of the S-phase duration to the cell cycle length. Therefore, the observed differences in labeled cell numbers in experimental and control groups cannot be interpreted in terms of changes in proliferative activity. This hypothetical situation demonstrates ambiguousness of data interpretation after pulse-chase labeling and imposes necessity on conducting proliferation assay in a combination with the evaluation of the cell cycle parameters to gain valid labeling results.
Cumulative labeling via the repeated pulse delivery of a label is frequently used to mark all cells in the proliferative population or a large cohort of dividing cells to enhance accuracy of quantitative readouts. In most studies, a choice of time intervals between the repeated pulse deliveries of a label is random and reasonless. To perform cumulative labeling rationally, preliminary estimation of the S-phase duration and the cell cycle length is necessary. Evidently, there is no reason to deliver pulse labels at time intervals shorter than the S-phase duration because some cells that have already incorporated a label are remaining in the S-phase and, therefore, receive additional doses of a label that can exert the cytotoxic effect (see below) in labeled cells. Hence, time intervals near the S-phase duration enable labeling distinct cohorts of cells within the same proliferative population, and, moreover, labeled cells in these cohorts receive a single dose of a label. For instance, neural progenitors in the hippocampal dentate gyrus of the adult mouse brain represent a relatively homogeneous cell population with the approximate S-phase duration of 10 to 13 h and the approximate cell cycle length of 23 to 28 h (
). Hence, two pulse label deliveries at the 12 h interval will enable marking almost 90% of the proliferative population with low risk of implementation of two label doses into the same cells.
These speculations indicate that application of any labeling scheme and interpretation of labeling results should be based on the preliminary determined or at least hypothetical cell cycle kinetics of the examined cell population. Otherwise, compromised results or misused labeling can be expected.
Bioavailability of modified nucleotides
When we label cultured cells with a thymidine analogue in a Petri dish, we are able to precisely control the time interval during which the label incorporates into newly synthesized DNA. The label incorporation can be interrupted by withdrawal of the thymidine analogue via simple washing. Optionally, cells can be washed followed by incubation with culture medium supplemented with regular thymidine. Regular thymidine competes with the residual thymidine analogue, thus blocking its incorporation into the replicating DNA. Therefore, loading of the label into cell culture can be as short as 1 min, if necessary. The same scenario of interruption of label incorporation can be applied to marine invertebrate species. In sharp contrast to cultured cells and marine invertebrates, incorporation of a thymidine analogue into the replicating DNA after a single pulse delivery (via either an intraperitoneal injection or an intravenous injection) into vertebrate species cannot be precisely controlled and is determined by the bioavailability of the label. Bioavailability is an integrative concept that describes both the fraction of a drug that reaches its site of action and the rate at which the drug becomes available at its site of action. Here, we will primarily refer to bioavailability time, which is determined as the time interval during which the thymidine analogue is completely metabolized by an organism after a single pulse labeling (an intraperitoneal injection or an intravenous injection). Beyond this time interval, the thymidine analogue is no longer available for incorporation into replicating DNA. The bioavailability time of a thymidine analogue depends on the dose injected, the species used, the stage development of the organism, diffusion into the blood circulatory system from the peritoneal cavity or digestive system (if a thymidine analogue is delivered nonintravenously), distribution with the blood stream, penetration into tissues (in particular, penetration through various blood-tissue barriers), and active pyrimidine transport in the cells.
3H-thymidine, which is a counterpart to natural thymidine, has been found to rapidly eliminate from the blood plasma after intravenous injection in rodents, monkeys, and humans (
). This elimination occurs via two phases: (i) a rapid phase with an approximate half-time of 1 min and (ii) a slow phase with an approximate half-time varying from 10 to 20 min depending on the species (
). Minute amounts of radiolabeled thymidine remained detectable in the blood plasma up to 60 min after intravenous injection. Autoradiographic analysis of blood cell concentrates or tumor tissue sections revealed that the bulk of 3H-thymidine was incorporated into DNA within 15 to 25 min after a pulse dose and that the uptake of 3H-thymidine achieved a plateau at approximately 40 to 60 min (
Only a portion of the injected BrdU incorporates into the replicating DNA after a single intravenous injection. The bulk of the injected BrdU is rapidly degraded with the formation of bromouracil and bromide ions (
). Similarly, the bioavailability time of BrdU estimated by measuring BrdU content by high-performance liquid chromatography assay in blood serum after an intravenous injection in dogs was approximately 2 h (
). When radiolabeled BrdU or 3H-thymidine was intraperitoneally injected into pregnant mice, the half-life of the nucleotide analogues in the embryos was found to be longer than in various maternal tissues, 60 to 80 min versus 30 min, respectively. Thus, the longer bioavailability time of the thymidine analogues must be taken into account when labeling dividing cells in embryos of mammalian species.
The bioavailability time for the rest of the nucleotide analogues used for labeling replicating DNA has not yet been evaluated. Our indirect observations on the progression of hippocampal neural progenitors through the cell cycle using triple S-phase labeling suggest that the bioavailability time of CldU, IdU, and EdU is shorter than 2 h (
). We sequentially injected mice with the three labels with 2-h intervals and counted the cells that incorporated the labels. The observed labeling pattern (numbers of cells with definite combinations of the labels) was consistent with the hypothetical linear progression of neural progenitors through the cell cycle (see Fig. 2, A–C in (
)). We repeated the experiment, injecting labels in a different order, and observed that the labeling pattern was independent of the order of label delivery (unpublished data). If there was at least one nucleotide analogue with a bioavailability time over 2 h, the observed labeling pattern should have differed from the expected pattern.
Bioavailability of a nucleotide analogue can be neglected in the following in vivo labeling situations: (i) pulse-chase labeling where the chase period is quite long (longer than a day) and (ii) prolonged (longer that a day) cumulative labeling when a thymidine analogue is delivered with drinking water, by repeated injections, or via an implanted osmotic pump. However, bioavailability must be always taken into consideration when the S-phase or cell cycle kinetics is analyzed using pulse-chase labeling (
) labeling. Inadequate labeling may occur when the time intervals between label pulses or between label pulses and euthanasia do not exceed the bioavailability time of thymidine analogues.
Cytotoxicity of modified nucleotides
Cytotoxicity of modified nucleotides is a serious pitfall for the study of cell proliferation and retrospective birth dating of cells. Toxic manifestation of modified nucleotides depends on the dose used, administration technique (single, multiple, or continuous administration), and time elapsed after treatment.
Toxic effects of the most widely used nucleotide analogue BrdU were examined in various experimental situations. BrdU was initially demonstrated to affect growth of mammalian cancer cells (
). It has been demonstrated that transient exposure of various cancer cell lines to BrdU at a dose of 1 μM or higher suppresses the rate of cell expansion. This effect becomes detectable within several days of exposure. However, BrdU-treated cells do not die. Instead, they alter the timing of the cell cycle, primarily increasing the duration of the G1 and G2 phases, and upregulate senescent-associated proteins. Both intraperitoneal injections of BrdU (six pulse doses 300 mg/kg within 2 days) and oral delivery of BrdU (0.8 mg/ml, for 7 days) suppressed progression of grafted tumors in rats (
). However, their toxic effects were primarily studied in the context of anticancer therapy and have not been yet evaluated in the context of labeling dividing cells. Therefore, it is difficult to judge whether they are more or less toxic than BrdU.
Virtually all studies that have addressed adverse effects of EdU indicate that this substance is highly toxic. The toxic effect of EdU was found to be more pronounced than the toxic effect caused by an equimolar dose of BrdU in various cells both in vitro and in vivo (