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Telomerase Activity Is Elevated in Early S Phase in Hamster Cells
BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS ARTICLE NO.
233, 717–722 (1997)
RC976549
Telomerase Activity Is Elevated in Early S Phase in Hamster Cells Patricia A. Kruk,*,†,1 David K. Orren,† and Vilhelm A. Bohr† *Department of Pathology, University of South Florida, Tampa, Florida 33612; and †Laboratory of Molecular Genetics, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224
Received March 26, 1997
Telomerase is a ribonucleoprotein that elongates telomeric repeats de novo. We examined the possibility that telomerase activity is cell cycle regulated by examining telomerase activity in cell cycle synchronized Chinese hamster ovary (CHO) B11 cells. Overall telomerase activity was similar in growing and quiescent cells. Further, cells synchronized in G1 , S, or G2/M showed similar levels of telomerase activity. However, a detailed analysis of cells within S phase showed that there was a higher level of telomerase activity in early S phase when compared with other points in the cell cycle. These results suggest a relationship between telomerase activity and cell cycle regulation. q 1997 Academic Press
Telomeres consist of tandem hexanucleotide (TTAGGG)n repeats that cap the ends of vertebrate chromosomes. Telomeres are essential for chromosomal stability and integrity. They protect chromosomes from illegitimate recombination, degradation by nuclease attack, and solve the end replication problem (1,2). Telomeric repeats are synthesized by telomerase, a ribonucleoprotein that elongates telomeric repeats de novo (2). Telomerase activity is generally detected in germ cells, cancer cells, and cell lines, but only occasionally in somatic cells (3-7). Very little is known about the regulation of telomerase activity in mammalian cells. There have been reports of telomerase activation associated with cellular proliferation (5,8) and tumor progression (3,5) as well as telomerase repression associated with cellular differentiation (8,9) and cellular quiescence (8). A relationship exists, then, between telomerase activity and the end replication problem as well as an association between telomerase activity and high cellular proliferative capacity. In addition, the re1 Corresponding author at Department of Pathology, MDC 11, University of South Florida, 12901 Bruce B. Downs Blvd., Tampa, FL 33612-4799. Fax: (813) 974-5536. E-mail: [email protected].
port that G-rich telomeric overhangs appear in late S phase in Saccharomyces cerevisiae (10) would suggest that telomerase activity is not only associated with cellular proliferation, but that the ability to synthesize telomeric repeats may be regulated in a cell cycle dependent manner. Yet information on the regulation of telomerase activity as it relates to cellular proliferation and, in particular, the cell cycle is sparse and often contradictory (8, 11-13). We initiated studies on telomerase activity as a function of cell growth and cell cycle stage in immortalized CHO B11 cells. A synchronization method developed previously (14) allowed us to measure telomerase activity in each cell cycle phase as well as at defined points within S phase. On the basis of equivalent cell numbers, we found no difference in telomerase activity between growing and quiescent cells, although the proportion of cellular protein comprising telomerase activity increased in quiescent CHO B11 cells. Though we observed telomerase activity in these cells throughout the cell cycle, our data indicate that there was a transient increase in telomerase activity in early S phase. These results indicate that telomerase is constitutively active in CHO B11 hamster cells, but may be additionally regulated in a cell cycle phase-specific manner. MATERIALS AND METHODS Cell cycle synchronization. CHO B11 cells were routinely grown as previously described (14) at 377C in 5% CO2 in complete medium consisting of Ham’s F12 medium (without glycine, hypoxanthine, and thymidine) supplemented with 10% dialyzed fetal bovine serum (dFBS). Cell cycle synchronized CHO B11 cells were obtained as described earlier (14). Briefly, G1 enriched cells were obtained following 48 hr incubation of exponentially growing CHO B11 cells in starvation medium. Starvation medium contained only 0.2% dFBS, but was otherwise as stated above. Cells arrested in early S phase were obtained by 48 hr serum starvation followed by a 14 hr incubation in complete medium plus 50 mM mimosine. Recent studies have shown that mimosine inhibits elongation at replication forks shortly after DNA synthesis has begun, probably by depletion of nucleotide pools (15). The effects of mimosine on replicative DNA synthesis are short-lived. After removal of the drug from the medium, mimosine 0006-291X/97 $25.00
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is rapidly cleared from the cells, and the cells subsequently proceed through S phase. The duration of S phase after mimosine release appears to be identical to the time that it normally takes to traverse S phase. Thus, mimosine appears to have no lingering effects on DNA metabolism making it an ideal drug for use in synchronization studies. Replacement of medium containing mimosine with fresh complete medium allowed the cells to pass synchronously through S phase and G2 phase (14). Cells synchronized at various points during the cell cycle were harvested for both flow cytometry and telomerase activity analyses. Flow cytometry. The cell cycle phase distribution of CHO B11 cells was determined by measuring the DNA content of individual cells by flow cytometry as described previously (14). Briefly, 1-5 1 106 cells were trypsinized, resuspended in complete medium, and washed in PBS. The cells were then resuspended, fixed in 5 ml cold ethanol (70%), and stored at 47C. Just prior to flow cytometric analysis, individual samples were treated with RNase then stained with propidium iodide according to instructions (and using reagents) provided with the Cellular DNA Flow Cytometric Analysis Reagent Set (Boehringer Mannheim). Propidium iodide-stained cell suspensions were then analyzed using a Becton-Dickinson FACScan. Histograms (relative fluorescence vs. cell number) were generated from the flow cytometric data using the Lysis II software. Telomerase assay. CHO B11 cells were assayed for telomerase activity as described previously (3,16). Cells were washed with PBS, trypsinized, counted, and spun by centrifugation at 100g. The pellets were resuspended in ice-cold wash buffer containing 10 mM HEPESKOH (pH 7.5)/1.5 mM MgCl2/10 mM KCl/1 mM DTT and spun at 10,000 1 g at 47C for 20 min. The cells were then resuspended at a density of 103 to 105 cells in 20 ml ice-cold lysis buffer containing 10 mM Tris-HCl (pH 7.5)/1 mM MgCl2/1 mM EGTA/ 0.1 mM PMSF/5 mM b-mercaptoethanol/0.5% CHAPS/ 10 % glycerol for 30 min at 47C. The lysates were then centrifuged at 100,000 1 g for 45 min at 47C. After centrifugation, the supernatants were stored at 0807C. The protein concentration of each extract was determined by the BCA protein assay (Pierce). Because protein content varies within cells with progression through the cell cycle, the level of telomerase activity was measured using volumes of cellular extracts equivalent to 103, 104 and 105 cells. However, in one set of experiments examining telomerase activity among cells in different growth states, telomerase activity was also measured using a serial dilution of volumes of extracts based on protein concentration (10, 1.0 and 0.1mg protein). Extracts were incubated with 20 mM Tris-HCl (pH 8.3)/1.5 mM MgCl2/63 mM KCl/0.005% Tween 20/1 mM EGTA/100 mM each dATP, dGTP, dTTP/50 mM dCTP/20 mCi 32P-dCTP/0.1 mg of upstream primer Tel I (5*-AATCCGTCGAGCAGAGTT-3*, Paragon Biotech)/1 mg of T4g32 protein (Boehringer Mannheim)/0.1 mg per ml BSA for 60 min at room temperature (RT). The reaction mixture was then heated to 907C to inactivate the telomerase activity. During this step, 0.1 mg of the downstream primer Tel II (5*-CCCTTACCCTTACCCTTACCCTAA-3*, Paragon Biotech) and 2 ml Taq polymerase (Boehringer Mannheim) were added to the reaction mixture and the reaction mixture was subjected to 30 PCR cycles at 947C for 30 sec, 507C for 30 sec, 727C for 90 sec. As a control, parallel reactions were pretreated with 10 mg/ml RNase A for 20 min at 377C to degrade the RNA component of telomerase and indicate telomerase specificity in the assay. Positive controls for telomerase activity consisted of cell extracts from the ovarian carcinoma cell line, CAOV3. PCR products were analyzed by electrophoresis in Tris-borate EDTA buffer on 8% polyacrylamide gels. The gels were dried and telomerase activity was analyzed using a PhosphoImager system from Molecular Dynamics. The signal intensity from each lane of the gel was defined by a rectangular boundary established around all visible bands in the lane of least activity. A rectangle of equivalent size and shape was used for quantifying the signal from the remaining lanes of the gels. Enzyme activity was expressed as the fold difference of the activity determined in the RNase-sensitive reactions products compared to the lane with the least activity. Extracts were considered negative for
FIG. 1. Telomerase activity in growing, confluent, and starved CHO B11 cells. Decreasing equivalent cell numbers of CHAPS extracted CHO B11 cells (105, 104, 103 cell number equivalents) from subconfluent exponentially growing cells (Sub), cells that had just reached confluence (Confl), and confluent cells maintained in a quiescent state for 5 days (5 Da) and 14 days (14 Da) were analyzed for telomerase activity as described in Materials and Methods. In control lanes (/), 105 cell equivalents were pre-treated with 10 mg/ml RNase A.
telomerase activity if no products were detected following prolonged exposure.
RESULTS AND DISCUSSION In the present study, we examined telomerase activity in immortalized, undifferentiated hamster cells in various growth states and different stages of the cell cycle. We compared telomerase activity in exponentially growing subconfluent CHO B11 cells, cells that had just reached confluence, and confluent cells maintained in a quiescent state for 5 and 14 days respectively. Telomerase activity was demonstrated as the characteristic ladder pattern in cellular extracts from growing cells (Fig. 1). Serum deprivation of CHO B11 cells for 5 and 14 days post confluence, which results in growth inhibition and withdrawal from the cell cycle, did not reduce or abolish telomerase activity as deter-
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Comparison of Cell Number and Protein Content in Growing and Quiescent CHO B11 Cells Growth state
Cell number (1107)
Total protein (mg)
ng Protein/cell
Subconfluent Confluent 5 day starvation
1.188 3.28 3.16
0.722 1.881 1.354
607.7 573.5 428.6
mined from volumes of cellular extracts corresponding to an equivalent number of cells (Fig. 1). Further, there was no ascertainable difference in the amount of telomerase activity in growing and quiescent cells even upon serial dilution of these cellular extracts (Fig. 1). Pretreatment of the extracts with RNase abolished the telomerase activity, indicating telomerase specificity in this assay. Since induction of cellular quiescence by serum starvation did not affect telomerase activity, we sought to determine whether serum starvation altered cellular protein levels and telomerase activity. We compared telomerase activity in growing and quiescent cells based on equivalent protein contents of cellular extracts. As illustrated in Table 1, an increased cellular protein content was seen in proliferating CHO B11 cells compared to the confluent cells. Following 5 days of starvation the cell number remained constant in confluent cultures, but the cellular protein levels decreased by approximately 0.4 fold. This indicates that there were proliferation-dependent changes in cellular protein content. Interestingly, a discernible telomerase ladder pattern was detected from 0.1 mg protein of extracts from starved cells while telomerase activity was undetectable from 0.1 mg protein extract from growing and confluent cells (Fig. 2). These results suggest that a greater proportion of cellular protein was composed of telomerase activity in starved, quiescent cells. Incubation of cells in medium with limiting amounts of growth factors (starvation) results in the eventual inhibition of cell division, an accumulation of cells in a quiescent state, and a reduction in cellular protein levels. This increase in telomerase activity may be due to the accumulation of telomerase relative to other cellular proteins or may reflect increased telomerase half life or stability (8). To determine if there were cell cycle-specific variations in telomerase activity, we collected CHO B11 cells synchronized at various points in the G1 , S, and G2/M phases of the cell cycle and compared their levels of telomerase activity. The levels of cell cycle synchrony achieved at the time of release from mimosine and at defined time points post mimosine release are presented in figure 3A. A typical asynchronous population of CHO B11 cells (Fig. 3A, panel A) was made up of
about 40-50% G1 phase, 40% S phase, and 10-20% G2/ M phase cells. Confluent cultures consisted primarily of cells in G1 (Fig. 3A, Panel B). Incubation of cells in 0.2% serum-containing medium led to an accumulation of cells in a quiescent state (Fig. 3A, Panel C). When CHO B11 cells were starved for at least 48 hr and then incubated for 6 hr in complete medium, most of the cells were in G1 (Fig. 3A, panel D). Serum starvation followed by a 14 hr period of incubation in 50 mM mimosine resulted in the accumulation of cells at early S (Fig. 3A, Panel E). A high level of synchronization of CHO B11 cells in either S or G2 phase was achieved by using a 48 hr period of serum starvation followed by a 14 hr incubation in complete medium supplemented with 50 mM mimosine, and finally incubation in mimosine-free complete medium. After the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase as evidenced by the increase in DNA content by 4 hr after release from mimosine (Fig. 3A, panel F). Over the next few hours, the cells continued synchronously through S phase (Fig. 3A, panel G) and into G2 by approximately 7.5 hr after release from mimosine (Fig. 3A, panel H). By 8.5 hr after release from mimosine, many of the cells had pro-
FIG. 2. Accumulation of telomerase in starved CHO B11 cells. Decreasing amounts of protein of CHAPS extracted CHO B11 cells (10mg, 1mg, 0.1mg protein extract) from subconfluent, exponentially growing cells (Sub), cells that had just reached confluence (Confl), and confluent cells maintained in a quiescent state for 5 days (5 Da) were analyzed for telomerase activity as described in Materials and Methods. In control lanes (/), 10 mg of protein extract were pretreated with 10 mg/ml RNase A.
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gressed into the subsequent G1 phase (Fig. 3A, panel I). Telomerase activity was detected in asynchronously growing CHO B11 cells (Fig. 3B, lane A) as well as from cells that had reached confluence (Fig. 3B, lane B). Telomerase activity was also found at all discrete stages of the cell cycle (Fig. 3B, lanes D-H). Further, serum-starved quiescent cells retained telomerase activity (Fig. 3B, lane C) equivalent to the levels seen at the specific stages of the cell cycle as well as to growing and confluent cells. There also was no substantial difference in telomerase activity in serial dilutions of cell extracts based on equivalent cell numbers collected during the cell cycle. Further, serum-deprived cultures were capable of re-entering the cycle while maintaining telomerase activity with progression through subsequent cell cycles (Fig. 3B, lane I). Single-stranded telomeric overhangs have been observed as intermediates of telomeric replication during late S phase (10) suggesting that it is specifically during early S phase that telomerase is functional. Because of the potential correlation between the S phase telomeric tails and telomerase activity we conducted a detailed examination of telomerase activity in CHO B11 cells within S phase. Post-starvation (P.S.) G1-arrested CHO B11 cells were obtained by growth to confluence followed by serum starvation (Fig. 4A, 0 hr P.S.). Following release from serum starvation, CHO B11 cells progressed into late G1 (Fig. 4A, 8 hr P.S.) and continued to progress through the cell cycle (Fig. 4A, 15 hr P.S.). In mimosine treated cultures, after the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase (Fig. 4A, panels 1-6 hr post mimosine, P.M.). By 7 hr P.M. the cells were at the S/ G2 boundary (Fig. 4A, Panel 7 hr P.M.). By 8 hr after release from mimosine, most of the cells were in G2/M phase still maintaining a high degree of synchrony (Fig. 4A, panel 8 hr P.M.). From 9 to 14 hr post mimosine increasing proportions of the cells had progressed into the subsequent G1 phase (Fig. 4A, panels 9 and 14 P.M.). While telomerase activity was present throughout the cell cycle-specific phases (Fig. 4B), an increase in telomerase activity was detected in diluted extracts during early S phase (Fig. 4B, lanes 1 and 2 hr P.M.). A 1.6 to 2 fold and 2 to 2.6 fold increase in telomerase activity relative to the other phases examined was detected 1 and 2 hr P.M. respectively. This increase in
FIG. 3. Telomerase activity in mimosine-synchronized CHO B11 cells. A, Panel A, Flow cytometric analysis of exponentially growing, asynchronous CHO B11 cells. Panels B-I, CHO B11 cells (21105 cells in 100-mm-diameter plates) were grown to confluence (Panel B), incubated in low-serum medium for 48hr (Panel C, quiescent cells), then in complete medium (10% dFBS) plus 50mM mimosine for 14hr (Panels E-I). Cells in Panel D represent cells mostly in late G1 collected following 6 hr incubation in complete medium without
mimosine following serum starvation. The cell cycle distribution was measured by flow cytometry as indicated in times corresponding to Panels E through I and 0, 4, 6, 7.5 and 8.5 hr after the mimosinecontaining medium was replaced with complete medium without mimosine. These cells were distributed in early S (Panel E), S (Panel F,G), G2/M (Panel H), and the subsequent G1 (Panel I). B, Decreasing equivalent numbers of cells of CHAPS extracted CHO B11 cells (105103 cells) corresponding to the cell cycle stages in part A were analyzed for telomerase activity as described in Materials and Methods.
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FIG. 4. Telomerase activity in CHO B11 cells during progression through S phase. A, Quiescent CHO B11 cells were obtained following incubation for 48 hr in low serum and collected immediately post starvation (P.S.). The cells entered late G1 8 hr P.S. and by 15 hr P.S. had progressed through the cell cycle (Panel 15 hr P.S.). In mimosine treated cultures, after the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase (Panels 1-6 hr post mimosine). By 7 hr post mimosine (P.M.) the cells were at the S/G2 boundary (Panel 7 hr P.M.). By 8 hr after release from mimosine, the cells entered G2 phase still maintaining a high degree of synchrony (Panel 8 hr P.M.). By 9 and 14 hr post mimosine most of the cells had progressed into the subsequent G1 phase (Panels 9 and 14 P.M.). B, 105-103 equivalent cell numbers of CHAPS extracted CHO B11 cells corresponding to the cell cycle stages in A were analyzed for telomerase activity as described in Materials and Methods.
telomerase activity was not detected in later S phase (Fig. 4A, lanes 3-7 P.M.). Our work is in agreement with another report (12) finding a relative increase in telomerase activity in S phase. Heightened telomerase activity during S phase, then, would precede the appearance of the telomeric tails seen in late S phase (10) and may serve as a mechanism regulating the addition of telomeric repeats in an S phase-specific manner. In all telomerase assays, CAOV3 cells served as positive controls and telomerase activity was abolished upon pretreatment of the cell extracts with RNase (data not shown). Like others (8,12) we have shown sustained telomerase activity throughout the cell cycle in immortal-
ized, undifferentiated cell lines. This supports the hypothesis, that at least in some immortalized cells lines, telomerase is constitutively active and that its regulation is not tightly bound to the cell cycle. However, using a protocol which allows synchronization at discrete points during the S phase, we have reproducibly demonstrated elevated telomerase activity in early S phase. This may be due to additional telomerase activity induced in late G1 or S phase, as observed in nontransformed cells (11), overlying the basal level of activity measured in all cell cycle phases in immortalized cell lines. This elevated telomerase activity in early S phase may be regulated by the cyclin-dependent kinase activity necessary for progression of cells from G1 into
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S phase. Further, the constitutive expression of telomerase in rodent cells, like the ones used in the present study, may in part be related to the high rate of spontaneous immortalization in rodent cells. However, constitutive telomerase expression does not necessarily imply that telomeric repeats are being continuously added to telomeric ends. Other mechanisms may be involved in telomerase regulation. Telomeric binding proteins (17) may sequester telomeric repeats manufactured by telomerase or regulate the addition of these repeats to telomeres during the appropriate period of S phase. The addition of telomeric repeats to telomeric ends may also be regulated at the level of the chromatin structure. Alterations in telomeric structure and nuclear localization (18,19) may render telomeric DNA inaccessible to the addition of telomeric repeats, except when in a particular conformational state. Clearly, telomerase regulation is complex and further studies are needed to resolve the multiple mechanisms regulating telomerase activity with respect to the addition of telomeric repeats and its relation to cellular growth. Such studies may provide useful information on the design, use, and limitations of telomerase inhibitors as potential chemotherapeutic agents. ACKNOWLEDGMENTS We thank Mr. Bob Pyle for his assistance in flow cytometric analysis and Drs. R. Brosh and N. Lipinski for critically reading the manuscript.
REFERENCES 1. Blackburn, E. H. (1991) Nature 350, 569–573. 2. Blackburn, E. H. (1992) Annu. Rev. Biochem. 61, 113–129.
3. Kim, N. W., Piatyszek, M. A., Prowse, K. R., Harley, C. B., West, M. D., Ho, P. L. C., Coviello, G. M., Wright, W. E., Weinrich, S. L., and Shay, J. W. (1994) Science 266, 2011–2015. 4. Wright, W. E., Piatyszek, M. A., Rainey, W. E., Byrd, W., and Shay, J. W. (1996) Dev. Genet. 18, 173–179. 5. Counter, C. M., Gupta, J., Harley, C. B., Leber, B., and Bacchetti, S. (1995) Blood 85, 2315–2320. 6. Yasumoto, S., Kunimura, C., Kikuchi, K., Tahara, H., Ohji, H., Yamamoto, H., Ide, T., and Utakoji, T. (1996) Oncogene 13, 433– 439. 7. Vaziri, H., Dragowska, W., Allsopp, R. C., Thomas, T. E., Harley, C. B., and Lansdorp, P. M. (1994) Proc. Natl. Acad. Sci. (USA) 91, 9857–9860. 8. Holt, S. E., Wright, W. E., and Shay, J. W. (1996) Molec. Cell. Biol. 16, 2932–2939. 9. Sharma, H. W., Sokoloski, J. A., Perez, J. R., Maltese, J. Y., Sartorelli, A. C., Stein, C. A., Nichols, G., Khaled, Z., Telang, N. T., and Narayanan, R. (1995) Proc. Natl. Acad. Sci. (USA) 92, 12343–12346. 10. Wellinger, R. J., Wolf, A. J., and Zakian, V. A. (1993) Cell 72, 51–60. 11. Buchkovich, K. J., and Greider, C. W. (1996) Molec. Biol. Cell 7, 1443–1454. 12. Zhu, X., Kumar, R., Mandal, M., Sharma, N., Sharma, H. W., Dhingra, U., Sokoloski, J. A., Hsiao, R., and Narayanan, R. (1996) Proc. Natl. Acad. Sci. (USA) 93, 6091–6095. 13. Weng, N., Levine, B. L., June, C. H., and Hodes, R. J. (1996) J. Exp. Med. 183, 2471–2479. 14. Orren, D. K., Petersen, L. N., and Bohr, V. A. (1995) Molec. Cell. Biol. 15, 3722–3730. 15. Hughes, T. A., and Cook, P. R. (1996) Exp. Cell Res. 222, 275– 280. 16. Kruk, P. A., Balajee, A. S., Rao, K. S., and Bohr, V. A. (1996) Biophys. Biochem. Res. Commun. 224, 487–492. 17. Cardenas, M. E., Bianchi, A., and de Lange, T. (1993) Genes Dev. 7, 883–894. 18. Funabiki, H., Hagan, I., Uzawa, S., and Yanagida, M. (1993) J. Cell Biol. 121, 961–976. 19. Vourc’h, C., Taruscio, D., Boyle, A. L., and Ward, D. C. (1993) Exp. Cell Res. 205, 142–151.
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233, 717–722 (1997)
RC976549
Telomerase Activity Is Elevated in Early S Phase in Hamster Cells Patricia A. Kruk,*,†,1 David K. Orren,† and Vilhelm A. Bohr† *Department of Pathology, University of South Florida, Tampa, Florida 33612; and †Laboratory of Molecular Genetics, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224
Received March 26, 1997
Telomerase is a ribonucleoprotein that elongates telomeric repeats de novo. We examined the possibility that telomerase activity is cell cycle regulated by examining telomerase activity in cell cycle synchronized Chinese hamster ovary (CHO) B11 cells. Overall telomerase activity was similar in growing and quiescent cells. Further, cells synchronized in G1 , S, or G2/M showed similar levels of telomerase activity. However, a detailed analysis of cells within S phase showed that there was a higher level of telomerase activity in early S phase when compared with other points in the cell cycle. These results suggest a relationship between telomerase activity and cell cycle regulation. q 1997 Academic Press
Telomeres consist of tandem hexanucleotide (TTAGGG)n repeats that cap the ends of vertebrate chromosomes. Telomeres are essential for chromosomal stability and integrity. They protect chromosomes from illegitimate recombination, degradation by nuclease attack, and solve the end replication problem (1,2). Telomeric repeats are synthesized by telomerase, a ribonucleoprotein that elongates telomeric repeats de novo (2). Telomerase activity is generally detected in germ cells, cancer cells, and cell lines, but only occasionally in somatic cells (3-7). Very little is known about the regulation of telomerase activity in mammalian cells. There have been reports of telomerase activation associated with cellular proliferation (5,8) and tumor progression (3,5) as well as telomerase repression associated with cellular differentiation (8,9) and cellular quiescence (8). A relationship exists, then, between telomerase activity and the end replication problem as well as an association between telomerase activity and high cellular proliferative capacity. In addition, the re1 Corresponding author at Department of Pathology, MDC 11, University of South Florida, 12901 Bruce B. Downs Blvd., Tampa, FL 33612-4799. Fax: (813) 974-5536. E-mail: [email protected].
port that G-rich telomeric overhangs appear in late S phase in Saccharomyces cerevisiae (10) would suggest that telomerase activity is not only associated with cellular proliferation, but that the ability to synthesize telomeric repeats may be regulated in a cell cycle dependent manner. Yet information on the regulation of telomerase activity as it relates to cellular proliferation and, in particular, the cell cycle is sparse and often contradictory (8, 11-13). We initiated studies on telomerase activity as a function of cell growth and cell cycle stage in immortalized CHO B11 cells. A synchronization method developed previously (14) allowed us to measure telomerase activity in each cell cycle phase as well as at defined points within S phase. On the basis of equivalent cell numbers, we found no difference in telomerase activity between growing and quiescent cells, although the proportion of cellular protein comprising telomerase activity increased in quiescent CHO B11 cells. Though we observed telomerase activity in these cells throughout the cell cycle, our data indicate that there was a transient increase in telomerase activity in early S phase. These results indicate that telomerase is constitutively active in CHO B11 hamster cells, but may be additionally regulated in a cell cycle phase-specific manner. MATERIALS AND METHODS Cell cycle synchronization. CHO B11 cells were routinely grown as previously described (14) at 377C in 5% CO2 in complete medium consisting of Ham’s F12 medium (without glycine, hypoxanthine, and thymidine) supplemented with 10% dialyzed fetal bovine serum (dFBS). Cell cycle synchronized CHO B11 cells were obtained as described earlier (14). Briefly, G1 enriched cells were obtained following 48 hr incubation of exponentially growing CHO B11 cells in starvation medium. Starvation medium contained only 0.2% dFBS, but was otherwise as stated above. Cells arrested in early S phase were obtained by 48 hr serum starvation followed by a 14 hr incubation in complete medium plus 50 mM mimosine. Recent studies have shown that mimosine inhibits elongation at replication forks shortly after DNA synthesis has begun, probably by depletion of nucleotide pools (15). The effects of mimosine on replicative DNA synthesis are short-lived. After removal of the drug from the medium, mimosine 0006-291X/97 $25.00
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is rapidly cleared from the cells, and the cells subsequently proceed through S phase. The duration of S phase after mimosine release appears to be identical to the time that it normally takes to traverse S phase. Thus, mimosine appears to have no lingering effects on DNA metabolism making it an ideal drug for use in synchronization studies. Replacement of medium containing mimosine with fresh complete medium allowed the cells to pass synchronously through S phase and G2 phase (14). Cells synchronized at various points during the cell cycle were harvested for both flow cytometry and telomerase activity analyses. Flow cytometry. The cell cycle phase distribution of CHO B11 cells was determined by measuring the DNA content of individual cells by flow cytometry as described previously (14). Briefly, 1-5 1 106 cells were trypsinized, resuspended in complete medium, and washed in PBS. The cells were then resuspended, fixed in 5 ml cold ethanol (70%), and stored at 47C. Just prior to flow cytometric analysis, individual samples were treated with RNase then stained with propidium iodide according to instructions (and using reagents) provided with the Cellular DNA Flow Cytometric Analysis Reagent Set (Boehringer Mannheim). Propidium iodide-stained cell suspensions were then analyzed using a Becton-Dickinson FACScan. Histograms (relative fluorescence vs. cell number) were generated from the flow cytometric data using the Lysis II software. Telomerase assay. CHO B11 cells were assayed for telomerase activity as described previously (3,16). Cells were washed with PBS, trypsinized, counted, and spun by centrifugation at 100g. The pellets were resuspended in ice-cold wash buffer containing 10 mM HEPESKOH (pH 7.5)/1.5 mM MgCl2/10 mM KCl/1 mM DTT and spun at 10,000 1 g at 47C for 20 min. The cells were then resuspended at a density of 103 to 105 cells in 20 ml ice-cold lysis buffer containing 10 mM Tris-HCl (pH 7.5)/1 mM MgCl2/1 mM EGTA/ 0.1 mM PMSF/5 mM b-mercaptoethanol/0.5% CHAPS/ 10 % glycerol for 30 min at 47C. The lysates were then centrifuged at 100,000 1 g for 45 min at 47C. After centrifugation, the supernatants were stored at 0807C. The protein concentration of each extract was determined by the BCA protein assay (Pierce). Because protein content varies within cells with progression through the cell cycle, the level of telomerase activity was measured using volumes of cellular extracts equivalent to 103, 104 and 105 cells. However, in one set of experiments examining telomerase activity among cells in different growth states, telomerase activity was also measured using a serial dilution of volumes of extracts based on protein concentration (10, 1.0 and 0.1mg protein). Extracts were incubated with 20 mM Tris-HCl (pH 8.3)/1.5 mM MgCl2/63 mM KCl/0.005% Tween 20/1 mM EGTA/100 mM each dATP, dGTP, dTTP/50 mM dCTP/20 mCi 32P-dCTP/0.1 mg of upstream primer Tel I (5*-AATCCGTCGAGCAGAGTT-3*, Paragon Biotech)/1 mg of T4g32 protein (Boehringer Mannheim)/0.1 mg per ml BSA for 60 min at room temperature (RT). The reaction mixture was then heated to 907C to inactivate the telomerase activity. During this step, 0.1 mg of the downstream primer Tel II (5*-CCCTTACCCTTACCCTTACCCTAA-3*, Paragon Biotech) and 2 ml Taq polymerase (Boehringer Mannheim) were added to the reaction mixture and the reaction mixture was subjected to 30 PCR cycles at 947C for 30 sec, 507C for 30 sec, 727C for 90 sec. As a control, parallel reactions were pretreated with 10 mg/ml RNase A for 20 min at 377C to degrade the RNA component of telomerase and indicate telomerase specificity in the assay. Positive controls for telomerase activity consisted of cell extracts from the ovarian carcinoma cell line, CAOV3. PCR products were analyzed by electrophoresis in Tris-borate EDTA buffer on 8% polyacrylamide gels. The gels were dried and telomerase activity was analyzed using a PhosphoImager system from Molecular Dynamics. The signal intensity from each lane of the gel was defined by a rectangular boundary established around all visible bands in the lane of least activity. A rectangle of equivalent size and shape was used for quantifying the signal from the remaining lanes of the gels. Enzyme activity was expressed as the fold difference of the activity determined in the RNase-sensitive reactions products compared to the lane with the least activity. Extracts were considered negative for
FIG. 1. Telomerase activity in growing, confluent, and starved CHO B11 cells. Decreasing equivalent cell numbers of CHAPS extracted CHO B11 cells (105, 104, 103 cell number equivalents) from subconfluent exponentially growing cells (Sub), cells that had just reached confluence (Confl), and confluent cells maintained in a quiescent state for 5 days (5 Da) and 14 days (14 Da) were analyzed for telomerase activity as described in Materials and Methods. In control lanes (/), 105 cell equivalents were pre-treated with 10 mg/ml RNase A.
telomerase activity if no products were detected following prolonged exposure.
RESULTS AND DISCUSSION In the present study, we examined telomerase activity in immortalized, undifferentiated hamster cells in various growth states and different stages of the cell cycle. We compared telomerase activity in exponentially growing subconfluent CHO B11 cells, cells that had just reached confluence, and confluent cells maintained in a quiescent state for 5 and 14 days respectively. Telomerase activity was demonstrated as the characteristic ladder pattern in cellular extracts from growing cells (Fig. 1). Serum deprivation of CHO B11 cells for 5 and 14 days post confluence, which results in growth inhibition and withdrawal from the cell cycle, did not reduce or abolish telomerase activity as deter-
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Comparison of Cell Number and Protein Content in Growing and Quiescent CHO B11 Cells Growth state
Cell number (1107)
Total protein (mg)
ng Protein/cell
Subconfluent Confluent 5 day starvation
1.188 3.28 3.16
0.722 1.881 1.354
607.7 573.5 428.6
mined from volumes of cellular extracts corresponding to an equivalent number of cells (Fig. 1). Further, there was no ascertainable difference in the amount of telomerase activity in growing and quiescent cells even upon serial dilution of these cellular extracts (Fig. 1). Pretreatment of the extracts with RNase abolished the telomerase activity, indicating telomerase specificity in this assay. Since induction of cellular quiescence by serum starvation did not affect telomerase activity, we sought to determine whether serum starvation altered cellular protein levels and telomerase activity. We compared telomerase activity in growing and quiescent cells based on equivalent protein contents of cellular extracts. As illustrated in Table 1, an increased cellular protein content was seen in proliferating CHO B11 cells compared to the confluent cells. Following 5 days of starvation the cell number remained constant in confluent cultures, but the cellular protein levels decreased by approximately 0.4 fold. This indicates that there were proliferation-dependent changes in cellular protein content. Interestingly, a discernible telomerase ladder pattern was detected from 0.1 mg protein of extracts from starved cells while telomerase activity was undetectable from 0.1 mg protein extract from growing and confluent cells (Fig. 2). These results suggest that a greater proportion of cellular protein was composed of telomerase activity in starved, quiescent cells. Incubation of cells in medium with limiting amounts of growth factors (starvation) results in the eventual inhibition of cell division, an accumulation of cells in a quiescent state, and a reduction in cellular protein levels. This increase in telomerase activity may be due to the accumulation of telomerase relative to other cellular proteins or may reflect increased telomerase half life or stability (8). To determine if there were cell cycle-specific variations in telomerase activity, we collected CHO B11 cells synchronized at various points in the G1 , S, and G2/M phases of the cell cycle and compared their levels of telomerase activity. The levels of cell cycle synchrony achieved at the time of release from mimosine and at defined time points post mimosine release are presented in figure 3A. A typical asynchronous population of CHO B11 cells (Fig. 3A, panel A) was made up of
about 40-50% G1 phase, 40% S phase, and 10-20% G2/ M phase cells. Confluent cultures consisted primarily of cells in G1 (Fig. 3A, Panel B). Incubation of cells in 0.2% serum-containing medium led to an accumulation of cells in a quiescent state (Fig. 3A, Panel C). When CHO B11 cells were starved for at least 48 hr and then incubated for 6 hr in complete medium, most of the cells were in G1 (Fig. 3A, panel D). Serum starvation followed by a 14 hr period of incubation in 50 mM mimosine resulted in the accumulation of cells at early S (Fig. 3A, Panel E). A high level of synchronization of CHO B11 cells in either S or G2 phase was achieved by using a 48 hr period of serum starvation followed by a 14 hr incubation in complete medium supplemented with 50 mM mimosine, and finally incubation in mimosine-free complete medium. After the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase as evidenced by the increase in DNA content by 4 hr after release from mimosine (Fig. 3A, panel F). Over the next few hours, the cells continued synchronously through S phase (Fig. 3A, panel G) and into G2 by approximately 7.5 hr after release from mimosine (Fig. 3A, panel H). By 8.5 hr after release from mimosine, many of the cells had pro-
FIG. 2. Accumulation of telomerase in starved CHO B11 cells. Decreasing amounts of protein of CHAPS extracted CHO B11 cells (10mg, 1mg, 0.1mg protein extract) from subconfluent, exponentially growing cells (Sub), cells that had just reached confluence (Confl), and confluent cells maintained in a quiescent state for 5 days (5 Da) were analyzed for telomerase activity as described in Materials and Methods. In control lanes (/), 10 mg of protein extract were pretreated with 10 mg/ml RNase A.
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gressed into the subsequent G1 phase (Fig. 3A, panel I). Telomerase activity was detected in asynchronously growing CHO B11 cells (Fig. 3B, lane A) as well as from cells that had reached confluence (Fig. 3B, lane B). Telomerase activity was also found at all discrete stages of the cell cycle (Fig. 3B, lanes D-H). Further, serum-starved quiescent cells retained telomerase activity (Fig. 3B, lane C) equivalent to the levels seen at the specific stages of the cell cycle as well as to growing and confluent cells. There also was no substantial difference in telomerase activity in serial dilutions of cell extracts based on equivalent cell numbers collected during the cell cycle. Further, serum-deprived cultures were capable of re-entering the cycle while maintaining telomerase activity with progression through subsequent cell cycles (Fig. 3B, lane I). Single-stranded telomeric overhangs have been observed as intermediates of telomeric replication during late S phase (10) suggesting that it is specifically during early S phase that telomerase is functional. Because of the potential correlation between the S phase telomeric tails and telomerase activity we conducted a detailed examination of telomerase activity in CHO B11 cells within S phase. Post-starvation (P.S.) G1-arrested CHO B11 cells were obtained by growth to confluence followed by serum starvation (Fig. 4A, 0 hr P.S.). Following release from serum starvation, CHO B11 cells progressed into late G1 (Fig. 4A, 8 hr P.S.) and continued to progress through the cell cycle (Fig. 4A, 15 hr P.S.). In mimosine treated cultures, after the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase (Fig. 4A, panels 1-6 hr post mimosine, P.M.). By 7 hr P.M. the cells were at the S/ G2 boundary (Fig. 4A, Panel 7 hr P.M.). By 8 hr after release from mimosine, most of the cells were in G2/M phase still maintaining a high degree of synchrony (Fig. 4A, panel 8 hr P.M.). From 9 to 14 hr post mimosine increasing proportions of the cells had progressed into the subsequent G1 phase (Fig. 4A, panels 9 and 14 P.M.). While telomerase activity was present throughout the cell cycle-specific phases (Fig. 4B), an increase in telomerase activity was detected in diluted extracts during early S phase (Fig. 4B, lanes 1 and 2 hr P.M.). A 1.6 to 2 fold and 2 to 2.6 fold increase in telomerase activity relative to the other phases examined was detected 1 and 2 hr P.M. respectively. This increase in
FIG. 3. Telomerase activity in mimosine-synchronized CHO B11 cells. A, Panel A, Flow cytometric analysis of exponentially growing, asynchronous CHO B11 cells. Panels B-I, CHO B11 cells (21105 cells in 100-mm-diameter plates) were grown to confluence (Panel B), incubated in low-serum medium for 48hr (Panel C, quiescent cells), then in complete medium (10% dFBS) plus 50mM mimosine for 14hr (Panels E-I). Cells in Panel D represent cells mostly in late G1 collected following 6 hr incubation in complete medium without
mimosine following serum starvation. The cell cycle distribution was measured by flow cytometry as indicated in times corresponding to Panels E through I and 0, 4, 6, 7.5 and 8.5 hr after the mimosinecontaining medium was replaced with complete medium without mimosine. These cells were distributed in early S (Panel E), S (Panel F,G), G2/M (Panel H), and the subsequent G1 (Panel I). B, Decreasing equivalent numbers of cells of CHAPS extracted CHO B11 cells (105103 cells) corresponding to the cell cycle stages in part A were analyzed for telomerase activity as described in Materials and Methods.
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FIG. 4. Telomerase activity in CHO B11 cells during progression through S phase. A, Quiescent CHO B11 cells were obtained following incubation for 48 hr in low serum and collected immediately post starvation (P.S.). The cells entered late G1 8 hr P.S. and by 15 hr P.S. had progressed through the cell cycle (Panel 15 hr P.S.). In mimosine treated cultures, after the medium containing mimosine was replaced with fresh complete medium, the cells quickly and synchronously progressed through S phase (Panels 1-6 hr post mimosine). By 7 hr post mimosine (P.M.) the cells were at the S/G2 boundary (Panel 7 hr P.M.). By 8 hr after release from mimosine, the cells entered G2 phase still maintaining a high degree of synchrony (Panel 8 hr P.M.). By 9 and 14 hr post mimosine most of the cells had progressed into the subsequent G1 phase (Panels 9 and 14 P.M.). B, 105-103 equivalent cell numbers of CHAPS extracted CHO B11 cells corresponding to the cell cycle stages in A were analyzed for telomerase activity as described in Materials and Methods.
telomerase activity was not detected in later S phase (Fig. 4A, lanes 3-7 P.M.). Our work is in agreement with another report (12) finding a relative increase in telomerase activity in S phase. Heightened telomerase activity during S phase, then, would precede the appearance of the telomeric tails seen in late S phase (10) and may serve as a mechanism regulating the addition of telomeric repeats in an S phase-specific manner. In all telomerase assays, CAOV3 cells served as positive controls and telomerase activity was abolished upon pretreatment of the cell extracts with RNase (data not shown). Like others (8,12) we have shown sustained telomerase activity throughout the cell cycle in immortal-
ized, undifferentiated cell lines. This supports the hypothesis, that at least in some immortalized cells lines, telomerase is constitutively active and that its regulation is not tightly bound to the cell cycle. However, using a protocol which allows synchronization at discrete points during the S phase, we have reproducibly demonstrated elevated telomerase activity in early S phase. This may be due to additional telomerase activity induced in late G1 or S phase, as observed in nontransformed cells (11), overlying the basal level of activity measured in all cell cycle phases in immortalized cell lines. This elevated telomerase activity in early S phase may be regulated by the cyclin-dependent kinase activity necessary for progression of cells from G1 into
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S phase. Further, the constitutive expression of telomerase in rodent cells, like the ones used in the present study, may in part be related to the high rate of spontaneous immortalization in rodent cells. However, constitutive telomerase expression does not necessarily imply that telomeric repeats are being continuously added to telomeric ends. Other mechanisms may be involved in telomerase regulation. Telomeric binding proteins (17) may sequester telomeric repeats manufactured by telomerase or regulate the addition of these repeats to telomeres during the appropriate period of S phase. The addition of telomeric repeats to telomeric ends may also be regulated at the level of the chromatin structure. Alterations in telomeric structure and nuclear localization (18,19) may render telomeric DNA inaccessible to the addition of telomeric repeats, except when in a particular conformational state. Clearly, telomerase regulation is complex and further studies are needed to resolve the multiple mechanisms regulating telomerase activity with respect to the addition of telomeric repeats and its relation to cellular growth. Such studies may provide useful information on the design, use, and limitations of telomerase inhibitors as potential chemotherapeutic agents. ACKNOWLEDGMENTS We thank Mr. Bob Pyle for his assistance in flow cytometric analysis and Drs. R. Brosh and N. Lipinski for critically reading the manuscript.
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