Telomerase is a ribonucleoprotein complex that maintains the stability of chromosome ends and regulates replicative potential. Telomerase is upregulated in over 85% of human tumors, but not in adjacent normal tissues and represents a promising target for anticancer therapy. Most telomerase-based therapies rely on the inhibition of telomerase activity and require extensive telomere shortening before inducing any antiproliferative effect. Disturbances of telomere structure rather than length may be more effective in inducing cell death. Telomerase RNA subunits (hTRs) with mutations in the template region reconstitute active holoenzymes that incorporate mutated telomeric sequences. Here, we analysed the feasibility of an anticancer approach based on the combination of telomere destabilization and conventional chemotherapeutic drugs. We show that a mutant template hTR dictates the synthesis of mutated telomeric repeats in telomerase-positive cancer cells, without significantly affecting their viability and proliferative ability. Nevertheless, the mutant hTR increased sensitivity to anticancer drugs in cells with different initial telomere lengths and mechanisms of telomere maintenance and without requiring overall telomere shortening. This report is the first to show that interfering with telomere structure maintenance in a telomerase-dependent manner may be used to increase the susceptibility of tumor cells to anticancer drugs and may lead to the development of a general therapy for the treatment of human cancers.
Telomerase is a ribonucleoprotein essential for the maintenance of the structure and the length of telomeres, the nucleoprotein complexes at the ends of eucaryotic chromosomes that cap and protect them from degradation, fusions and recombination. Telomerase contains two essential subunits, a catalytic component with reverse transcriptase activity, called human telomerase reverse transcriptase (hTERT) in humans, and an RNA moiety, called hTR, which functions as a template for the synthesis of T2AG3 repeats onto the chromosome termini. In humans, telomerase is expressed during embryonic development, but is repressed in adult tissues, with the exception of germ line and stem cells, which remain positive for the enzyme and maintain telomere length (Collins and Mitchell, 2002). Some proliferating somatic cells also have low levels of telomerase activity sufficient for the maintenance of telomere structure but not length (Masutomi et al., 2003). In contrast, telomerase is detected in most immortalized cell lines and in over 85% of human cancers, indicating that its reactivation is an essential step for unlimited proliferation and cancer progression (Shay and Bacchetti, 1997). Telomerase may therefore represent an excellent anticancer target. Anti-telomerase approaches rely on the loss of telomerase activity and result in erosion of telomeric DNA, which causes genomic instability and cell death. These effects are predicted to be relatively specific to cancer cells with minor side effects for somatic cells and stem cells, which have longer telomeres, no or low levels of telomerase and slower duplication time. The major limitation of telomerase inhibition is the time necessary for the telomeres to become critically short before the antiproliferative effects are observed. Several reports have shown, in fact, that inhibition of telomerase by either a dominant-negative protein or antisense oligonucleotides against hTR results effectively in death of tumor cells with short but not long telomeres (Hahn et al., 1999; Herbert et al., 1999; Zhang et al., 1999). A complementary strategy to block tumor cell growth is to use telomerase inhibition as a means to sensitize cancer cells to chemotherapeutic drugs or angiogenesis inhibitors (Kondo et al., 1998; Ludwig et al., 2001; Misawa et al., 2002; Chen et al., 2003; Tentori et al., 2003). However, the responsiveness to such treatments appears to vary in a cell- and drug-type-dependent fashion (Folini et al., 2000; Tentori et al., 2003) and in most cases requires prior telomere shortening (Kondo et al., 1998; Misawa et al., 2002; Chen et al., 2003; Ward and Autexier, 2005), indicating that even combination approaches based on telomerase inhibition may not be clinically suitable for cancer treatment.
Telomere uncapping, independent of shortening, represents a way to eliminate the lag phase associated with telomerase inhibition. Expression of hTRs with mutations in the template sequences results in the synthesis of mutated repeats that cause telomere structure disturbances and reduced cell viability. These effects are most likely owing to the inability of telomeric proteins to bind to the mutated repeats and do not require telomere shortening (Marusic et al., 1997; Kim et al., 2001). However, unless the mutant hTR is highly overexpressed or the endogenous hTR is absent (Guiducci et al., 2001; Li et al., 2004), the endogenous telomerase complex adds wild-type sequences to mutant ones, partially restoring telomere function and mitigating the deleterious effects of the mutant telomerase. Hence, the expression of mutant template RNAs may only be partially effective in inducing cell death of telomerase-positive cancer cells and human tumors.
The goal of the current work is to validate a different approach for anticancer treatment based on the combination of telomere destabilization induced by the introduction of mutant template hTRs and anticancer drugs. The advantage of this strategy is that telomere disturbances are achieved in a telomerase-dependent manner, thereby maintaining tumor selectivity, but without the lag phase associated with telomerase-based therapies. We found that telomere destabilization induced by the introduction of mutant hTRs increased the susceptibility of human breast cancer cells to chemotherapeutic drugs independently of their initial telomere length and without requiring bulk telomere shortening. Moreover, we showed that ALT (alternative lengthening of telomeres) GM847 cells, engineered to express both hTERT and the mutant hTR, became more sensitive to drug treatments compared to controls. These data indicate that interfering with the maintenance of telomere structure in combination with anticancer drugs may be exploitable as a general anticancer approach to target cancer cells independently of telomere length and mechanisms of telomere maintenance.
Establishment of stable clonal populations expressing a mutant template hTR
We investigated whether telomere destabilization induced by mutant template hTRs could increase the susceptibility of tumor cells to anticancer drugs without the lag phase associated with telomerase inhibition. For this purpose, we screened several human breast cancer cell lines and chose three telomerase-positive cell lines with different telomere lengths and p53 status (Figure 1): YCC-B1 cells with short telomeres (average terminal restriction fragment (TRF) 3.2 kb), MCF-7 cells with intermediate telomere length (average TRF 7 kb) and YCC-B2 cells with long telomeres (average TRF 11 kb). YCC-B1 and MCF-7 cells had wild-type p53, which was induced upon doxorubicin treatment, a known p53 activator; YCC-B2 cells had undetectable levels of p53, which was not induced upon doxorubicin treatment and did not induce p21 expression (Figure 1c and data not shown).
We disturbed telomere maintenance in these cells through the expression of an hTR with a point mutation in the template region (MuA-hTR). This RNA reconstitutes a mutant holoenzyme that adds mutant repeats (T3G3) onto the telomeres (Marusic et al., 1997; Guiducci et al., 2001; Kim et al., 2001). YCC-B1, MCF-7 and YCC-B2 cells were transfected with the vector alone or a plasmid encoding MuA-hTR and clonal populations were selected. For each cell line, one vector clone and three clonal populations expressing the mutant RNA were selected for further studies (Figure 2 and Table 1). The presence of the mutant hTR was monitored by telomere repeat amplification protocol (TRAP) assays under conditions that specifically detect mutant telomerase activity (Figure 2a–c). Wild-type telomerase activity was monitored using standard TRAP conditions and was not affected by the presence of the mutant RNA (data not shown; Feng et al., 1995; Marusic et al., 1997; Kim et al., 2001). The mutant hTR was able to reconstitute an active telomerase that synthesized mutated telomeric repeats (Figure 2d and data not shown). The presence of mutant repeats had only a mild effect on viability and proliferative ability of the mutant hTR clones compared to vector cells both in mass culture and in colony-forming assays, most likely because telomere function could be restored by the addition of wild-type repeats to mutant ones by the endogenous telomerase (Supplementary Figure 1S; Marusic et al., 1997). However, as previously reported, cells expressing the mutant RNA formed smaller colonies than the control cells when plated at low density (data not shown). In addition, no difference in the cell cycle profile of the mutant hTR clones compared to the control cells was observed (Supplementary Figure 1SE and data not shown). The mutant hTR-dependent effects require the presence of a biologically active telomerase enzyme (Guiducci et al., 2001). To confirm the dependence of MuA-hTR action on a functional telomerase, we used ALT VA13 cells that do not express hTERT, transfected them with MuA-hTR and selected clonal populations (Supplementary Figure 3SA).
The presence of MuA-hTR increases drug sensitivity without requiring overall telomere shortening
In order to investigate whether telomere destabilization following mutant hTR expression could rapidly sensitize tumor cells to anticancer drugs, we analysed the response of cells expressing the mutant template RNA to two commonly used DNA-damaging agents, etoposide and doxorubicin. The response to the drugs was evaluated by colony-forming assays, which measure the ability of each cell in the population to recover from drug treatment and proliferate. Vector- and MuA-hTR-expressing cells were treated for 24 h with either etoposide or doxorubicin at concentrations ranging between the IC15 and IC25 of the parental cells and subsequently plated at low density and allowed to proliferate (Figure 3). YCC-B1 cells expressing the mutant hTR showed a significant reduction in the number of colonies compared to the vector-transfected cells upon treatment with either drug, indicating that telomere disturbances induced by the mutant RNA enhanced their sensitivity to the drugs (Figure 3a). Similar results were obtained with the MuA-hTR clones derived from MCF-7 cells and most interestingly with the YCC-B2 derivatives (Figure 3b and c). The observation that cell lines with different telomere lengths responded similarly to drug treatments in the presence of the mutant RNA indicated that the reconstituted mutant holoenzymes affected telomere structure independently of the initial telomere lengths, most likely by disturbing the binding of telomeric proteins and the formation of a proper cap. The mutant hTR-dependent antiproliferative effects were present immediately after the isolation of the clonal derivatives, without any lag phase (Figure 3 and data not shown). TRF analyses showed that telomere length was maintained over time in all clonal populations (Figure 2e and data not shown), indicating that the increased sensitivity to the drugs did not require overall telomere shortening. Quantitative fluorescence in situ hybridization (Q-FISH) analyses at the single cell level revealed an increase in telomere length heterogeneity in the clones expressing MuA-hTR compared to the control cells (Figure 4 and Supplementary Figure 2S). However, no signal-free ends or significant increase in the frequency of chromosome fusions were observed (Supplementary Figure 2S and data not shown). Finally, the lengths of the G-tails were not affected in cells expressing the mutant RNA (data not shown).
In contrast with the results in telomerase-positive cells, the introduction of the mutant hTR in VA13 cells that do not express hTERT did not have any significant effect on the response to doxorubicin treatment, demonstrating that the effects induced by the mutant template hTR require a functional telomerase enzyme (Supplementary Figure 3SB).
Etoposide and doxorubicin are both DNA-damaging agents. Some studies have suggested that telomerase may play an active role in the DNA damage response in human cells and therefore its inhibition results in increased sensitivity specifically to DNA-damaging compounds (Lee et al., 2001; Sharma et al., 2003; Shin et al., 2004; Masutomi et al., 2005; Nakamura et al., 2005). To investigate whether the enhanced response to drug treatment in cells expressing MuA-hTR was specific for DNA-damaging drugs, such as etoposide and doxorubicin, we treated MuA-hTR- and vector-transfected YCC-B2 cells with paclitaxel, a commonly used drug that exerts its antitumor activity primarily by stabilizing the microtubules (Jordan et al., 1993). We found that MuA-hTR clones were significantly more sensitive to paclitaxel than control cells (Figure 3d). These results indicate that the incorporation of mutated telomeric repeats makes YCC-B2 cells more susceptible to different harmful stimuli, and not specifically to DNA-damaging agents.
Mutant template hTR induces the formation of DNA damage foci and exacerbates the antiproliferative effects of anticancer drugs
We also analysed the cell cycle profile in the vector and the mutant hTR clones derived from the YCC-B2 cells after drug treatment. Vector cells showed a marked decrease in the percentage of cells in G1 and a slight increase in G2/M after 24 h of doxorubicin treatment as indicated by a reduction of the G1/G2–M ratio compared to untreated cells (Figure 5a and data not shown). Also, we detected a slight increase in the sub-G1 fraction after drug treatment (data not shown). These characteristics are indicative of mitotic catastrophe, a process of cell death that occurs during or after mitosis in response to several anticancer agents, such as anthracyclines and taxanes (Castedo et al., 2004; Brown and Attardi, 2005). Mitotic catastrophe is characterized by aberrant mitoses and polyploidy, which may be followed by apoptosis (Castedo et al., 2004; Brown and Attardi, 2005). Interestingly, in the clonal populations expressing the mutant hTR, the effects of doxorubicin treatment were more pronounced than in the vector cells. The percentage of cells in G1 was further decreased compared to the vector cells, whereas the percentage of cells in G2/M was similar, resulting in a greater reduction of the G1/G2–M ratio (Figure 5a). After 72 h, the main effect of doxorubicin treatment was a marked increase in the percentage of cells in sub-G1 in the vector clone compared to untreated cells, with an even larger increase in the mutant clones (Figure 5b). Thus, the reduction in the number of colonies following drug treatment in the mutant populations could be attributed to an exacerbation of the antiproliferative effects of doxorubicin in the presence of the mutant hTR, resulting in an alteration of the cell cycle profile associated with an initial G2/M accumulation and high levels of cell death.
A previous study has shown that expression of high levels of mutant template hTRs induces a DNA damage response with the formation of nuclear foci at the telomeres that colocalize with the DNA damage protein 53BP1 (Xu and Blackburn, 2004). In order to understand whether that was the case also in cells expressing low levels of the mutant hTR, we monitored the presence of 53BP1 foci in vector and MuA-hTR YCC-B2 clones. We found that the mutant populations had a significantly higher percentage of cells containing 53BP1 foci compared to the vector cells, although only few foci/cell were detected (1–4 foci/cell; Figure 5c and data not shown). As expected, following doxorubicin treatment both the percentage of foci-containing cells and the number of foci/cell were increased in all populations analysed, whereas treatment with paclitaxel did not affect either parameter (data not shown). These results suggest that the presence of the mutant template hTR causes a slight disturbance of the telomere structure, which results in the formation of DNA damage foci; given the low level of MuA-hTR expression and the presence of the wild-type enzyme, these effects are mild and do not affect cell viability. When cells are treated with anticancer drugs, however, these disturbances of the telomere cap exacerbate the antiproliferative effects of the drugs.
Mutant hTR increases drug sensitivity in immortal cells that maintain telomeres via the ALT pathway
A total of 10–15% of human tumors do not rely on telomerase for telomere maintenance but use an alternative mechanism most likely based on recombination (Bryan et al., 1997; Dunham et al., 2000). We sought to investigate whether drug treatment combined with telomere destabilization caused by the mutant hTR could be exploited as a general approach for various tumor cells regardless of the mechanism used to maintain their telomeres. For this purpose, we chose an ALT cell line, GM847, which does not use telomerase to maintain telomere length and expresses only hTR. Telomerase can be reconstituted in these cells by reintroducing the catalytic subunit hTERT (Wen et al., 1998). GM847 cells were transfected with a plasmid encoding hTERT alone or hTERT and MuA-hTR together to reconstitute a wild-type or a mutant telomerase, respectively (Figure 6). We reasoned that if the synthesis of mutant telomeric repeats disturbed the telomere cap, the mutant cells should be more sensitive to drug treatment than the controls although they can maintain telomere length through the ALT pathway. Indeed colony-forming assays of doxorubicin-treated GM847 cells expressing hTERT and MuA-hTR yielded fewer colonies compared to both the vector clone and clones expressing wild-type telomerase (Figure 6d), indicating that the presence of hTERT and the mutant RNA sensitizes ALT GM847 cells to doxorubicin treatment. Moreover, these effects did not require changes in overall telomere length (Figure 6c).
Telomere maintenance is an essential requisite for cell proliferation. Disturbing telomere integrity results in impairment of cell proliferation and loss of viability. Several reports have validated telomerase as a possible therapeutic target for cancer treatment (reviewed by Kelland, 2005). However, targeting telomerase alone or in combination with anticancer drugs is not sufficient to trigger rapid death of all tumor cells owing to the lag phase necessary for telomeres to become critically short and dysfunctional (Kelland, 2005). In contrast, disturbances of the telomeric capping induce cell growth arrest without significant telomere shortening and therefore may act more rapidly (Karlseder et al., 1999; Takai et al., 2003). In this study, we analysed the effects of drug treatment in cells in which we interfered with telomere structure by the introduction of a mutant template hTR. We show that disturbing telomere structure significantly increased the sensitivity of human tumor cells to a variety of anticancer drugs.
We report that cells in which telomere disturbances were induced through the introduction of a mutant template hTR did not undergo overall telomere shortening, although broader telomere length distributions within cells were apparent. The presence of the mutant hTR did not significantly affect the ability of the cells to proliferate both in mass culture and when plated at limiting dilutions nor did induce any change in the cell cycle profile. Nonetheless, cells expressing the mutant RNA were more sensitive to treatment with anticancer drugs compared to the controls. The response to drug treatment required an active telomerase enzyme and did not depend on the p53 status of the cells. Cell cycle analyses of YCC-B2 vector and mutant template hTR clones showed an alteration of the cell cycle profile after drug treatment, with a marked decrease in the percentage of the cells in G1 and an increase in the sub-G1 fraction, which was more pronounced in the mutant clones than in the vector cells. These changes are indicative of cell death by mitotic catastrophe. Previous studies in human ALT cells and in lower eucaryotes have shown that mutated telomeres have deleterious effects on cell viability and result in cell cycle abnormalities and cell death by mitotic catastrophe (Yu et al., 1990; Kirk et al., 1997; Smith and Blackburn, 1999; Guiducci et al., 2001; Lin et al., 2004). Thus, although the expression of MuA-hTR does not affect cell viability in normal growth conditions, it may enhance the antiproliferative effects of doxorubicin, resulting in higher drug sensitivity compared to control cells.
Although the aim of this work was not to characterize the mechanism by which mutated telomeric repeats increase cell sensitivity to anticancer drugs, we speculate that the presence of mutant sequences affects the binding of the shelterin complex, disturbs telomere capping and induces a DNA damage response. A recent study has shown that high levels of mutant template hTRs induce the formation of nuclear foci at the telomeres that colocalize with DNA damage proteins, such as 53BP1 (Xu and Blackburn, 2004). Similar to these findings, we found a higher percentage of cells containing 53BP1 foci in the mutant clones compared to the vector cells. However, we did not observe any effect of the mutant hTR on cell viability and proliferation, possibly owing to the lower levels of MuA-hTR expression. Nevertheless, the presence of mutated repeats is likely to disturb the telomere capping thereby exacerbating the antiproliferative effects of anticancer drugs. It has recently been proposed that telomerase has other yet uncharacterized telomere-independent functions besides telomere length maintenance (Chung et al., 2005). Although we cannot exclude the possibility that the mutant hTR may interfere with these alternative functions of telomerase, it seems very likely that the ability to synthesize new telomeric repeats is essential for the mutant hTR-dependent effects.
Our results obtained using the combination of mutant template hTR and chemotherapeutic drugs differ significantly from those reported with telomerase inhibition-based approaches described to date. The main difference is that the increased sensitivity to anticancer drugs imparted by the mutant hTR does not require overall telomere shortening, eliminating the lag phase associated with telomerase inhibition. The observation that all the clonal derivatives expressing the mutant RNA were more sensitive to drug treatment than the controls independently of their initial telomere lengths supports this conclusion. Indeed, cells with long telomeres (YCC-B2) and cells with much shorter telomeres (YCC-B1) responded similarly to the drugs. Moreover, we found that reconstitution of a mutant telomerase enzyme in immortal ALT cells with extremely long telomeres resulted in greater sensitivity to chemotherapeutic drugs compared to control cells, without any obvious effect on telomere length maintenance or cell proliferation. These results confirm that disturbing the telomeres in a telomerase-dependent manner can be used to sensitize immortal ALT cells to anticancer drugs. Another important difference between our study and previous telomerase-based anticancer approaches is that cells expressing the mutant hTR became more sensitive to anticancer drugs with different mechanisms of action. Several reports have indicated that telomerase inhibition sensitizes cells specifically to agents that induce DNA breaks, but has no effect with drugs that act through other mechanisms (Lee et al., 2001; Sharma et al., 2003; Shin et al., 2004; Masutomi et al., 2005; Nakamura et al., 2005). However, we found that YCC-B2 cells expressing MuA-hTR are also more susceptible than control cells to treatment with paclitaxel, a widely used chemotherapeutic that acts primarily by stabilizing the microtubules of the mitotic spindle, excluding specificity for DNA-damaging agents at least in the context of the mutant hTR.
Altogether, our data indicate that interfering with telomere structure in cancer cells through the introduction of mutant template hTR could be an effective and general strategy to block tumor cell growth. More importantly this approach may lead to the development of a clinical therapy for the treatment of all tumors independently of their initial telomere lengths and mechanisms to maintain them. This could allow the use of lower levels of chemotherapeutics or shorter treatment time thereby reducing systemic cytotoxicity.
Materials and methods
Cell lines and plasmids
YCC-B1 and YCC-B2 breast cancer cells were kindly provided by Dr Sun Young Rha of the Cancer Metastasis Center, Yonsei University College of Medicine, Korea (Park et al., 1998) and were grown in minimum essential medium (MEM) with 10% heat-inactivated fetal bovine serum (FBS, Wisent). MCF-7 cells were obtained from Dr Pollack (Lady Davis Institute, Montreal, Canada) and were grown in RPMI with 10% FBS. GM847 and VA13 cells obtained from Dr Silvia Bacchetti (Regina Elena Cancer Institute, Rome, Italy) were grown in α-MEM with 10% FBS.
phTR and phTR-MuA containing, respectively, wild-type hTR and a mutant hTR specifying TTTGGG telomeric sequences (Marusic et al., 1997; Guiducci et al., 2001) driven by the endogenous hTR promoter, and phTERT containing wild-type hTERT and the puromycin resistance gene were obtained from Dr Silvia Bacchetti. phTERT/hTR-MuA was obtained by subcloning hTR-MuA into the phTERT plasmid.
Transfections of breast cancer cells were performed by DNA-calcium phosphate with the vector or phTR-MuA. Stable clonal populations were selected with 0.25 μg/ml puromycin for 10–14 days. GM847 cells were transfected with either phTERT or phTERT/hTR-MuA and selected in 0.2 μg/ml puromycin. VA13 cells were transfected with either the vector or phTR-MuA and selected in 0.3 μg/ml puromycin. All cell populations were routinely subcultured at a 1:4 split ratio as they reached confluence. Population doublings (PDs) were calculated taking as PD 0 the time when clones first reached confluence in a 60 mm plate.
Telomerase assay and Western blot analysis
Whole-cells extracts were prepared by detergent lysis and assayed by the PCR-based TRAP (Kim et al., 1994) using serial dilutions of the extracts. For the MuA-hTR-expressing clones, TRAP assay was performed using 5–7 μg of protein extracts with PCR conditions and reverse primers specific for the mutant hTR as described previously (Feng et al., 1995). Western blot analyses were performed using 40 μg of protein extracts. The following antibodies were used: anti-mouse p53 (Ab6, Oncogene Science, kindly provided by Dr Koromilas, Lady Davis Institute, Montreal, Canada), anti-mouse p21 (Upstate, Lake Placid, NY, USA) and anti-mouse β-actin (MA1501, Chemicon, Temecula, CA, USA). Secondary antibodies were purchased from Sigma, Oakville, ON, USA and used according to the manufacturer's instructions.
For TRF analysis, DNA was extracted using standard procedures, digested with HinfI/RsaI and separated by pulse field gel electrophoresis (PFGE; Bryan et al., 1995). The gel was then denatured, neutralized and partially dried. Wild-type telomeric sequences were detected by hybridization with a [γ-32P]dATP 5′-end-labeled telomeric probe (C3TA2)3, whereas mutant telomeric sequences were detected using a [γ-32P]dATP 5′-end-labeled mutant-specific probe (C3A3)4. pBSdT34, containing nine TTTGGG repeats, was used as a positive control for hybridization with the mutant probe. After hybridization, gels were exposed to Phosphoimager. Hybridization signals were quantified with ImageQuant (Molecular Dynamics, Sunnyvale, CA, USA) as described (Harley et al., 1990; Bryan et al., 1995).
G-tail length was analysed with the T-OLA assay (telomere-oligonucleotide ligation assay) as described previously (Cimino-Reale et al., 2001; Stewart et al., 2003). Genomic DNA (5 μg) was hybridized with 0.5 pmol of [γ-32P]dATP 5′-end-labeled telomeric probe at 50°C overnight, ligated for 5 h, precipitated and separated on 5% acrylamide, 6 M urea gel. As a loading control, 10 ng of the ligated DNA was used for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) amplification.
RNA was isolated using TRIzol (Invitrogen, Burlington, ON, USA) according to the manufacturer's instructions. The following primers were used:
For wt-TR and MuA-hTR: LX, 5′-IndexTermGAGAGAGTGACTCTCACGAGAGCC-3′; F3B, 5′-IndexTermTCTAACCCTAACTGAGAAGGGCGTAG-3′; and R3C, 5′-IndexTermGTTTGCTCTAGAATGAACGGTGGA-3′. The LX and R3C primers recognize the transfected hTR, whereas F3B and R3C primers recognize the endogenous hTR.
For hTERT: hT1, 5′-IndexTermAAGTTCCTGCAGTGGCTGATGAG-3′ and hT5, 5′-IndexTermTCGTAGTTGAGCACGCTGAACAG-3′.
For human GAPDH: RT11, 5′-IndexTermCGGAGTCAACGGATTTGGTCGTAT-3′ and RT12, 5′IndexTermTGCTAAGCAGTTGGTGGTGCAGGA-3′.
Q-FISH and immunofluorescence
Metaphase chromosome spreads were prepared from vector- or MuA-hTR-expressing YCC-B2 cells as described previously (Cerone et al., 2001). Fixed cells were hybridized with a telomeric (C3TA2)-Cy3 PNA probe and counterstained with 4,6,diamidino-2-phenylindole and fluorescent signals were captured using a CCD camera (Photometrics-Sensys, Tucson, AZ, USA). Original black and white Cy3 images were used for quantitative analysis using the Iplab Spectrum P Software. To obtain telomere relative intensities, the mean pixel value of each telomere was divided by the mean telomere intensity of the metaphase.
For immunofluorescence, vector and MuA-hTR YCC-B2 cells were fixed with 2% paraformaldehyde for 15 min, permeabilized with 0.25% Triton X-100 for 5 min and blocked in phosphate-buffered saline (PBS) with 5% FBS for 1 h at room temperature. 53BP1 foci were detected with a mouse monoclonal antibody against 53BP1 (Upstate Cell Signaling Solutions, clone BP13) at 1:100 dilution, followed by fluorescein isothiocynate-conjugated goat anti-mouse secondary antibody (Jackson Immuno Research, West Grove, PA, USA) at 1:200 dilution. Nuclei were stained with 0.5 μg/ml Hoechst 22358 and cells were analysed using an Olympus BX51 fluorescence microscope. Between 35 and 150 cells were counted in each experiment. Statistical differences were analysed by the unpaired t-test using the online GraphPad QuickCalcs software and statistical significance is expressed as *P<0.05 and **P<0.01. The experiments were repeated at least three times.
Drug treatments and cell viability
Doxorubicin, etoposide and paclitaxel were purchased from Sigma. For the 3-[4,5] dimethylthiazole-2,5-diphenyltetrazolium bromide (MTT) assay, cells were seeded at 5 × 103−104/well in 0.2 ml in 96-well plates. After 24 h, cells were treated with increasing concentrations of the drugs for 48 h. At the end of the experiment, the MTT assay was performed as described (Christodoulopoulos et al., 1999). The IC50 was calculated as the concentration of the drugs that resulted in 50% reduction of cell viability compared to untreated controls.
For colony-forming assays in the absence of drug, 2.5–5 × 103 cells/6-well plates were seeded, incubated for 48 h and plated at low density in 10 cm plates in triplicate to allow colony formation. After 14–21 days, the colonies were stained with crystal violet and counted. For drug treatment, YCC-B1, YCC-B2, GM847 and VA13 derivatives were seeded at 105 cells/6-well plates, whereas MCF-7 derivatives were seeded at 2.5 × 105 cells/6-well plates. The next day, cells were treated with the indicated concentrations of drugs for 24 h and then plated at low density in 10 cm plates in triplicate until colonies were visible and could be stained with crystal violet and counted. For accuracy, experiments resulting in less than 25 and more than 1500 colonies in the controls were not used. Relative numbers of colonies were calculated as a ratio between the numbers of colonies in the mutant hTR clones and the number of colonies in the vector clones. Comparisons between vector cells and each derivative were analysed by the unpaired t-test using the online GraphPad QuickCalcs software, and statistical significance is expressed as *P<0.05, **p<0.01 and ***P<0.001. All experiments were repeated at least three times.
Cell cycle analysis
For cell cycle analysis, cells were either left untreated or treated with doxorubicin as described above. Samples were collected after 1 and 3 days of treatment and fixed with ice-cold ethanol. Before analysis, cells were centrifugated, resuspended in 500 μl of cold PBS containing 200 μg/ml of RNAse A and incubated overnight at 4°C. The following day, cells were incubated for 15 min with 25 μg of propidium iodide and the DNA content was measured by flow cytometry (Becton Dickinson, San Jose, CA, USA, fluorescence activated cell sorter, FACS), followed by quantification with CellQuest software. For each sample, at least 20 000 events were collected. The experiments were repeated at least three times. Statistical differences between vector cells and each mutant hTR clone were analysed by the unpaired t-test using the online GraphPad QuickCalcs software and are expressed as *P<0.05 and **P<0.01.
Brown JM, Attardi LD . (2005). Nat Rev Cancer 5: 231–237.
Bryan TM, Englezou A, Dalla Pozza L, Dunham MA, Reddel RR . (1997). Nat Med 3: 1271–1274.
Bryan TM, Englezou A, Gupta J, Bacchetti S, Reddel RR . (1995). EMBO J 14: 4240–4248.
Castedo M, Perfettini JL, Roumier T, Andreau K, Medema R, Kroemer G . (2004). Oncogene 23: 2825–2837.
Cerone MA, Londono-Vallejo JA, Bacchetti S . (2001). Hum Mol Genet 10: 1945–1952.
Chen Z, Koeneman KS, Corey DR . (2003). Cancer Res 63: 5917–5925.
Christodoulopoulos G, Malapetsa A, Schipper H, Golub E, Radding C, Panasci LC . (1999). Clin Cancer Res 5: 2178–2184.
Chung HK, Cheong C, Song J, Lee HW . (2005). Curr Mol Med 5: 233–241.
Cimino-Reale G, Pascale E, Battiloro E, Starace G, Verna R, D'Ambrosio E . (2001). Nucleic Acids Res 29: E35.
Collins K, Mitchell JR . (2002). Oncogene 21: 564–579.
Dunham MA, Neumann AA, Fasching CL, Reddel RR . (2000). Nat Genet 26: 447–450.
Feng J, Funk WD, Wang S-S, Weinrich SL, Avilion AA, Chiu C-P et al (1995). Science 269: 1236–1241.
Folini M, De Marco C, Orlandi L, Daidone MG, Zaffaroni N . (2000). Eur J Cancer 36: 2137–2145.
Guiducci C, Cerone MA, Bacchetti S . (2001). Oncogene 20: 714–725.
Hahn WC, Stewart SA, Brooks MW, York SG, Eaton E, Kurachi A et al (1999). Nat Med 5: 1164–1170.
Harley CB, Futcher AB, Greider CW . (1990). Nature 345: 458–460.
Herbert B-S, Pitts AE, Baker SI, Hamilton SE, Wright WE, Shay JW et al (1999). Proc Natl Acad Sci USA 96: 14276–14281.
Jordan MA, Toso RJ, Thrower D, Wilson L . (1993). Proc Natl Acad Sci USA 90: 9552–9556.
Karlseder J, Broccoli D, Dai Y, Hardy S, de Lange T . (1999). Science 283: 1321–1325.
Kelland LR . (2005). Eur J Cancer 41: 971–979.
Kim MM, Rivera MA, Botchkina IL, Shalaby R, Thor AD, Blackburn EH . (2001). Proc Natl Acad Sci USA 98: 7982–7987.
Kim NW, Piatyszek MA, Prowse KR, Harley CB, West MD, Ho PLC et al (1994). Science 266: 2011–2015.
Kirk KE, Harmon BP, Reichardt IK, Sedat JW, Blackburn EH . (1997). Science 275: 1478–1481.
Kondo Y, Kondo S, Tanaka Y, Haqqi T, Barna BP, Cowell JK . (1998). Oncogene 16: 2243–2248.
Lee K-H, Rudolph KL, Ju Y-J, Greenberg RA, Cannizzaro L, Chin L et al (2001). Proc Natl Acad Sci USA 98: 3381–3386.
Li S, Rosenberg JE, Donjacour AA, Botchkina IL, Hom YK, Cunha GR et al (2004). Cancer Res 64: 4833–4840.
Lin J, Smith DL, Blackburn EH . (2004). Mol Cell Biol 15: 1623–1634.
Ludwig A, Saretzki G, Holm PS, Tiemann F, Lorenz M, Emrich T et al (2001). Cancer Res 61: 3053–3061.
Marusic L, Anton M, Tidy A, Wang P, Villeponteau B, Bacchetti S . (1997). Mol Cell Biol 17: 6394–6401.
Masutomi K, Possemato R, Wong JM, Currier JL, Tothova Z, Manola JB et al (2005). Proc Natl Acad Sci USA 102: 8222–8227.
Masutomi K, Yu EY, Khurts S, Ben-Porath I, Currier JL, Metz GB et al (2003). Cell 114: 241–253.
Misawa M, Tauchi T, Sashida G, Nakajima A, Abe K, Ohyashiki JH et al (2002). Int J Oncol 21: 1087–1092.
Nakamura M, Masutomi K, Kyo S, Hashimoto M, Maida Y, Kanaya T et al (2005). Hum Gene Ther 16: 859–868.
Park KH, Rha SY, Kim CH, Kim TS, Yoo NC, Kim JH et al (1998). Int J Oncol 13: 489–495.
Sharma GG, Gupta A, Wang H, Scherthan H, Dhar S, Gandhi V et al (2003). Oncogene 22: 131–146.
Shay JW, Bacchetti S . (1997). Eur J Cancer 33: 787–791.
Shin KH, Kang MK, Dicterow E, Kameta A, Baluda MA, Park NH . (2004). Clin Cancer Res 10: 2551–2560.
Smith CD, Blackburn EH . (1999). J Cell Biol 145: 203–214.
Stewart SA, Ben-Porath I, Carey VJ, O'Connor BF, Hahn WC, Weinberg RA . (2003). Nat Genet 33: 492–496.
Takai H, Smogorzewska A, de Lange T . (2003). Curr Biol 13: 1549–1556.
Tentori L, Portarena I, Barbarino M, Balduzzi A, Levati L, Vergati M et al (2003). Mol Pharm 63: 192–202.
Ward RJ, Autexier C . (2005). Mol Pharm 68: 779–786.
Wen J, Cong YS, Bacchetti S . (1998). Hum Mol Genet 7: 1137–1141.
Xu L, Blackburn EH . (2004). J Cell Biol 167: 819–830.
Yu G-L, Bradley JD, Attardi LD, Blackburn EH . (1990). Nature 344: 126–132.
Zhang X, Mar V, Zhou W, Harrington L, Robinson MO . (1999). Genes Dev 13: 2388–2399.
We thank Dr Silvia Bacchetti for cell lines, constructs and for comments on the manuscript, Dr Sun Young Rha for cell lines, Dr Koromilas for the anti-p53 antibody and Graeme Nimmo for technical help with the FACS and critical discussion. This work was funded by Canadian Institute of Health Research IAO-64655 and Cancer Research Society Inc. grants to CA and by a Grant from l'Association pour la Recherche contre le Cancer to AL-V. CA is a chercheur-boursier of the Fonds de la Recherche on Santé du Quebec. MAC is supported by a US Army Department of Defense Breast Cancer Research Program Award.
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Cerone, M., Londoño-Vallejo, J. & Autexier, C. Mutated telomeres sensitize tumor cells to anticancer drugs independently of telomere shortening and mechanisms of telomere maintenance. Oncogene 25, 7411–7420 (2006). https://doi.org/10.1038/sj.onc.1209727
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