During metastases, cancer cells are temporarily exposed to the condition in which interactions with extracellular environment can be restricted (anchorage-independence). We demonstrate that the sensitivity of prostate cancer cell lines, DU145 and PC-3, to genotoxic treatment (cisplatin and γ-irradiation) increased several folds when cells were forced to grow in anchorage-independence. This enhanced drug sensitivity was associated with a severe impairment of homologous recombination-directed DNA repair (HRR). The mechanism involves Rad51, which is the major enzymatic component of HRR. The protein level of Rad51 and its recruitment to DNA double-strand breaks (DSBs) were both attenuated. Rad51 deficiency in anchorage-independence was not associated with Rad51 promoter activity and was not compensated by a constitutive overexpression of Rad51 cDNA. Instead, Rad51 protein level and its ability to colocalize with DSBs were restored in the presence of proteosome inhibitors, or when cells from the suspension cultures were allowed reattachment. Presented results indicate that anchorage-independence sensitizes prostate cancer cells to genotoxic agents; however, it also attenuates faithful component of DNA repair by targeting stability of Rad51. This temporal attenuation of HRR may contribute to the accumulation mutations after DNA damage and possibly the selection of new adaptations in cells, which survived genotoxic treatment.
A transition from androgen-dependent benign prostatic hyperplasia to a highly invasive and untreatable metastatic disease happens in about 30% of prostate cancer patient (Karan et al., 2003). This deadly transition relies on newly acquired cellular properties that allow prostate cancer cells to escape from the primary site, get into circulation, invade distant tissues and finally to re-establish cell proliferation in a new, tissue-specific environment. The question, how prostate cancer cells develop these multiple cellular adaptations in a time frame, which represents only a fraction of a lifespan, still remains to be answered.
A compromise in the stability of the genome may increase rate of mutations. In fact, genomic instability is frequently present in cancer cell lines (Sieber et al., 2003). There are several ways how genomic instability could develop: (i) telomere dysfunction; (ii) aberrant chromosomal segregation; and finally (iii) dysfunction of cellular mechanisms responsible for DNA repair. This brings us to a problem in which disproportion between faithful and unfaithful DNA repair mechanisms may result in the accumulation of spontaneous mutations, possibly leading to the selection of a malignant phenotype (Mills et al., 2003). Prostate cancer cells have already been characterized by multiple defects at the level of postreplicative inspection of DNA. In particular, mismatch repair (MMR) proteins such as MLH1, PMS1, PMS2, MSH2 and MSH6 are either strongly downregulated or are not detectable in PC-3, DU145 and LNCaP cells (Chen et al., 2001, 2003; Yeh et al., 2001). Although it has been postulated that insufficient MMR may contribute to genomic instability in prostate cancer, the question remains as to how these mutations are formed. One possibility involves unfaithful repair of double-strand breaks (DSBs). DSBs are lethal when left unrepaired in cells replicating DNA. They may arise from ionizing radiation, reactive oxygen species, a variety of genotoxic chemicals including cisplatin and mitomycin and also when replication forks encounter other DNA lesions (Hoeijmakers, 2001; Khanna and Jackson, 2001). To support cell survival after DSBs, at least one of the two major DNA repair mechanisms has to be activated: (i) usually error-free homologous recombination DNA repair (HRR); or (ii) usually error-prone nonhomologous end joining (NHEJ). The choice between DNA repair mechanisms can be controlled, at least partially, by the availability of a DNA template. Proliferating cells utilize newly synthesized template supplied during DNA replication. Alternatively, cells may simply link the ends of a DSB without any template using the end binding KU70/KU80 complex and DNA-PK, followed by DNA ligation with XRCC4 - DNA ligase 4 complex (Pierce and Jasin, 2001; Lundin et al., 2002). This rapid repair of DNA by NHEJ, however, may cost the cell a gain or loss of small fragments of DNA (Hoeijmakers, 2001).
When a DNA template is available, cells have a chance of utilizing DNA repair mechanism, which involves homologous recombination. Rad51 is a structural and functional eukaryotic homologue of the bacterial RecA recombinase, which is considered the key enzyme for HRR (Baumann and West, 1998; Tombline and Fishel, 2002; Van Komen et al., 2002). Following the detection of DSBs, which involves activation of a series of DNA-dependent kinases, such as ATM, ATR and DNA PK, Rad51 is translocated to the sites of damaged DNA (Davies et al., 2001). In parallel, the ATM-activated 5′–3′ endonuclease complex (Rad50–MRE11–NBS1) exposes both 3′ ends of DNA at the DSB (Petrini, 2000; Zhu et al., 2000). The ends are initially protected by the single-stranded DNA (ssDNA)-binding protein RPA, which is subsequently replaced by Rad51 in a process involving initial binding of Rad52 into the ssDNA–RPA complex (Chen et al., 1999; Essers et al., 2002). As a result, newly formed Rad51 nucleoprotein filaments are directly involved in homology search and strand invasion (Sigurdsson et al., 2002; Tombline and Fishel, 2002; Tombline et al., 2002).
Our results indicate a severe impairment of HRR in prostate cancer cells exposed to anchorage-independent culture conditions. The mechanism involves downregulation of Rad51 protein level and its absence at the site of damaged DNA. This Rad51 deficiency was not associated with Rad51 promoter activity and was not compensated by a constitutive overexpression of Rad51 cDNA. Instead, Rad51 protein stability was efficiently restored in the presence of proteosome inhibitors, epoxomycin and lactacystin, or when cells from suspension cultures were allowed to reattach. These data indicate that prostate cancer cells have a tendency of loosing faithful component of DNA repair when forced to survive anchorage-independence.
Our initial observations indicate that two metastatic prostate cancer cell lines, PC-3 and DU145, become sensitive to genotoxic agents (cisplatin, mitomycin and γ-irradiation), when forced to grow in anchorage-independence – the condition in which interactions with extracellular matrix proteins are restricted (Valentinis et al., 1998). For example, cisplatin applied to the suspension cultures, at 0.75 μg/ml for 48 h, caused a dramatic 5.2-fold and 2.5-fold decrease in survival of DU145 and PC-3 cells, respectively (Figure 1a). In comparison, control cells from monolayer cultures were much more resistant, showing only a marginal (not significant) decreases in cell survival following the same treatments. Although at higher doses (1.5 and 3.0 μg/ml), cisplatin was also toxic for cells kept in monolayer, it was consistently more effective when applied to cells in anchorage-independence (Figure 1a). Similar results were obtained following mitomycin treatment used at concentrations ranging from 0.0625 to 1 μg/ml (not shown) and following γ-irradiation, in the experiment in which clonogenic abilities of DU145 and PC-3 cells were evaluated (Figure 1b).
Although prostate cancer cell lines are capable of proliferating in anchorage-independent culture condition, both cell cycle distribution studies (panel c) and BrdU pulse labeling (panel d) indicated that in suspension cultures the fraction of cells, which replicate DNA, is significantly lower than in monolayer. Considering that cytotoxic effects of cisplatin depend mostly on the formation of DSBs, when replication forks collapse on cisplatin-induced primary lesions (guanidine intrastrand cross-links) (Hoeijmakers, 2001; Wozniak and Blasiak, 2002), cells that proliferate faster should be, at least in theory, more sensitive to the treatment with this genotoxic agent. However, in our experimental setting, slower proliferating cells in anchorage-independence (Figure 1c and d) are more sensitive to genotoxic treatments (Figure 1a and b). To address this discrepancy, first we had to determine whether observed drug sensitivity in anchorage-independence relays on enhanced DNA damage following genotoxic treatments. We have measured the effectiveness of cisplatin to induce DNA damage first indirectly by immunodetection of the phosphorylated histone H2AX (γ-H2AX) (Figure 2a) and later by the comet assay following γ-irradiation (Figure 2b). After cisplatin treatment, both DU145 and PC-3 cells showed a slightly lower percentage of cells positive for immunolabeling with anti γ-H2AX antibody in anchorage-independence (Figure 2a). Quantitatively, 35 and 34% decreases in γH2AX labeling calculated for DU145 and PC-3 cells in suspension did not reach statistical significance. The comet assay (Figure 2b), which in its alkaline version reveals both single and double DSBs, indicated that the initial extent of DNA damage after γ-irradiation (20 Gy) was quite similar when suspension cultures were compared to monolayer cultures (0 min of DNA repair). Surprisingly, DNA repair evaluated at 5, 15 and 30 min after γ-irradiation was also similar between these two conditions, indicating that neither the initial extent of DNA damage nor the overall DNA repair could explain drug sensitivity acquired by prostate cancer cells in anchorage-independence.
Since the results presented so far did not explain why the cells in anchorage-independence are more sensitive to DNA damage, we asked whether mechanisms known to repair DSBs, that is, HRR and NHEJ, contribute to DNA repair when prostate cancer cells are forced to grow in suspension.
DNA repair of DSBs in anchorage-independence
DSBs are the most lethal of all DNA lesions caused by genotoxic agents. Since prostate cancer cell lines are particularly sensitive to genotoxic treatments when kept in suspension (Figure 1a and b), we asked whether mechanisms involved in the repair of DSBs are fully active when cells are deprived of attachment. NHEJ is considered as a quick way to ligate DSBs, which operates mostly during the G1 phase of the cell cycle (Richardson and Jasin, 2000). We have used well-documented cell-free assay to evaluate NHEJ (Labhart, 1999). The results in Figure 3a show a slight although reproducible reduction of NHEJ, measured in the presence of nuclear extracts from the suspension cultures in comparison to monolayer cultures. Densitometric analysis of the secondary bands, which were formed after the in vitro ligation of the linear plasmid showed 9 and 11% reduction in suspension cultures (not significant), when compared to corresponding monolayer cultures of DU145 and PC-3 cells, respectively. Considering that NHEJ was only slightly affected by anchorage-independence, it was quite unlikely that observed drug sensitivity in anchorage-independence could rely solely on this inhibition.
The assay to evaluate repair of DSBs by HRR is based on reconstruction of the wild-type green fluorescent protein (GFP) from two nonfunctional heteroallelic fragments of GFP cDNA delivered into cells by the pDRGFP expression vector (Pierce et al., 1999). The DRGFP construct was introduced into PC-3 and DU145 cells. Two stable clones of PC-3/DRGFP cells (clone #5 and #15) carrying a single copy of the DRGFP construct were transiently transfected with two additional expression vectors; the first containing rare cutting endonuclease, I-SceI, to generate DSBs in GFP cDNA and the second containing red fluorescent protein with a mitochondrial localization signal, to monitor efficiency of transfection (see Material and methods). Figure 3c illustrates that in monolayer both clones of PC-3/DRGFP are characterized by a similar level of GFP reconstitution, which in this detection system is possible only when homologous recombination is involved in repair of I-SceI -inflicted DSBs. Quantitatively, an average of 1 and 1.23% of cells showed nuclear green fluorescence detected in monolayer cultures from clone #5 and clone #15, respectively. In contrast, levels of GFP reconstitution in anchorage-independent cultures were practically undetectable (Figure 3c). Similarly, a severe inhibition of HRR in anchorage-independence was observed in two clones of DU145/DRGFP cells (clone #9 and #12), which happened despite of the fact that the reconstitution of GFP function in DU145 cells was several-folds higher than in PC-3 cells.
Although the inhibition of HRR in suspension cultures was remarkably high (Figure 3c), at least part of this inhibition could depend on the fact that in suspension cultures the fraction of cells found in G1 phase of the cell cycle increased from 62 to 75% in DU145 cells and from 52 to 65% in PC-3 cells. However, independently from this partial inhibition of the cell cycle progression, both BrdU incorporation and flow cytometric data indicate that a substantial number of cells still replicate DNA in suspension (Figures 1c and d). Therefore, from the cell cycle point of view, HRR should be still detectable.
Expression and subcellular localization of DNA repair proteins
Having established that HRR is impaired and NHEJ is only slightly attenuated by the condition of anchorage-independence, we have evaluated levels and subcellular localization of several major DNA repair proteins, which could contribute to DNA repair in our experimental setting. In comparison to monolayer cultures (M), Rad51 protein, which is the major enzymatic component of the HRR, decreased dramatically in both PC-3 and DU145 cells cultured in suspension (S) for 48 h (Figure 4a). Other partners of Rad51 in the process of homologous recombination, Rad52 and Rad54, were not affected by anchorage-independence, although their levels detected in PC-3 cells were significantly lower than in DU145 cells. For NHEJ, proteins like KU70 and KU80 were also not affected by the conditions of anchorage-independence, showing similar levels in both DU145 and PC-3 cells.
We have evaluated also mismatch repair proteins (MSH2, MSH6, PMS2 and MLH1), since these proteins are often downregulated in prostate cancer cell lines (Chen et al., 2001). Our results partially confirmed previous findings, by showing that MLH1 and PMS2 proteins were strongly downregulated in DU145 cells. These changes were not confirmed, however, in our PC-3 cells (Figure 4b). In respect to anchorage-independence, MSH6 was the only protein among the mismatch repair system, which showed a moderate decrease in suspension cultures (Figure 4b).
A dramatic decrease of Rad51 protein in anchorage-independence was additionally confirmed by immunocytofluorescence. We took advantage of the fact that DNA repair proteins, including Rad51, form functional complexes at DNA lesions (nuclear foci), which can be visualized in cells with DSBs (Trojanek et al., 2003). As shown in Figure 4c, abundant Rad51 nuclear foci (detected in about 25% of DU145 and in about 15% PC-3 cells) were present in monolayer cultures of DU145 and PC-3 cells at 6 h following the cisplatin treatment. In contrast, Rad51 nuclear foci were not detectable in cells from the corresponding suspension cultures (Figure 4c, green fluorescence). Importantly, for the comparison of Rad51 nuclear foci formation, we have selected cells, which were characterized by comparable levels of DNA damage (γH2AX nuclear staining; red fluorescence). Therefore, independently from the fact that cells from suspension and monolayer cultures had similar levels and patterns of γH2AX immunolabeling (indication of DNA DSBs), cells cultured in suspension were unable to form Rad51 nuclear foci. Superimposition of γH2AX images (red fluorescence) with Rad51 images (green fluorescence) revealed expected partial colocalization (yellow fluorescence) between these two proteins, detected exclusively in monolayer cultures (Overlap).
Anchorage-independence and the stability of Rad51
We have attempted to overcome the loss of the Rad51 protein in anchorage-independence. First, we transfected DU145 cells with the expression vector containing human Rad51 cDNA under the control of cytomegalovirus (CMV) constitutive promoter (pcDNA3/Rad51). In monolayer cultures, two stable clones of DU145 cells, DU/Rad51#2 and DU/Rad51#3, revealed two-fold and three-fold higher levels of Rad51 protein in comparison to parental DU145 cells (Figure 5a). When these new cell lines were forced to grow in suspension, the levels of Rad51 protein dropped dramatically and independently from the fact that CMV promoter was responsible for Rad51 expression in this setting. Next, we have evaluated Rad51 promoter activity. The histogram illustrated in Figure 5b shows, surprisingly, that Rad51 promoter is 41% more active in suspension cultures when compared to monolayer culture (P=0.052). This almost significant difference was calculated from four independent experiments performed in triplicate (n=12). Also unexpected was the result from Northern blot analysis, showing, in contrast to elevated Rad51 promoter activity, slightly lower level of Rad51 mRNA in suspension cultures (inset to Figure 5). These results could reflect unsuccessful compensatory effort of cells against Rad51 downregulation in anchorage-independence, which seem to take place on both protein stability and mRNA stability levels. To address the possibility of Rad51 protein degradation, the monolayer and suspension cultures of DU145 and PC-3 cells were exposed to proteosome inhibitors, epoxomycin and lactacystin (Rodgers and Dean, 2003). Figure 6a shows that epoxomycin applied at 100 nM for 48 h reversed the process of Rad51 downregulation in both DU145 and PC-3 cells cultured in suspension. The same treatment applied to monolayer cultures did not affect the level of Rad51 protein. The blots were subsequently probed with anti-Grb-2 antibody to monitor equal loading condition and with antiubiquitin antibody to visualize accumulation of ubiquitinated proteins following epoxomycin -mediated proteosome inhibition. Similar results were obtained with another proteosome inhibitor, lactacystin.
The reversibility of Rad51 downregulation in anchorage-independence was further illustrated by the experiment in which DU145 cells kept in suspension cultures for 48 h were replated on regular culture dishes. After reconstituting attachment, DU145 cells were capable again of forming Rad51 nuclear foci (Figure 6b, lower panel), whose morphology was comparable to that of Rad51 foci detected in regular monolayer cultures (Figure 6b, upper panel). Importantly, Rad51 nuclear foci formed after replating of cells from suspension cultures into adhesive surfaces were functional, since multiple colocalization centers between Rad51 and γH2AX were also detected (Figure 6c). In summary, our results indicate that the lose of Rad51 in anchorage-independence is reversible; is controlled mostly on the level of Rad51 proteosomal degradation; and that cell adhesion plays an important role in maintaining Rad51 protein stability and function (Figure 6).
The results presented here prompted us to speculate how new mutations could accumulate during the course of metastatic invasion from the prostate. The mechanism involves a severe impairment of HRR in prostate cancer cells exposed to anchorage-independence. Further analysis revealed that Rad51 protein levels decline dramatically in cells cultured in suspension; and that proteosome -mediated degradation of Rad51 rather than Rad51 promoter activity contributed to this inhibition. Importantly, this downregulation of Rad51 protein in cells devoid of the attachment was efficiently restored by allowing reattachment, or by the treatment of suspension cultures with proteosome inhibitors.
During metastatic spread, cancer cells are exposed to the condition of anchorage-independence (restricted interaction with extracellular matrix). This may happen when cells break out from the primary tumor, digest their way into the circulation and survive within the blood stream until successful reinstatement of new extracellular connections within newly invaded tissues. However, to complete this transition, the cells are pressured to develop new adaptations and often became quite different from those which were left at the primary site of the tumor. From this perspective, our results indicate that in anchorage-independence prostate cancer cells are forced to repair DSBs without homologous recombination, almost exclusively by less faithful NHEJ.
Rad51, which is considered the key enzyme for HRR and is shown here as the primary target in the attenuation of HRR, forms nucleoprotein filaments on ssDNA and mediates homologous pairing and strand exchange between homologous DNA duplexes (Baumann and West, 1998; Tombline and Fishel, 2002; Van Komen et al., 2002). In normal circumstances, the expression of Rad51 is cell cycle regulated, with a pick of expression at the S/G2 phase of the cell cycle (Xia et al., 1997). Deregulations of Rad51 expression, both inhibition and elevated expression, compromises integrity of the genome. First reports came from immortalized cell lines and tumor cells in which elevated expression of Rad51 was associated with an aberrant recombination between short repetitive elements and homologous sequences (Xia et al., 1997; Flygare et al., 2001; Raderschall et al., 2002). Further analyses indicated elevated Rad51 protein levels in cells from chronic myelogenous leukemia (CML) and even higher Rad51 expression in cells from the blast crisis -stage of the disease characterized by a profound genome instability (Slupianek et al., 2001). The evidence for abnormal recombination was provided by Maria Jasin, who demonstrated the presence of a novel class of recombination product in cells that overexpressed Rad51 (Richardson et al., 2004).
Genomic aberrations are even more pronounced in cells with a severe downregulation Rad51, when the shift from HRR towards less faithful NHEJ could be anticipated (Valerie and Povirk, 2003; Bindra et al., 2004). In one study, downregulation of Rad51 and the attenuation of HRR were observed in several cancer cell lines forced to grow in prolonged hypoxia. The decrease of Rad51 was initially detected by RNA microarray and was associated with downregulation of Rad51 promoter activity in hypoxia (Bindra et al., 2004). Chromosomal damage and early embryonic lethality were detected in mice knockouts of Rad51 (Lim and Hasty, 1996). Multiple chromosomal aberrations and radiosensitivity were found in chicken cells with inducible loss of Rad51 expression (Sonoda et al., 1998) and in prostate cancer cells with ribozyme -mediated downregulation of Rad51 (Collis et al., 2001).
In our study, downregulation of Rad51 observed in anchorage-independence (Figures 3 and 4) did not involve transcriptional suppression, instead was mainly controlled by proteosome -mediated Rad51 degradation (Figure 5). It has been already proposed that protein stability could play a role in Rad51 regulation. For instance, caspase 3 -mediated degradation of Rad51 has been shown in cells committed to apoptosis after the exposure to ionizing radiation (Huang et al., 1999). In a different report, the binding between Rad51 and ubiquitin-conjugating enzyme Ubc9 could suggest a possibility of ubiquitination of Rad51 (Kovalenko et al., 1996).
We have demonstrated that anchorage-independence sensitizes prostate cancer cells to different genotoxic agents (Figure 1) and in parallel, we found that HRR is severely impaired. However, in our experimental setting, the extent of DNA repair by HRR ranges from about 1% in PC-3 cells to about 7% in DU145 cells and NHEJ is only slightly downregulated in both cell lines. Owing to these discrepancies, one could consider the presence of an additional mechanism, which together with attenuated HRR, could explain observed drug sensitivity in anchorage -independence (Figure 1). One well-established signaling pathway known to protect prostate cancer cells from apoptosis in anchorage-independence is the pathway from insulin-like growth factor I receptor (IGF-IR) (Reiss et al., 1998a, 2000). The IGF-IR sends multiple antiapoptotic signal including: PI3-kinase -mediated activation of AKT; IRS-1 -mediated recruitment of Grb-2 and subsequent activation of the Ras-MAP kines pathway; and finally IGF-I -mediated translocation of Raf to mitochondria, which depend on IGF-IR C-terminal domain responsible for 14-3-3 binding (Peruzzi et al., 1999). Different molecular manipulations targeting the function of IGF-IR were quite effective in challenging the survival of prostate cancer cells, but only when the treatments were applied to cells growing in suspension (Reiss et al., 1998a, 1999). Beside that, recent attempts to explain resistance of prostate cancer cells to anoikis (apoptosis induced by anchorage-independence) excluded BCL-2 component in this process (Bondar and McConkey, 2002), further supporting a possible role of IGF-IR in the protection of cells in anchorage-independence. From this perspective, one could speculate that the attenuation of IGF-IR -dependent and Bcl-2 -independent survival pathway could work in synergy with the impaired HRR, causing drug sensitivity in anchorage- independence. Importantly, in cells capable of surviving genotoxic treatments, anchorage-independence could create a temporal mutagenic environment. With mismatch repair proteins being often attenuated (Chen et al., 2001) and with severely compromised HRR, the mistakes could accumulate and propagate to the next generation of prostate cancer cells, when DNA repair machinery is forced to operate in anchorage-independence.
Materials and methods
DU145 (ATCC #HTB-81) and PC-3 (ATCC #CRL-1435) are human prostate cancer cell lines derived from brain and bone metastases, respectively. In monolayer cultures, both cell lines grow in DMEM (GIBCO BRL, Grand Island, NY, USA) supplemented with 50 U/ml penicillin, 50 ng/ml streptomycin and 10% FBS (SIGMA, St Louis, MO, USA). Anchorage-independent growth of DU145 and PC-3 cells was evaluated in 60 mm culture dishes coated with poly(2-hyrdoxyethyl methacrylate) (PolyHEMA) (Aldrich), by the methodology routinely used in our laboratory (Reiss et al., 1998b; Valentinis et al., 1998; Wang et al., 2001). Cell survival was evaluated in both monolayer and suspension cultures by trypan blue exclusion test at 48 h following cisplatin (ranging from 0 to 3 μg/μl) or mitomycin (ranging from 0 to 1 μg/ml) treatments. For the clonogenic assay, cells were plated at clonal density (1 × 103) either on uncoated six-well plates (Costar, Corning, NY, USA), or on plates coated with PolyHEMA (to prevent cell attachment). After 24 h, adherent cells or cells in suspension were irradiated with 0, 5, 10 and 20 Gy by utilizing γ-irradiator (J.L Shepherd Mark I, Model 30-1). Colonogenic potential was evaluated by the inspection of the resulting clones (more than 50 cells/clone) at 10 days after irradiation and following 0.25% Brilliant Blue staining. The SF was calculated as a ratio of the number of colonies and the product of the number of cells plated and plating efficiency (Chendil et al., 2004).
Cell cycle analysis
Aliquots of cells, 1x106/ml, from both monolayer and suspension cultures growing in 10% FBS, were fixed in 70% ethanol at 4°C for 30 min. The cells were centrifuged at 1600 r.p.m. and the resulting pellets were resuspended in 1 ml of freshly prepared propidium iodide/RNaseA solution. Cell cycle distribution was analysed by FACSCalibur (Becton-Dickinson, Franklin Lakes, NJ, USA) using the CellQuest Program (Skorski et al., 1995). BrdU pulse labeling (60 min) and accumulative labeling (24 h) of newly synthesized DNA was performed in monolayer and suspension cultures of PC-3 and DU145 cells by utilizing In Situ Cell Proliferation Kit, FLUOS (Roche, Molecular Biochemicals).
To inflict DNA damage, cells received 20 Gy of γ-irradiation. The DNA damage was analysed by alkaline single-cell gel electrophoresis (comet assay), as previously described (Schindewolf et al., 2000), with some modifications. Briefly, an aliquot of 1 × 105 cells was suspended in 0.75% LMP agarose and spread on microscopic slides precoated with 0.5% agarose (Sigma). The cells were lysed for 1 h at 4°C in a buffer containing 2.5 M NaCl, 100 mM EDTA, 1% Triton X-100 and 10 mM Tris, pH 10. The slides were placed in an electrophoresis unit and DNA was allowed to unwind for 40 min in the running buffer (300 mM NaOH, 1 mM EDTA, pH>13). Electrophoresis was conducted for 30 min at 0.73 V/cm. The slides were neutralized with 0.4 M Tris, pH 7.5, stained with 2 mg/ml 4′ 6′ diamedino-2-phenylindole (DAPI) and covered with cover slips. A total of 100 images were randomly selected from each sample and the Olive tail moment was calculated by Comet 6.0 (Kinetic Imaging, Liverpool, UK) image analysis system.
The cell-free NHEJ assay was followed (Labhart, 1999) and nuclear lysates were prepared according to the protocol previously described (Trojanek et al., 2003). Briefly, NHEJ reactions were performed in the following conditions: 10 μg of nuclear lysate; 1 mM ATP, 0.25 mM dNTPs, 25 mM Tris-Acetate (pH 7.5), 100 mM potassium acetate, 10 mM magnesium acetate and 1 mM DTT. After 5 min of preincubation at 37°C, the reaction mixture was supplemented with the substrate (400 ng of XhoI–XbaI linearized pBluescript KS+). The reaction was incubated for 1 h at 37°C to ligate the plasmid and treated with proteinase K to digest DNA bound proteins. Products of NHEJ reaction were resolved in 0.5% agarose gel containing 0.5 μg/ml of ethidium bromide. For each experiment, the following control reactions were performed: (i) different quantities of the substrate (linearized plasmid DNA); (ii) different concentrations of the nuclear extracts; (iii) elimination of samples in which nuclear extracts were contaminated with genomic DNA; (iv) presence of the inhibition of DSBs joining by neutralizing antibody against KU70.
The plasmid pDR-GFP (generously provided by Dr M Jasin, Sloan-Kettering Cancer Center, NY, USA) (Pierce et al., 1999) was stably introduced, by calcium phosphate reagent (Promega, Madison WI, USA), into PC-3 and DU145 cells. Stable clones were selected in puromycin (2 μg/ml) and characterized by Southern blot to estimate DR-GFP copy number. pDRGFP contains a nonactive gene for GFP (SceGFP) and a fragment of the GFP gene as a donor for homologous repair. The SceGFP cassette has an inactivating insertion, which consists of two STOP codons and a restriction site for the rare cutting endonuclease, I-SceI. When I-SceI is expressed in DR-GFP-expressing clones, it inflicts DSBs within the SceGFP fragment providing a signal for homologous recombination and the reconstruction of functional GFP (Pierce et al., 1999). To compare HRR in monolayer versus anchorage-independent culture conditions, DR-GFP-containing cells were transiently transfected with 3 μg of pCβA-Sce and 1 μg of pDsRed1-Mito (Clontech, Palo Alto, CA, USA) utilizing FuGENE 6 reagent (Roche, Indianapolis, IN, USA). At 24 h after transfection, cells were trypsinized and replated to normal or PolyHEMA precoated 60 mm culture dishes. DNA repair by HRR was evaluated by counting cells with both green nuclear fluorescence and red mitochondrial fluorescence versus all positively transfected cells (only red cells+red and green cells) at 96 h after replating. Cells showing green nuclear fluorescence only were sporadically detected; however, these cells were not considered in the calculation.
Southern blot was utilized to screen for DR-GFP and to determine DR-GFP copy number in selected clones. Genomic DNA was extracted by Puregene DNA isolation kit (Gentra Systems), according to the manufacturer's recommendations. Southern blots were carried out according to the methodology previously described (Ludwig et al., 1996). The GFP probe (800 bp) was obtained from pDR-GFP by SalI/HindIII digestion.
To evaluate levels of DNA repair proteins, monolayer and suspension cultures were lysed with 400 μl of lysis buffer (50 mM HEPES; pH 7.5; 150 mM NaCl; 1.5 mM MgCl2; 1 mM EGTA; 10% glycerol; 1% Triton X-100; 1 mM phenylmethylsulfonyl fluoride (PMSF)); 0.2 mM Na-orthovanadate and 10 μg/ml aprotinin). To improve recovery of DNA bound proteins, DNAse was added to the lysis buffer. In total, 50 μg protein aliquots were separated on a 4–15% gradient SDS–PAGE (BioRad) and transferred onto nitrocellulose membranes. The resulting blots were probed with following antibodies: anti-Rad51 (Ab-1, Oncogene); anti-Rad52 (H-300; Santa Cruz), anti-Rad54 (H-152 Santa Cruz), anti-Ku70 (Serotec, UK); and anti-Ku80 (Serotec, UK), anti-MSH6 (H-141, Santa Cruz), anti-PMS2 (#556415, Pharmingen), anti-MSH2 (NA27, Oncogene) and anti-MLH1 (PC56, Oncogene). Anti Grb-2 antibody (Transduction Laboratories, Lexington, KY, USA), was used as a control to monitor equal loading conditions (Del Valle et al., 2002). For experiments in which epoxomycin and lactacystin treatments were applied, Western blots were additionally probed with antiubiquitin antibody (U5379, Sigma) to evaluate accumulation of ubiquitinated proteins following proteosome inhibitions.
Before immunolabeling, cells were cultured in the presence of 10% FBS, either as monolayer or suspension cultures. The treatment with cisplatin (1 μg/ml) was applied for 6 h in both culture conditions. For monolayer experiments, cells were plated on poly D-lysine-coated Lab-Tek Chamber Slides (Nalge Nunc International). For the experiments in anchorage-independence, cells from suspension cultures were attached to poly D-lysine-coated slides by utilizing cytospin 3 centrifuge (Thermo Shandon). Subsequently, all cells were fixed and permeablized with the buffer containing 0.02% Triton X-100 and 4% formaldehyde in PBS. Fixed cells were washed 3 × in PBS and blocked in 1% BSA for 30 min at 37°C. RAD51 was detected by mouse anti-RAD51 monoclonal antibody (UBI, Lake Placid, NY, USA) followed by FITC-conjugated goat anti-mouse secondary antibody (Molecular Probes). Phosphorylated form of histone H2AX (γH2AX) was detected by a mouse monoclonal antibody, which recognizes phosphorylated serine within the 134–142 aa fragment of human histone H2A.X (UBI) and rhodamine-conjugated goat anti-mouse secondary antibody (Molecular Probes). Negative controls were performed in the presence of irrelevant antibody replacing primary antibody, or in the absence of primary antibody. In all cases, DNA was counterstained with DAPI. Immunofluorescent images were visualized with an inverted Olympus 1X70 fluorescence microscope equipped with a Cook Sensicom ER camera (Olympus America, Inc., Melville, NY, USA). In some cases, a series of three-dimensional images of each individual picture were deconvoluted into one two-dimensional picture and resolved by adjusting the signal cutoff to near maximal intensity, to increase resolution. Final pictures were prepared with Adobe Photoshop to demonstrate subcellular localization and possible colocalization between detected proteins.
Total RNA was extracted by RNeasy kit according to the manufacturer's protocol (Quagen, Valencia, CA, USA). Aliquots (20 μg) of total RNA were gel separated in denaturing conditions and transferred to nitrocellulose membrane according to standard procedures (Reiss et al., 1998b). A 1020 bp Rad51 cDNA fragment was released from Pet11d/Rad51 plasmid by NcoI/BamH1 digestion. The fragment was labeled by the Random Primed DNA Labeling Kit (Amersham, Arlington Heights) and (3000 Ci/mmol α-32P dCTP) and used for hybridization.
Rad51 expression and Rad51 promoter activity
Human Rad51 cDNA (gift of Dr Warren Pear, University of Pennsylvanian) was cloned into pMX-flag-Rad51 expression vector. Following transfection, DU145 cells were selected in 1 μg/ml of puromycin and stable clones expressing different quantities of Rad51 were isolated. pGL3-luc-Rad51 reporter plasmid (a gift of Dr R Fishel, Tomas Jefferson University, Philadelphia, PA, USA) and pCMV-β-gal were used to determine Rad51 promoter activity. DU145 cells were initially plated at 0.5 × 106 cells per 100 mm dish and monolayer cultures were cotransfected with 1 μg of pGL3-luc-Rad51 plasmid and 0.5 μg of pCMV-β-gal using FuGENE 6 reagent. On day 1 post-transfection, cells were replated either into uncoated or PolyHEMA-coated 60 mm dishes. Following 24 h of exposure to monolayer or anchorage-independent culture conditions, cells were lysed according to the manufacturer's instructions (Luciferase assay, Promega, Madison, WI, USA) and Rad51 promoter -mediated luciferase activity was evaluated by Femtomaster FB12 luminometer (Zylux Corporation). All samples were additionally analysed for β-galactosidase to normalize obtained luciferase activities with the corresponding efficiencies of transfection (Promega, Madison, WI, USA).
Baumann P and West SC . (1998). Trends Biochem. Sci., 23, 247–251.
Bindra RS, Schaffer PJ, Meng A, Woo J, Maseide K, Roth ME, Lizardi P, Hedley DW, Bristow RG and Glazer PM . (2004). Mol. Cell. Biol., 24, 8504–8518.
Bondar VM and McConkey DJ . (2002). Prostate, 51, 42–49.
Chen G, Yuan SS, Liu W, Xu Y, Trujillo K, Song B, Cong F, Goff SP, Wu Y, Arlinghaus R, Baltimore D, Gasser PJ, Park MS, Sung P and Lee EY . (1999). J. Biol. Chem., 274, 12748–12752.
Chen Y, Wang J, Fraig MM, Henderson K, Bissada NK, Watson DK and Schweinfest CW . (2003). Int. J. Oncol., 22, 1033–1043.
Chen Y, Wang J, Fraig MM, Metcalf J, Turner WR, Bissada NK, Watson DK and Schweinfest CW . (2001). Cancer Res., 61, 4112–4121.
Chendil D, Ranga RS, Meigooni D, Sathishkumar S and Ahmed MM . (2004). Oncogene, 23, 1599–1607.
Collis SJ, Tighe A, Scott SD, Roberts SA, Hendry JH and Margison GP . (2001). Nucleic Acids Res., 29, 1534–1538.
Davies AA, Masson JY, McIlwraith MJ, Stasiak AZ, Stasiak A, Venkitaraman AR and West SC . (2001). Mol. Cell, 7, 273–282.
Del Valle L, Enam S, Lassak A, Wang JY, Croul S, Khalili K and Reiss K . (2002). Clin. Cancer Res., 8, 1822–1830.
Essers J, Houtsmuller AB, van Veelen L, Paulusma C, Nigg AL, Pastink A, Vermeulen W, Hoeijmakers JH and Kanaar R . (2002). EMBO J., 21, 2030–2037.
Flygare J, Falt S, Ottervald J, Castro J, Dackland AL, Hellgren D and Wennborg A . (2001). Exp. Cell Res., 268, 61–69.
Hoeijmakers JH . (2001). Nature, 411, 366–374.
Huang Y, Nakada S, Ishiko T, Utsugisawa T, Datta R, Kharbanda S, Yoshida K, Talanian RV, Weichselbaum R, Kufe D and Yuan ZM . (1999). Mol. Cell. Biol., 19, 2986–2997.
Karan D, Lin MF, Johansson SL and Batra SK . (2003). Int. J. Cancer, 103, 285–293.
Khanna KK and Jackson SP . (2001). Nat. Genet., 27, 247–254.
Kovalenko OV, Plug AW, Haaf T, Gonda DK, Ashley T, Ward DC, Radding CM and Golub EI . (1996). Proc. Natl. Acad. Sci. USA, 93, 2958–2963.
Labhart P . (1999). Eur. J. Biochem., 265, 849–861.
Lim DS and Hasty P . (1996). Mol. Cell. Biol., 16, 7133–7143.
Ludwig T, Eggenschwiler J, Fisher P, D’Ercole AJ, Davenport ML and Efstratiadis A . (1996). Dev. Biol., 177, 517–535.
Lundin C, Erixon K, Arnaudeau C, Schultz N, Jenssen D, Meuth M and Helleday T . (2002). Mol. Cell. Biol., 22, 5869–5878.
Mills KD, Ferguson DO and Alt FW . (2003). Immunol. Rev., 194, 77–95.
Peruzzi F, Prisco M, Dews M, Salomoni P, Grassilli E, Romano G, Calabretta B and Baserga R . (1999). Mol. Cell. Biol., 19, 7203–7215.
Petrini JH . (2000). Curr. Opin. Cell Biol., 12, 293–296.
Pierce AJ and Jasin M . (2001). Mol. Cell, 8, 1160–1161.
Pierce AJ, Johnson RD, Thompson LH and Jasin M . (1999). Genes Dev., 13, 2633–2638.
Raderschall E, Stout K, Freier S, Suckow V, Schweiger S and Haaf T . (2002). Cancer Res., 62, 219–225.
Reiss K, D’Ambrosio C, Tu X, Tu C and Baserga R . (1998a). Clin. Cancer Res., 4, 2647–2655.
Reiss K, Valentinis B, Tu X, Xu SQ and Baserga R . (1998b). Exp. Cell Res., 242, 361–372.
Reiss K, Wang JY, Romano G, Furnari FB, Cavenee WK, Morrione A, Tu X and Baserga R . (2000). Oncogene, 19, 2687–2694.
Reiss K, Yumet G, Shan S, Huang Z, Alnemri E, Srinivasula SM, Wang JY, Morrione A and Baserga R . (1999). J. Cell Physiol., 181, 124–135.
Richardson C and Jasin M . (2000). Mol. Cell. Biol., 20, 9068–9075.
Richardson C, Stark JM, Ommundsen M and Jasin M . (2004). Oncogene, 23, 546–553.
Rodgers KJ and Dean RT . (2003). Int. J. Biochem. Cell Biol., 35, 716–727.
Schindewolf C, Lobenwein K, Trinczek K, Gomolka M, Soewarto D, Fella C, Pargent W, Singh N, Jung T and Hrabe de Angelis M . (2000). Mamm. Genome., 11, 552–554.
Schmutte C, Tombline G, Rhiem K, Sadoff MM, Schmutzler R, von Deimling A and Fishel R . (1999). Cancer Res., 59, 4564–4569.
Sieber OM, Heinimann K and Tomlinson IP . (2003). Nat. Rev. Cancer, 3, 701–708.
Sigurdsson S, Van Komen S, Petukhova G and Sung P . (2002). J. Biol. Chem., 277, 42790–42794.
Skorski T, Nieborowska-Skorska M, Campbell K, Iozzo RV, Zon G, Darzynkiewicz Z and Calabretta B . (1995). J. Exp. Med., 182, 1645–1653.
Slupianek A, Schmutte C, Tombline G, Nieborowska-Skorska M, Hoser G, Nowicki MO, Pierce AJ, Fishel R and Skorski T . (2001). Mol. Cell, 8, 795–806.
Sonoda E, Sasaki MS, Buerstedde JM, Bezzubova O, Shinohara A, Ogawa H, Takata M, Yamaguchi-Iwai Y and Takeda S . (1998). EMBO J., 17, 598–608.
Tombline G and Fishel R . (2002). J. Biol. Chem., 277, 14417–14425.
Tombline G, Shim KS and Fishel R . (2002). J. Biol. Chem., 277, 14426–14433.
Trojanek J, Ho T, Del Valle L, Nowicki M, Wang JY, Lassak A, Peruzzi F, Khalili K, Skorski T and Reiss K . (2003). Mol. Cell. Biol., 23, 7510–7524.
Valentinis B, Reiss K and Baserga R . (1998). J. Cell Physiol., 176, 648–657.
Valerie K and Povirk LF . (2003). Oncogene, 22, 5792–5812.
Van Komen S, Petukhova G, Sigurdsson S and Sung P . (2002). J. Biol. Chem., 277, 43578–43587.
Wang JY, Del Valle L, Gordon J, Rubini M, Romano G, Croul S, Peruzzi F, Khalili K and Reiss K . (2001). Oncogene, 20, 3857–3868.
Wozniak K and Blasiak J . (2002). Acta Biochim. Pol., 49, 583–596.
Xia SJ, Shammas MA and Shmookler Reis RJ . (1997). Mol. Cell. Biol., 17, 7151–7158.
Yeh CC, Lee C and Dahiya R . (2001). Biochem. Biophys. Res. Commun., 285, 409–413.
Zhu XD, Kuster B, Mann M, Petrini JH and Lange T . (2000). Nat. Genet., 25, 347–352.
We gratefully acknowledge editorial assistance of Dr Sidney Croul and Jessica Otte, technical manager of the Center for Neurovirology and Cancer Biology, for her operational efforts in our laboratory. TS is a scholar of the Leukemia and Lymphoma Society.
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Wang, J., Ho, T., Trojanek, J. et al. Impaired homologous recombination DNA repair and enhanced sensitivity to DNA damage in prostate cancer cells exposed to anchorage-independence. Oncogene 24, 3748–3758 (2005) doi:10.1038/sj.onc.1208537
- prostate cancer
- homologous recombination
- DNA repair
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