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Department of Biochemistry and New York University Cancer Institute, New York University School of Medicine, New York, New York1
Received 19 May 2004/ Returned for modification 14 June 2004/ Accepted 13 December 2004
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ABSTRACT |
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INTRODUCTION |
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RPA is composed of three distinct subunits of 
70 (RPA1), 30 (RPA2), and 14 (RPA3) kDa and is an essential factor in many DNA processing reactions. Genetic and biochemical studies demonstrate that RPA has required roles both in the initiation and in the elongation stages of DNA replication (31, 57). Similarly, RPA is necessary for homologous recombination and for DNA repair events that use the recombination machinery (for example, see reference 53 and references therein). It is also indispensable for nucleotide excision repair (1). Along with stabilizing DNA in its single-stranded form, RPA supports the activity of other factors through obligate interactions. For example, simian virus 40 (SV40) DNA replication can be reconstituted with RPA of a metazoan origin but not with Saccharomyces cerevisiae RPA (6, 39). RPA is intimately involved in the cellular checkpoint response as RPA recruits the ATR-ATRIP complex to sites of DNA damage and supports activation of the ATR kinase (59). RPA also recruits the replication factor C-like Rad17 complex to various DNA structures and assists the binding of the Rad9-Rad1-Hus1 complex (60).
As would be expected of a protein with multiple roles in DNA metabolism and in the response to DNA damage, RPA activity is regulated at various levels. The RPA2 subunit of RPA becomes phosphorylated in response to genotoxic stress by phosphatidylinositol 3-kinase-related kinases, including ATM and DNA-PK (see citations within references 5 and 52). Mutational analysis of the RPA2 phosphorylation sites indicates that RPA phosphorylation prevents recruitment of RPA to replication centers while having no effect on localization to sites of DNA damage (52). Downregulation of RPA activity also occurs by apparent phosphorylation-independent mechanisms. The most clearly identified pathway involves the inhibition of RPA activity by association with the nucleolar factor nucleolin (13, 54).
Nucleolin is an abundant protein that is required for the first step of pre-rRNA processing (22). Mutation of the genes encoding nucleolin homologues in budding and fission yeast disrupts balanced production of the small and large ribosomal subunits (24, 34, 35). Nucleolin has many other diverse activities, including regulation of transcription (20, 23, 26, 45, 58), modulation of mRNA stability (9, 48), and acting as a low-affinity receptor for human immunodeficiency virus on the cell surface (7, 41). In response to DNA damage conditions or heat shock, a significant fraction of the nucleolin pool relocalizes from the nucleolus to the nucleoplasm in a process stimulated by physical association with p53 (13, 14, 54). After heat shock, nucleolin-RPA complex formation is greatly stimulated, and formation of this complex is inhibitory to DNA replication in vitro (13, 54). In vivo, the mobilized nucleolin sequesters RPA at sites distinct from replication centers (13). The mobilization of nucleolin in response to heat shock thus represents a novel pathway for regulating DNA replication.
We examined the interaction of nucleolin and RPA in response to DNA damage. We found that, like heat shock, genotoxic stress strongly induces nucleolin-RPA complex formation. The RPA-interacting domain was localized to the 63-amino-acid (aa) glycine-arginine-rich (GAR) domain at the extreme C terminus of nucleolin. Expression of GAR or a nucleolin mutant with constitutive association with RPA causes a block in the cellular transit from G1 into S phase. The nucleolin-mediated inhibition of chromosomal DNA replication could be prevented by overexpression of RPA2 to increase the cellular level of RPA. These data demonstrate a novel intra-S-phase checkpoint response in response to genotoxic stress through target of RPA by mobilized nucleolin.
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MATERIALS AND METHODS |
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For in vivo studies, nucleolin or nucleolin derivatives were expressed with N-terminal green fluorescent protein (GFP), cyan fluorescent protein (CFP), or Myc epitope tags. GFP and CFP fusion proteins were constructed by using PCR cloning into the pEGFP-C1 or pECFP-C1 vectors (Clontech). Similarly, Myc-tagged nucleolin (FL or mutants) was expressed from the pEF6/Myc-HisA plasmid (Invitrogen), as modified by Vassin et al. (52) to prevent expression of the His tag or, for proliferation studies, from a modified pEGFP-C1 vector in which the GFP tag was replaced by the Myc tag. Human RPA2 containing an N-terminal yellow fluorescent protein (YFP) tag was generated by excising the RPA2 coding sequence from pENeGFP RPA34 (kindly provided by M. C. Cardoso) (50) into pEYFP-C1 vector (Clontech). The construction of the Myc-RPA2 expression vector was described previously (52). The pECFP-C1-H-Ras61L and pEYFP-N1-RasBD expression vectors were kindly provided by Trever Bivona of Mark Philips laboratory (New York University [NYU] School of Medicine). All fusion constructs were sequenced and shown to be faithful copies of the corresponding genes.
Purification of proteins. GST-tagged nucleolin proteins were purified by the protocol of Haluska et al. (25). After transformation of S. cerevisiae JEL1 strain with the appropriate plasmid, cells were grown in synthetic defined (SD) medium under selection in 2% raffinose, and protein expression was induced by 2% galactose. Extracts from these cultures were made by disruption of the yeast cells by using 25- to 50-µm glass beads in uracil RIPA buffer (50 mM Tris-HCl [pH 7.2], 150 mM NaCl, 0.1% sodium dodecyl sulfate [SDS], 1% Triton X-100, 1% sodium deoxycholate) with protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 0.5 µg of leupeptin/ml, 1 µg of pepstatin/ml), 1 mM EDTA, and 1 mM dithiothreitol (DTT). Glutathione-Sepharose beads (Pharmacia) were then added to the clarified yeast extract, followed by incubation to bind the GST-nucleolin proteins. After three washes with a 10x bead volume of RIPA buffer, the GST-tagged proteins were eluted with 10 mM reduced glutathione and 50 mM Tris-HCl (pH 7.5). After overnight dialysis at 4°C against phosphate-buffered saline (PBS) and 20% glycerol, eluates were assayed for purity by SDS-polyacrylamide gel electrophoresis (PAGE) and Coomassie blue staining.
The human RPA heterotrimer was produced in Escherichia coli BL21 transformed with the p11dtRPA vector and purified as described previously (29, 30).
Far-Western analysis. Far-Western blotting was carried out basically as described by Jayaraman et al. (32). Purified GST-tagged nucleolin (FL and mutants) proteins were subjected to SDS-PAGE and then transferred to a nitrocellulose membrane. After two incubations in denaturation buffer (6 M guanidine-HCl in PBS) for 5 min at 4°C, the membrane was incubated six times in serial dilutions (1:1 [vol/vol]) of denaturation buffer, each dilution being with PBS containing 1 mM DTT. The membrane was blocked with PBS containing 0.1% Tween 20 (PBS-T) and 5% nonfat dry milk (NFDM) for 45 min at room temperature and washed twice with PBS-T and 0.25% NFDM. The membrane was then incubated with purified human RPA (0.2 µg/ml) in PBS-T, 0.25% NFDM, 1 mM DTT, and 2.5 mM phenylmethylsulfonyl fluoride for 2 h at room temperature and subsequently washed four times in PBS-T and 0.25% NFDM. The presence of bound RPA was probed by using a mouse anti-RPA2 monoclonal antibody (SSB34A; NeoMarkers) and horseradish peroxidase-conjugated sheep anti-mouse antibody as the primary and secondary antibodies, respectively, and detected by using enhanced chemiluminescence (Amersham Biosciences).
In vitro DNA replication assay.
The SV40-based in vitro DNA replication assay was described previously (52) and utilized a pBluescript SK+ phagemid (Stratagene) containing a 90-bp SV40 origin region segment (positions 5186 to 32) subcloned into the BamHI and XhoI sites (pBS-ori). Reaction mixtures (25 µl) contained the following: 40 mM HEPES (pH 7.5); 40 mM creatine phosphate; 7 mM MgCl2; 0.5 mM DTT; 4 mM ATP; 200 µM concentrations each of CTP, GTP, and UTP; 100 µM concentrations each of dATP, dGTP, and dTTP; 40 µM [
-32P]dCTP (3,000 cpm/pmol; Perkin-Elmer Life Sciences); 1.25 µg of creatine phosphokinase; 150 ng of pBS-ori; 100 µg of AS65 protein fraction prepared from HeLa cells; 200 ng of RPA; 200 to 400 ng of the GST fusion proteins; and 500 ng of SV40 large T antigen. The reaction mixtures were first preincubated on ice for 30 min without the addition of plasmid DNA, deoxynucleoside triphosphates, ATP, and creatine phosphokinase. After the addition of the remaining factors, the complete reaction mixture was further incubated at 37°C for 2 h. The replication activity was determined by precipitating the high-molecular-weight DNA with trichloroacetic acid and quantitating the amount of incorporated radioactivity in the precipitate by liquid scintillation counting.
Immunoprecipitation and immunoblotting. Plated U2-OS cells were transfected with 1 µg of specified expression plasmids by using Effectene transfection reagent (Qiagen). The transfection efficiencies of each construct were similar when visualized at 24 h posttransfection. When required, cells were either treated with 1 µM CPT or 2.5 mM hydroxyurea or exposed to 10 Gy of ionizing radiation or 30 J of UV light m–2. The immunoprecipitation reaction was carried out by using the IMMUNOcatcher kit (CytoSignal) according to the manufacturer's instructions. Immunoprecipitated proteins were separated by using SDS-10% PAGE and transferred to a nitrocellulose membrane (Schleicher & Schuell). After incubation with the appropriate primary antibody, the membrane incubated with an horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit secondary antibody, and the presence of bound proteins was detected with ECLplus (Amersham Pharmacia Biotech). The following antibodies were used for both detection and immunoprecipitation: RPA2, mouse monoclonal antibody SSB34A (NeoMarkers); RPA1, mouse monoclonal antibody Ab-1; nucleolin, either the MS3 mouse monoclonal or the H-250 rabbit polyclonal antibody (Santa Cruz Biotechnology); GFP, rabbit polyclonal antibody (Molecular Probes); Myc, rabbit polyclonal antibody (Upstate Biotechnology); p53, DO-1 mouse monoclonal antibody (Santa Cruz Biotechnology); (pSer15)p53, rabbit polyclonal antibody (Cell Signaling Technology); and p21, mouse monoclonal Cip1/WAF1 antibody (BD Biosciences/Pharmingen).
Immunofluorescence microscopy. To prepare for imaging, U2-OS cells grown on fibronectin-coated coverslips (BD Biotechnology) were treated as described previously (15). Cells were fixed for 20 min at room temperature with 4% (wt/vol) formaldehyde in PBS, permeabilized with 0.5% Triton X-100, rinsed with PBS, and then incubated with PBS containing 0.5% Nonidet P-40. Coverslips were incubated with 1:100 dilution of the appropriate primary antibody for 1 h at room temperature. After three rinses with PBS containing 0.5% Tween 20, coverslips were incubated for 1 h at room temperature with 1:100 dilution of Texas Red- or fluorescein isothiocyanate-conjugated secondary antibody (Jackson Immunoresearch Laboratories). Coverslips were then rinsed three times with PBS containing 0.5% Tween 20 and mounted onto glass slides. Fluorescent signals were detected by using either epifluorescence or confocal microscopy.
FRET. U2-OS cells were grown and cotransfected with the appropriate YFP- and CFP-tagged expression constructs in 35-mm uncoated glass bottom cell culture dishes (MatTek). Live cell images were obtained with a Zeiss LSM510 Meta laser scanning confocal microscope with a Plan-Apochromat x63 objective lens and a 30-mW Argon laser set at 50% of total output. CFP as the donor channel was excited with a 458-nm laserline, and CFP fluorescence was collected with a band-pass filter of 475 to 525 nm. YFP, the acceptor channel, was excited at 514 nm, and YFP emission was collected with a long-pass filter of 530 nm. The fluorescence resonance energy transfer (FRET) channel consisted of CFP excited at 458 nm and YFP fluorescence collected with a long-pass filter of 530 nm. Photobleaching was performed with the 514-nm laser line set at 100% power with an average bleach time of 5 s. Specific regions of interest (ROIs) were chosen, and positive FRET was determined graphically based on the decrease of YFP signal, and the subsequent increase in the CFP fluorescence postbleaching. Although transfection of any combination of YFP-RPA2 and CFP-nucleolin (or nucleolin derivative) did not have notable deleterious effects on cell viability, only cells with a normal appearance and relatively low expression levels were tested.
BrdU incorporation assay and FACS. U2-OS cells were plated at 30% confluency in 60-mm dishes. Plates were mock transfected, transfected with 1 µg of the Myc tag (empty) vector, or 1 µg of the appropriate N-terminal Myc-tagged nucleolin expression construct. At 24 h posttransfection, the cells were incubated for 20 min with 10 µM bromodeoxyuridine (BrdU). Cells were then washed twice with ice-cold PBS and collected by centrifugation at 180 x g for 5 min at 4°C. Pelleted cells were carefully resuspended into 300 µl of 4% (wt/vol) formaldehyde in PBS, fixed for 15 min at room temperature, and washed with PBS twice. Cells were then permeabilized for 15 min on ice with PBS containing 0.2% (vol/vol) Triton X-100 and 1% (wt/vol) bovine serum albumin (BSA), washed once with PBS, and then treated with PBS containing 0.25 mg of DNase/ml for 1 h at 37°C. Cells were incubated with 100 µl of PBS containing rat anti-BrdU (Harlan Sera-Lab) and rabbit anti-Myc (Upstate Biotechnology) polyclonal antibodies and 2% (wt/vol) BSA for 40 min at 37°C. Cells were washed twice with PBS and incubated for 40 min at room temperature with 100 µl of PBS containing anti-rabbit phycoerythrin-conjugated and anti-rat fluorescein isothiocyanate-conjugated antibodies (Jackson Laboratories) and 2% BSA. After preincubation of cells with 4 mM sodium citrate, 30 U of RNase A/ml, and 0.1% (vol/vol) Triton X-100 for 10 min at 37°C, the DNA was stained with 7-aminoactinomycin D (Sigma), and the cells were subjected to fluorescence-activated cell sorting (FACS) analysis.
[3H]thymidine uptake assay. U2-OS cells were plated into 24-well tissue culture plates in complete McCoy's media containing 10% fetal bovine serum (FBS). The cells were transfected with plasmids (100 ng) expressing one of the following proteins: Myc-tag, Myc-nucleolin TM, or Myc-nucleolin GAR. As indicated, cells were also cotransfected with various amounts of a Myc-RPA2 expression vector. After 6 to 8 h, the medium was changed to a low serum (0.1% FBS) condition and further incubated for 18 h. After recovery in complete medium for 8 to 10 h, the cells were incubated with [3H]thymidine (1 µCi/well) for 10 h. Cells were then washed with ice-cold PBS extensively and treated with 5% trichloroacetic acid for 30 min on ice. After further washes with ice-cold PBS, cells were solubilized in 0.5 N NaOH-0.5% (wt/vol) SDS and harvested, and the amount of incorporated radiolabel was determined with a scintillation counter.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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5 to 10% of the RPA pool is coimmunoprecipitated with nucleolin at the peak level of complex formation, although this value would be an underestimate if the complex were transient or unstable under immunoprecipitation conditions. A similar induction of nucleolin-RPA complex formation was observed after treatment with hydroxyurea (to cause replicative stress; Fig. 1B) or exposure to ionizing radiation (10 Gy; Fig. 1C). Nucleolin was not seen to form a complex with RPA after exposure to UV radiation (Fig. 1D) similar to previous observations finding a lack of induced nucleolin-p53 complex and nucleolin relocalization after UV irradiation (14). Induction of nucleolin-RPA complex formation was observed in p53-null H1299 cells after CPT treatment (Fig. 1E). Therefore, although nucleolin relocalization from the nucleolus to the nucleoplasm is p53 dependent (14), this dependence does not extend to nucleolin-RPA complex formation. Note that previous studies from our laboratory indicated that complex formation is not mediated by the presence of DNA and can also be detected by precipitation of RPA rather than nucleolin (13). In general, enhanced nucleolin-RPA complex formation is not restricted to heat shock but is also detected after genotoxic stress.
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Effect of the GAR domain on SV40 DNA replication in vitro. We previously showed that SV40 DNA replication in vitro was inhibited by the addition of nucleolin, purified from human cells, which interfered with RPA action (13). Because our data indicate that the nucleolin GAR domain interacts with RPA, we similarly tested the effect of this peptide on SV40 DNA replication. GST-tagged nucleolin or nucleolin derivatives were purified, and titrated into a T-antigen-dependent SV40 DNA replication reaction (Fig. 2C). In reactions containing nucleolin FL or GAR, DNA synthesis was significantly inhibited as a function of the amount of nucleolin protein added. In contrast, no obvious inhibition was seen by addition of nucleolin NT/RBD1-4 or GST. Thus, nucleolin molecules that are capable of binding RPA also inhibit DNA replication in vitro.
Stress-dependent formation of the nucleolin FL-RPA complex. We examined the interaction of nucleolin and the nucleolin mutants with RPA in vivo. Because the cellular localization of nucleolin may be a determinant affecting its interaction with RPA (13), we first examined the localization of the different nucleolin derivatives. GFP-tagged nucleolin FL, NT/RBD1-4, and GAR were expressed in U2-OS cells, and the localization of the fusion proteins captured by indirect immunofluorescence microscopy. Nucleolin FL localized exclusively to nucleolar regions (Fig. 3B), as determined by colocalization with endogenous nucleolin and upstream binding factor (necessary for RNA polymerase I-mediated transcription of rRNA [27]) (data not shown). The NT/RBD1-4 and GAR proteins had primary localization in the nucleolus (Fig. 3C and D, respectively), although the level of nucleolar staining was higher for the NT/RBD1-4 mutant. A significant fraction of each mutant protein pool was located in the nucleoplasm, and both mutants showed a weak but clear cytoplasmic signal. As expected, GFP alone was localized throughout the cell (Fig. 3A). These data are consistent with previous findings that the nucleolin RBD and GAR domains each contribute to nucleolar localization (12, 28, 40, 47).
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We next examined the effect of CPT treatment on GFP-tagged nucleolin FL localization and on the interaction of RPA with nucleolin FL and the nucleolin derivatives. Although nucleolin FL localized to the nucleolus in the absence of stress (Fig. 4A [see also Fig. 3B above]), incubation with CPT caused a significant fraction of the nucleolin FL pool to move to the nucleoplasm (Fig. 4B), similar to the behavior of endogenous nucleolin (13). In testing the interactions, RPA was observed to associate with nucleolin FL but only after CPT treatment (Fig. 4C, upper panel). In contrast, the NT construct lacking the GAR domain did not form a detectable complex with RPA irrespective of stress (Fig. 4C, middle panel). The CT construct that contains the GAR domain coprecipitated with RPA both in the presence of CPT and in its absence, thus revealing a constitutive interaction (Fig. 4C, lower panel). The localizations of these truncated proteins were not affected by prior CPT treatment (data not shown). These data indicate that although the presence of the GAR is necessary to support detectable complex formation with RPA in vivo, detectable interaction of RPA with the full-length nucleolin also requires stress conditions such as caused by CPT treatment.
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Cell cycle arrest upon overexpression with either nucleolin GAR or TM. We found previously that heat shock mobilizes nucleolin to move to the nucleoplasm, whereupon it binds RPA at sites distinct from the DNA replication centers (13). These data predict that expression of nucleolin derivatives that bind RPA in nonstressed cells will cause a G1/S arrest. The effect of nucleolin TM and GAR expression on cell cycle progression were therefore investigated by FACS. Both the nontransfected control and the vector control showed a similar distribution, indicating that transfection alone did not inhibit cell cycle transit (Fig. 7A). Expression of nucleolin FL led to only a slight increase in G1-phase cells. However, much more significant effects were observed in cells transfected with nucleolin TM or GAR. The expression of nucleolin GAR resulted in an increase in the G1 population to 52% of cells compared to 36% of vector-transfected cells. We also detected a decrease in S-phase cells from 39% in vector-transfected cells to 23% in GAR-transfected cells. Expression of nucleolin TM had a similar influence on cell cycle progression with G1- and S-phase cells contributing 53 and 25%, respectively, of the total cell pool. In each case, the fraction of G2 cells remained constant. Thus, the expression of nucleolin GAR or TM was sufficient to elicit an arrest in the cell cycle, leading to the accumulation of cells in G1 and a decrease in cells in S phase. We note that the overall degree of replication inhibition in the GAR- or TM-transfected cells (a >40% decrease) is similar to that observed after ionizing irradiation (42).
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To test this method, U2-OS cells were transfected with either nucleolin GAR or TM, and the level of DNA replication was measured by determining thymidine incorporation (Fig. 7C). Corroborating the results of the FACS analysis, expression of either nucleolin construct inhibited DNA synthesis by ca. 50%. In parallel reactions, these cells were cotransfected with increasing levels of RPA2. We observed that the degree of replication inhibition caused either by nucleolin GAR or TM expression was progressively reduced by transfection of the Myc-RPA2 expression vector. The stimulatory effect of RPA2 overexpression was somewhat more pronounced in nucleolin TM-transfected cells compared to GAR-transfected cells, for unknown reasons. Transfecting higher levels of RPA2 vector (i.e., 100 ng) caused some toxic effects on cell viability (data not shown). These data strongly indicate that nucleolin can inhibit DNA synthesis by direct interaction with RPA.
Nucleolin GAR expression does not activate p53. It is possible that the expression of the GAR domain causes a cellular stress response and therefore only inhibits cell cycle progression indirectly. As a test of this possibility, we examined the effect of GAR expression on p53 activation in U2-OS cells (which express wild-type p53). Expression of nucleolin GAR did not increase p53 levels (Fig. 7D, upper panel) or the level of p53 phosphorylated on Ser15, a site modified by the ATM and ATR kinases in response to genotoxic stress (49, 51). The lack of Ser15 phosphorylation demonstrates that p53 and, indirectly, ATM and ATR do not become activated in response to GAR expression (Fig. 7D, second panel). Aliquots of these lysates were probed for the presence of p21waf1, a key stress-induced inhibitor of cyclin-dependent kinases whose expression results in a G1/S arrest. No changes in the level of p21waf1 were detected in response to GAR expression (Fig. 7D, third panel). In contrast, treatment of cells with CPT was found to simultaneously stimulate the levels of p53, (pSer15)p53, and p21waf1. The block in cell cycle progression caused by expression of nucleolin GAR is therefore unrelated to p53 activation, induction of p21, or, likely, activation of ATM or ATR. Instead, our data indicate that nucleolin can itself inhibit DNA replication by binding to RPA and inhibiting RPA activity.
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DISCUSSION |
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What is the mechanism by which nucleolin inhibits cell cycle progression? We find that GST-tagged nucleolin FL and the GAR domain each can inhibit SV40 DNA replication in vitro, recapitulating similar inhibitory effects that were observed when endogenous nucleolin purified from human cells was tested (13). Although our studies did not find an inhibitory effect of nucleolin on RPA binding to single-stranded DNA, we do find that nucleolin can inhibit the binding of RPA to duplex molecules containing a central nonpaired region (data not shown). Such data suggest that the nucleolin-RPA complex is selectively inhibitory to the initiation stages of replication. However, we have recently presented data indicating that RPA does not randomly bind to single-stranded DNA at a chromosomal DNA replication fork but is instead actively loaded by a component of the replication machinery (52). Thus, complex formation with nucleolin has the potential to prevent RPA from productive loading onto single-stranded DNA at a replication fork in vivo. Both of these processes could inhibit DNA replication in vivo and cause a reduction in DNA synthesis. Overall, our data indicate that a direct interaction between the GAR domain of nucleolin and RPA is sufficient for replication inhibition in vivo.
Our data lead to the model that genotoxic stress activates nucleolin, such that the GAR domain becomes exposed for complex formation with RPA. In support of this model, consider the following data. First, although the GAR domain is required to bind RPA, nucleolin FL also requires stress conditions to bind RPA. Second, the nucleolin TM molecule that constitutively binds RPA was mutated at three N-terminal positions, whereas the RPA-interacting GAR domain is located on the extreme C terminus of nucleolin. Third, nucleolin relocalization to the nucleoplasm, although an outcome of genotoxic stress and heat shock, is not required for RPA complex formation because our FRET data show interaction in the nucleolus, as well as in the nucleoplasm. Fourth, a requirement for changes in RPA modification does not appear to be required as the GAR domain binds RPA in a stress-independent fashion. Along these same lines, preliminary evidence obtained from test of a hyperphosphorylation mimic of RPA (RPA2D) (52) showed no significant effects on nucleolin complex formation compared to RPA2wt (data not shown). We postulate that changes in nucleolin modification promote conformational changes which remove steric constraints preventing RPA complex formation. Although expression of nucleolin TM or GAR do not cause apparent ATM or ATR activation, it is quite possible that activation of these kinases by genotoxic stress facilitates nucleolin-RPA complex formation, a possibility under investigation.
Our FRET data indicate that nucleolin-RPA complex formation occurs both in the nucleoplasm and in the nucleolus and hence nucleolin relocalization is not required for these two proteins to interact. Even so, nucleolin relocalization probably facilitates interaction with RPA. The nucleolus comprises ca. 10 to 15% of the nuclear volume in human cells (e.g., see reference 18) and a nucleoplasmic localization would provide a larger volume in which complex formation can occur. Although p53 is not required for nucleolin-RPA complex formation, nucleolin relocalization is strongly dependent on p53 (14) (see also below), suggesting that p53 might stimulate the nucleolin-RPA interaction. Testing the ability of p53-positive (U2-OS) and negative (H1299) cells to induce nucleolin-RPA complex formation after stress did not reveal any obvious differences. That said, these cells have genetic differences other than p53 that preclude our drawing firm conclusions on the potential role of p53 in facilitating complex formation at this time.
The mechanism of nucleolin relocalization remains somewhat unclear. Previous study has found that movement of a portion of the nucleolin pool to the nucleoplasm is greatly facilitated by p53 (14). Because genotoxic stress transiently induces nucleolin and p53 complex formation (14), increased nucleoplasmic levels of appropriately modified p53 and nucleolin may lead to more complex formation and hence a net nucleoplasmic flow of nucleolin. The lack of requirement for p53 in supporting nucleolin-RPA complex formation would indicate that an event(s), such as changes to the nucleolin modification state, occurs prior to nucleolin-RPA and nucleolin-p53 complex formation. This event would lead to the apparently independent increase in the association of nucleolin with either p53 or RPA. Along with a p53 requirement in supporting nucleolin mobilization from the nucleolus in response to stress, it has also been recently proposed that p53 activation by stress itself involves nucleolar disruption (ND) (46). In this model, ND interrupts a requisite nucleolar export pathway for p53 destined for degradation. If this model is correct, ND initiates p53 activation, which itself leads to increased ND.
We identified the nucleolin GAR domain as being necessary for interaction with RPA in vitro and in vivo. The GAR domain is contained within 63 residues and includes more than 10 RGG or FGG repeats. Similar RGG/FGG repeat sequences are found in other RNA-binding proteins, including hnRNP A1, hnRNP U, and fibrillarin (3). The RGG region forms a ß-spiral structure and binds nonspecifically to single- and double-stranded RNA and DNA (21). The GAR domain of nucleolin interacts with various ribosomal subunits, including L3 (22), and, along with its ability to bind RNA, presumably explains the role of the nucleolin GAR domain in supporting efficient nucleolar localization (12, 28, 40, 47). The nucleolin GAR domain also contains a 12-residue unique lysine-rich element at the extreme C terminus. Our far-Western analysis indicates that nucleolin molecules with small C-terminal deletions do not support RPA binding, suggesting that RPA may bind this unique C-terminal end. Additional studies will be needed to determine the relative contributions of the RGG region and the unique element in supporting complex formation with RPA.
It is becoming clear that the nucleolus is a critical cellular body whose components regulate cell cycle progression. For example, p19ARF (p14ARF in humans) localizes to the nucleolus, where it can bind and sequester the p53 antagonist MDM2 and thereby cause p53 stabilization (56). Similarly, the binding of the human MDM2 RING domain to ATP stimulates nucleolar localization in the absence of p14ARF (44). The yeast Yph1p protein is a BRCT domain-containing nucleolar factor whose depletion causes both G1 and G2 arrest (17). With regard to mitotic progression, it has been found that exit from mitosis is controlled by the Cdc14 protein phosphatase that is sequestered in the nucleolus until anaphase (2). These and other observations, combined with our findings that nucleolus also serves a dual role in ribosome biogenesis and inhibiting S-phase progression in response to genotoxic stress, highlights the importance of the nucleolus in serving to integrate cell growth and cell stress pathways.
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ACKNOWLEDGMENTS |
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This study was supported by NIH grant AI29963, DOD Breast Cancer Research Program DAMD17-03-1-0299, Philip Morris grant 15-B0001-42171, the NYU Cancer Institute, and the Rita J. and Stanley Kaplan Comprehensive Cancer Center (NCI P30CA16087). Purchase of the confocal microscope was funded by the Shared Instrumentation Grant Program of the NIH (S10 RR017970).
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FOOTNOTES |
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