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Cyclophilin A (CypA) is a member of a family of cellular proteins that share a peptidyl prolyl cis-trans isomerase (PPIase) activity. CypA was previously reported to be required for the biochemical stability and function (specifically, induction of G2 arrest) of the human immunodeficiency virus type 1 (HIV-1) protein R (Vpr). In the present study, we examine the role of the Vpr-CypA interaction on Vpr-induced G2 arrest. We find that Vpr coimmunoprecipitates with CypA and that this interaction is disrupted by substitution of proline-35 of Vpr as well as incubation with the CypA inhibitor cyclosporine A (CsA). Surprisingly, the presence of CypA or its binding to Vpr is dispensable for the ability of Vpr to induce G2 arrest. Vpr expression in CypA−/− cells leads to induction of G2 arrest in a manner that is indistinguishable from that in CypA+ cells. CsA abolished CypA-Vpr binding but had no effect on induction of G2 arrest or Vpr steady-state levels. In view of these results, we propose that the interaction with CypA is independent of the ability of Vpr to induce cell cycle arrest. The interaction between Vpr and CypA is intriguing, and further studies should examine its potential effects on other functions of Vpr.
Cyclophilin A is a member of a family of cellular proteins that shares a peptidyl prolyl cis-trans isomerase (PPIase) activity. To date, 16 cyclophilin genes and numerous cyclophilin pseudogenes have been identified in the human genome (11, 26, 32). The PPIase activity of cyclophilins appears to be important for the maintenance of proper protein conformation through cis-trans interconversion of N-terminal peptide bonds aminoterminal to proline (10, 31). This enzymatic activity contributes to cyclophilin involvement in cell signaling, mitochondrial function, molecular chaperone activity, RNA splicing, stress responses, gene expression, and regulation of kinase activity (10, 31, 32).
Cyclophilins owe their name to their ability to bind to the immunosuppressive drug cyclosporine A (CsA) with high affinity (17). CsA binding to CypA potently inhibits CypA isomerase activity. The immunosuppressive effects of CsA result from binding of the CypA/CsA complex to calcineurin, resulting in calcineurin inhibition (20, 24).
A role for CypA in the life cycle of primate lentiviruses emerged in 1993 with the isolation of CypA as a yeast two-hybrid partner of human immunodeficiency virus type 1 (HIV-1) core protein, p24 (26). CypA binding to p24 is necessary for the infectivity of HIV-1, and blocking this binding, either through site-directed mutations in p24 or competitive inhibitors such as CsA, severely impairs the infectivity of HIV-1 particles (reviewed in reference 14). It is thought that the binding of CypA to p24 is a necessary event for the efficient uncoating of viral particles following viral entry (25). Recent evidence indicates that the effects of CsA on HIV-1 replication are not solely derived from CsA inhibition of CypA binding to p24 in target cells, as treating HIV-1-producing cells with CsA also reduces the infectivity of progeny viral particles, albeit in a CypA-independent fashion (9, 18, 30).
HIV-1 Vpr exerts several deleterious effects when expressed in human cells, including induction of cell cycle arrest in G2 and apoptosis. Vpr induces a DNA damage-like signal that triggers known downstream checkpoint responses involving certain cell cycle-related kinases and phosphatases, such as ATR, Chk1, Wee1, and Cdc25 (5, 15, 19, 21, 22, 28, 29, 33, 37). Recent evidence suggests that Vpr interacts with the chromatin in a unique manner, which results in activation of the G2 checkpoint without causing double-strand breaks (22). Vpr is packaged into virus particles, and its expression has been shown to induce a plethora of other effects in target cells, including transactivation of the viral promoter, modulation of the accuracy of the reverse transcription process, induction of apoptosis, and disruption of nuclear envelope integrity (reviewed in references 4, 23, and 35).
The Vpr gene product is a small 96-amino-acid protein. The N terminus of Vpr contains four conserved proline residues (positions 5, 10, 14, and 35). These proline residues were shown to undergo cis-trans isomerism to varying degrees in studies that used a synthetic peptide encompassing residues 1 to 40 of Vpr (8). The activity of CypA was shown to be required for the biochemical stability and function (specifically, induction of G2 arrest) of Vpr, and proline-35 was shown to be essential for both interaction with CypA and activity of Vpr (34).
In the present study, we examine the impact of CypA interaction on Vpr-induced G2 arrest and Vpr levels of expression. In agreement with the studies by Zander et al. (34), we find that Vpr can be coimmunoprecipitated with CypA and that this interaction is disrupted by mutation of the proline-35 residue. However, substitution of proline-35 by alanine or asparagine results in stable proteins that fail to bind to CypA and are still capable of inducing cell cycle arrest. Cells in which CypA had been genetically ablated are sensitive to Vpr-induced G2 arrest. In addition, incubation with CsA abolished the Vpr-CypA interaction but failed to inhibit Vpr-induced G2 arrest. We propose that the ability of Vpr to interact with CypA is independent of its ability to induce cell cycle arrest.
Human embryonic kidney (HEK) cell line 293FT (Invitrogen, Carlsbad, Calif.) cells were cultured in Dulbecco's modified Eagle's medium with 10% fetal bovine serum (FBS), 1% l-glutamine solution. One milligram of G418 per milliliter was added when maintaining the cells. CypA−/− Jurkat cells (7) and CypA+ cells were cultured in RPMI 1640 medium (Invitrogen, Carlsbad, CA) supplemented with 10% FBS, 2 mM l-glutamine (Invitrogen). Exponentially growing HeLa cells were cultured in Dulbecco's minimal essential medium (Invitrogen) supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin-l-glutamate (Invitrogen). Where indicated, cells were incubated with 2.5 μM cyclosporine A (CsA; Sigma Aldrich, St. Louis, MO) for 24 h.
Lentivirus vectors were produced by transient transfection of HEK293FT cells. For defective lentivirus vector production, plasmids pHR-GFP and pHR-VPR and the indicated mutants were cotransfected with pCMVΔR8.2ΔVpr (3) and pHCMV-VSVG (2) by calcium phosphate-mediated transfection (36). Virus supernatants were collected at 48, 72, and 96 h posttransfection. Harvested supernatants were cleared by centrifugation at 2,000 rpm (828 × g). Cleared supernatants were concentrated by ultracentrifugation at 25,000 rpm (115,889 × g) for 1.5 h at 4°C. Concentrated virus was allowed to resuspend overnight at 4°C, and the suspension was frozen at −80°C for storage. Vector titers were measured by infection of HeLa cells as described below, followed by flow cytometric analysis of cells that were positive for the reporter molecule, green fluorescent protein (GFP). Vector titers were calculated with the equation [(F × C0)/V] × D, where F is the frequency of GFP-positive cells found by flow cytometry, C0 is the total number of target cells at the time of infection, V is the volume of inoculum, and D is the virus dilution factor. The virus dilution factor used for titrations was 10. The total number of target cells at the time of infection was 1 × 106. Infections were performed at a multiplicity of infection (MOI) of 2 with 10 μg of Polybrene/ml for 2 h.
HIV-1 molecular clones pNL4-3 and pNL4-3-VprX were transfected into 293FT cells by calcium phosphate transfection as described for lentiviral vector production. Twenty-four hours after transfection, virus-producing 293FT cells were cocultured with 1 × 107 MT-2 human T-cell leukemia virus-transformed CD4+ T cells for 5 h. MT-2 cells were then cultured alone until approximately 75% of cell blasts exhibited syncitia. Virus-containing supernatants were then cleared of cell debris by centrifugation at 828 × g for 10 min. Viral stocks were then frozen at −80°C. Cells were infected by spin infection as follows: 1 × 106 cells were diluted in viral stocks with 10 μg/ml Polybrene and centrifuged at 1,700 × g for 2 h at 25°C, after which cells were resuspended in normal growth medium and incubated at 37°C in 5% CO2.
HEK293FT cells were transfected using calcium phosphate-mediated transfection. Twenty-four hours posttransfection, cells were detached using trypsin, washed, and lysed in lysis-and-immunoprecipitation (IP) buffer (20 mM Tris, pH 7.5, 100 mM NaCl, 0.5% NP-40, 0.5 mM EDTA) containing protease inhibitors (Complete tablets; Roche, Indianapolis, IN) for 10 min on ice. Lysates were centrifuged at 2,000 rpm for 10 min and supernatants were collected. One microliter of rabbit anti-CypA (undiluted serum; EMD Biosciences, San Diego, CA) antibody was added, and tubes were incubated with an end-over-end mixing at 4°C for 1 h. Protein A/G agarose (Santa Cruz Biotechnology, Santa Cruz, CA) was added to immunoprecipitates and incubated overnight with an end-over-end mixing at 4°C. Immunoprecipitates were washed five times with IP buffer, followed by boiling in XT sample buffer (Bio-Rad, Hercules, CA).
Cell lysates or IP eluates were boiled for 5 min prior to being loaded on Criterion XT bis-tris gels (Bio-Rad, Hercules, Calif.) for electrophoretic separation. Proteins were transferred to Immobilon polyvinylidene difluoride membranes (Millipore, Bedford, MA) by a semidry transfer method (Bio-Rad) and then blocked for 45 min at room temperature in blocking solution (5% skim milk and 0.1% Tween 20 in phosphate-buffered saline [PBS]). Rabbit primary antibodies against CypA (1:10,000; EMD Biosciences, San Diego, CA), actin (1:1,000; Santa Cruz), or mouse anti-hemagglutinin (HA) (1:1,000; Covance, Berkeley, CA) were applied at indicated dilutions and incubated at 4°C overnight. Blots were washed three times in TPBS (0.1% Tween 20 in PBS) for 5 min, each time at room temperature. Secondary horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit immunoglobulin G antibodies were applied for 45 min at room temperature. Blots were washed again three times in TPBS before protein detection with enhanced chemiluminescence reagent (Amersham, Buckinghamshire, England).
At given times postinfection, Jurkat cells were collected, washed with fluorescence-activated cell sorting (FACS) buffer (2% FBS, 0.5 mM EDTA, and 0.02% sodium azide in PBS), fixed with 2% paraformaldehyde in PBS, and permeabilized with 0.1% Triton X-100 in PBS for 15 min. Cells were washed again with FACS buffer, incubated in DNA staining buffer (10 μg of propidium iodide/ml and 11.25 kU of RNase A/ml in FACS buffer) for 15 min, and analyzed by FACScan flow cytometry for GFP expression or DNA content (Beckton Dickinson, Franklin Lakes, N.J.). For detection of HIV-1 p24 antigen-expressing cells, we used a previously described protocol (6, 12). Briefly, 1 × 106 cells were washed twice in flow cytometry buffer and permeabilized using the Cytofix/Cytoperm kit (Pharmingen BD, San Jose, CA). The permeabilized cells were resuspended in 100 μl of intracellular staining buffer and incubated at 4°C with 5 μl of human anti-HIV p24 monoclonal antibody (obtained from the AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health; monoclonal antibody to HIV-1 p24, clone 71-31, was from Susan Zolla-Pazner ). The cells were washed twice in intracellular staining buffer, resuspended at 1 × 107 cells/ml in intracellular staining buffer, and incubated at 4°C with 10 μl fluorescein isothiocyanate-conjugated F(ab′)2 goat anti-human immunoglobulin G (Caltag, Burlingame, CA) for 30 min. The antibody-labeled cells were washed twice with intracellular staining buffer. The cells were then resuspended in DNA staining buffer as described above. Cell cycle profiles were modeled by using ModFit software (Verity Software, Topsham, ME).
Our previous studies have shown that infection with a lentiviral vector (pHR-VPR-IRES-GFP, referred to as pHR-VPR in this study) (29) that encodes a Vpr-IRES-GFP tandem cistron faithfully recapitulates the effects of Vpr, including induction of cell cycle arrest (37), apoptosis (5), and transactivation of the viral long terminal repeat (29). To test whether the presence of CypA is necessary for induction of cell cycle arrest, we infected CypA−/− (7, 34) or unmodified Jurkat cells (herein referred to as CypA+) with pHR-VPR or a control vector, pHR-GFP (29). Vector-infected cells were analyzed 24 h postinfection for DNA content and, separately, for GFP expression using flow cytometry (Fig. (Fig.1A).1A). CypA−/− Jurkat cells exhibited cell cycle arrest (68% cells in G2/M; 65% GFP+ cells) comparable to that of CypA+ Jurkat cells (77% cells in G2/M; 70% GFP+ cells). The CypA expression status of the cells was verified by Western blotting (rabbit anti-CypA; EMD Biosciences, San Diego, CA) and confirmed the lack of CypA expression in CypA−/− cells (Fig. (Fig.1B1B).
Since lentiviral vectors allow expression of high levels of heterologous proteins, it is possible that ectopic expression of Vpr in the previous experiment may have overcome a potential restriction (such as biochemical instability of Vpr or an inactive conformation) derived from the lack of CypA. To validate the above results in a more relevant expression system, we performed similar experiments in which Vpr was introduced by infection with replication-competent HIV-1NL4-3 (1). As a negative control, we utilized the isogenic, Vpr-negative mutant HIV-1NL4-3vprX, which contains a frameshift mutation that completely inactivates Vpr (5, 19, 27). We performed infections of CypA+ and CypA−/− cells at a multiplicity of infection (MOI) of 0.5. Infected cultures were analyzed at 96 h by flow cytometry after combined staining for intracellular p24 and DNA content. The p24-positive and -negative cells from each infection were then electronically gated and analyzed for cell cycle distribution (Fig. (Fig.2).2). Infections of CypA−/− cells were routinely low (between 0.5% and 2.5% at 96 h postinfection) and viruses failed to spread in these cells, consistent with a known requirement for CypA in target cells during the uncoating step (9, 18, 30). In contrast, infections of CypA+ cells were at a high level (between 25% and 60% at 96 h postinfection) and showed viral spread (data not shown). Despite the low frequency of infected cells in CypA−/− cultures, it was possible to use electronic gating to analyze the cell cycle of infected and uninfected cells separately. As expected, CypA+ cell infection with HIV-1NL4-3, but not with HIV-1 NL4-3VprX, resulted in a dramatic increase of the G2/M peak after 96 h (Fig. (Fig.2A;2A; 63.2% and 29.2% cells in G2/M, respectively). The lack of CypA in CypA−/− cells did not affect the sensitivity of cells to G2 arrest by infection with HIV-1NL4-3 (67.3% cells in G2/M). After 7 days of infection, the percentages of HIV-1NL4-3-infected cells exhibiting G2/M DNA content were 80.1% and 60.8% for CypA+ and CypA−/− cells, respectively (Fig. (Fig.2B2B).
The above results indicate that Vpr expression is associated with induction of G2 arrest in a manner that is indistinguishable between CypA+ and CypA−/− cells, whether Vpr is expressed from a lentiviral vector or from infectious, full-length HIV-1. The apparent discrepancy between our results and those reported by Zander et al. (34) prompted us to reexamine the Vpr-CypA interaction. Vpr binding to CypA was previously reported (8, 34), and this interaction was postulated to be required for Vpr stability and for its ability to induce G2 arrest (8). Bruns et al. indicated that the proline residue at position 35 of Vpr was essential for the interaction with CypA (8). In order to directly examine the interaction between CypA and Vpr, we performed coimmunoprecipitation studies with wild-type Vpr as well as with three Vpr mutants. Vpr(P35N) (8) and Vpr(P35A) disrupt proline-35. Vpr(R80A) is unable to induce G2 arrest or apoptosis (5, 13). 293FT cells were transfected with pHR-VPR, Vpr mutants, or GFP. Twenty-four hours posttransfection, cells were lysed and anti-CypA antibody was used for immunoprecipitation. Immunoprecipitates were then analyzed by sodium dodecyl-sulfate polyacrylamide gel electrophoresis followed by Western blotting, using an anti-HA antibody that recognizes an amino-terminal hemagglutinin epitope present in all Vpr vector constructs.
In agreement with the previous finding, we found that Vpr efficiently coprecipitated with CypA (Fig. (Fig.3A,3A, lane 5). In contrast, both Vpr(P35N) and Vpr(P35A) were impaired in their abilities to coprecipitate with CypA, as evidenced by extremely faint Vpr bands (Fig. (Fig.3A,3A, lanes 3 and 4). Vpr(R80A) also failed to coprecipitate with CypA, as indicated by the absence of a detectable Vpr band (Fig. (Fig.3A,3A, lane 2). The blot shown on the top panel of Fig. Fig.3A3A was then stripped and reprobed with anti-CypA antibody in order to verify that equal amounts of CypA had been immunoprecipitated in experiments 1 through 6 (Fig. (Fig.3A,3A, second blot). Analysis of the steady-state levels of Vpr by Western blotting of input cell lysates (Fig. (Fig.3A,3A, third blot from the top, lanes 2′ to 5′) revealed that all Vpr mutants were expressed at comparable or higher levels than wild-type Vpr.
The above data indicate that the inability of Vpr mutants to coprecipitate with CypA did not stem from a decrease in the mutant protein steady-state levels, which would have suggested a loss of protein stability. In addition, the fact that Vpr(R80A) failed to coprecipitate with CypA indicates that residues outside the Vpr (residues 1 to 40) domain (8) are also important for binding to CypA and that the presence of proline-35 is not sufficient for such binding.
To further explore the specificity of Vpr-CypA interaction, we performed parallel coimmunoprecipitation experiments in which cells were cultured in the presence of 2.5 μM CsA. CsA binds to CypA and competitively inhibits the interaction between p24 Gag and CypA (9, 18, 30). CsA incubation did not impair the steady-state levels of Vpr in the cells, as evidenced by Western blotting of input lysate (Fig. (Fig.3A,3A, third blot from the top, compare lanes 5′ and 6′). CsA incubation, however, abolished the interaction between CypA and Vpr (Fig. (Fig.3A,3A, top blot, compare lanes 5 and 6).
Expression of wild-type or Vpr mutants in CypA−/− Jurkat cells further demonstrated that CypA is dispensable for the steady-state levels of Vpr expression (Fig. (Fig.3B,3B, lanes 2, 4, and 5). CypA−/− Jurkat cells presumably contain all cyclophilins other than CypA. Thus, it is formally possible that Vpr may interact with other cyclophilins, and this, in turn, may compensate for the absence of CypA. Since CsA inhibits all known cyclophilins, we asked whether incubation of CypA−/− cells with CsA would affect the levels of Vpr expression. As shown in Fig. Fig.3B3B (compare lanes 2 and 3), CsA did not appreciably affect the steady-state level of Vpr. The Vpr expression level in CypA−/− cells was similar to that obtained in CypA+ cells (Fig. (Fig.3B,3B, compare lanes 2 and 6) when tested in parallel. Thus, we conclude that the presence of CypA is not required for efficient expression of Vpr. We also conclude that even though CsA potently inhibits the Vpr-CypA interaction, this interaction is dispensable for efficient expression of Vpr.
The ultimate goal of the present study was to establish the requirement of CypA toward Vpr-induced G2 arrest. The availability of Vpr mutants defective for CypA binding, as well as the pharmacological inhibitor CsA, allowed us to ask whether binding to CypA could be dissociated from induction of G2 arrest. Expression of wild-type Vpr and Vpr(P35N) induced dramatic G2 arrest in CypA+ cells as well as in CypA−/− Jurkat cells (Fig. (Fig.4;4; 85.8% and 60.1%, respectively). Addition of CsA to the cultures did not significantly change the G2 arrest levels (70.3% in CypA+ and 56.7% in CypA−/− cells, respectively). Vpr(P35N) also induced G2 arrest (52.7%) in CsA, although with slightly decreased efficiency compared with wild-type Vpr. In CypA−/−, Vpr(P35N) induced levels of G2 arrest (67.1%) comparable to those of wild-type Vpr. Incubation of cultures with CsA did not significantly affect G2 arrest by Vpr(P35N) (45% in CypA+ and 65% in CypA−/− cells). Therefore, binding to CypA is not required for the induction of G2 arrest by Vpr.
The sharp discrepancies between our results and those reported by Zander and colleagues could be reconciled, in part, if CsA treatment induced the loss of stability of Vpr (and, therefore, loss of function as well) in a CypA-dependent manner. However, our findings demonstrate that although CsA inhibits the interaction between Vpr and CypA, the steady-state level of Vpr in the cells remains the same in CsA-treated and untreated cells, and incubation with CsA does not affect Vpr function.
The concentration of CsA used in our experiments was 2.5 μM. We find that this concentration of CsA effectively inhibits CypA-Vpr binding. This concentration is well below the 50-μg/ml (equivalent to 41.6 μM) CsA concentration used by Zander and colleagues (34) when they observed loss of Vpr stability in cells treated with this CypA inhibitor. We reason that, since genetic removal of CypA failed to inhibit Vpr function or stability, the effects observed with 50 μg/ml CsA may be due to other effects of CsA which are CypA independent. For example, high concentrations of CypA may be cytotoxic. A precedent for the existence of an additional, ill-understood effect of CsA stems from the observation that CsA inhibits the infectivity of progeny HIV-1 virions in producer cells by a mechanism that is independent of CypA inhibition (18, 30).
Since CsA is thought to inhibit all known cyclophilins, our observations combining the presence of CsA and the genetic elimination of CypA suggest that cyclophilins other than CypA appear to also be dispensable for Vpr expression and induction of G2 arrest.
We conclude that while in vivo binding between Vpr and CypA is clearly detectable, this binding is not necessary for the stability of Vpr or its ability to induce G2 arrest. The interaction between Vpr and CypA is highly intriguing and should be further investigated as a potential modulator of virus-host interactions.
We are thankful to Michael J. Blackwell and Sam Campbell for excellent technical help and Warner Greene for useful discussions. The following reagents were obtained through the NIH AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH: Jurkat T-cells CypA−/− from D. Braaten and J. Luban and HIV-p24 human monoclonal antibody (71-31, catalog no. 530) from S. Zolla-Pazner.
This work was supported by NIAID grant AI49057 to V.P. J.L.A. is supported by a Training Program in Microbial Pathogenesis, NIAID T32 AI055434. E.S.Z. is supported by NIH Genetics Training Grant T32 GM07464.