Significance
In nondividing host cells, HIV is targeted by intrinsic antiviral defense mechanisms that introduce marks of damage into viral cDNA, thereby tagging it for processing by cellular DNA repair machinery. Surprisingly, our findings reveal that the two main types of HIV exhibit very different interactions with enzymes involved in DNA repair. HIV-1, but not HIV-2, efficiently removes select DNA repair enzymes, whereas HIV-2 increases dNTP supply in infected cells by removing SAMHD1 (a cell cycle-regulated dNTP triphosphohydrolase) dNTPase. Our findings imply that increasing dNTP supply during viral cDNA synthesis or repair, or blocking cDNA processing by DNA repair enzymes, are alternative strategies used by HIV-2 and HIV-1 to guard their DNA genomes and facilitate their replication/persistence in the host.
Keywords: HIV, Vpr, postreplication DNA repair, SAMHD1, restriction
Abstract
HIV replication in nondividing host cells occurs in the presence of high concentrations of noncanonical dUTP, apolipoprotein B mRNA-editing, enzyme-catalytic, polypeptide-like 3 (APOBEC3) cytidine deaminases, and SAMHD1 (a cell cycle-regulated dNTP triphosphohydrolase) dNTPase, which maintains low concentrations of canonical dNTPs in these cells. These conditions favor the introduction of marks of DNA damage into viral cDNA, and thereby prime it for processing by DNA repair enzymes. Accessory protein Vpr, found in all primate lentiviruses, and its HIV-2/simian immunodeficiency virus (SIV) SIVsm paralogue Vpx, hijack the CRL4DCAF1 E3 ubiquitin ligase to alleviate some of these conditions, but the extent of their interactions with DNA repair proteins has not been thoroughly characterized. Here, we identify HLTF, a postreplication DNA repair helicase, as a common target of HIV-1/SIVcpz Vpr proteins. We show that HIV-1 Vpr reprograms CRL4DCAF1 E3 to direct HLTF for proteasome-dependent degradation independent from previously reported Vpr interactions with base excision repair enzyme uracil DNA glycosylase (UNG2) and crossover junction endonuclease MUS81, which Vpr also directs for degradation via CRL4DCAF1 E3. Thus, separate functions of HIV-1 Vpr usurp CRL4DCAF1 E3 to remove key enzymes in three DNA repair pathways. In contrast, we find that HIV-2 Vpr is unable to efficiently program HLTF or UNG2 for degradation. Our findings reveal complex interactions between HIV-1 and the DNA repair machinery, suggesting that DNA repair plays important roles in the HIV-1 life cycle. The divergent interactions of HIV-1 and HIV-2 with DNA repair enzymes and SAMHD1 imply that these viruses use different strategies to guard their genomes and facilitate their replication in the host.
Nondividing memory T cells and myeloid cells are the main targets of primate lentiviruses during the initial weeks of the acute, in vivo infection (1–4). Infection of these cells is inhibited by intrinsic and innate antiviral mechanisms, several of which converge on reverse transcription of the viral RNA genome. One such restriction is imposed by SAMHD1, a cell cycle-regulated dNTP triphosphohydrolase that, in G1-phase leukocytes, maintains the concentrations of canonical dNTPs below the threshold required for efficient reverse transcription (5–8). Another is caused by a relatively high concentration of noncanonical deoxyuridine triphosphate compared with canonical TTP. dUTP is a substrate for HIV reverse transcriptase, which leads to uracil incorporation into viral cDNA. HIV reverse transcripts are heavily uracilated in macrophages (9, 10). Moreover, viral cDNA is a substrate for apolipoprotein B mRNA-editing, enzyme-catalytic, polypeptide-like 3 (APOBEC3)-family editing enzymes, which convert cytidine to uridine in the minus strand of HIV reverse-transcription intermediates (6, 7, 11, 12). The latter two mechanisms flag viral cDNA for processing by cellular DNA repair enzymes.
The restriction mechanisms that target lentivirus genome replication are counteracted by accessory virulence proteins Vif, Vpx, and Vpr, which usurp specific cellular E3 ubiquitin ligases to direct antiviral proteins for degradation by the proteasome (13). In particular, Vif loads antiviral APOBEC3 family proteins onto a CRL5 E3 ubiquitin ligase for polyubiquitination and subsequent proteasome-dependent degradation (14). Vpx, encoded by the HIV-2/simian immunodeficiency virus (SIV) SIVsm lineage of primate lentiviruses, and closely related Vpr proteins from a subset of SIV viruses isolated from various primate species, counteract SAMHD1-mediated restriction (6, 7, 11). Specifically, Vpx binds to the DCAF1 substrate receptor subunit of the CRL4DCAF1 E3 ubiquitin ligase and loads SAMHD1 onto this enzyme, thereby targeting it for degradation (15). Although HIV-1 does not counteract SAMHD1 directly, it was proposed to bypass the SAMHD1-imposed restriction as a result of a more efficient reverse transcriptase that can synthesize viral cDNA in a low-dNTP environment (16). The failure of the aforementioned viral countermeasures flags HIV cDNA for processing by DNA repair enzymes, which, if not successfully completed in a low-dNTP environment set up by SAMHD1, could lead to the initiation of an innate response to viral nucleic acids, increased HIV-1 mutation rate, and/or inhibition of HIV-1 infection (17–19).
Vpr, a paralogue of Vpx found in all primate lentiviruses, coordinates interactions with postreplication DNA repair machinery, whose role for the replication cycle of primate lentiviruses is not well understood. Early studies revealed that HIV-1 Vpr modulates mutation rates in plasmid shuttle vectors in model systems (20, 21). Vpr, like Vpx, binds the DCAF1 subunit of the CRL4DCAF1 E3 complex and hijacks this enzyme (22). This interaction is associated with the induction of DNA repair foci and activation of the serine/threonine kinase ATR-controlled DNA damage checkpoint; the latter usually reflects the presence of, or failure to repair, damaged DNA at replication forks (23–25). Although the exact mechanism by which Vpr mediates the induction of replication stress is unclear, it was recently linked to the Vpr-mediated recruitment of SLX4-SLX1/MUS81-EME1 structure-specific endonucleases to DCAF1, resulting in activation of the MUS81-EME1 endonuclease and, surprisingly, DCAF1- and proteasome-dependent MUS81 degradation (26). Separately, HIV-1 and HIV-2 Vpr were reported to bind a base excision repair enzyme, uracil DNA glycosylase (UNG2), which can restrict HIV-1 infection in cells with high concentrations of dUTP (19, 27–29). HIV-1 Vpr was shown to target UNG2 to the ubiquitin proteasome pathway via the CRL4DCAF1 E3 complex and thereby disrupt UNG2-initiated base excision repair in HIV-1–infected cells (30–32).
The fact that cellular restriction factors flag HIV cDNA for processing by DNA repair enzymes, taken together with the considerable complexity of cellular repair machineries, raises the possibility that Vpr coordinately engages DNA repair pathways. Unveiling additional targets in these pathways would lead to a more integrated model for the role of Vpr and DNA repair in the HIV life cycle. Here, we performed a proteomic screen for Vpr-interacting DNA repair proteins, aiming in particular to identify novel substrates of the reprogrammed by HIV-1 Vpr CRL4DCAF1 E3 ubiquitin ligase (CRL4DCAF1-H1.Vpr). We show that human HLTF DNA repair helicase is such a target of HIV-1 Vpr. Our findings reveal that HIV-1 Vpr impacts three distinct types of DNA repair transactions and illustrate the complexity of HIV-1 Vpr interactions with cellular postreplication DNA repair machinery. We also show that HLTF and UNG2 are common targets of Vpr proteins from all main groups of HIV-1, but, somewhat unexpectedly, not of those found in HIV-2 A or B. Thus, our findings support the possibility that the two main types of HIV use very different strategies to stabilize their DNA genomes. HIV-2 increases dNTP supply in infected cells by removing SAMHD1, whereas HIV-1 inactivates DNA repair enzymes, possibly to compensate for its inability to drive up dNTP concentrations in primary target cells.
Results
Proteomic Screen for DNA Repair Proteins Associated with HIV-1 Vpr.
We searched for candidate DNA repair proteins that are targeted by HIV-1 Vpr by purifying Vpr–protein complexes from T cells and characterizing their composition by multidimensional protein identification technology (MudPIT) (33). In brief, we constructed a CEM.SS T-cell line harboring a doxycycline-inducible HIV-1 NL4-3 Vpr allele tagged with a double HA-FLAG-epitope (CEM.SS-iH1.Vpr). Vpr expression was induced with doxycycline for 6 h, and Vpr, together with its associated proteins, was purified by successive immunoprecipitations via the HA and FLAG tags (6, 34). As expected, the most abundant cellular polypeptides found to be associated with Vpr were DCAF1, DDB1, and DDA1, subunits of the CRL4DCAF1 E3 complex, which Vpr binds via its DCAF1 subunit (Table S1) (34, 35. Analysis of the MudPIT datasets using the Database for Annotation, Visualization and Integrated Discovery bioinformatics resource (36, 37) identified 21 Vpr-associated proteins that have been linked to DNA replication and/or repair. None of these proteins was detected in MudPIT datasets obtained for purifications from control CEM.SS T cells that did not express Vpr. Among the proteins identified (Table S1) were several that mediate postreplication DNA repair, including UNG2, MSH6, a mismatch repair protein, RFC clamp loader, and PCNA, replication factors that are central to DNA replication and repair (38–40), and HTLF, one of two mammalian homologs of yeast RAD5 DNA helicase that controls the postreplication error-free DNA repair pathway (41, 42). The presence of multiple proteins involved with DNA repair was not surprising, as HIV-1 Vpr and DCAF1 were reported to colocalize with DNA repair foci in chromatin (23). The presence of UNG2, a known, specific substrate of the CRL4DCAF1-H1.Vpr E3, in MudPIT datasets indicates that our experimental approach detects cellular proteins that Vpr recruits to the CRL4DCAF1 E3 ubiquitin ligase and thereby directs for proteasome-dependent degradation.
Table S1.
Identification of HIV-1 Vpr-associated DNA repair proteins
Gene | DNSAF | Function/complex | |||
MudPITα-Vpr 1 | MudPITα-Vpr 2 | Average | MudPIT Mock | ||
HIV-1 Vpr | 0.068256 | 0.132352 | 0.100304 | ND | |
DDB1 | 0.095255 | 0.057099 | 0.076177 | ND | CRL4.DCAF1 |
DDA1 | 0.083479 | 0.043864 | 0.063672 | ND | CRL4.DCAF1 |
DCAF1 | 0.060501 | 0.039376 | 0.049939 | ND | CRL4.DCAF1 |
CUL4A | 0.001688 | 0.004629 | 0.003159 | ND | CRL4.DCAF1 |
CUL4B | 0.002013 | 0.005168 | 0.00359 | ND | CRL4.DCAF1 |
ESCO2 | 0.001924 | 0.003542 | 0.002733 | ND | Cohesin acetylase |
TRIM28 | 0.00277 | 0.002081 | 0.002425 | ND | DNA damage, retrovirus silencing |
UNG | 0.001567 | 0.001613 | 0.00159 | ND | BER |
MSH6 | 0.001520 | 0.001453 | 0.001487 | ND | MMR |
PCNA | 0.000537 | 0.001831 | 0.001184 | ND | DNA replication and repair |
ANKRD17 | 0.001386 | 0.000902 | 0.001144 | ND | DNA replication, innate immunity |
VCP | 0.000435 | 0.001401 | 0.000918 | ND | DSB repair |
RPS27L | 0.001669 | 0.000517 | 0.001093 | ND | p53 inducible, checkpoint signaling |
NONO | 0.000521 | 0.000415 | 0.000468 | ND | DSB repair, NHEJ |
RUVBL2 | 0.000076 | 0.000844 | 0.00046 | ND | DNA helicase, HR |
RFC3 | 0.000804 | 0.000071 | 0.000438 | ND | DNA replication, repair |
MMS19 | 0.000340 | 0.000527 | 0.000434 | ND | DNA replication, repair |
PRKDC | 0.000314 | 0.000395 | 0.000354 | ND | NHEJ |
RFC4 | 0.000579 | 0.00006 | 0.00032 | ND | DNA replication, repair |
TRIP13 | 0.000162 | 0.000452 | 0.000307 | ND | NHEJ |
HLTF | 0.000347 | 0.000237 | 0.000292 | ND | Error-free DNA repair |
KIF22 | 0.000474 | 0.000016 | 0.000245 | ND | Motor, spindle, cohesion |
RFC2 | 0.000198 | 0.000245 | 0.000222 | ND | DNA replication, repair |
RFC1 | 0.000336 | 0.000038 | 0.000187 | ND | DNA replication, repair |
RFC5 | 0.00022 | 0.000068 | 0.000144 | ND | DNA replication, repair |
PRPF19 | 0.000139 | 0.000086 | 0.000113 | ND | ATR, ATRIP recruitment |
MudPIT identification of cellular proteins involved with DNA replication and/or repair that specifically copurify with HIV-1 Vpr from CEM.SS-iH1.Vpr cells. Protein complexes were purified by two sequential immunoprecipitations via HA-tag and then via FLAG-tag, each followed by competitive elution with peptide epitope and analyzed by MudPIT. Upper rows indicate dNSAF values for DDB1, DDA1, DCAF1, Cul4A, and Cul4B subunits of CRL4DCAF1 E3 complex. Lower rows (from ESCO2 down) indicate dNSAF values for DNA replication/repair proteins that copurified with HIV-1 Vpr in two independent experiments (α-Vpr 1 and 2) and their means. None of these Vpr-associated proteins was detected in control immunopurifications from CEM.SS cells that did not express Vpr (Mock). ATR, ataxia telangiectasia related; ATRIP, ataxia telangiectasia-related interacting protein; BER, base excision repair; dNSAF, distributed normalized spectral abundance factor; DSB, double-stranded break; HR, homologous recombination; MMR, mismatch repair; ND, not detected; NHEJ, nonhomologous end-joining.
HIV-1 Vpr Down-Regulates HLTF, a Postreplication DNA Repair Helicase.
To assess whether any of the 21 identified DNA repair proteins is a potential substrate of CRL4DCAF1-H1.Vpr E3, we first tested their levels in CEM.SS-iH1.Vpr and/or U2OS-iH1.Vpr, the latter also harboring a doxycycline-inducible HIV-1 NL4-3 Vpr transgene (Fig. S1). Of note, U2OS cells retain many of the cell cycle regulation characteristics of normal cells and are commonly used for cell cycle/DNA repair/replication studies. Interestingly, the levels of endogenous HLTF were much lower in CEM.SS-iH1.Vpr and U2OS-iH1.Vpr cells that had been arrested by Vpr at the DNA damage checkpoint in the G2 phase of the cell cycle compared with control asynchronously dividing cells that did not express Vpr (Fig. S1). Significantly, HLTF was not depleted in control cells arrested in late S/G2 phase by etoposide or in early M phase by nocodazole treatments. These observations are consistent with the possibility that HLTF, a DNA repair protein expressed in natural target cells of HIV-1 infection (Fig. S2), is a specific target of HIV-1 Vpr.
Fig. S1.
Search for proteins down-modulated by Vpr among Vpr-associated DNA repair proteins. (A) Cell-cycle profiles of CEM.SS-iH1.iVpr and U2OS-iH1.iVpr cells treated with doxycycline for 24 h or control cells cultured with etoposide (0.4 μM, 18 h) or nocodazole (100 ng/mL, 18 h). Percentage fractions of cells in G1, S, and G2/M phases are indicated in each panel. (B) Whole-cell extracts prepared from cells shown in A were immunoblotted with antibodies to the indicated proteins. Asynchronously dividing cells (indicated by “A”) were used as an additional control.
Fig. S2.
HLTF is expressed in HIV natural target cells. Whole-cell extracts prepared from the indicated human leukocyte populations were immunoblotted for HLTF, MUS81, and TFIID loading control.
HIV-1 Vpr Down-Regulates HLTF Independently of Cell Cycle Position.
Vpr activates the ATR-controlled DNA damage checkpoint, thereby arresting cells in G2 phase (24). The possibility existed that HLTF down-regulation is an indirect consequence of Vpr-induced cell cycle perturbations. Hence, to demonstrate that HLTF depletion by Vpr is independent of cell cycle phase and ATR activation, additional experiments were performed.
First, we asked whether Vpr can deplete HLTF in U2OS-iH1.Vpr cells outside of the G2 phase. U2OS-iH1.Vpr were synchronized in late G1/early S phase by double-thymidine block, and Vpr expression was induced at 8 h into the second thymidine treatment (Fig. 1A). Cells were harvested at 0, 6, 12, and 24 h after induction for flow cytometry analysis of cell cycle position and for immunoblot analysis of HLTF levels. As shown in Fig. 1B, U2OS-iH1.Vpr cells remained at the G1/S border through the duration of the experiment. Significantly, endogenous HLTF levels became severely depleted within 6 h of the induction of Vpr expression (Fig. 1C).
Fig. 1.
HIV-1 Vpr depletes HLTF outside of G2 phase. (A) Experimental strategy. U2OS cells harboring the indicated HIV-1 NL4-3 Vpr transgenes were synchronized by double thymidine block and then kept arrested at the G1/S border for the duration of the experiment. HIV-1 Vpr expression was induced by doxycycline during the second thymidine treatment, and cells were collected at the indicated time points for FACS analysis of cell cycle position (B) and immunoblot analysis of the indicated proteins (C). Transcription factor II D (TFIID) was used as a loading control. The cell cycle profile of an asynchronously dividing cell population that did not undergo double thymidine block is shown in B and labeled with “A”.
To assess whether the Vpr effect on HLTF was linked to its interaction with the CRL4DCAF1 E3 ubiquitin ligase, we next tested the Vpr(H71R) variant that does not bind DCAF1 (32). Significantly, this mutant did not detectably modulate HLTF levels even at the late 24-h time point. These findings link the ability of Vpr to deplete HLTF to its interaction with CRL4DCAF1 E3 Ub ligase.
Excess thymidine stresses replication forks (43), potentially contributing to the observed Vpr-mediated HLTF depletion. To exclude this possibility, we characterized HLTF levels across the cell cycle in asynchronously dividing U2OS-iH1.Vpr cells. The cells were cultured in the presence or absence of doxycycline for 6 h, stained with a vital stain, Vybrant DyeCycle Green, to reveal their DNA content, and then sorted into highly enriched G1, S, and G2/M populations (Fig. 2A). Whole-cell extracts prepared from the sorted cells were analyzed by immunoblotting for CyclinA2, HLTF, and UNG2, the previously identified specific substrate of the CRL4DCAF1-H1.Vpr E3 ubiquitin ligase (Fig. 2B). CyclinA2 was detected in only the S and G2/M extracts, as expected, thus confirming the purity of the sorts. HLTF levels were similar in G1, S, and G2/M extracts from uninduced U2OS-iH1.Vpr cells, indicating that HLTF is expressed in all cell cycle phases. Significantly, HLTF levels were much lower in the induced U2OS-iH1.Vpr cells, regardless of their cell cycle position. The levels of UNG2 were similarly depleted following induction of Vpr expression. Finally, HLTF depletion by Vpr was not inhibited by culturing the cells in the presence of caffeine, indicating that the depletion was not a G2/M DNA damage checkpoint-mediated response (Fig. S3).
Fig. 2.
HIV-1 Vpr depletes HLTF independently of cell cycle position. (A) Purity of G1, S, and G2/M cell populations isolated by cell sorting. The DNA profiles of U2OS (mock) and U2OS-iH1.Vpr (HIV-1 Vpr) G1-, S-, and G2/M-phase cells isolated by sorting for DNA content are shown overlaid with that of the parental asynchronously growing unsorted population (labeled “A”). (B) Cell extracts prepared from asynchronously dividing (“A”), G1, S, and G2/M populations were analyzed by immunoblotting with antibodies reacting with HLTF, UNG2, MUS81, CyclinA2, FLAG-tagged Vpr, and TFIID loading control. Asterisk indicates a nonspecific background band revealed by the α-UNG2 antibody.
Fig. S3.
Vpr-mediated depletion of HLTF and MUS81 levels is independent of activation of DNA damage checkpoint responses. U2OS-iH1.Vpr and parental U2OS cells were cultured in the presence of doxycycline (100 ng/mL) and caffeine (0.75 mM, 1.5 mM) for 9 h or 24 h. Then, the cells were (A) stained with 7AAD and the cell cycle profiles were analyzed by FACS or (B) processed for analysis of HLTF, MUS81, Vpr, and TFIID loading control levels by immunoblotting.
Together, the aforementioned findings demonstrate that Vpr down-regulates HLTF levels independently of the cell cycle position and of G2 DNA damage checkpoint, thus further supporting the possibility that HLTF is a substrate for the HIV-1 Vpr-modified CRL4DCAF1 E3 Ubiquitin ligase.
HIV-1 Vpr Targets HLTF Independently of MUS81.
HIV-1 Vpr was recently reported to down-modulate the MUS81 subunit of a structure-specific endonuclease in a DCAF1-dependent manner (26). Therefore, we characterized MUS81 levels in control and Vpr-expressing U2OS-iH1.Vpr sorted cell populations. Strikingly, the endogenous MUS81 levels were not detectably altered following a brief, 6-h-long pulse of Vpr expression. This contrasted with the robust depletion of HLTF in the same time frame (Fig. 2B). These findings implied that the effects of HIV-1 Vpr on HLTF and MUS81 are likely to be independent of each other. We further explored this by carefully comparing the time course of HLTF and MUS81 depletion following induction of Vpr expression in U2OS-iH1.Vpr cells. As shown in Fig. 3A, depletion of HLTF levels was almost complete at the 6-h time point. In contrast, MUS81 levels were down-regulated at a much slower rate, and 24–48 h were required for complete depletion, which coincides with the accumulation of Vpr-expressing cells at the DNA damage checkpoint in late S/G2 phase.
Fig. 3.
Vpr selectively targets the HLTF homolog of RAD5, and this effect is independent from interactions with UNG2 and MUS81. (A and B) Time course of HLTF, SHPRH, MUS81, and UNG2 depletion by HIV-1 Vpr. U2OS-iH1.Vpr and control cells were treated with doxycycline, and cell extracts prepared at the indicated times were analyzed by immunoblotting. Lamin B and TFIID provided loading controls. (C) HLTF, MUS81, and UNG2 levels are not coregulated. U2OS cells were subjected to control nontargeting RNAi (scr) or RNAi targeting HLTF or MUS81, and cell extracts were immunoblotted for HLTF, MUS81, UNG2, and TFIID.
A similar analysis was performed with the HIV-1 Vpr(W54R) variant, which is defective for UNG2 loading onto the CRL4DCAF1 complex but retains binding to DCAF1 (28, 30). This variant retained full ability to deplete HLTF and MUS81 levels (Fig. 3B). We conclude that Vpr exerts its effects on HLTF and MUS81 in a manner independent of UNG2, even though all three require the interaction with DCAF1 (26, 30).
The possibility still remained that lower MUS81 levels in Vpr-expressing cells reflected a cellular response to HLTF depletion, rather than a direct effect of Vpr. Therefore, we asked whether HLTF and MUS81 levels are correlated in the absence of Vpr. Parental U2OS cells were subjected to RNAi targeting of HLTF or MUS81, and cell extracts were prepared 48 h later and analyzed by immunoblotting. Notably, HLTF knockdown led to elevated MUS81 levels, suggesting that the latter was to compensate for the loss of HLTF (Fig. 3C). In contrast, MUS81 depletion did not have a detectable effect on HLTF levels. This evidence supports the possibility that HLTF and MUS81 are specific and independent targets of the CRL4DCAF1-H1.Vpr E3 ubiquitin ligase.
HIV-1 Selectively Depletes Only the HLTF Homolog of Yeast RAD5 DNA Repair Protein.
Mammalian cells possess two Rad5 homologs, HLTF and SHPRH, which serve distinct roles in postreplication DNA repair (41). Experiments were performed to assess whether Vpr selectively depletes HLTF or, instead, targets both Rad5 homologs. As shown in Fig. 3, SHPRH levels remained largely unaltered with a small, at most approximately twofold, decrease at the 24- and 48-h time points. We conclude that Vpr selectively targets the HLTF homolog of yeast Rad5 helicase.
HIV-1 Vpr Targets HLTF for Proteasome-Dependent Degradation via CRL4DCAF1 E3.
We next tested whether Vpr directs HLTF for proteasome-dependent degradation. U2OS-iH1.Vpr cells were treated with doxycycline in the absence or presence of MG132 proteasome inhibitor for 9 h. As shown in Fig. 4A, proteasome inhibition stabilized HLTF levels in Vpr expressing cells while having no detectable effect on HLTF in control cells. We conclude that Vpr activates a proteasome-dependent mechanism to down-modulate HLTF levels.
Fig. 4.
HIV-1 Vpr recruits HLTF to the DDB1-DCAF1 module of the CRL4DCAF1 E3 complex for proteasome-dependent degradation. (A) Vpr targets HLTF for proteasome-dependent degradation. U2OS-iH1.Vpr cells (HIV-1 Vpr) and control U2OS cells (mock) were treated with doxycycline or not treated in the absence or presence of MG132 proteasome inhibitor (1 μg/mL) for 9 h. HLTF, MUS81, and Vpr levels in cell lysates were revealed by immunoblotting. TFIID provided a loading control. (B) Schematic representation of HIV-1–mediated recruitment of HLTF onto the CRL4DCAF1 E3 complex. The placement of HA and FLAG epitope tags on Vpr and HLTF, respectively, is indicated. (C) Vpr recruits HLTF to the DCAF1-DDB1 module of the CRL4DCAF1 E3 complex. FLAG-HLTF was transiently coexpressed with myc-tagged HIV-1 Vpr or Vpr(H71R) in HEK293T cells as indicated. Endogenous DCAF1, DDB1, and ectopic Vpr and HLTF were revealed in HLTF immune complexes and in detergent extracts by immunoblotting.
Next, we asked whether HIV-1 Vpr can recruit HLTF to the endogenously expressed CRL4DCAF1 E3 complex (Fig. 4 B and C). FLAG-tagged HLTF was expressed alone or coexpressed with HIV-1 NL4-3 Vpr or the Vpr(H71R) variant in HEK 293T cells. Detergent extracts were prepared from the transfected cells, and HLTF immune complexes were analyzed by immunoblotting for HLTF, Vpr, and the DCAF1 and DDB1 subunits of the CRL4DCAF1 E3. We found that HIV-1 Vpr directed the assembly of a protein complex containing HLTF as well as DCAF1 and DDB1. In contrast, neither the Vpr(H71R) variant, which retained a reduced binding to HLTF, nor HLTF alone, nucleated such a DCAF1-containing complex. These observations indicate that Vpr mediates specific recruitment of HLTF to the CRL4 E3 complex that uses the DCAF1 substrate receptor, to which Vpr binds. Of note, HLTF complexes formed in the absence of Vpr contained low levels of DDB1, which suggests that HLTF may physiologically participate in DDB1 complexes that do not contain DCAF1.
Overall, these findings indicate that HIV-1 Vpr specifically binds to HLTF and recruits it to the DCAF1-DDB1 module of the CRL4DCAF1 E3 complex, thereby directing HLTF for degradation by proteasome.
HIV-1 Vpr Interaction with HLTF Defines Previously Unidentified Functional Elements in both Proteins.
HLTF comprises an N-terminal DNA-binding HIRAN domain and a C-terminal RING domain possessing E3 Ubiquitin ligase activity (Fig. 5A). To map the HLTF region that mediates the effect of Vpr, we analyzed the activity of HIV-1 NL4-3 Vpr toward a set of HLTF deletion mutants by using a transient coexpression assay in HEK 293T cells. This assay faithfully reproduced the Vpr-induced proteasome-dependent HLTF down-modulation (Fig. S4). As shown in Fig. 5B, an HLTF fragment lacking the N-proximal 151 residues, including the HIRAN domain [HLTF(152–1009)], as well as those lacking the C-distal ∼700 residues comprising the RING domain and HELICc/DEXDc helicase components, were fully responsive to HIV-1 Vpr, thus tentatively mapping the Vpr target site to HLTF residues 152–299. This region appears to serve as a linker between the HIRAN domain and the N-proximal component of the helicase ATP-binding domain, and, as such, is not known to possess another function.
Fig. 5.
HIV-1 Vpr interaction with HLTF does not involve known functional elements in both proteins. (A) Summary of mutations in HLTF and their effects on Vpr-mediated down-modulation of HLTF levels. Schematic representation of the HLTF protein and HLTF deletion mutants shows the location of the HIRAN, RING, SNF2, and discontinuous DEXDc and HELICc helicase domains. The HLTF region mediating Vpr sensitivity is boxed. (B) HIV-1 Vpr does not act on known functional domains of HLTF. FLAG-HLTF and its variants, indicated in A, were transiently coexpressed with increasing doses of HA-tagged HIV-1 Vpr in HEK 293T cells, and HLTF and Vpr levels in cell extracts were revealed by immunoblotting. α-Tubulin provided a loading control. (C) The Vpr N terminus is required for depletion of HLTF levels. HIV-1 Vpr or Vpr(24R,36P) variant was coexpressed at increasing doses with HLTF (Upper) or UNG2 (Lower) in HEK293T cells, and cell extracts were analyzed as described earlier. (D) The E24R,R36P mutation selectively disrupts Vpr binding to HLTF. HA-Vpr and its variants were coexpressed with FLAG-HLTF in HEK 293T cells. The Vpr(H71R) variant that does not bind DCAF1 was used as a negative control. Endogenous DCAF1, DDB1, and ectopic Vpr and HLTF were revealed in Vpr immune complexes and detergent extracts by immunoblotting.
Fig. S4.
Transient dose-dependent assay for Vpr-induced DCAF1- and proteasome-dependent HLTF degradation. (A) FLAG-HLTF was transiently coexpressed with HIV-1 HA-Vpr in HEK 293T cells. Cells were cultured overnight in the presence or absence of MG132 (0.62 μM, 1.25 μM) or lactacystin (2.5 μM, 5 μM), proteasome inhibitors, and HLTF, Vpr, and α-tubulin loading control levels were revealed in whole-cell extracts by immunoblotting. (B) The ability of HIV-1 Vpr to deplete HLTF levels requires binding to DCAF1. The effects on HLTF levels of HIV-1 Vpr (NL4-3) and two variants, Vpr(H71R) or Vpr(W54R), that do not bind DCAF1 or do not degrade UNG2, respectively, were tested in transient coexpression assays.
We next mapped the residues in HIV-1 Vpr that are required for the effect on HLTF levels. Because Vpr and its paralogue Vpx use their N-terminal regions to recruit novel protein substrates to CRL4DCAF1 E3 (15, 30, 44), we constructed a set of HIV-1 Vpr point mutants in the N-proximal region and screened them by using the previously described transient, dose-dependent HLTF down-modulation assay. As shown in Fig. 5C, a double amino acid substitution E24R,R36P [Vpr(E24R,R36P)] diminished the ability of Vpr to down-modulate HLTF, but not UNG2. These observations revealed that the E24R,R36P mutation selectively disrupts Vpr binding to HLTF, but does not grossly interfere with binding to DCAF1 or recruitment of UNG2 to the CRL4DCAF1 E3 complex.
To corroborate this possibility, we characterized the binding of the Vpr(E24R,R36P) variant to HLTF and DCAF1. FLAG-HLTF was coexpressed with HA-Vpr or HA-Vpr(E24R,R36P) in HEK 293T cells, and Vpr immune complexes were analyzed by immunoblotting. The Vpr and Vpr(E24R,R36P) variants efficiently coprecipitated the endogenously expressed DCAF1-DDB1 module of CRL4DCAF1 E3 complex (Fig. 5D). Vpr(H71R), which was analyzed in parallel as a negative control, did not associate with DDB1-DCAF1, and only weakly associated with HLTF, as expected. Significantly, only WT Vpr, but not the Vpr(E24R,R36P) variant, efficiently coprecipitated HLTF. Notably, to our knowledge, the E24 and R36 residues have not been reported to be important for any other known Vpr interaction with cellular DNA repair machinery. These findings together confirm that Vpr recruits HLTF to DCAF1 independent of its previously reported interactions with other DNA repair proteins.
HLTF, MUS81, and the Induction of G2 Arrest by Vpr.
We asked whether Vpr-mediated depletion of HLTF disrupts postreplication DNA repair sufficiently to modulate the DNA damage checkpoint such that cells are arrested in the G2 phase of the cell cycle. To this end, we tested the HIV-1 Vpr(E24R,R36P) variant for its ability to arrest cells in G2 phase and determined whether HLTF and/or MUS81 are required for the induction of G2 arrest by Vpr. U2OS-iH1.Vpr(E24R,R36P) cells were induced to express HLTF degradation-defective Vpr(E24R,R36P) variant, and their cell cycle profile were determined 2 d later. As shown in Fig. 6A, HIV-1 Vpr(E24R,R36P) retained the ability to arrest U2OS cells in G2 phase. Similar experiments performed with U2OS HLTF-KO cells [U2OS.HLTF.KO-iH1.Vpr (45)] revealed that WT NL4-3 Vpr arrested these cells in G2 phase (Fig. 6B, panel 12). Thus, HLTF deficiency per se is not sufficient to trigger DNA-damage checkpoint, as reported in previous studies (45, 46), nor is HLTF required to mediate cell cycle arrest in G2 phase by Vpr.
Fig. 6.
The effects of HLTF and MUS81 depletion on HIV-1 Vpr-induced G2 arrest. (A) Vpr(E24R,R36P) arrests U2OS cells in G2 phase. Cell cycle profiles of U2OS cells induced with doxycycline [Dox (+)], or not [Dox (−)] to express HIV-1 NL4-3 Vpr, Vpr(E24R,R36P), or Vpr(H71R) for 48 h. The percentage fraction of cells in G1, S, and G2 phase is indicated, and panel numbers are shown in upper right corner. The abscissa is DNA content shown on a linear scale. The ordinate is DNA synthesis shown on a logarithmic scale (B) HIV-1 Vpr arrests U2OS.HLTF.KO cells in G2 phase. Cell-cycle profiles of U2OS.HLTF.KO cells induced with doxycycline [Dox (+)] or not [Dox (−)] to express HIV-1 NL4-3 Vpr for 48 h. (C) Vpr arrests MUS81-depleted U2OS and U2OS.HLTF.KO cells in G2 phase. Schematic representation of cell cycle profiles of parental (mock) U2OS (HLTF), U2OS.HLTF.KO cells (HLTF.KO), or their derivative cell lines carrying doxycycline-inducible HIV-1 Vpr transgenes (HIV-1 Vpr). The cells transfected with siRNA targeting MUS81 (MUS81) or nontargeting (scr) and induced (or not) with doxycycline 2 d later as indicated. Cell cycle profiles were determined 2 d after induction, and cells in G1, S, and G2/M phases were quantified as shown in A. Each bar represents averaged results from four replicates. The significance of the observed differences in G2-phase populations between the indicated conditions shown above the bars was calculated using an unpaired two-tailed Student’s t test with Welch’s correction (n = 4; *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). Representative results of three independent experiments are shown.
As the data shown in Fig. 3C suggest a possible compensatory interaction between MUS81 and HLTF, we next tested the effect of RNAi-mediated MUS81 knockdown on cell cycle distribution of U2OS.HLTF.KO cells in the absence of Vpr expression (Fig. 6C and Fig. S5). Interestingly, MUS81 knockdown in U2OS.HLTF.KO cells was associated with an altered cell cycle profile with an increase in the G2-phase population compared with that for cells subjected to nontargeting siRNA, even in the absence of Vpr (P < 0.01) (Fig. 6C). This observation indicated that a combined MUS81 and HLTF deficiency alone could lead to the accumulation of cells in G2 phase.
Fig. S5.
RNAi-mediated depletion of MUS81 in U2OS and U2OS.HLTF.KO cells (relevant to Fig. 6). U2OS [HLTF(+/+)] and U2OS.HLTF.KO [HLTF(−/−)] cells were subjected to RNAi targeting MUS81, and whole-cell extracts analyzed by immunoblotting for the indicated proteins.
Next, we investigated whether MUS81 is required for the induction of G2 arrest by HIV-1 Vpr in U2OS-iH1.Vpr cells. MUS81 levels were depleted by RNAi 48 h before induction of Vpr expression, and cell-cycle profiles were determined 2 d after induction. Strikingly, MUS81 depletion did not diminish the ability of Vpr to arrest cells in G2. To the contrary, it slightly exacerbated the Vpr phenotype in these cells, but the difference was not statistically significant. Notably, a similar effect was seen in HLTF-KO cells, in which the difference conferred by the MUS81 knockdown was statistically significant (P < 0.01). These observations reveal that the presence of MUS81 is probably not required for the induction of G2 arrest by Vpr. This evidence supports the model in which replication stress resulting from combined depletion of MUS81 and HLTF contributes to the ability of Vpr to arrest cells at the DNA damage checkpoint in G2 phase.
HLTF and UNG2 Are Common Targets of HIV-1 and SIVcpz Vpr Proteins.
Experiments were performed to establish whether HLTF is a common target of primate lentiviral Vpr proteins. We focused on Vpr proteins from HIV-1 and HIV-2, which represent two evolutionary divergent branches of primate lentiviruses that adapted to replicate in human cells. Fig. 7A shows an amino acid sequence alignment for consensus Vpr proteins derived from the main groups of HIV-1 (M, N, O) and SIVcpz viruses, isolated from two chimpanzee subspecies (Ptt, Pts) from which HIV-1 originated after cross-species transmissions (47), as well as Vpr proteins from HIV-2 groups A and B. Of note, to account for the considerable amino acid sequence divergence at the C terminus of HIV-2 group A Vpr proteins, two distinct consensus Vpr proteins, termed A1 and A2, were derived. Inspection of the alignment revealed that residues E24 and R36, which we found to be important for HLTF depletion by HIV-1 NL4-3 Vpr, are conserved in all consensus HIV-1 and SIVcpz Vpr sequences (Fig. 7A). Similarly, residue W54, which is required for UNG2 loading onto CRL4DCAF1 E3, was also present in all HIV-1/SIVcpz consensus Vpr sequences. These observations predicted that HLTF as well as UNG2 are common targets of HIV-1 and SIVcpz Vpr proteins.
Fig. 7.
HLTF and UNG2 are depleted by HIV-1 and SIVcpz but not by HIV-2 Vpr proteins. (A) Alignment of consensus sequences of Vpr proteins from main HIV-1 (M, N, O), SIVcpz (Ptt, Pts), and HIV-2 (A1, A2, B) groups. Sequences are shown in one letter code. Residues required for binding to HLTF (E24, R36), UNG2 (W54), and DCAF1 (H71) are boxed. (B) HIV-1 and SIVcpz Vpr programs HLTF and UNG2 for degradation. FLAG-HLTF or FLAG-UNG2 were transiently coexpressed with increasing doses of the indicated consensus HA-epitope–tagged HIV-1 Vpr proteins. Levels of HLTF and Vpr in cell extracts were revealed by immunoblotting. α-Tubulin provided loading control. (C and D) Consensus HIV-2 Vpr and Vpx proteins do not modulate HLTF or UNG2 levels. FLAG-HLTF or FLAG-UNG2 were coexpressed with the indicated HA-epitope–tagged consensus HIV-2 Vpr or Vpx proteins. HLTF, UNG2, and Vpr/Vpx were analyzed as indicated earlier.
To test this prediction, we synthesized vpr genes for the consensus Vpr proteins shown in Fig. 7A and characterized their ability to deplete HLTF and UNG2. As shown in Fig. 7B, all tested HIV-1 and SIVcpz Vpr proteins down-modulated HLTF and UNG2. Their activities were comparable to those of the NL4-3 Vpr that we used in the studies described earlier. Remarkably, even the SIVcpzPts Vpr, which is ∼30% divergent from HIV-1 M Vpr in the core region, was quite active against HLTF.
HIV-2 Vpr and Vpx Do Not Modulate HLTF Nor UNG2 Levels.
Sequence alignment, shown in Fig. 7A, revealed that the key residues required for robust interaction with HLTF, as well as UNG2, are not conserved in consensus HIV-2 Vpr proteins. Indeed, the consensus HIV-2 groups A and B Vpr proteins were inactive toward HLTF and UNG2 despite being expressed to levels much higher than that of the NL4-3 Vpr (Fig. 7C). Importantly, the HIV-2 consensus Vpr proteins bound DCAF1 in coimmunoprecipitation assays, similar to HIV-1 NL4-3 Vpr, and therefore appeared to be functional (Fig. S6A). We conclude that only the HIV-1 and SIVcpz, but not HIV-2, Vpr proteins efficiently direct HLTF and UNG2 for proteasome-dependent degradation via CRL4DCAF1-H1.Vpr E3 ubiquitin ligase.
Fig. S6.
Consensus HIV-2 Vpr and Vpx proteins are functional. (A) Consensus HIV-2 subtype A and B Vpr proteins bind the endogenous DDB1-DCAF1 module of CRL4DCAF1 E3 complex. Consensus HIV-2 HA-Vpr proteins were transiently expressed in HEK293T cells. HIV-1 NL4-3 Vpr and its H71R-substituted variant that does not bind DCAF1 were used as controls. Endogenous DCAF1, DDB1, and ectopic Vpr in α-HA-Vpr immune complexes and detergent extracts were revealed by immunoblotting. (B) Vpx proteins deplete SAMHD1 levels. HIV-2 HA-Vpr proteins were transiently coexpressed with SAMHD1 in HEK 293T cells. HIV-1 NL4-3 Vpr was expressed in parallel as a negative control. Cells were cultured overnight in the presence or absence of MG132 proteasome inhibitor (1 μg/mL), and SAMHD1, Vpr, Vpx and α-tubulin loading control levels were revealed in whole-cell extracts by immunoblotting.
HIV-2/SIVsm lineage viruses encode a Vpr paralogue termed Vpx, which was lost during adaptation of ancient SIV to chimpanzees and is absent in the present HIV-1/SIVcpz branch of primate lentiviruses (48). Vpr and Vpx proteins usurp CRL4DCAF1 E3 and are highly adaptable, as revealed by their shared ability to reprogram this ubiquitin ligase toward a common SAMHD1 substrate in many SIV lineages (11). Therefore, we asked whether HIV-2 Vpx has acquired the ability to program HLTF or UNG2 degradation. As shown in Fig. 7D, neither the HIV-2 A nor B group consensus Vpx protein decreased HLTF or UNG2 levels. Of note, the Vpx proteins were functional as judged by their ability to deplete SAMHD1 (Fig. S6B). We conclude that HIV-2 lineage Vpr/Vpx proteins do not endow CRL4DCAF1 E3 with specificity toward HLTF or UNG2.
HLTF and UNG2 Are Depleted in HIV-1–Infected Cells.
To corroborate the aforementioned findings, we characterized HLTF and UNG2 levels in the context of HIV infection. As shown in Fig. 8 A and B, challenge of Jurkat T cells with VSV-G pseudotyped, Vpr-loaded HIV-1 virus-like particles (VLPs) or infection with a VSV-G pseudotyped single-cycle HIV-1 NL4-3.GFP.R+ carrying an intact vpr gene led to the depletion of HLTF and UNG2. Importantly, infection of primary CD4+ T cells with NL4-3.GFP.R+ virus was also associated with a profound depletion of HLTF levels, indicating that endogenous Vpr expression is sufficient to remove HLTF from natural HIV-1 target cells (Fig. 8C). In contrast, these effects were not seen in cells infected with control capsid-normalized HIV-1 VLP produced in the absence of Vpr, or with vpr-deficient NL4-3.GFP.R– HIV-1 (Fig. 8 A and B).
Fig. 8.
Divergent effects of HIV-1 and HIV-2 on HLTF and UNG2 levels. (A and B) HIV-1 infection depletes HLTF and UNG2 levels. (A) Jurkat T cells were infected with HIV-1 VLP transcomplemented (Vpr) or not (−) with HIV-1 Vpr (HIV-1 VLP; Left) or a single-cycle HIV-1 NL4-3.GFP.R+ (vpr+), or R− (vpr−) virus (HIV-1; Right). The levels of HLTF, UNG2, and α-tubulin loading control in extracts prepared from infected and control (marked “c”) cells 48 h post infection were revealed by immunoblotting. (B) HIV-1 VLP and HIV-1 NL4-3.GFP.R+ and R– viruses used in A were normalized by immunoblotting for p24 capsid. Vpr was revealed with an α-HA antibody (HIV-1 VLP) or α-Vpr antibody (HIV-1). (C) HIV-1 depletes HLTF and MUS81 in primary CD4+ T cells. CD4+ T cells were activated with α-CD3/α-CD28 beads and transduced with HIV-1 NL4-3.GFP.R+ (or R–), and GFP-positive cells were isolated by cell sorting 2 d after infection and analyzed for expression of the indicated proteins by immunoblotting. Twofold serial dilutions of control CD4+ T whole-cell lysates provided quantification standards. (D and E) Divergent effects of HIV-1 and HIV-2 on HLTF and UNG2 levels. Jurkat T cells were infected with HIV-1 NL4-3.GFP.R+ (HIV-1) or HIV-2 ROD with intact vpr and vpx genes expressing an RFP marker. Two days after infection, reporter gene expression was revealed by FACS. The abscissa is GFP/RFP fluorescence shown on a logarithmic scale. The ordinate is forward scatter shown on a linear scale (D), and UNG2, SAMHD1, HLTF, and Lamin B loading control in cell extracts were visualized by immunoblotting (E). (F and G) THP-1 cells were infected with HIV-1 NL4-3.GFP.R+ [or R− (−)] or HIV-2 ROD expressing Vpr and Vpx (vpr/vpx) or only Vpr [vpr/(−) or not infected (marked as “c”)] (F), or with SIVmac251 VLP or SIVmac239 (G) as indicated, and extracts were analyzed by immunoblotting. Asterisk indicates a nonspecific band revealed by the α-SAMHD1 antibody.
Next, we asked whether HIV-1 and HIV-2 indeed exert divergent effects on HLTF and UNG2. Jurkat T cells were infected with two doses of normalized, single-cycle HIV-1 NL4-3.GFP.R+, and HIV-2 Rod having intact vpr and vpx genes (Fig. 8 D and E). HIV-1 infection led to a dose-dependent depletion of HLTF and UNG2 levels, as expected. In contrast, HIV-2 did not exert a comparable robust effect, even though the levels of both proteins appeared to be somewhat lower in cells infected with the highest dose of HIV-2. In parallel, similar experiments were performed in THP-1 cells (Fig. 8F). Of note, these cells express SAMHD1 at high levels, but HLTF levels were not detectable by immunoblotting. We found that HIV-1 readily depleted UNG2 while having no effect on SAMHD1 levels. In contrast, HIV-2 depleted SAMHD1 in a Vpx-dependent manner, as expected, whereas UNG2 levels were not affected. To corroborate these findings, we tested two ancestor SIVmac251 and SIVmac239 viruses, each encoding an uninterrupted vpr gene. As shown in Fig. 8G, infection of THP-1 cells resulted in almost complete depletion of SAMHD1 but had no detectable effect on UNG2 levels in these cells. We conclude that HIV-1/SIVcpz lineage viruses counteract HLTF and UNG2 DNA repair proteins via the CRL4DCAF1 E3, whereas the HIV-2/SIVmac lineage viruses tested do not possess a comparable ability.
Discussion
When replicating in primary, nondividing, target cells, primate lentiviruses are targeted by innate immune mechanisms that introduce marks of damage into viral cDNA, thereby flagging it for processing by host-cell DNA repair enzymes. Hence, it is not surprising that lentiviral accessory proteins intersect with cellular mechanisms that modulate DNA repair, presumably to moderate the negative impact of DNA repair on virus genome stability and/or priming an innate response to viral nucleic acids. In HIV-1, the interactions with postreplication DNA repair processes are coordinated via the CRL4DCAF1 E3 ubiquitin ligase, which is hijacked by Vpr (22, 26, 34). The present study aimed to gain a more comprehensive description of the interactions between HIV-1 Vpr and cellular DNA repair pathways. To this end, we performed a proteomic screen, which led to the identification of HLTF DNA helicase as a specific target of the CRL4DCAF1-H1.Vpr E3 ubiquitin ligase. Of note, HLTF was also identified as HIV-1 Vpr target by Lahouassa et al. (49).
Our evidence supports the possibility that HLTF is a direct target of HIV-1 Vpr for recruitment to the CRL4DCAF1 E3 and proteasome-dependent degradation. First, Vpr rapidly depletes HLTF independently of cell cycle position, and this is not a downstream effect of DNA damage checkpoint activation. Second, coimmunoprecipitation experiments show that Vpr bridges HLTF to the CRL4DCAF1 E3 complex. Third, Vpr-mediated depletion does not require the HLTF RING domain, which was shown to mediate proteasome-dependent HLTF degradation in response to DNA damage (41). These latter findings indicate that HLTF depletion is not a consequence of other Vpr interactions with DNA repair pathways.
Overall, our studies reveal a striking complexity of Vpr interactions with cellular DNA repair machinery. This and previous reports together demonstrate that HIV-1/SIVcpz Vpr uses the CRL4DCAF1 E3 to remove, from infected cells, three key enzymes involved in distinct DNA repair pathways (15, 26, 32). Significantly, the recruitment of HLTF and UNG2, as well as Mus81 and UNG2, to CRL4DCAF1 E3 is mediated via different surfaces of the HIV-1 Vpr molecule. Although we have not yet separated HLTF and MUS81 binding surfaces on Vpr, RNAi studies revealed that expression of these two DNA repair proteins is not coordinated, thereby implying that Vpr depletes their levels independently of each other. Thus, the effects on UNG2, HLTF, and MUS81 most likely reflect three independently selected functions of HIV-1 Vpr.
Viral accessory virulence factors usually redirect cellular E3 ubiquitin ligases to remove cellular proteins that exhibit direct or indirect antiviral activity (50). Hence, the concerted removal of several postreplication DNA repair enzymes by Vpr is consistent with the notion that host cell DNA repair machinery restricts steps in HIV-1 replication in host cells. Indeed, the key role for a dUTP-mediated antiretroviral host defense pathway is evident from the fact that many retrovirus families captured a host dUTPase gene during viral evolution (51, 52). Although HIV-1 does not encode dUTPase, its Vpr protein effectively prevents UNG2-initiated uracil excision repair (31). This alternative mechanism may allow HIV-1 to alleviative negative consequences of UNG2-mediated uracil excision and downstream pathways for the integrity of HIV-1 cDNA (19).
The finding that HIV-1/SIVcpz Vpr specifically removes HLTF from infected cells suggests that it negatively impacts HIV-1 replication. Possible scenarios are suggested by known HLTF functions. In particular, HLTF remodels replication forks that are stalled by DNA lesions into four-way branched DNA structures, thereby providing an undamaged DNA template that allows for error-free bypass of the lesion and resumption of DNA replication. Interestingly, fork-like branched DNA structures resembling replication forks are transient intermediates in plus-strand displacement synthesis that is carried out by retroviral reverse transcriptase (53, 54). Recognition and processing of such intermediates by HLTF could impede an ordered synthesis of the cDNA copy of the HIV-1 RNA genome. Although integration competent HIV-1 preintegration complexes form in the cytoplasm, the completion of reverse transcription does not appear to be a prerequisite for HIV-1 entry to the nucleus (55), where partially reverse-transcribed viral cDNA can be accessed by DNA repair proteins. Alternatively, HLTF could modulate processing of the integrated proviral DNA, as it is known to control the choice between two pathways of postreplication repair of DNA lesions at the replication fork: error-free, by template switching, and error-prone, through recruitment of mutagenic translesion DNA polymerases (45, 56). As these two distinct modes of DNA repair, error-free and error-prone, have different mutagenic outcomes upon repair of chromosomal DNA, they would also have different consequences for the fidelity of HIV-1 provirus repair, which provides another tentative rationale for the removal of HLTF by HIV-1 Vpr.
Notably, Vpr also disrupts one of the Holliday junctions resolvases by depleting the MUS81 component of SLX1-SLX4/MUS81-EME1 (2) complex. This interaction was proposed to be associated with untimely activation of MUS81-EME1 (2) nuclease activity to prevent an innate response to products of abortive HIV-1 reverse transcription (26). Somewhat unexpectedly, our data show that depletion of MUS81 levels does not alleviate the ability of Vpr to arrest cells in G2. Moreover, MUS81 and HLTF depletion, although each insufficient on its own, together appear to facilitate G2 arrest, consistent with the previously reported roles of these proteins in replication fork maintenance and restart, and with recent genetic studies of the Vpr–MUS81 interaction (45, 57, 58). Thus, it appears that HIV-1 Vpr must perturb additional pathways to trigger DNA damage checkpoint leading to G2 arrest.
The findings described here provide compelling evidence that HIV-1 Vpr disrupts select DNA repair pathways via CRL4DCAF1 E3 ubiquitin ligase. This in turn prompts speculation that Vpr acts to delay the repair until after integration of the HIV-1 provirus into the host cell chromosome, an environment in which repair is conventionally handled by alternative postreplication DNA repair pathways. Alternatively, Vpr may block the repair through these pathways altogether and thereby evade DNA repair-mediated restriction. Such scenarios, as well as the timing of the action of HLTF, UNG2, and other aspects of DNA repair impacted by Vpr, will be assessed in future studies.
Our comparative studies of the two main types of HIV unexpectedly revealed that HIV-1/SIVcpz Vpr proteins potently deplete HLTF and UNG2, whereas HIV-2 and their ancestral HIV-2sm lineage Vpr do not possess such robust abilities. Significantly, HIV-2/HIVsm viruses antagonize SAMHD1 dNTPase via their Vpx proteins (6, 7) whereas HIV-1/SIVcpz do not possess this ability. We propose that HIV-2 does not need to antagonize cellular DNA repair mechanisms in a manner comparable to that seen with HIV-1, because it effectively counteracts SAMHD1, which leads to an increase in cellular dNTP concentrations. Through this mechanism, HIV-2 avoids marking its cDNA for processing by DNA repair enzymes and the resulting repair-mediated restriction and/or provides sufficient dNTP supply to support faithful repair in nondividing G1 cells, unlike HIV-1. Regardless, the evidence presented here implies that distinct lineages of primate lentiviruses use different strategies to manage their genomes, protecting them from the acquisition of DNA damage and/or from processing by DNA repair enzymes when replicating in dNTP-poor cellular environments.
The emergence of SIVcpz is thought to have involved recombination between two SIV lineages from Old World Monkey (OWM) species (59), resulting in a deletion of the vpx gene and the ensuing loss of SAMHD1 antagonism by Vpx. SAMHD1 antagonism by Vpx is well-conserved in distinct SIV strains isolated from OWM (11, 60), indicating the importance of SAMHD1-exerted restriction on replication of primate lentiviruses, and thereby raising the question of how the loss of SAMHD1 counteraction was compensated for in the HIV-1/SIVcpz lineage. One such mechanism was suggested by the previous finding that cross-species transmission of SIV to chimpanzees was associated with an expansion of the range of ABOBEC3 proteins targeted by Vif (48). Our studies reveal that the loss of Vpx in the HIV-1/SIVcpz lineage was also associated with the acquisition/enhancement of an ability to disrupt DNA repair by HIV-1 Vpr via HLTF and UNG2, and suggest that the latter was to compensate for the inability to antagonize SAMHD1. This possibility makes sense mechanistically, as reverse transcription, taking place in the dNTP-poor environment of nondividing target cells, leads to incorporation of uracil and possibly other marks of damage into viral cDNA, thereby flagging it for processing and restriction by UNG2 and DNA repair enzymes.
Materials and Methods
Cell Lines.
HEK293T, U2OS, and U2OS.HLTF.KO cells (45) were maintained in DMEM (Life Technologies) supplemented with 10% (vol/vol) FBS, 2 mM l-glutamine, and penicillin/streptomycin in 5% (vol/vol) CO2 at 37 °C. CEM.SS (61) (NIH AIDS Reagent Program), Jurkat, and THP-1 cells were maintained in RPMI 1640 medium supplemented as described earlier. CEM.SS-iH1.Vpr, U2OS-iH1.Vpr, U2OS-iH1V.Vpr(E24R,R36P), and U2OS.HLTF.KO-iH1.Vpr cells harboring doxycycline-inducible HIV-1 NL4-3 vpr or other variant vpr transgenes were engineered by using lentiviral Tet-On 3G Inducible Expression System (Clontech) and maintained in the aforementioned media supplemented with G418 (200 μg/mL) and puromycin (2 μg/mL). Expression was induced by addition of doxycycline (Sigma-Aldrich) to the culture medium.
Immunoblotting and Antibodies.
Typically, whole-cell (34), cytoplasmic, or chromatin extracts (62) were separated by SDS/PAGE and transferred to PVDF membranes for immunoblotting. Proteins were detected with appropriate primary antibodies and immune complexes revealed with HRP-conjugated antibodies specific for the Fc fragment of mouse or rabbit IgG (Jackson ImmunoResearch Laboratories) and enhanced chemiluminescence (GE Healthcare), or with fluorescent antibodies to mouse or rabbit IgG (KPL) and Odyssey Infrared Imager (Licor). SI Materials and Methods includes the list of antibodies used.
Transfections, Immunoprecipitations, and DDB1-DCAF1 Recruitment Assay.
Transfections of HEK 293T cells and immunoprecipitations were performed as described previously (34, 63). For HLTF recruitment assays, HEK 293T cells, at 2 × 107 cells in four 10-cm plates per condition, were cotransfected with pCG plasmids expressing FLAG-tagged HLTF and appropriate HA-tagged HIV-1 Vpr proteins in combinations. Whole-cell extracts were immunoprecipitated with FLAG-M2 beads (Sigma-Aldrich), and immune complexes were eluted by competition with FLAG-peptide under native conditions. HIV-1 Vpr immune complexes for MudPIT analyses were purified as we described previously (6, 63).
Cell Synchronization and Cell Cycle Analyses.
U2OS-iH1.Vpr cells were synchronized in early S phase by double thymidine block. To reveal cell-cycle profiles, aliquots of 1 × 105 cells were pulse-labeled with 5-ethynyl-2′-deoxyuridine (EdU; 10 μM) for 60 min, and the incorporated EdU was detected by using a Click-iT Plus EdU Alexa Fluor 647 or Alexa Fluor 488 Flow Cytometry Assay Kit (Life Technologies). Cells were then stained with 2 μg/mL 7AAD (Life Technologies) to reveal their DNA content, and analyzed with an LSRFortessa flow cytometer (BD Biosciences) and FlowJo software. A total of 10,000 events were collected for each sample. Statistical analyses of cell cycle profiles based on quantification of cell populations in G1, S, and G2/M phases in FlowJo were performed in GraphPad Prism software, with P values calculated by unpaired two-tailed Student’s t test with Welch’s correction.
Isolation of Primary HIV-1–Infected CD4+ T Cells by Cell Sorting.
CD4+ T cells obtained from human peripheral blood mononuclear cells by negative selection using EasySep hCD4+ T Cell Enrichment Kit (Stemcell Technologies) were plated in 96-well plates at 1 × 106 cells per well and activated with Dynabeads Human T-Activator CD3/CD28 (Invitrogen) in the presence of IL-2 (30 U/mL) in full RPMI 1640 medium [supplemented with 10% (vol/vol) heat-inactivated FBS and antibiotics (63)]. Two days later, the cells were infected with HIV-1 NL4-3.GFP.R+ or NL4-3.GFP.R− virus and transferred into wells of a 24-well plate in a final volume of 500 μL per well of full RPMI 1640 medium with IL-2 (30 U/mL). Two days post infection, cells were pooled and resuspended in PBS + 1% BSA at 8 × 106/mL. Dynabeads were removed and live GFP-positive cells were isolated by sorting on a FACSAria. Whole-cell lysates were prepared from sorted and from uninfected control cells.
Isolation of G1, S, and G2M Cells by Sorting.
Cells (8 × 106) were stained with 30 μM Vybrant DyeCycle Green stain (Life Technologies), and G1, S, and G2/M populations (∼1 × 106 cell each) were sorted based on DNA content on a FACSAria. Aliquots (105 cells) of the purified populations were reanalyzed by FACS to assess their purity. The cells were lysed (62, 64), and cytoplasmic or chromatin extracts were analyzed by immunoblotting. Detailed experimental procedures are described in SI Materials and Methods.
SI Materials and Methods
Plasmid Expression Vectors and Synthetic Genes.
pCG plasmids expressing epitope-tagged Vpr proteins UNG2, DDB1, and DCAF1 were previously described (34). The human HLTF protein coding sequence was amplified by PCR from an EST clone (MHS6278-202759242; Open Biosystems) and subcloned into pCG and retroviral MSCV(puro) vector encoding N-terminal myc-, HA-, or FLAG-epitope tags. Vpr and HLTF deletion and point mutants were constructed by using QuikChange II mutagenesis kit, and mutations were confirmed by DNA sequencing. HIV-1 group M, N, O, SIVcpzPtt, SIVcpzPts, and HIV-2 group A1, A2, and B consensus vpr genes were designed based on sequence information available at Los Alamos National Laboratory databases (www.hiv.lanl.gov). Codon-optimized genes were synthesized by Genscript and Life Sciences.
Lentiviral Expression Vectors and Viruses.
HIV-1 NL4-3.GFP.R+ and NL4-3.GFP.R− proviral clones were constructed by replacing the luciferase gene with GFP gene in pNL4-3.Luc.R+ and R− constructs (65). HIV-1taqRFP (vif-, vpr-, nef-, env-) proviral construct was provided by Nicolas Manel, Institut Curie, Paris. The psi packaging signal in HIV-1tagRFP was inactivated by deleting the sequence “TACGCCAAAAATTTTGACTAGCGGAGGCTAGAAGGAGAG” [HIV-1(psi−)]. HIV-1 VLP was produced from HEK293T cells transiently cotransfected with HIV-1(psi–), VSV-G expression vector, and/or pCG plasmid expressing HA-tagged NL4-3 Vpr. HIV-2.RFP (env-, Rod9) proviral clone with tagRFP reporter residing in nef locus was provided by Michael Emerman, Fred Hutchinson Cancer Research Center, Seattle, and Nicolas Manel. HIV-2 vpr gene was inactivated by M1E,E3* mutation and vpx by M1T substitution, using standard molecular biology techniques. SIVmac251 VLP was produced from pSIV3+ construct possessing intact vpr and vpx genes (66). Virus particles were produced and concentrated as described previously (6, 63). Virus preparations were normalized by immunoblotting for p24/p27 CA and/or Vpr, followed by quantification of fluorescent signals relative to a recombinant HIV-1 CA standard, using Odyssey imaging. Retroviral MSCV-based expression vectors were produced from HEK 293T cells (34, 64). For inducible expression, cDNAs encoding various epitope-tagged Vpr proteins were subcloned into pLVX-TRE3G vector (Lenti-X Tet-On 3G Inducible Expression System; Clontech), and viruses were produced as described previously (34).
Isolation of G1, S, and G2M Cells by Sorting.
Cells (8 × 106) were stained with 30 μM Vybrant DyeCycle Green stain (Life Technologies), and G1, S, and G2/M populations (∼1 × 106 cells each) were sorted based on DNA content on a FACSAria. Aliquots (105 cells) of the purified populations were reanalyzed by FACS to assess their purity. The cells were lysed (62, 64), and cytoplasmic or chromatin extracts were analyzed by immunoblotting.
List of Antibodies Used for Immunoblotting.
α-HLTF (A300-230A), α-AKAP8 (A301-062A), α-ESCO2 (A301-689A), α-PWP1 (A303-635A), α-RMBX2 (A302-222A), α-RuvBL2 (A302-536A-T), α-PRP19 (A300-101A-T), α-NONO (A300-587A-T), α-TRIP13 (A300-607A-T), and α-VCP (A300-588A-T) were from Bethyl Laboratories; α-TIF1β (4124), α-HP1α (2616), α-KU80 (2180), α-RAD18 (9040), α-MSH6 (5424), and PCNA (2586S) were from Cell Signaling; α-MUS81 (sc-53382, sc-376661), α-TFIID (sc-204), α-DCAF1 (sc-376850), α-Cyclin A2 (sc-751), α-SHPRH (sc-69324), and α-tubulin (sc-5286) were from Santa Cruz Biotechnology; α-tubulin (T5168) and α-Flag (M2) were from Sigma-Aldrich; α-DDB1 (37-6200) was from Invitrogen; α-ZC3HAV1 (PA5-20986) was from Thermo Scientific; α-Lamin B1 (ab16048) was from Abcam; and α-UNG2 (TA503563) was from OriGene. Concentrated α-HA [12CA5 (67)], α-Myc [9E10 (68)], and α-HIV-1 CA [183-H12-5C (69)] were produced from hybridomas by using CELLine Bioreactor flasks (Argos). The HIV-1 p24 Hybridoma 183-H12-5C was obtained through the NIH AIDS Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, NIH, from Bruce Chesebro.
Immunoaffinity Purifications of Protein Complexes and MudPIT Analyses.
CEM.SS-iH1.Vpr cells expressing a tandem HA-FLAG-epitope–tagged HIV-1 NL4-3 Vpr (hfa-H.Vpr) under control of the Lenti-X Tet-On 3G Inducible Expression System (Clontech) and control CEM.SS cells (3 × 105 cells per milliliter; 4 L) were induced with doxycycline (0.1 μg/mL) for 9 h. Whole-cell extracts were prepared, and tandem affinity purifications of Vpr and its associated proteins, from ∼5 g of wet cell mass, were performed as described previously (6). MudPIT analyses of purified protein complexes and calculation of distributed normalized spectral abundance factors were previously described (70).
RNAi.
U2OS cells were plated in 12-well plates (2 × 104 cells per well; Becton Dickinson) in antibiotic-free DMEM supplemented with 10% (vol/vol) FBS. One day later, cells were transfected with siRNA to a 10–25 nM final concentration in 1 mL per well, using Dharmafect (Invitrogen). siRNAs targeting MUS81 (AUGGUCACCACUUCUUAAC, CAGCCCUGGUGGAUCGAUA) (26, 71), HLTF (L-006448-00-005), and control nontargeting siRNA pools “NT1” (D-001206-14-05) and “NT2” (D-001810-10), were purchased from Dharmacon.
Acknowledgments
We thank Jinwoo Ahn, Michael Emerman, Nicolas Manel, and Karlene Cimprich for sharing reagents; Chuanping Wang for technical assistance; Karlene Cimprich for stimulating discussions; Teresa Brosenitsch for critical reading of the manuscript and editorial help; Angela Gronenborn for support; and Tomek Swigut for his help with statistical analyses. CEM.SS cells were obtained through the NIH AIDS Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, NIH, from Dr. Peter L. Nara. This work was supported by NIH Grants AI077459 and AI100673 and a P50GM082251 subcontract (to J.S.). The Flow Cytometry Facility at Case Western Reserve University is supported by Center for AIDS Research Grant P30 AI036219.
Footnotes
The author declares no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The complete MudPIT mass spectrometry dataset (raw files, peak files, search files, as well as DTASelect result files) can be obtained from the MassIVE database via ftp://MSV000079605@massive.ucsd.edu with password KHJS60144.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1605023113/-/DCSupplemental.
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