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Hina Z. Ghory,1 and
Alan Engelman1*
Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston Massachusetts1
Received 12 July 2004/ Accepted 12 October 2004
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Partial proteolysis (24), deletion mutagenesis (9, 70), and functional complementation (23, 69) analyses revealed that HIV-1 IN is comprised of three domains, the N-terminal domain, catalytic core domain (CCD), and C-terminal domain (CTD) (reviewed in reference 29). The CCD (residues 50 to 212 of the 288-residue protein) harbors a triad of invariant carboxylate amino acids (HIV-1 residues Asp-64, Asp-116, and Glu-152, which form the D,D-35-E motif) that comprises the enzyme active site (7, 20, 21, 24, 46, 49, 67). Results of IN-DNA photo-cross-linking (28, 40, 43), footprinting (19), and in vitro enzyme assays (4, 26, 33, 38, 43, 67) revealed other conserved CCD residues that likely contact viral (28, 33, 40, 41, 43) and target (4, 19, 38, 40, 67) DNA during integration. Specifically, Lys-159, Tyr-143, and Gln-148 cross-linked to viral DNA substrates (28, 43) and IN enzymes with alterations at Lys-159 or Gln-148 were defective in in vitro integration assays (33, 43, 67, 68). Peptides spanning residues 49 to 69 (40) and 51 to 64 (28) cross-linked to viral DNA and IN proteins altered at Gln-62 were sensitive to the concentration of salt in in vitro assays (26, 33), suggesting that Gln-62 also binds HIV-1 DNA (13, 28, 33). Since IN enzymes carrying substitutions of Ser-119 (38) or Asn-120 (67) displayed altered patterns of DNA strand transfer, these residues likely contact target DNA during integration. Since Lys-159, Gln-148, and Tyr-143 interacted with 1-(5-chloroindol-3-yl)-3-hydroxy-(2 H-tetrazol-5-yl)-propenone (5CITEP) in an IN-inhibitor cocrystal structure, it was suggested that 5CITEP might inhibit IN activity by interfering with critical IN-DNA contacts that occur during integration (36).
Mixtures of two defective IN mutant proteins can display 3' processing and DNA strand transfer activities, suggesting that IN normally functions as a multimer (23, 69). Since mutants carrying changes in two different active-site residues failed to complement each other, the D,D-35-E active-site residues belong to the same complementation group (69). In other words, integration requires the same protomer within the active multimer to donate all three active-site residues. The organization of the active site to viral and target DNA-binding functions within the CCD, however, has not been probed by complementation. Although three-dimensional structures of IN-DNA complexes have not been solved, models based on protein fragment structures (14, 71), photo-cross-linking (31, 41), and/or computer simulations (2, 18, 32, 58, 59, 76) have been proposed. Most models posit that the CCD that donates the active site also binds the viral end and the target DNA to which that end will integrate; however, other arrangements are plausible (32, 41). To probe the relationships between these CCD functions, mutants altered for catalytic and presumed viral and target DNA binding activities were tested in complementation assays. This was done in the context of HIV-1 infection by fusing one mutant partner to the virion accessory protein Vpr and testing that fusion in trans to defective mutant virus (30, 73). A prerequisite for this strategy was the identification of viral mutants that supported relatively low levels of inherent infectivity. Thus, we began by characterizing a relatively large panel of CCD mutants in assays for HIV-1 spread.
Numerous investigators have analyzed HIV-1 mutants altered at IN active-site residues (3, 10, 25, 27, 39, 47, 48, 54, 55, 57, 60, 62, 63, 65, 72, 74, 75). The resulting class I replication-defective mutants were specifically blocked at integration as evidenced by (i) wild-type (WT) levels of cDNA synthesis in acutely infected cells and (ii) two long terminal repeat (2-LTR)-containing DNA circles that were elevated compared to WT levels, likely due to transient increases in unintegrated DNA pools (3, 25, 39, 48, 54, 57, 60, 62, 72, 75; see reference 22 for a review). In contrast, CCD mutants with changes in potential DNA binding residues have not been systematically analyzed, and unlike active site mutants, wherein any of a number of different missense mutations rendered HIV-1 inactive, the identity of the amino acid substitution had a relatively large impact on the virus replication phenotype. For example, whereas HIV-1K159Q displayed WT replication kinetics (63) and HIV-1E157A/K159A transduced cells 46% as efficiently as the WT (72), HIV-1K159P replicated with an approximate 2-week delay compared to the WT (10), and HIV-1K159E and HIV-1K156E/K159E failed to replicate over a 20-day observation period (43). As another example, HIV-1N120L and HIV-1N120E transduced cells at only about 0.06% of the WT level under conditions where HIV-1N120G supported about 3.3% of the WT transduction activity (48). Cells infected with HIV-1K159E, HIV-1K156E/K159E (43), or HIV-1N120L (48) contained elevated levels of 2-LTR circles, which defined these mutants as class I (reviewed in reference 22). In contrast to the integration-specific class I phenotype, class II IN mutant viruses display particle assembly and/or reverse transcription defects (22).
In this study, 22 CCD residues were targeted by site-directed mutagenesis, yielding 35 mutant viruses. In addition to their potential role in DNA binding, residues were targeted due to 5CITEP binding (36), degree of sequence conservation and surface exposure in crystal structures, and/or previous determination as essential for integration. Our results categorize conserved CCD residues as either dispensable or important for HIV-1 replication and classify defective viruses as either class I or class II. Of note, defective viruses altered at putative DNA contacts were class I mutants, highlighting the integration-specific roles of these residues in HIV-1 replication. Results of Vpr-IN complementation assays indicated that the active CCD protomer within the IN multimer interacts with both viral and target DNA during integration. Revertant mutant viruses were screened for the presence of second-site mutations, and E138K was characterized as an effective suppressor of Gln-62 mutant viruses and IN. Our results help to define the residues and regions of the IN CCD that engage viral and target DNA during integration.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The envelop (Env)-deleted single-round vector pNLX.Luc(R–) was derived from pNLX
envCAT (52) as follows. Plasmid pNLX.Luc was generated by amplifying the gene for firefly luciferase (Luc) from pGL3-Basic (Promega Corp., Madison, Wis.) with ClaI- and XhoI-tagged primers, digested, and ligated to ClaI, XhoI-digested pNLXenvCAT. To make pNLX(Vpr–), pNL43/XmaI was digested with EcoRI, filled in with Klenow, and religated. Plasmid pNLX.Luc(R–) was built by swapping the 1.6-kb PflMI-NheI fragment from pNLX(Vpr–) for the corresponding fragment in pNLX.Luc. IN mutant derivatives of pNLX.Luc(R–) were built by swapping altered 1.8-kb AgeI-PflMI fragments from pNL4-3 or pNL43/XmaI-based plasmids for the corresponding fragment in pNLX.Luc(R–). The HIV-1NL4-3 Env expression vector pNLXE7 was previously described (52).
Plasmid pRL2P-Vpr-IN (73) encoding HIV-1YU2-derived Vpr fused to HIV-1SG3 IN was a generous gift from J. Kappes (University of Alabama). Mutations were introduced into pRL2P-Vpr-IN using QuikChange mutagenesis as recommended by the manufacturer (Stratagene, La Jolla, Calif.). DNA sequencing was used to verify the regions of all plasmids that were generated by PCR.
Cells, viruses, and infections. HeLa and 293T cells were grown in Dulbecco's modified Eagle's medium supplemented to contain 10% fetal calf serum (FCS), 100 IU of penicillin per ml, and 0.1 µg of streptomycin sulfate per ml. Jurkat, CEM-12D7, and C8166 T-cells were grown in RPMI 1640 containing 10% FCS, 100 IU of penicillin/ml, and 0.1 µg of streptomycin/ml (RPMI-FCS).
Viruses for spreading-infection and 3' processing assays were derived from HeLa or 293T cells by transfection in the presence of calcium phosphate as previously described (11, 25, 57). The concentration of virus in cell supernatants was determined using a 32P-based assay for reverse transcriptase (RT) activity (25, 57).
Single-round viruses carrying the HIV-1NL4-3 Env for real-time quantitative PCR (RQ-PCR) assays were derived by cotransfecting 293T cells with pNLX.Luc(R–) and pNLXE7 at the ratio of 30:1 using FuGENE 6 (Roche Molecular Biochemicals, Indianapolis, Ind.). Resulting cell supernatants were treated with 40 U of TURBO DNase (Ambion, Austin, Tex.)/ml for 1 h at 37°C to degrade the bulk of plasmid DNA remaining after transfection. Single-round viruses for Vpr-IN complementation studies were assembled by cotransfecting 293T cells with pNLX.Luc(R–), pRL2P-Vpr-IN, and pNLXE7 at the ratio of 30:15:1, using calcium phosphate. Background infectivity measurements of IN mutant viruses in the absence of added Vpr-IN were made by substituting pcDNA3 (Invitrogen, Carlsbad, Calif.) for pRL2P-Vpr-IN during transfection. All single-round viral stocks were normalized for RT content prior to infection. To analyze Vpr-IN incorporation, 293T cells were transfected with pNLX(Vpr–) and pRL2P-Vpr-IN in a 2:1 ratio by using FuGENE6.
For assays of viral spread, Jurkat or CEM-12D7 cells (2 x 106) were infected with WT or IN mutant HIV-1NL4-3 (106 RT cpm) for 18 h at 37°C in 0.5 ml of medium unless otherwise noted. The following day, cells were washed with serum-free RPMI 1640 and plated in 5 ml of RPMI-FCS. Cells were split at regular intervals, at which time aliquots of cell supernatants were stored at –20°C for RT assays. CEM-12D7 cells (5 x 106) were infected with 106 RT cpm of WT or IN mutant HIV-1NL4-3 for 2-LTR circle PCR analyses as previously described (6).
For RQ-PCR assays, Jurkat cells (4 x 106 per well of a six-well plate) were infected with DNase-treated WT or IN mutant single-round HIV-1NLX.Luc(R-) (5 x 106 RT cpm) by spinoculation for 2 h as previously described (52). Spun cells were washed three times with serum-free RPMI 1640 before plating in 4 ml of RPMI-FCS. For Vpr-IN complementation assays, Jurkat cells (2 x 106) were infected with single-round viruses (5 x 105 RT cpm in 0.5 ml) for 18 h at 37°C, at which time culture volumes were expanded by the addition of 5 ml of RPMI-FCS.
Analysis of HIV-1 proteins. Transfected HeLa cells were radiolabeled with 35S-Met and 35S-Cys as previously described (26). Labeled cells and viruses were lysed and processed for immunoprecipitation, and the resulting immunoprecipitates were analyzed by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) and fluorography as previously described (25, 26).
For Western blot analysis of viral particles, 293T cell supernatants were pelleted through 20% sucrose cushions for 120 min at 4°C and 27,000 rpm in a Beckman SW28 rotor. Pellets were lysed in 25 µl of radioimmunoprecipitation assay buffer (140 mM NaCl, 8 mM Na2HPO4, 2 mM NaH2PO4, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.05% SDS) and normalized for p24 protein content (Alliance HIV-1 p24 ELISA kit; New England Nucleotides, Boston, Mass.), and 2.5 µg of total p24 was analyzed by Western blotting using polyclonal anti-IN antibodies (15).
Analysis of HIV-1 DNA synthesis. (i) 2-LTR circle PCR. Total DNA was extracted from CEM-12D7 cells 18 h postinfection (hpi), and 10 µl of cell extract was analyzed for levels of HIV-1 2-LTR circles by nested PCR as previously described (6).
(ii) RQ-PCR. Total DNA was extracted from infected Jurkat cells 7 and 24 hpi by using the DNeasy kit (QIAGEN, Valencia, Calif.), and 10 µl was amplified by PCR in a 30-µl volume as described previously (51, 54). The PCR primers and Taqman probe were designed to detect total viral cDNA after the second template switch of reverse transcription, and HIV-1 signals were normalized to total cellular DNA using endogenous retrovirus 3 as described previously (51, 54). Parallel infections with Env– viruses were performed to control for residual levels of plasmid DNA that may have resisted DNase treatment, and the resulting PQ-PCR values were subtracted from those obtained with Env+ viruses.
Analysis of intracellular 3' processing activity. To determine 3' processing levels, WT and IN mutant preintegration complexes were obtained from cytoplasmic extracts of infected C8166 cells as described previously (11-13). Following deproteinization and precipitation with ethanol, HIV-1 cDNA was cleaved with HaeIII and HindIII and separated by denaturing PAGE. Following electrotransfer, the blot was probed by indirect end labeling as described previously (11, 12). Results were detected by autoradiography and quantified by densitometry (IS-1000 Digital Imaging System; Alpha Innotech Corp., San Leandro, Calif.).
Cloning and sequencing of revertant viruses. Supernatants were prepared from Jurkat cells infected with WT and revertant viruses by Hirt fractionation as previously described (57). Viral sequences amplified from Hirt supernatants by PCR were molecularly cloned as described previously (57).
Expression and purification of recombinant IN protein and in vitro integration assays. INWT and INQ62A proteins were purified from insoluble fractions of bacterial lysates as previously described (26). INQ62A/E138K and INE138K were similarly purified and refolded into CN buffer {20 mM HEPES (pH 7.6), 1 mM EDTA, 0.2 M NaCl, 10% (wt/vol) glycerol, 1 mM dithiothreitol, 15 mM [(3-cholamidopropyl)-dimethylammonio]-l-propanesulfonate [CHAPS]}. The N-terminal hexahistidine tag used for protein purification was cleaved from IN using thrombin as previously described (26).
The 5' end of oligonucleotide AE144 (5'-TTTTAGTCAGTGTGGAAAATCTCTAGCAGT) was radiolabled with T4 polynucleotide kinase as described previously (17). After heat inactivation of the kinase, the complementary single strand AE143 (5'-ACTGCTAGAGATTTTCCACACTGACTAAAA) was annealed and the 30-bp DNA substrate was separated from unincorporated nuclide as described previously (17).
In vitro integration reaction mixtures (16 µl) contained 25 mM morpholinepropanesulfonic acid (pH 7.2), 0.1 mg of bovine serum albumin/ml, 10 mM ß-mercaptoethanol, 10% glycerol, 25 nM substrate DNA, 0.4 mM CHAPS, 7 mM NaCl, 7.5 mM MnCl2, and 0.3 µM IN. The concentration of NaCl in the reactions was varied as appropriate. Reactions were terminated after 60 min at 37°C and analyzed by electrophoresis on denaturing polyacrylamide sequencing gels as previously described (17). Results were visualized by autoradiography and quantitated using densitometry.
Vpr-IN complementation. Cells harvested 48 hpi were washed with phosphate-buffered saline (Mediatech, Hamilton, Va.) and lysed with 75 µl of passive lysis buffer (Promega Corp., Madison, Wis.). Lysates frozen on dry ice and thawed at 37°C were centrifuged at 18,730 x g for 15 min at 4°C. Supernatants (20 µl) were tested in duplicate for Luc activity using the Luciferase Assay system (Promega Corp.) and an EG&G Berthold Microplate LB 96-V luminometer and Microlite 1 flat-bottom microtiter plate (Thermo Labsystems, Franklin, Mass.). Luc activity was normalized to the total cellular protein concentration as determined by the Bio-Rad protein assay kit (Bio-Rad Laboratories, Hercules, Calif.). Background levels of activity derived from mock infections with Env– HIV-1NLX.Luc(R–) were subtracted from WT and IN mutant values.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Replication profiles of IN mutant viruses. Jurkat and CEM-12D7 T-cells infected with 106 RT cpm (an approximate multiplicity of infection [MOI] of 0.04; see references 52 and 57) supported peak levels of HIV-1NL4-3 replication 4 to 6 days postinfection (dpi) (Fig. 1). Since substituting Ala for Asp-55 rendered HIV-1R7-3 inactive (72), we were a bit surprised to find that NL4-3-based HIV-1D55A replicated similarly to the WT (Fig. 1A). Because of this, a similar substituent, Ser, as well as nonconservative Lys, were tested at this position. Whereas HIV-1D55S grew like the WT, HIV-1D55K was defective (Fig. 1A).
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(ii) Mutant virus assembly. In addition to the common reverse transcription defect, certain class II IN mutants are defective for the relatively late steps of particle assembly and release (8; reviewed in reference 22). To address whether any of the DNA synthesis-defective mutants displayed late event defects, virus-producing cells were radiolabeled with [35S]Met and [35S]Cys, cell and viral lysates were immunoprecipitated with AIDS patients' antisera, and HIV-1 proteins were analyzed after SDS-PAGE and fluorography. Assembly-release mutants were identified as viruses containing levels of HIV-1 proteins that were significantly lower than their intracellular levels of gene expression.
As previously reported (26, 42), the HIV-1F185K class II mutant was released poorly from cells (Fig. 3B, compare lane 5 to lane 1). As predicted from previous analyses of IN deletion mutants (3, 8, 25), the CTD truncation mutant HIV-11-212 also released poorly in this assay (Fig. 3A, lane 6 and Fig. 3B, lane 2). Despite these disproportionately low levels of HIV-1F185K and HIV-11-212 virion proteins, the mutants were expressed in cells as efficiently as WT HIV-1NL4-3 (Fig. 3A, upper panel, lanes 1 and 6; upper panel of Fig. 3B, lanes 1, 2, and 5). Whereas HIV-1E152K (Fig. 3A lower panel, lane 5) and HIV-1N184L (Fig. 3B, lane 3) also displayed release defects, HIV-1D55K, HIV-1R199A, HIV-1R199E, and HIV-1N184D were released from cells similarly to the WT (Fig. 3). We note that assembly-defective class II mutants expressed an increase in the ratio of the Gag processing intermediate capsid-spacer peptide 1 (p25) to mature capsid (p24) (Fig. 3A and B, upper panels), a phenotype associated with a variety of maturation-defective HIV-1 mutants (reference 50 and references therein).
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As predicted from their poor performances in spreading-infection assays, HIV-1Q62K.Luc(R–), HIV-1K156E/K159E.Luc(R–), and HIV-1N120K.Luc(R–) infectivities were near the detection limit of the Luc assay, similar to the case with active-site mutant HIV-1D64N/D116N.Luc(R–) [approximately 0.05 to 0.3% of that for WT HIV-1NLX.Luc(R–)] (Table 3). Vpr-INWT restored 10 to 30% of HIV-1NLX.Luc(R–) function to HIV-1Q62K.Luc(R–), HIV-1K156E/K159E.Luc(R–), HIV-1N120K.Luc(R–), and HIV-1D64N/D116N.Luc(R–) (Table 3). Mutant Vpr-IN activity was reported as percent Vpr-INWT function. As expected (5, 53), Vpr-INV165A efficiently complemented active-site mutant HIV-1D64N/D116N.Luc(R–), and HIV-1Q62K.Luc(R–) was also efficiently complemented by Vpr-INV165A (Table 3). Although Vpr-INV165A also complemented HIV-1K156E/K159E.Luc(R–) and HIV-1N120K.Luc(R–), these levels were approximately 5 to 10% of the levels observed with HIV-1D64N/D116N.Luc(R–) and HIV-1Q62K.Luc(R–) (Table 3). In contrast to the relatively robust activity of Vpr-INV165A, Vpr-INQ62K did not complement HIV-1D64N/D116N.Luc(R–), HIV-1Q62K.Luc(R–), HIV-1K156E/K159E.Luc(R–), or HIV-1N120K.Luc(R–) (Table 3). Vpr-INN120K behaved similarly to Vpr-INQ62K: although Vpr-INN120K weakly complemented HIV-1Q62K.Luc(R–) (approximately 3% of Vpr-INWT), we noted that the reciprocal arrangement yielded negligible activity (Table 3). Based on this, we concluded that Q62K, D64N/D116N, K156E/K159E, and N120K belong to the same functional complementation group. We therefore infer that D,D-35-E active site, viral DNA binding, and target DNA binding functions of the CCD are supplied by the same protomer within the active IN multimer during integration.
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Our results demonstrate that CCD class II mutant Vpr-INs effectively complemented CCD class I mutant viruses under conditions where analogous class I mutant proteins failed to function (Table 3). Our ability to assign various CCD functions to the same IN protomer during postentry integration therefore relied on efficient incorporation of class I and class II Vpr-IN proteins during virus assembly. Although there was little reason to expect differential packaging of different mutant classes, virus particles generated by cotransfecting Vpr-IN expression vectors with HIV-1NLX(Vpr–) were pelleted through sucrose and normalized for p24 content, and Western blots were probed with anti-IN antibodies. Whereas the only reactive HIV-1NLX(Vpr–) band comigrated with recombinant IN protein (Fig. 5, compare lane 2 to lane 1), trans-complemented particles contained an additional 
45-kDa product indicative of uncleaved Vpr-IN (lanes 3 to 8) (30, 61, 73). Although class II mutants Vpr-IND55K and Vpr-INR199E were incorporated somewhat more efficiently than class I Vpr-INQ62K and Vpr-INN120K, the level of class I mutant incorporation was indistinguishable from that of Vpr-INV165A and Vpr-INWT (Fig. 5). Based on this, we conclude that CCD class I mutant Vpr-INs do not function with CCD class I mutant viruses due to the lack of postviral entry phenotypic complementation.
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Although some class I mutants, such as HIV-1H67E and HIV-1N120K, showed growth that was significantly delayed compared to that of the WT, their titers eventually approached those of the WT (Fig. 1C and E). We considered two possibilities for these observations. First, the replication delay was a direct consequence of the activity of the mutant IN protein. In other words, replication occurred in the face of the original mutation, and the delay was due to the time required to amass sufficient infection events to drive replication throughout the culture. In the second case, the original mutation, although sufficient to support an initial low level of integration, was insufficient to drive virus spread, and thus, other mutations were required to revert the IN to a more active form. In this case the delay was due to the time required for the initial infection to generate the required mutation(s) through subsequent rounds of reverse transcription and integration. To distinguish between these two scenarios, viruses derived from the peaks of HIV-1H67E (Fig. 1C) and HIV-1K159E (Fig. 1I) replication were passed onto fresh cells. If the first model was true, the passaged viruses would grow with delays similar to those seen in the original infections. If on the other hand additional mutations had occurred, the passaged viruses would replicate at increased rates. Since HIV-1Q62A also grew with an approximate 2-week delay compared to the WT (26), it was included in this analysis. Using a similar selection scheme, we previously determined that a second CCD substitution (T125A) effectively suppressed the deleterious effects of P109S (64).
Jurkat cells infected with passaged HIV-1H67E and HIV-1K159E revealed cytopathic evidence of viral spread 9 dpi (data not shown). Since HIV-1H67E and HIV-1K159E reached their original growth peaks 21 and 16 dpi, respectively (Fig. 1C and I), we reasoned that the passaged viruses had accumulated new changes. To investigate this, infected cells were lysed by Hirt extraction, and HIV-1 DNA in Hirt supernatants was sequenced after PCR amplification. Whereas HIV-1NL4-3 uses an AAG codon for Lys-159, GAA was engineered in HIV-1K159E. The revertant virus harbored AAA at this position, indicating back reversion of the mutation to restore WT HIV-1NL4-3. This was not unexpected for a substitution such as Lys
Glu wherein a single nucleotide change would restore INWT function. HIV-1H67E contained GAG in place of CAT, and the revertant virus altered this to AAG, indicating that a Lys at position 67 was sufficient for near-INWT function. Due to what were considered relatively uninformative back reversions of K159E and H67E changes, these revertant viruses were not studied further.
Similar to the results with HIV-1H67E and HIV-1K159E, passaged HIV-1Q62A(r) (r for revertant) spread significantly faster than the starting mutant strain (Fig. 6A). Hirt supernatant DNA prepared from HIV-1Q62A(r)-infected cells at day 14 postinfection was PCR amplified, and IN coding regions were molecularly cloned prior to sequencing. In one of four cases, the mutant GCG codon underwent two reversional substitutions back to the WT CAG. However, in the other three cases, the mutant GCG codon persisted and a G-to-A transition occurred at nucleotide 4641 of HIV-1NL4-3. This altered the GAA encoding IN residue Glu-138 to the same E138K change that we had included in our starting mutant pool (Table 1 and Fig. 1E). The entire IN coding regions of two of the HIV-1Q62A(r) molecular clones were sequenced, which revealed the G-to-A change as the only new alteration in each case.
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The phenotype of E138K-containing virus was next analyzed. Since HIV-1Q62K was completely defective (Fig. 1B), E138K was tested in the stringent background of HIV-1Q62K rather than the HIV-1Q62A strain from which it originated. HIV-1Q62K/E138K was observed to replicate, although with an approximately 4-day delay from WT replication in repeat experiments (Fig. 6C; also data not shown). Thus, E138K is a bona fide second site suppressor mutation that can revert the deleterious effects of different changes (Ala or Lys) at Gln-62.
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DISCUSSION |
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Our results help in understanding conserved residues and structural features of the CCD Phe139-Gln146 flexible loop and abutting Ser147-Val165 amphipathic alpha-helix, which are likely important for DNA binding during HIV-1 replication. In terms of the alpha-helix, previous in vitro work indicated that the polar face of the amphipathic structure interacted with viral DNA (76), and our work with mutant viruses supports these findings. For example, Gln-148, Glu-152, Asn-155, and Lys-159 reside on the same side of the helix (Fig. 7, upward face of helix 4 as drawn).
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The Phe139-Gln146 flexible loop cross-linked to DNA substrates (40), and Tyr-143 was subsequently shown to cross-link to the adenosine at the 5' end of the nontransferred U5 minus-strand (28). The other loop residues targeted here, Asn-144, Pro-145, and Gln-146, were similar to Tyr-143 in that they were well conserved among HIV-1 strains and divergent retroviruses (Table 1). However, out of these four highly conserved residues, only Asn-144 played a significant role in HIV-1 replication (Fig. 1D and F). Since HIV-1N144K synthesized more 2-LTR circles than the WT (Fig. 2B), our results indicate that Asn-144 plays a specific role in integration. However, it is unclear if this includes DNA binding. From the structure-based pattern of required residues, one can deduce an extended pocket that encompasses the approximate lower half of the molecule, as drawn in Fig. 7, to which viral DNA might bind. Asn-144 seems to jut away from this pocket. However, it should be noted that due to its flexibility, the loop is generally not observed in CCD structures, and the precise locations of loop side chains differ dramatically among the few structures where they are observed (7, 35, 37). Despite these limitations, Gln-146 may be dispensable for HIV-1 replication because it points down toward the underside of helix 4 (Fig. 7).
Although results of an in vitro footprinting assay indicated that DNA strongly protected Glu-138 from proteolysis (19), HIV-1E138K grew similarly to the WT, indicating that Glu-138 does not play an important role in HIV-1 replication under these conditions (Fig. 1E). Interestingly, E138K was identified as an intramolecular mutation that suppressed the adverse affects of Q62A and Q62K changes (Fig. 6). In the CCD structure, Glu-138 is positioned toward oppositely charged Lys-136, noticeably away from Gln-62 (Fig. 7). Because of this, we speculate that substituting Lys for Glu-138 would redirect the mutant side chain away from Lys-136, which might orient it toward Gln-62 and in doing so directly compensate for a loss of DNA binding incurred through Gln-62 substitutions.
The roles of other conserved CCD residues in HIV-1 replication. Five residues, Gln-53, Asp-55, Lys-71, Asn-184, and Arg-199, that were neither implicated in DNA binding nor part of the Phe139-Gln146 flexible loop, were also targeted in this study. Although Gln-53 was invariant among HIV-1 strains (Table 1), HIV-1Q53K grew as did the WT (Fig. 1B), and thus, our assay system did not reveal a role for Gln-53 in HIV-1 replication. The same can be said for highly conserved Lys-71 (Table 1 and Fig. 1L). In contrast, HIV-1D55K, HIV-1N184D, HIV-1N184L, and HIV-1R199E were replication defective (Fig. 1A, K, and L). Since each of these mutants synthesized significantly less cDNA than the WT (Fig. 2 and data not shown) and HIV-1N184L was released less efficiently than the WT from HeLa cells (Fig. 3B), these viruses were typed as pleiotropic class II mutants. The results of Asp-55 mutagenesis are reminiscent of our previous findings with Phe-185 (26). In that paper we determined that HIV-1F185H replicated as did the WT and that HIV-1F185K, HIV-1F185A, and HIV-1F185L were replication-defective class II mutants (26; see reference 22 for a review). Somewhat surprisingly, HIV-1E152K was also a class II mutant (Fig. 2A and 3A; Table 2). Since HIV-1E152D and HIV-1E152Q were class I viruses (Table 1), these results demonstrate that different substitutions of the same residue can yield either the class I or class II mutant phenotype.
Functional organization of the HIV-1 IN CCD. Vpr-IN complementation was used to probe the functional relationship between catalysis and viral and target DNA binding that occurs during integration. For this we needed viral mutants for each of these functions that supported relatively low levels of inherent infectivity. By characterizing a large collection of CCD mutants in assays for viral spread, HIV-1Q62K, HIV-1K156E/K159E, and HIV-1N120K were determined to meet these requirements. Since Vpr-INQ62K complemented HIV-1V165A.Luc(R–) (53) but failed to complement either HIV-1K156E/K159E.Luc(R–) or HIV-1D64N/D116N.Luc(R–), our results indicate that different DNA-binding residues, such as Gln-62 and Lys-159, originate from the same functional protomer and that the active CCD binds to the viral DNA end that it functionally integrates. This result enforces previous IN-DNA models, which invariably predicted that the active CCD also bound to the viral DNA that it processed and joined to target DNA (2, 14, 18, 31, 32, 41, 58, 59, 71). Despite this "cis" requirement for active-site and CCD viral DNA-binding functions to reside within the same protomer, we note that the CTD's viral DNA end binding function is donated by a separate IN molecule (14, 31).
HIV-1N120K was a class I mutant (Fig. 2A, lane 4) that processed significantly more of its 3' ends than other class I mutants, including active-site mutant HIV-1D116N and viral DNA-binding mutants HIV-1Q62K and HIV-1K156E/K159E (Fig. 4). Based on these results and previous observations that recombinant IN proteins altered at Asn-120 or the adjacent Ser-119 displayed altered patterns of DNA strand transfer (38, 67), we inferred that HIV-1N120K was in part defective for target DNA binding in vivo. Since Vpr-INQ62K did not complement HIV-1N120K.Luc(R-) and Vpr-INN120K failed to appreciably complement HIV-1Q62K.Luc(R-), HIV-1K156E/K159E.Luc(R–),or HIV-1D64N/D116N.Luc(R–), our results indicate that the active protomer within the IN multimer that binds and processes viral DNA also binds the chromosomal DNA into which it integrates. Although we inferred from in vitro studies that Gln-62, Asn-120, and Lys-159 directly contact DNA, we have not extensively characterized biochemical properties of recombinant proteins and because of this cannot rule out that disruption of other IN functions (for example, multimerization) contributed to the class I mutant phenotypes described here.
In conclusion, our results prioritized the importance of a number of conserved CCD residues that were previously implicated in DNA binding for their roles in HIV-1 replication. Together with the results of Vpr-IN complementation, our results were used to refine the regions of the active CCD protomer that interact with viral and target DNA during integration (Fig. 7). Precise structural delineation of the active IN-DNA complex is expected to aid the design of inhibitors that block HIV-1 replication before the virus has the chance to integrate into chromosomal DNA.
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ACKNOWLEDGMENTS |
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This work was supported by National Institutes of Health grant AI39394. Core facilities were supported by a Center for AIDS Research grant (AI28691) and the Dana-Farber Cancer Institute/Harvard Cancer Center.
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FOOTNOTES |
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Present address: Department of Medical Oncology, Dana-Farber Cancer Institute, Boston, MA 02115.
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REFERENCES |
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helices within the core domain of retroviral integrase in selecting target DNA sites for integration. J. Biol. Chem. 276:45848-45855.
) transiently transfected HeLa cells (