Wild-Type Levels of Nuclear Localization and Human Immunodeficiency Virus Type 1 Replication in the Absence of the Central DNA Flap (original) (raw)

J Virol. 2002 Dec; 76(23): 12078–12086.

Ana Limón

Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115

Noriko Nakajima

Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115

Richard Lu

Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115

Hina Z. Ghory

Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115

Alan Engelman

Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02115

*Corresponding author. Mailing address: Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, 44 Binney St., Boston, MA 02115. Phone: (617) 632-4361. Fax: (617) 632-3113. E-mail: ude.dravrah.icfd@namlegne_nala.

†Present address: Department of Pathology, National Institute of Infectious Diseases, Shinjuku-ku, Tokyo 162-8640, Japan.

Received 2002 Feb 19; Accepted 2002 Aug 27.

Abstract

Numerous factors have been implicated in the nuclear localization of retroviral preintegration complexes. Whereas sequences in human immunodeficiency virus type 1 (HIV-1) matrix, Vpr, and integrase proteins were initially reported to function specifically in nondividing cells, other recently identified sequences apparently function in dividing cells as well. One of these, the central DNA flap formed during reverse transcription, is specific to lentiviruses. It was previously reported that flap-negative (F−) HIV-1LAI was completely defective for viral spread in the MT-4 T-cell line, yet F− HIV-1 vectors were only 2- to 10-fold defective in various single-round transduction assays. To address these different findings, we analyzed the infectivity and nuclear localization phenotypes of two highly related T-cell-tropic strains, HIV-1NL4-3 and a derivative of HIV-1HXBc2 deficient for both Vpr and Nef. In stark contrast to the previous report, F− derivatives of both strains replicated efficiently in MT-4 cells. F− HIV-1NL4-3 also spread like wild-type HIV-1NL4-3 in infected Jurkat and primary T-cell cultures. In contrast, F− HIV-1HXBc2 was replication defective in primary T cells. Results of real-time quantitative PCR assays, however, indicated that F− HIV-1HXBc2 entered primary T-cell nuclei as efficiently as its wild-type counterpart. Thus, the F− HIV-1HXBc2 growth defect did not appear to correlate with defective nuclear import. Consistent with this observation, wild-type nef restored replication to F− HIV-1HXBc2 in primary T cells. Our results indicate that the central DNA flap does not play a major role in either preintegration complex nuclear import or HIV-1 replication in a variety of cell types.

Two enzymes, reverse transcriptase (RT) and integrase, carry out essential tasks during the early phase of the retroviral life cycle. Soon after entry, RT copies viral RNA into linear double-stranded cDNA. The integrase then catalyzes the insertion of this cDNA into a cell chromosome. Reverse transcription and integration take place in the context of large nucleoprotein complexes called reverse transcription complexes and preintegration complexes, respectively (reviewed in reference 22).

Retroviral preintegration complexes must enter cell nuclei, where the chromosomal targets of integration reside. The nuclear envelope contains numerous nuclear pore complexes, which regulate the transport of macromolecules into and out of the nucleus. Nuclear pore complexes permit passive diffusion of macromolecules with diameters of less than about 9 nm, equating to proteins in the 40- to 60-kDa range (reviewed in reference 34). The size of human immunodeficiency virus type 1 (HIV-1) and Moloney murine leukemia virus preintegration complexes, estimated to be near or greater than the size of the eukaryotic ribosome (3, 18), precludes their passive transport through nuclear pore complexes (reviewed in references 21 and 22).

Lentiviruses like HIV-1 productively infect nondividing target cells (6, 33, 47). In contrast, Moloney murine leukemia virus requires cell division to establish a productive infection (24, 32, 43). Whereas HIV-1 enters nondividing cell nuclei by an energy-dependent process (6), Moloney murine leukemia virus requires the dissociation of nuclear membranes during mitosis, which in theory leads to preintegration complex-chromosome interactions in the absence of active transport (32, 43). One interpretation of these results is that HIV-1 preintegration complexes contain nuclear localization signals that govern their active transport through intact nuclear pore complexes, and Moloney murine leukemia virus fails to infect nondividing cells due to the lack of appropriate signals (21, 22).

Recently, replication-defective Moloney murine leukemia virus (49) and HIV-1 (2, 50) mutants were identified as being blocked at the nuclear import step in dividing target cells. This suggested that retroviral preintegration complexes might interact with specific host cell factors to achieve nuclear localization even in cells that are actively dividing (15). One of the two HIV-1 factors implicated in this novel type of nuclear localization is the _cis_-acting central DNA flap formed during reverse transcription (50). The flap is a 99-base overlap of the plus-strand formed by the central polypurine tract (cPPT) and central termination sequence (8-10, 50). Flap-negative (F−) strain HIV-1LAI was replication defective in peripheral blood mononuclear cells (PBMC) and the MT-4 T-cell line (50). The complete replication block in MT-4 cells suggested that F− HIV-1LAI was at least 104-fold defective in its ability to initiate a spreading viral infection (37, 50).

One model for the flap in HIV-1 replication was that it allowed preintegration complexes to adopt a folded structure required for efficient transport through nuclear pore complexes (50). This model was based on the central location of the flap in the HIV-1 genome, which in two dimensions places it diametrically opposed to the integrase-containing intasome that synapses the two ends of the retroviral cDNA together and mediates integration (11, 12, 46). In contrast to the results with replication-competent HIV-1, F− viral vectors were only 2- to 10-fold defective in single-round infection assays (16, 20, 39, 40, 44, 50, 51). Because of this, it was suggested that the smaller size of vector genomes precluded the absolute requirement for the flap during nuclear import (50).

In this study, we investigated the role of the flap in preintegration complex nuclear import by analyzing the infectivity and nuclear localization of two different HIV-1 strains. Our results show that the flap is not required for efficient HIV-1 spread and preintegration complex nuclear import in a variety of cell types.

MATERIALS AND METHODS

Plasmids.

The cPPT-D mutation (50) was introduced into pNL43/_Xma_I (4) and pHXBH10-vpu+ (23), encoding wild-type HIV-1NL4-3 (1) and a Vpu-positive derivative of HIV-1HXBc2 (19), respectively, by PCR mutagenesis (17, 37). The presence of desired base changes as well as the absence of off-site changes was confirmed by DNA sequencing. To generate a Nef+ F− derivative of HIV-1HXBc2, the 1.8-kb _Pfl_M-I fragment from F− pHXBH10-vpu+ was swapped for the corresponding fragment in pHXB/Nef+ (5).

Plasmid pHXBH10ΔenvCAT-vpu+ (42) encoded envelope (Env)-deleted (Env−) HIV-1HXBc2 carrying the bacterial chloramphenicol acetyltransferase (CAT) gene in the viral nef position. Env− F− HIV-1HXBc2 was made by swapping the 1.8-kb _Pfl_M-I fragment from F− pHXBH10-vpu+ for the corresponding fragment in pHXBH10ΔenvCAT-vpu+. Plasmid pNLXΔenv was made by digesting pNL43/_Xma_I with _Nhe_I and _Stu_I, treating with Klenow fragment, and religating. Plasmid pNLXΔenvCAT was made by exchanging the CAT gene for nef region nucleotides 8785 to 9017 (1, 35). F− pNLXΔenv and F− pNLXΔenvCAT were made by swapping the 1.8-kb _Age_I-_Pfl_M-I fragment from F− pNL43/_Xma_I for the corresponding fragments in pNLXΔenv and pNLXΔenvCAT, respectively.

Env expression vectors pSVIII-env (25) and pHCMV-G (48) encoded HIV-1HXBc2 and vesicular stomatitis virus G (VSV-G) glycoproteins, respectively. An HIV-1NL4-3 Env expression vector was made by swapping the 2.1-kb _Kpn_I-_Bam_HI fragment from pNL43/_Xma_I for the corresponding pSVIII-env fragment. The resulting pNLXE7 plasmid encoded a chimeric Env with amino acid residues 42 to 750, comprising most of gp120 and gp41, derived from HIV-1NL4-3.

Plasmid pUC19.2LTR was built by amplifying Hirt supernatant from HIV-1NL4-3-infected CEM-12D7 cells (37) with _Eco_RI-tagged primer AE761 (5′-ACTGACGAATTCGAGCTTGCTACAAGG-3′) and _Bam_HI-tagged AE762 (5′-CATGCAGGATCCCAGGGTGTAACAAGCTGG-3′), digestion, and ligation to _Eco_RI- and _Bam_HI-digested pUC19.

Cells, viruses, and infections.

Virus stocks for replication assays were prepared by transfecting 293T cells with calcium phosphate as described previously (17, 37). Single-round viruses for real-time quantitative (RQ)-PCR and in vitro CAT assays were prepared by transfecting 293T cells with FuGENE 6 (Roche Molecular Biochemicals, Indianapolis, Ind.) and treating the filtered supernatants with DNase I as described previously (12). Viruses were counted with an exogenous 32P-based RT assay (17, 37).

Activated PBMC were prepared from human blood as described previously (37). PBMC, Jurkat cells, and MT-4 cells (2 × 106) were infected with equal RT counts per minute (cpm) of wild-type and mutant virus for spreading infection assays essentially as described previously (37). For in vitro CAT assays, cells (2 × 106) were infected with 5 × 106 RT cpm of HIV-1 pseudotypes. In vitro CAT activity was determined as previously described (37).

For RQ-PCR assays, cells (6 × 106) were infected with 6 × 106 RT cpm of HIV-1NL4-3 or 3 × 107 to 6 × 107 RT cpm of HIV-1HXB2 pseudotypes by spinoculation (38) at 480 × g for 2 h. For Southern blotting, SupT1 cells (2 × 107) were infected with 10 ml of DNase I-treated HIV-1NL4-3 essentially as described previously (12), and MT-4 cells (2 × 106) were infected with 107 RT cpm of HIV-1HXBc2 by spinoculation. All infection assays were performed a minimum of two times.

Flap detection assay.

HIV-1NL4-3 cDNA was purified from SupT1 cell (2 × 107) cytoplasmic extracts (0.5 ml) prepared at 7 h postinfection essentially as described previously (12). HIV-1HXBc2 cDNA was purified from Hirt supernatant fractions of MT-4 cells (4 × 106) lysed (37) 2 to 3 days postinfection. Purified cDNA was treated with S1 nuclease (1.5 U/μg of total DNA) (Amersham Pharmacia Biotech Inc., Piscataway, N.J.) for 30 min at 37°C (27). The minus strand of HIV-1 was detected by Southern blotting with a nef/U3-specific riboprobe as described previously (4, 12).

RQ-PCR assays.

DNA from infected cells was extracted with the DNeasy tissue kit as recommended by the manufacturer (Qiagen, Valencia, Calif.). DNA concentration was determined spectrophotometrically, and equal levels of total cellular DNA (250 ng or 500 ng) were analyzed by RQ-PCR essentially as described previously (7, 13, 29, 31). Infections were performed in duplicate, and each infection was analyzed in duplicate by RQ-PCR. Virus particles lacking Env glycoprotein were used in most infections to control for plasmid DNA carryover from transfections.

HIV-1 primers, probes, and PCR conditions were as previously described (7) except that the minus-strand primer for late reverse transcription HIV-1NL4-3 products was AE331 (5′-CCTGCGTCGAGAGATCTCCTCTGG-3′). Plasmids p83-2/_Xma_I and pSVC21/5′-LTR (4) were used to generate standard curves for HIV-1NL4-3 and HIV-1HXBc2 late reverse transcription products, respectively. Plasmid pUC19.2LTR was used as the standard for the two-long-terminal-repeat (2-LTR)-containing HIV-1 circle.

RESULTS

F− HIV-1NL4-3 displays wild-type replication kinetics under a variety of infection conditions.

Retroviral polypurine tracts are preceded by an upstream uridine-rich sequence known as the U-box (28), and a 4-base U-box abuts the 16-bp HIV-1 cPPT (Fig. 1). The cPPT-D mutation, which replaced two U-box bases with cytosine and replaced 6 of the 16 purines with pyrimidines (Fig. 1), inhibited synthesis of the central DNA flap during reverse transcription of HIV-1LAI (50). The resulting F− mutant was unable to initiate a spreading infection in either PBMC or MT-4 cells (50). We found it surprising that F− HIV-1LAI, which expressed a functional integrase protein (Fig. 1), was unable to initiate a spreading infection in MT-4 cells because we recently reported that HIV-1NL4-3 lacking functional integrase replicated in these cells, albeit delayed approximately 1 week compared with the wild-type (37). Because of the complete block in MT-4 cells, we inferred that F− HIV-1LAI was at least 104-fold defective in its ability to initiate a spreading viral infection (37, 50). In contrast to this large defect in viral spread, F− HIV-1 vectors were only 2- to 10-fold defective in single-round infectivity assays (16, 20, 39, 40, 44, 50, 51).

Wild-type and mutant sequences. The central U-box and cPPT form part of the integrase coding region. Mutant sequences that differ from the wild-type (WT) sequence are highlighted by black boxes. Note that the cPPT-D mutation substitutes Arg for Lys-188 in the integrase. This change alone does not appear to impact integrase function, as HIV-1LAI carrying just the K188R change replicated like the wild-type virus under conditions in which the HIV-1LAI cPPT-D mutant was replication defective (50). Numbering is that for HIV-1NL4-3 (1, 35).

To further investigate the role of the flap in HIV-1 infection, the cPPT-D mutation was introduced into two related T-cell-tropic strains, HIV-1NL4-3 and HIV-1HXBc2 (1, 19, 35). HIV-1NL4-3 is a chimera of HIV-1NY5′ and HIV-1LAI (1). Because HIV-1LAI is highly related to HIV-1HXB2 (35), HIV-1NL4-3 and HIV-1HXBc2 are most related in their 3′ halves; the strains are 96.7% and 98.5% identical upstream and downstream of the _Eco_RI site that determined the chimera junction in HIV-1NL4-3, respectively (1, 19, 35). HIV-1NL4-3 and HIV-1HXBc2 differ in the functionality of accessory gene products. Whereas all genes in HIV-1NL4-3 (1) and HIV-1LAI (41) are intact, the HIV-1HXBc2 strain used in the majority of the spreading replication assays here was defective for vpr and nef (19, 23).

Since it was determined that F− HIV-1LAI was completely defective in MT-4 cells (50), we first assayed wild-type and F− virus replication in MT-4 cells infected at a relatively high multiplicity of infection (MOI). Cells infected with 106 RT cpm of wild-type HIV-1NL4-3, which equated to an MOI of approximately 0.5 (37), supported peak virus replication 3 days postinfection (Fig. 2A). Unexpectedly, F− HIV-1NL4-3 displayed the wild-type replication profile (Fig. 2A). Diluting virus 25-fold again yielded similar wild-type and F− replication kinetics in MT-4 cells (Fig. 2B). A further 625-fold dilution, equating to an MOI of just 3.3 × 10−5, again yielded identical replication profiles for the wild-type and F− HIV-1NL4-3 viruses (Fig. 2C). We note that on average, only 65 virions initiated infection under these conditions (37).

F− HIV-1NL4-3 displays wild-type replication kinetics. (A) MT-4 cells infected with 106 RT cpm of wild-type (WT) HIV-1NL4-3 (▪), F− HIV-1NL4-3 (□), or supernatant from mock-transfected cells (×). (B) MT-4 cells infected with 4 × 104 RT cpm of the viruses indicated in panel A. (C) MT-4 cells infected with 64 RT cpm of virus. (D through F) Jurkat cells infected with the levels of wild-type and F− viruses indicated for MT-4 cells in panels A to C, respectively. (G and H) PBMC infected with 106 and 4 × 104 RT cpm, respectively, of the indicated viruses. Numbers in panels A though F are multiplicities of infection for wild-type HIV-1NL4-3 based on previously reported TCID50 values (37). Dpi, days postinfection.

Since Jurkat cells were approximately 11-fold less permissive than MT-4 cells in their ability to replicate HIV-1NL4-3 (37), we next analyzed wild-type and F− virus replication in Jurkat cells to see if a defective F− phenotype might be visible under these somewhat more stringent growth conditions. Jurkat cells infected with 106 and 4 × 104 RT cpm, which correlated to approximate MOIs of 0.04 and 0.002, respectively (37), supported very similar replication profiles for the wild-type and F− HIV-1NL4-3 viruses (Fig. 2D and E). Jurkat cells infected at an MOI of 2.8 × 10−6 supported peak wild-type HIV-1NL4-3 replication 9 days postinfection (Fig. 2F). In this case, the growth of F− HIV-1NL4-3 was delayed approximately 3 days compared with the wild-type virus (Fig. 2F). Under these conditions, only about six wild-type virions initiated infection.

Wild-type and F− HIV-1NL4-3 virus replication was also tested in primary human PBMC. PBMC infected with 106 and 4 × 104 RT cpm once again supported very similar profiles of wild-type and F− HIV-1NL4-3 replication (Fig. 2G and H). We note that the precise MOIs for these infections are unknown because we have not determined 50% tissue culture infectious doses (TCID50) with PBMC. Nonetheless, we conclude that an intact cPPT sequence confers very little if any growth advantage on HIV-1NL4-3 under a variety of conditions of viral spread (Fig. 2).

HIV-1HXBc2 cPPT-D mutant is replication defective in primary T cells.

We next analyzed the replication profiles of the wild-type and F− HIV-1HXBc2 viruses under the infection conditions described for Fig. 2. In this case, precise MOIs were unknown, as we did not determine HIV-1HXBc2 titers by TCID50. MT-4 cells infected with 106 RT cpm of wild-type HIV-1HXBc2 supported peak virus replication 2 days postinfection, and F− HIV-1HXBc2 replicated with an apparent 1-day delay under these conditions (Fig. 3A). Diluting HIV-1HXBc2 prior to infection revealed a subtle replication defect: the F− virus was delayed about 3 days compared with the wild-type virus in cells infected with 4 × 104 and 64 RT cpm (Fig. 3B and C).

F− HIV-1HXBc2 displays a target cell type-dependent replication defect. Cells in panels A though H were infected with the same levels of wild-type (WT, •) and F− (○) HIV-1HXBc2 RT cpm as indicated for HIV-1NL4-3 in Fig. 2, panels A to H. Other labeling is the same as in Fig. 2.

The slight defect observed in MT-4 cells was enhanced in less permissive Jurkat cells and PBMC. Jurkat cells infected with 106 RT cpm supported similar replication profiles of the wild-type and F− HIV-1HXBc2 viruses (Fig. 3D). However, F− HIV-1HXBc2 growth was delayed about 6 days compared with the wild-type virus following infection with 4 × 104 RT cpm (Fig. 3E) and 64 RT cpm (Fig. 3F). Most notably, F− HIV-1HXBc2 was almost completely replication defective in PBMC (Fig. 3G and H). Whereas PBMC infected with 106 RT cpm supported weak F− HIV-1HXBc2 growth (Fig. 3G), replication was barely detected following infection with 4 × 104 RT cpm in repeated experiments (Fig. 3H). Unlike the results obtained with HIV-1NL4-3, we conclude that the cPPT-D mutation can alter the replication profile of HIV-1HXBc2. However, the ability to detect a complete replication defect depended on MOI and target cell type (Fig. 3).

Wild-type levels of HIV-1 replication in the absence of the central DNA flap.

The results of the previous experiments revealed the cPPT-D mutation differentially affected the replication capacities of HIV-1HXBc2 and HIV-1NL4-3. Whereas F− HIV-1NL4-3 displayed wild-type replication kinetics under a variety of conditions (Fig. 2), 4 × 104 RT cpm of F− HIV-1HXBc2 failed to initiate a spreading infection in PBMC (Fig. 3). Because of this, we considered the possibility that the cPPT-D mutation might differentially affect the synthesis of the central DNA flap in the two strains. Although this seemed unlikely because the cPPT and central termination sequence lie in a region that is highly conserved between HIV-1LAI, HIV-1HXBc2, and HIV-1NL4-3 (1, 19, 35, 41), we nonetheless characterized wild-type and F− HIV-1NL4-3 and HIV-1HXBc2 cDNAs purified from infected cells to address the possibility of differential flap DNA synthesis.

The flap detection assay (Fig. 4A) was based on previous observations that HIV-1 contains a hypersensitive site for S1 nuclease near the middle of the minus strand (9, 26). The flap is a 99-base overlap of the lentiviral plus strand, and this plus strand discontinuity yields the minus-strand cleavage site (Fig. 4A). Southern blots were probed with a riboprobe complementary to the downstream region of the minus strand. In this way, wild-type cDNA should yield a 4.9-kb minus-strand S1 digestion product. F− digests, in contrast, should lack this product (Fig. 4A).

cPPT-D mutation inhibits synthesis of the central DNA flap in HIV-1NL4-3 and HIV-1HXBc2. (A) Flap detection assay. The predicted structures of the plus and minus strands in the central region of wild-type (WT) and cPPT-D mutant cDNAs are shown. The discontinuous wild-type plus strand yields the minus-strand S1 nuclease-sensitive site. The thick line indicates the relative position of the minus-strand-specific riboprobe. CTS, central termination sequence. (B) Southern blot. Purified cDNAs were either digested with S1 nuclease or left untreated, as indicated. The migration positions of the full-length and half-genome minus strands are marked cDNA and 4.9 kb, respectively. Molecular mass standards were loaded in lanes 3 and 9. M, lysate prepared from mock-infected cells.

Similar levels of full-length minus-strand wild-type and F− HIV-1NL4-3 cDNAs were synthesized in the cytoplasm of acutely infected SupT1 cells (Fig. 4B, lanes 1 and 2). Digestion with S1 nuclease yielded a readily detectable 4.9-kb wild-type product (Fig. 4B, lane 4) that was absent from the F− digest (Fig. 4B, lane 5). Thus, as predicted, the cPPT-D mutation abolished the centrally located S1-hypersensitive site in the HIV-1NL4-3 minus strand and, by extension, the synthesis of the central DNA flap. We noted, however, the appearance of several novel bands in the F− HIV-1NL4-3 digest (Fig. 4B, lane 5). The potential significance of these minor cleavage products is addressed in the Discussion.

SupT1 cells infected with the wild-type and F− HIV-1HXBc2 viruses yielded barely detectable levels of cytosolic cDNA synthesis. To circumvent this, infected MT-4 cells were propagated for 2 to 3 days to allow viral spread and cDNA amplification, and total cell lysates were prepared by the Hirt method. The wild-type and F− HIV-1HXBc2 viruses yielded similar levels of cDNA synthesis under these conditions (Fig. 4B, lanes 6 and 7). As witnessed for HIV-1NL4-3, S1-digested wild-type HIV-1HXBc2 yielded a subgenomic 4.9-kb product that was absent from the F− digest (Fig. 4B, lanes 10 and 11). Thus, as predicted, the cPPT-D mutation effectively inhibited the synthesis of the HIV-1HXBc2 flap.

F− mutants are at most twofold defective for nuclear import.

We next analyzed the role of the flap in the nuclear localization of HIV-1NL4-3 and HIV-1HXBc2 cDNA. The linear duplex product of reverse transcription is the cDNA substrate for integrase-mediated integration into chromosomal DNA (reviewed in references 14 and 22). In addition to integrated proviruses, unintegrated linears are converted into 1- and 2-LTR-containing DNA circles in the nuclei of infected cells (14, 21, 22). The 2-LTR circle has been used extensively as a marker for nuclear localization because the LTR-LTR circle junction affords a unique sequence for PCR amplification (14).

In this study, we used RQ-PCR to quantify levels of total HIV-1 and 2-LTR circles in single-round infections and used the fraction of 2-LTR circles as an indicator of preintegration complex nuclear import. Although this technique did not account for total nuclear HIV-1 DNA, it nonetheless afforded quantitative comparison of wild-type and mutant nuclear import activities. Env− derivatives of HIV-1NL4-3 and HIV-1HXBc2 were used in RQ-PCR assays to restrict reverse transcription and nuclear import to single-round infections.

Jurkat cells infected with HIV-1NL4-3 supported similar levels of wild-type and F− cDNA synthesis in repeated experiments (Fig. 5A and data not shown). By 24 h postinfection, 0.7% to 0.9% of wild-type cDNA was present as 2-LTR circles (Fig. 5A and B and data not shown). In four different experiments, the fraction of F− HIV-1NL4-3 circles ranged from 48% to 89% of the wild-type value (Fig. 5A and B and data not shown), with an average value of 63% ± 16.2%. Thus, the flap affected HIV-1NL4-3 2-LTR circle formation at most twofold under these assay conditions.

cPPT-D mutation reduces the formation of HIV-1NL4-3 2-LTR circles approximately twofold in Jurkat cells. (A) Cells infected with wild-type (WT) HIV-1NL4-3 (▪), F− HIV-1NL4-3 (□), or supernatant from Env− transfections (×) were lysed at the indicated times, and 250 ng of total cellular DNA was assayed for total HIV-1 cDNA by RQ-PCR. (B) Cells infected with wild-type HIV-1NL4-3 (▪), F− HIV-1NL4-3 (□), or mock-treated supernatant (×) were analyzed for levels of 2-LTR circles by RQ-PCR. Error bars represent variations between duplicate RQ-PCR assays. LRT, late reverse transcription; Hpi, hours postinfection.

Wild-type HIV-1HXBc2 yielded only 10% to 20% of the level of wild-type HIV-1NL4-3 cDNA under identical infection conditions (data not shown). To improve on this relatively low cDNA yield, the apparent MOI was increased 10-fold by (i) increasing the level of virus inoculum fivefold and (ii) analyzing twice as much total cellular DNA by RQ-PCR. In repeated experiments, Jurkat cells supported three- to fivefold less F− HIV-1HXBc2 cDNA synthesis than wild type, and F− HIV-1HXBc2 formed about half as many 2-LTR circles as the wild type did (Fig. 6A and B and data not shown). However, due to the lower amount of total cDNA synthesis, the fraction of F− HIV-1HXBc2 cDNA converted into 2-LTR circles actually exceeded the wild-type value in repeated experiments (Fig. 6A and B; Table 1).

F− HIV-1HXBc2 supports wild-type level of 2-LTR circle formation in different cell types. (A) Jurkat cells infected with wild-type (WT) HIV-1HXBc2 (•) or F− HIV-1HXBc2 (○) virus were lysed at the indicated times, and 500 ng of total cellular DNA was assayed for total HIV-1 cDNA by RQ-PCR. (B) Cell lysates from panel A were analyzed for 2-LTR circle levels. (C) PBMC infected and processed as indicated for Jurkat cells in panel A. (D) 2-LTR circle levels of wild-type and F− HIV-1HXBc2 in PBMC. Other labeling is the same as in Fig. 5.

TABLE 1.

Nuclear import activity of F− HIV-1HXBC2 in Jurkat cells and PBMC

Cells	Expt	2-LTR circle formation_a_ (% of control) for infections mediated by:
HIV-1 Env	VSV-G
Jurkat	1	185_b_	112_c_
2	274	249
PBMC	1	75_b_	113_c_
2	192	93

Since the flap was more important for efficient spread in PBMC than in Jurkat cells (Fig. 3), we next analyzed F− HIV-1HXBc2 cDNA synthesis and 2-LTR circle formation in PBMC. PBMC supported about twofold less F− HIV-1HXBc2 than wild-type cDNA synthesis in repeated experiments (Fig. 6C and data not shown). Wild-type HIV-1HXBc2 converted only a minor fraction of its total cDNA into 2-LTR circles under these infection conditions (Fig. 6D). Although the fraction of total F− HIV-1HXBc2 converted into 2-LTR circles was similar to that of the wild type (Fig. 6C and D; Table 1), infections were repeated with VSV-G pseudotypes in hopes of increasing the efficiencies of cDNA synthesis and 2-LTR circle formation.

As anticipated, VSV-G significantly enhanced wild-type HIV-1HXBc2 cDNA synthesis and 2-LTR circle formation in Jurkat cells (compare Fig. 7A and B with Fig. 6A and B). Wild-type and F− HIV-1HXBc2 VSV-G pseudotypes synthesized similar cDNA levels in repeated experiments (Fig. 7A and data not shown). The flap did not appear to affect 2-LTR circle formation under these conditions, as similar levels of wild-type and F− cDNA were converted into 2-LTR circles in repeated experiments (Fig. 7A and B; Table 1). PBMC infected with VSV-G pseudotypes also supported similar levels of wild-type and F− HIV-1HXBc2 cDNA synthesis (Fig. 7C). And, in repeated experiments, both viruses formed similar levels of 2-LTR circles (Fig. 7C and D; Table 1).

Wild-type and F− HIV-1HXBc2 VSV-G pseudotypes form similar levels of 2-LTR circles in Jurkat cells and PBMC. (A) Jurkat cells infected with VSV-G-pseudotyped wild-type (WT) and F− HIV-1HXBc2 viruses were lysed at the indicated times, and 500 ng of total cellular DNA was assayed for total HIV-1 cDNA by RQ-PCR. (B) Lysates from panel A were assayed for 2-LTR circles by RQ-PCR. (C) PBMC infected with VSV-G pseudotypes were processed as described for panel A. (D) 2-LTR circle levels in PBMC infected with VSV-G pseudotypes. Other labeling is the same as in Fig. 5.

Strain-dependent flap function.

F− HIV-1HXBc2 failed to replicate in PBMC (Fig. 3H) under conditions in which F− HIV-1NL4-3 replication was readily detected (Fig. 2H). Yet the results in the previous section failed to detect a defect in F− HIV-1HXBc2 2-LTR circle formation relative to total cDNA synthesis in Jurkat cells or PBMC (Fig. 6 and 7; Table 1). To further probe the relationship between strain-specific sequences and flap function, wild-type and F− HIV-1NL4-3 and HIV-1HXBc2 viruses were analyzed in side-by-side single-round transduction assays. Jurkat cells were infected with equal RT cpm of Env− vectors carrying the bacterial CAT gene in place of nef, and the levels of wild-type and F− virus required for 50% activity were calculated following endpoint dilution analyses of infected cell lysates in in vitro CAT assays (37).

Wild-type HIV-1NL4-3 was about sixfold more active than wild-type HIV-1HXBc2 in these assays, as about 3 × 107 RT cpm and 5 × 106 RT cpm of HIV-1HXBc2 and HIV-1NL4-3, respectively, were required for 50% conversion. Whereas the cPPT-D mutation reduced the infectivity of HIV-1HXBc2 two- to threefold, it had no discernible effect on the transduction activity of HIV-1NL4-3 (Fig. 8). Thus, the slight nuclear import defect observed in Fig. 5 did not appear to impact the activity of F− HIV-1NL4-3 in either spreading (Fig. 2) or single-round (Fig. 8) infection assays.

Single-round infectivities of wild-type and F− HIV-1NL4-3 and HIV-1HXBc2. Levels of wild-type (WT) HIV-1NL4-3 and wild-type HIV-1HXBc2 required for 50% CAT activity, approximately 5 × 106 RT cpm and 3 × 107 RT cpm, respectively, were normalized to 1.0. The levels of F− HIV-1NL4-3 and F− HIV-1HXBc2 required for 50% activity are shown in comparison to their wild-type counterparts. Error bars represent the standard deviation of three independent infections.

Since Nef has been shown to play an important role in primary cell infectivity (30, 36, 45) and the HIV-1HXBc2 strain used in the spreading infection assays in Fig. 3 was a vpr nef double mutant, we next analyzed Nef-positive derivatives of wild-type and F− HIV-1HXBc2 to determine if nef contributed to the strain-dependent replication phenotypes of F− HIV-1NL4-3 (Fig. 2) and F− HIV-1HXBc2 (Fig. 3) in PBMC.

PBMC infected with 4 × 104 RT cpm of Nef− HIV-1HXBc2 supported peak virus growth 11 days postinfection (Fig. 9). Although Nef− HIV-1HXBc2 replication was delayed approximately 5 days compared with its growth in Fig. 3H, we note that the two experiments used PBMC derived from different blood donors. Similar to the results in Fig. 3H, F− Nef− HIV-1HXBc2 failed to initiate a spreading infection in PBMC (Fig. 9).

Replication defect of F− HIV-1HXBc2 in PBMC is partially restored by wild-type nef. PBMC infected with 4 × 104 RT cpm of Nef− HIV-1HXBc2 (•), Nef− F− HIV-1HXBc2 (○), Nef+ HIV-1HXBc2 (▴), Nef+ F− HIV-1HXBc2 (▵), or supernatant from a mock transfection (×) were analyzed for RT activity at the indicated times. Dpi, days postinfection.

Functional Nef significantly enhanced the replication of both wild-type and F− HIV-1HXBc2 in PBMC. Whereas Nef+ HIV-1HXBc2 reached its peak growth 9 days postinfection, Nef+ F− HIV-1HXBc2 displayed peak growth at 11 days postinfection (Fig. 9). Since the results of RQ-PCR assays indicated that Nef− wild-type and F− Nef− HIV-1HXBc2 entered PBMC nuclei with similar efficiencies (Fig. 6 and 7; Table 1), we conclude that the nef defect, and not nuclear import, was primarily responsible for the spreading replication defect of Nef− F− HIV-1HXBc2 in PBMC (Fig. 3H and 9).

DISCUSSION

The central DNA flap is an approximately 100-base plus-strand overlap in de novo-synthesized lentiviral cDNA. Previously, it was shown that the cPPT-D mutation, which altered a total of 10 bp in the central U-box and cPPT (Fig. 1), destroyed the synthesis of the flap during reverse transcription of HIV-1LAI (50). Consistent with this finding, we found that the cPPT-D mutation abolished the synthesis of the central DNA flap in HIV-1NL4-3 and HIV-1HXBc2 (Fig. 4). In stark contrast to the previous report, however, we found that F− HIV-1NL4-3 and F− HIV-1HXBc2 replicated efficiently in MT-4 cells, even when assayed at extremely low MOIs (Fig. 2C and 3C). We note that our description of efficient F− replication did not depend on the HIV-1NL4-3 and HIV-1HXBc2 strains studied here, as Dvorin and coworkers have described efficient F− HIV-1YU-2 and F− HIV-1LAI replication under a variety of infection conditions (see the companion paper by Dvorin et al. [16a] in this issue).

Virus strain- and target cell type-dependent flap function.

Our results did reveal a role for the flap in HIV-1 spreading replication, but this was highly strain and target cell type dependent (Fig. 2 and 3). F− HIV-1NL4-3 was basically a wild-type virus (Fig. 2). Although F− HIV-1HXBc2 replicated poorly in PBMC, we failed to detect a nuclear import defect under these infection conditions (Fig. 6 and 7; Table 1), and instead determined that the nef mutant phenotype of this strain played a large part in the observed replication defect (Fig. 9).

Since several previously reported F− vectors incorporated HIV-1NL4-3 sequences (16, 20, 44, 50, 51), we were somewhat surprised that our F− HIV-1NL4-3 vector transduced Jurkat cells as efficiently as the wild type did (Fig. 8). One possible explanation may be the genome sizes of the vectors in the different studies. Since most of HIV-1NL4-3 was removed from the previously analyzed vector constructs, it is possible that sequences present in our nearly full-length HIV-1NL4-3 vector helped to counteract the loss of the central DNA flap. We speculate that minor discontinuities observed throughout the F− HIV-1NL4-3 plus strand (Fig. 4, lane 5) could potentially fulfill this role. Although the cPPT-D mutation reduced the single-round infectivity of our HIV-1HXBc2 vector two- to threefold, we emphasize that this did not correlate with reduced nuclear import (Fig. 6 and Table 1). We speculate that reduced cDNA synthesis (Fig. 6A) and/or delayed nuclear import (Fig. 6B) may account for the observed reduction in F− HIV-1HXBc2 activity in single-round infections (Fig. 8). A delay in 2-LTR circle formation in infected HeLa cells was previously noted for a small F− HIV-1NL4-3 vector (20).

Single-round infectivity versus spreading HIV-1 replication.

Our finding that F− HIV-1HXBc2 was two- to threefold defective in single-round transduction assays (Fig. 8) agrees with numerous previous reports that the flap imparted an infection advantage of between 2- and 10-fold on single-round HIV-1 vectors (16, 20, 39, 40, 44, 50, 51). However, we emphasize that this slight defect in single-round infectivity did not correlate with a severe defect in spreading HIV-1 replication (Fig. 3D).

We previously determined by endpoint dilution that HIV-1NL4-3 lacking functional integrase was approximately 58,000-fold defective in its ability to initiate a spreading infection in MT-4 cells (37). Yet, when analyzed in bulk cultures infected at a relatively high MOI, growth of the integrase-negative HIV-1NL4-3 was delayed only about 1 week compared with the wild-type virus (37). Considering these results, one would predict that mutants displaying just 2- to 10-fold defects in single-round infectivity assays, like the F− HIV-1HXBc2 strain studied here, would spread with close to wild-type efficiency in bulk-infected cell cultures (Fig. 3 and 9).

Although the flap can influence the single-round transduction activity of certain HIV-1 vectors as much as 10-fold and is therefore likely an important regulatory sequence for lentivirus-based gene therapy vectors, the results presented here cast doubt on the prior claim that the central DNA flap is an essential determinant of preintegration complex nuclear import and spreading HIV-1 infectivity (50). Experiments aimed at elucidating the major player(s) in the nuclear import of HIV-1 preintegration complexes in nondividing and dividing cells are currently under way in our laboratory.

Acknowledgments

We thank D. Gabuzda, H. Göttlinger, and J. Sodroski for gifts of plasmid DNAs and valuable discussions and J. D. Dvorin and M. H. Malim for sharing results prior to publication.

This work was supported by NIH grants AI39394 (A.E.), AI45313 (A.E.), and AI28691 (Center for AIDS Research); by a gift from the G. Harold and Leila Y. Mathers Charitable Foundation (A.E.); and by the Japanese Foundation for AIDS Prevention (N.N.).

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