Published online 1 August 2005
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Transcription factor binding sites in the pol gene intragenic regulatory region of HIV-1 are important for virus infectivity
Laboratoire de Virologie Moléculaire, Institut de Biologie et de Médecine Moléculaires (IBMM), Service de Chimie Biologique, Université Libre de Bruxelles Rue des Profs Jeener et Brachet 12, 6041 Gosselies, Belgium 1Faculté de Médecine, Laboratoire de Virologie Moléculaire, Université Libre de Bruxelles 808 Route de Lennik, 1070 Bruxelles, Belgium 2Institut de Biologie de Lille, Institut Pasteur de Lille, Université de Lille 1, UMR 8117 CNRS BP 447, 1 Rue Calmette, 59021 Lille Cedex, France 3INSERM U119 27 Boulevard Lei Roure, 13009 Marseille, France
*To whom correspondence should be addressed. Tel: +32 2 650 9807; Fax: +32 2 650 9800; Email: cvlint{at}ulb.ac.be
Received June 21, 2005. Accepted July 4, 2005.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We have previously identified in the pol gene of human immunodeficiency virus type 1 (HIV-1) a new positive transcriptional regulatory element (nt 4481–4982) containing recognition sites for nuclear proteins (sites B, C, D and a GC-box) [C. Van Lint, J. Ghysdael, P. Paras, Jr, A. Burny and E. Verdin (1994) J. Virol. 68, 2632–2648]. In this study, we have further physically characterized each binding site and have shown that the transcription factors Oct-1, Oct-2, PU.1, Sp1 and Sp3 interact in vitro with the pol region. Chromatin immunoprecipitation assays using HIV-infected cell lines demonstrated in the context of chromatin that Sp1, Sp3, Oct-1 and PU.1 are recruited to the HS7 region in vivo. For each site, we have identified mutations abolishing factor binding to their cognate DNA sequences without altering the underlying amino acid sequence of the integrase. By transient transfection assays, we have demonstrated the involvement of the pol binding sites in the transcriptional enhancing activity of the intragenic region. Our functional results with multimerized wild-type and mutated pol binding sites separately (i.e. in the absence of the other sites) have demonstrated that the PU.1, Sp1, Sp3 and Oct-1 transcription factors regulate the transcriptional activity of a heterologous promoter through their respective HS7 binding sites. Finally, we have investigated the physiological role of the HS7 binding sites in HIV-1 replication and have shown that these sites are important for viral infectivity.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The expression of human immunodeficiency virus type 1 (HIV-1), the etiological agent of AIDS (1–3), is regulated both at the transcriptional and the post-transcriptional levels (4). Control of HIV-1 transcription is mediated by cis-acting elements located in the viral 5'-long terminal repeats (5'-LTRs), by the viral trans-regulatory protein Tat and by cellular transcription factors. In addition to the enhancer located in the 5'-LTR, a 12-O-tetradecanoylphorbol-13-acetate (TPA)-inducible intragenic enhancer has been identified in the pol gene of HIV-1 (5). This element is composed of two functional subdomains encompassing nt 4079–4342 and nt 4781–6026, both exhibiting TPA-inducible enhancing activity, on the herpes simplex virus (HSV) thymidine kinase (TK) promoter in HeLa cells (5), but no significant activity in T-lymphoid and monocyte/macrophage cell lines. Nevertheless, analysis of the chromatin organization of integrated HIV-1 identified a single major nuclease-hypersensitive site in the 8 kb region located between the two LTRs (6,7). This hypersensitive site, centered around nt 4490–4766 [according to the numbering of the HIV-1 NY5 genome (where nt +1 is the start of U3 in the 5'-LTR)], is located in the part of the pol gene encoding the integrase protein, precisely between the two functional domains of the intragenic enhancer identified in HeLa cells (7). This constitutive hypersensitive site is present only in a cell line of monocytic origin (U1) and not in two cell lines of lymphoid origin (8E5 and ACH2), suggesting a cellular specificity associated with this intragenic element (6). A 500 bp fragment (nt 4481–4982) encompassing the pol gene hypersensitive site region (called HS7 region) acts ex vivo as a cis-acting positive regulatory element. Indeed, when cloned downstream and in the sense orientation relative to the HIV-1 promoter, the HS7 region reproducibly increases transcription mediated by the HIV-1 5'-LTR (7).
Physical characterization of the HS7 region identified several recognition sites for ubiquitous and cell-type-specific nuclear factors, suggesting their involvement in the control of the transcriptional activity of this region (7). By in vitro binding studies of the HS7 region, four distinct DNA-binding motifs for nuclear proteins (called from 5' to 3' site B, GC-box, site C and site D) were physically defined (Figure 1A). Site B (nt 4519–4545) specifically binds four distinct nuclear protein complexes: an ubiquitous factor (B1), a B-cell specific factor (B2), a T-cell specific factor (B3) and a protein(s) related to the transcription factor PU.1 (B4) (Figure 1B). The GC-box (nt 4623–4631) binds purified Sp1 protein in vitro. Site C (nt 4681–4701) specifically binds three nuclear protein complexes: an ubiquitous factor (C1), a B-cell specific factor (C2) and a T-cell specific factor (C3) (Figure 1C). These factors have a DNA-binding specificity similar to that of factors binding to site B (B1, B2 and B3, respectively). Site D (nt 4816–4851) specifically binds an ubiquitously expressed factor(s) (7). However, the physiological role of these HS7 binding sites in HIV-1 transcription and replication remains unknown to date.
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In this study, we have further characterized physically each of these four binding sites (sites B, C, D and the GC-box) located in the HS7 region, examined the functional transcriptional role of each site separately in a heterologous context, and investigated their biological significance in the HIV-1 replication cycle.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell culture
The monocytic cell lines U937 and U1 and the T-lymphoid cell lines JE6-1 (a clonal line of Jurkat cells), A3.01 (a clonal derivative of the CEM cell line), the clonal Jurkat JE6-1 cell lines stably expressing HIV-1 Tat proteins (Tat72 and Tat101) or the empty vector cassette (8), and ACH2 were maintained in RPMI 1640-Glutamax I medium (Invitrogen) supplemented with 10% fetal bovine serum (Myoclone Superplus). The HIV-1-infected U1 and ACH2 cell lines were obtained from AIDS Research and Reference Reagent Program, National Institute of Allergy and Infectious Diseases (NIAID), National Institutes of Health (NIH). The adherent cell line NIH3T3 (a murine fibroblastic cell line) and 293T cells (human embryonic kidney cell line) were cultured in Dulbecco's modified Eagle's-Glutamax I medium containing 10% calf serum. The cell line SL2 (a Drosophila cell line) was cultured in Schneider's Drosophila medium supplemented with L-glutamine and with 10% fetal bovine serum. All media (Invitrogen) also contained 50 U of penicillin/ml and 50 µg of streptomycin/ml (Invitrogen). All cells were grown at 37°C in a 5% CO2 atmosphere, except for SL2 cells which were grown at 28°C without CO2.
Plasmid constructs
An SmaI–XhoI fragment containing the HIV-1 LAI 5'-LTR (nt 1–789) was purified from the previously described pLTR(1-789)-luc construct (9). This fragment was cloned into the unique EcoICRI–XhoI sites of the reporter vector pGL3-Basic (Promega). The resulting plasmid was designated pLTR.
PCR was used to amplify the pol gene fragment (nt 4481–4982) corresponding to the HS7 region from the infectious proviral molecular clone pNL4-3 (reagent no. 114, received from the AIDS Research and Reference Reagent Program, NIAID, NIH). BamHI sites were introduced in the PCR primers, and the BamHI-restricted PCR fragment was cloned in the unique BamHI site of the pLTR, placing the fragment downstream of the 5'-LTR-luc transcriptional unit. The 5' primer oligonucleotide encompassed the coding strand sequence from nt 4481 to 4505 and contained an added BamHI restriction site (underlined) at the 5' end (5'-TCCCCCGGGATCC[nt 4481]GAAGCAGAAGTAATTCCAGCAGAG-3'). The 3' primer oligonucleotide encompassed the complementary sequence of the pol gene from nt 4957 to 4982, and contained an added BamHI site (underlined) at the 5' end (5'-TCCCCCGGGATCC[nt 4982]TATTACTACTGCCCCTTCACCTTTCC-3'). This plasmid was designated pLTR-HS7wt. Fragments containing the HS7 region mutated in the different binding sites (sites B, C, D and the GC-box) individually or in combination were PCR amplified from the pCV11, pCV14, pCV16, pCV19, pCV25, pCV1069 and pCV1107 plasmids (see below). The 5' and 3' primer oligonucleotides were as described above. The different amplified fragments were digested with BamHI and then ligated into the BamHI site of pLTR to generate pLTR-HS7totmut, pLTR-HS7mutB/Oct, pLTR-HS7mutB/PU, pLTR-HS7mutC, pLTR-HS7mutSp, pLTR-HS7mutD and pLTR-HS7mutCmutB/Oct, respectively.
The pTK reporter plasmid contains the HSV TK minimal promoter and was described previously (10).
The p(Cwt)3TK was generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats of the site C sequence (5'-TAGAATCTATGAATAAAGAAT-3') in the forward orientation into SmaI-digested pTK-luc. Similarly, the p(CmutOct)3TK was generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats of site C mutated in the Oct motif with the sequence 5'-TAGAATCCATGAACAAAGAAT-3' (the mutations are underlined) in the forward orientation into SmaI-digested pTK.
The p(Bwt)3TK was generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats of the site B sequence (5'-CAGCATACTTCCTCTTAAAATTAGCAG-3') in the forward orientation into SmaI-digested pTK. Similarly, the p(BmutOct)3TK and p(BmutPU.1)3TK were generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats either of site B mutated in the Oct motif with the sequence 5'-CAGCATACTTCCTCTTGAAGTTGGCAG-3', or of site B mutated in the PU box with the sequence 5'-CAGCATATTTTCTCTTAAAATTAGCAG-3' (the mutations are underlined) in the forward orientation into SmaI-digested pTK.
The p(Spwt)3TK was generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats of the Sp site sequence (5'-CCTGTTGGTGGGCGGGGATCAAG-3') in the forward orientation into SmaI-digested pTK. Similarly, the p(Spmut)3TK was generated by inserting, upstream of the TK promoter, a double-stranded oligonucleotide corresponding to three direct repeats of the Sp site mutated in the GC-box with the sequence 5'-CCTGTTGGTGGGCAGGAATCAAG-3' (the mutations are underlined) in the forward orientation into SmaI-digested pTK-luc.
The Oct-1 and Oct-2 expression vectors (pCG-Oct-1 and pCG-Oct-2) (kindly provided by Dr Winship Herr) contained the human Oct-1 and Oct-2 cDNAs cloned in the pCG parent plasmid and were described previously (11). The PU.1 expression vector pJ6-PU.1 was described previously (12). The Sp1 and Sp3 expression vectors (pPacSp1 and pPacSp3) (kindly provided by Dr Guntram Suske) contained the human Sp1 and Sp3 cDNAs cloned in the pPac parent plasmid and were described previously (13).
Electrophoretic mobility shift assays
Nuclear extracts were prepared by a rapid method described by Osborn et al. (14). All buffers contained the protease inhibitors antipain (10 µg/ml), aprotinin (2 µg/ml), chymostatin (10 µg/ml), leupeptin (1 µg/ml) and pepstatin (1 µg/ml). Protein concentrations were determined by the method of Bradford (15). The DNA sequences of the coding strand of the double-stranded oligonucleotides used for this study are listed in Figure 9 or in the figure legends. Electrophoretic mobility shift assays (EMSAs) were performed as described previously (7). Briefly, nuclear extract (15 µg of protein) was first incubated on ice for 10 min in the absence of probe and specific competitor DNA in a 16 µl reaction mixture containing 10 µg of DNase-free BSA (Amersham Biosciences), 2 µg of poly(dI–dC) (Amersham Biosciences) as non-specific competitor DNA, 50 µM ZnCl2, 0.25 mM DTT, 20 mM HEPES (pH 7.3), 60 mM KCl, 1 mM MgCl2, 0.1 mM EDTA and 10% (v/v) glycerol. 20 000 c.p.m. of probe (10–40 fmol) was then added to the mixture with or without a molar excess of an unlabeled specific DNA competitor, and the mixture was incubated for 20 min on ice. Samples were subjected to electrophoresis at room temperature on 6% polyacrylamide gels at 150 V for 2–3 h in 1x TGE buffer (25 mM Tris–acetate (pH 8.3), 190 mM glycine and 1 mM EDTA). Gels were dried and autoradiographed for 24–48 h at –70°C. For supershift assays, polyclonal antibodies against Oct-1 (sc-232X), Oct-2 (sc-233X), Sp1 (sc-059X), Sp2 (sc-643X), Sp3 (sc-644X), Sp4 (sc-645X), MEF-2 (sc-10794X) (Santa Cruz Biotechnology), PU.1 (16), MEF2A, -B or -D (kindly provided by Dr Ron Prywes) (17), anti-MEF2C (kindly provided by Dr John Schwarz) (18), or a purified rabbit immunoglobulin (IgG) were added to the reaction mixture and incubated for 30 min on ice before the addition of the radiolabeled probe.
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For analysis of PU.1 binding, whole-cell extracts of Spodoptera frugiperda SF9 cells infected with a recombinant Ac-NPV-PU.1 baculovirus were used as a source for PU.1 protein and extracts from uninfected insect cells were used as a control. Conditions for EMSAs were as described previously (19).
Chromatin immunoprecipitation assays
The Chromatin immunoprecipitation (ChIP) assays were performed by using the ChIP assay kit (Upstate Biotechnology) according to the manufacturer's recommendations. Formaldehyde cross-linking reactions from 5 x 106 HIV-infected U1 cells or ACH2 cells were quenched with 125 mM glycine. Cells were lysed, and chromatin was sonicated to obtain an average DNA length of 500 bp. Following centrifugation, the chromatin was diluted 10-fold and precleared with a protein A–agarose slurry containing salmon sperm DNA and BSA (Upstate Biotechnology). Precleared chromatin was incubated overnight at 4°C with no antibody or with 5 µg of either anti-Oct-1 antibody (sc-232X), anti-Sp1 (sc-59) antibody, anti-Sp3 (sc-644) antibody, anti-PU.1 antibody (sc-352) or normal rabbit IgG control antibody (Upstate Biotechnology, ref. no. 12-370) as control, followed by immunoprecipitation with protein A–agarose. Immunoprecipitated complexes were washed and eluted twice with 200 µl of elution buffer. The protein–DNA cross links were reversed by heating at 65°C overnight, and 10% of the recovered DNA was used for PCR amplification (35 cycles) with a primer set amplifying the HS7 region (nt 4497–4769): 5'-CCAGCAGAGACAGGGCAAGAA-3'/5'-ACTGCCATTTGTACTGCTGTCTT-3' or with an unrelated primer set amplifying a pol gene region located 
2 kb upstream of the HS7 region (nt 2326–2573): 5'-TACAGGAGCAGATGATACAG-3'/5'-CCTGGCTTTAATTTTACTGG-3'. PCR products from all reactions were resolved by agarose gel electrophoresis.
Transient transfection and luciferase reporter assays
NIH3T3 and SL2 cells were transfected using FuGENETM-6 (Roche Molecular Biochemicals) according to the manufacturer's protocol. Twenty-four hours before transfection, cells were seeded at a density of 2 x 105 cells/well in 6-well plates. FuGENETM-6 was added directly to serum-free medium 5 min before addition to the DNA. An aliquot of 100 µl of this FuGENETM-6/serum-free medium mixture was added to the DNA mixture in microcentrifuge tubes. The mixture was incubated for 15 min at room temperature and, finally added to each well of a 6-well plate. Transfected cells were grown in 2 ml of supplemented medium for 40–44 h. Cells were then lysed and assayed for luciferase activity (Promega). Luciferase activities derived from TK promoters were normalized with respect to protein concentrations using the Detergent-Compatible Protein Assay (Bio-Rad).
Jurkat and U937 cells were transfected using jetPEITM (POLYplus) according to the manufacturer's protocol. Briefly, on the day of transfection, cells were harvested at a density of 1 x 106/ml, resuspended in fresh complete medium and seeded at a density of 0.5 x 106 cells/well in 12-well plates. For each well, 1 µg of DNA was diluted into 50 µl of 150 mM NaCl and 4 µl of jetPEITM were diluted into 50 µl of 150 mM NaCl. Next, 50 µl of this jetPEITM solution were added to the 50 µl DNA solution. The 100 µl jetPEITM/DNA mixture was then incubated for 15 min at room temperature and, finally added dropwise to each well. Transfected cells were grown in 1.5 ml for 40–44 h, lysed and assayed for luciferase activity (Promega). All DNA solutions also contained 50 ng of pRL-TK (used as an internal control for transfection efficiency) in which a cDNA encoding Renilla luciferase is under the control of the HSV TK promoter region (Promega). Firefly luciferase activities derived from the HIV-1 LTR were normalized with respect to the Renilla luciferase activities by using the dual-luciferase reporter assay system (Promega).
Site-directed mutagenesis of the HS7 binding sites
An ApaI/EcoRI fragment containing nt 2011–5743 of the HIV-1 genome was obtained after digestion with ApaI and EcoRI of pNL4-3. This fragment was cloned in pBluescript II SK (Stratagene) digested with ApaI and EcoRI to generate pCV10. This plasmid was used as a substrate for mutagenesis of the HS7 binding sites by the transformer site-directed mutagenesis method (Clontech). The HS7 region was mutated in all the HS7 sites (site B, Sp site, site C and site D) with the following five mutagenic oligonucleotides (mutations are highlighted in boldface and underlined): CV1 (siteBmutPU.1), 5'-CCTGCTAATTTTAAGAGAAAATATGCTGTTTCTTGCC-3'; CV3 (siteBmutOct), 5'-CCTGCCAACTTCAAGAGGAAGTATGCTGTTTCTTGCC-3'; CV4 (siteBmutOct/PU.1), 5'-CCTGCCAACTTCAAGAGAAAATATGCTGTTTCTTGCC-3'; CV6 (siteCmutOct), 5'-CTTTAATTCTTTGTTCATGGATTCTATTACTCCTTGACTTTG-3'; CV7 (siteSpmut), 5'-CCTGCTTGATTCCTGCCCACCAAC-3'; and CV14 (siteDmut), 5'-GTATGTCTGTTGCTATTATATCGACTATTCTTTCCCCTGC-3'.
In addition, three individual mutations of the HS7 site B PU box, site C or Sp site alone and one double mutation of the octamer motif in both site B and site C were generated with the CV1, CV6 or CV7 and CV6/CV3, respectively. The oligonucleotide 5'-CTTTTGCTCCCATGGTCTTTCCTG-3', changing a unique AflIII restriction site in pCV10 into a NcoI site (highlighted in boldface) and the reverse oligonucleotide 5'-CTTTTGCTCACATGTTCTTTCCTG-3', changing the unique NcoI restriction site in a mutated pCV10 into a AflIII site (highlighted in boldface) were successively used for selection during mutagenesis. The five mutant resulting pCV10-derivative plasmids were designated as pCV14 (HS7totmut), pCV11 (HS7mutB/PU), pCV25 (HS7mutC), pCV19 (HS7mutSp) and pCV16 (HS7mutCmutB/Oct), respectively. In addition, two other individual mutations were generated by the QuikChange Site-Directed Mutagenesis Kit (Stratagene) using the pCV10 construct as a substrate with the following two pairs of mutagenic oligonucleotide primers (mutations are highlighted in boldface and underlined on the coding strand primer): CV1141-CV1142 (siteBmutOct), 5'-AGCATACTTCCTCTTGAAGTTGGCAGGAAGATGGCCA-3' and CV1147-CV1148 (siteDmut), 5'-CAGGGGAAAGAATAGTCGATATAATAGCAACAGAC-3'.
The resulting pCV10-derivative plasmids were designated as pCV1107 (HS7mutB/Oct) and pCV1069 (HS7mutD), respectively.
The mutated clones containing the HS7 binding sites mutated individually or in combination were fully resequenced between ApaI and EcoRI after identification (Applied Biosystems).
Construction of infectious proviruses containing the HS7 mutations
To eliminate the pUC8 MCS EcoRI site (nt 420) and to keep only the pol gene EcoRI site (nt 5743), the HIV-1 circularly permuted single-LTR-containing infectious molecular clone pEV46 (pHIV, a derivative of pILIC) (20) was partially digested with EcoRI, dephosphorylated and gel purified. The resulting DNA fragment was next ligated to a phosphorylated partially double-stranded oligonucleotide. The coding and non-coding sequences of this oligonucleotide were as follows: CV20, 5'-AATTAGTGGACGTCAC-3' and CV21, 5'-AATTGTGACGTCCACT-3', respectively. The resulting plasmid was designated pCV1. The ApaI/EcoRI mutagenized fragment from the pCV14 was introduced into the unique ApaI–EcoRI sites of pCV1 to generate pCV426 (named pHIV-1*-HS7totmut). As a control, an unmutated ApaI–EcoRI fragment was purified from pCV10 and cloned into the unique ApaI–EcoRI sites of pCV1. We refer to this construct as pCV422 (named pHIV-1*).
Another set of mutated proviral infectious clones were also constructed. To that end, the ApaI/EcoRI mutagenized fragments from pCV14, pCV1107, pCV11, pCV25, pCV19, pCV1069 and pCV16 were introduced into the unique ApaI–EcoRI sites of the two LTRs containing proviral clone pNL4.3 to generate pCV1102 (named pHIV-1-HS7totmut), pCV1106 (named pHIV-1-HS7mutB/Oct), pCV1101 (named pHIV-1-HS7mutB/PU), pCV1105 (pHIV-1-HS7mutC), pCV1104 (named pHIV-1-HS7mutSp), pCV1082 (named pHIV-1-HS7mutD) and pCV1103 (pHIV-1-HS7mutCmutB/Oct), respectively. As a control, an unmutated ApaI–EcoRI fragment was purified from pCV10 and cloned in an identical manner into the unique ApaI–EcoRI sites of pNL4.3. We refer to this construct as pHIV-1.
Generation of viral stocks
Wild-type and mutant HIV-1 infectious DNAs were generated from the single-LTR-containing proviral constructs pCV422 and pCV426, as described above, by BamHI digestion and self-ligation. These concatemerized proviral DNAs (10 µg) were transfected into 107 Jurkat cells using the DEAE–dextran procedure. At 24 h post-transfection, the cultures were cocultivated with 107 SupT1 cells to allow rapid and efficient recovery of progeny virus. HIV-1 wild-type (HIV-1*) and mutant (HIV-1*-HS7totmut) stocks were prepared at the peak of viral production from supernatants after filtration through a 0.45 µm pore-size membrane. The full-length molecular clones pHIV-1, pHIV-1-HS7totmut, pHIV-1-HS7mutB/Oct, pHIV-1-HS7mutB/PU, pHIV-1-HS7mutC, pHIV-1-HS7mutSp, pHIV-1-HS7mutD and pHIV-1-HS7mutCmutB/Oct described above were used to generate the other stocks of wild-type and mutant viruses (HIV-1, HIV-1-HS7totmut, HIV-1-HS7mutB/Oct, HIV-1-HS7mutB/PU, HIV-1-HS7mutC, HIV-1-HS7mutSp, HIV-1-HS7mutD and HIV-1-HS7mutCmutB/Oct). These DNAs (3 µg) were transfected into 293T cells by Lipofectamine 2000 (Invitrogen). At 48 h post-transfection, HIV-1 stocks were prepared from supernatants after filtration through a 0.45 µm pore-size membrane and stored at [–80°C. All viral stocks were quantified by determining p24 antigen concentration by an enzyme-linked immunosorbent assay (ELISA) (Innogenetics).
Viral infections
Infections were performed by incubating 0.5 x 106 cells with 50 ng of p24 protein of wild-type or mutant viruses (at 37°C for 2 h in 500 µl of culture medium). After infection, the cells were pelleted at 300 g, washed three times with 1 ml of culture medium, resuspended in 1 ml of standard medium, and grown under standard conditions. Every 2 or 3 days, aliquots of 200 µl were removed from the infected cultures and replaced by normal medium. The aliquots were assayed for p24 concentration in order to monitor the kinetics of viral replication.
Sequence analysis of HIV-1 genomic RNA
Viral particles from each stock were pelleted by ultracentrifugation (250 ng of p24 at 20 000 g for 2 h at 4°C) and digested with RNase-free DNase I (60 U/ml for 10 min at 4°C; Roche) in the presence of RNasin (40 U/ml; Promega) to remove contaminating DNA. HIV-1 genomic RNA was purified by using Trizol Ls reagent (Invitrogen). cDNA synthesis was performed by the Titan One Tube RT–PCR kit method (Roche Molecular Biochemicals). cDNAs were amplified by PCR with a 5' oligonucleotide primer corresponding to nt 4342–4364 (5'-CCAGCTGTGATAAATGTCAGCT-3') and a 3' primer corresponding to nt 4982–4961 (5'-TATTACTACTGCCCCTTCACC-3'). PCR fragments were subcloned into the vector pCR4 Blunt-TOPO (Zero Blunt TOPO PCR Cloning Kit; Invitrogen). After identification of recombinant clones, three inserts from each construct were sequenced with the BigDye terminator sequencing kit (Applied Biosystems). The nucleotide sequences of all three clones were identical and confirmed the presence of the original mutations.
RNase protection assays
HIV-1 genomic RNAs were detected by RNase protection analysis. An HIV-1-specific 32P-labeled antisense riboprobe was synthesized in vitro by transcription of pGEM23 (a gift from M. Laspia (21) with SP6 polymerase. For HIV-1 genomic RNA analysis, equivalent amounts of viral particles from each stock (6 µg of p24) were pelleted by ultracentrifugation (at 20 000 g for 2 h at 4°C) and viral RNAs were prepared using Trizol Ls reagent (Invitrogen). The RNase protection assays were performed with the RPA II kit (Ambion) according to the manufacturer's recommendations by using the HIV-1 riboprobe as described above. The protected RNA fragments were analyzed by electrophoresis through 6% urea polyacrylamide gels.
Western immunoblot analysis
HIV-1 lysates were prepared by ultracentrifugation of each virus stock and the pellets were resuspended in Laemmli buffer at a concentration of 37.5 ng of p24/µl. Lysates were heated at 95°C for 5 min, separated by electrophoresis on an 8% polyacrylamide gel, transferred onto a polyvinylene difluoride membrane, and probed with a 1:2000 dilution of purified human anti-HIV-1 IgG (NIH AIDS Research and Reagent Program, reagent no. 192 donated by Alfred Prince). A second antibody, horseradish peroxidase-conjugated goat anti-human IgG (Pierce) (diluted 1:3300) was used for enhanced chemiluminescence detection (Cell Signaling).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Localization of the DNA sequences required for factor binding to the HS7 sites B and C by substitution mutant analysis
In order to identify the nuclear factors leading to the formation of complexes B1/C1, B2/C2 and B3/C3, we performed an in silico search (using the TESS program) for potential transcription factor binding sites in the HS7 sites B and C. The numerous potential sites revealed by this in silico analysis prompted us to perform a systematic substitution-scanning mutational analysis of sites B and C in order to localize by in vitro studies the nucleotides important in these sites for formation of the retarded complexes B1/C1, B2/C2 and B3/C3.
Nine mutant site B oligonucleotides (named m1–m9) were prepared in which consecutive 3 bp regions of the wild-type site B were sequentially replaced with a 3 bp C/G-C/G-C/G sequence (Figure 2A). To examine the effects of these mutations on specific factor binding to site B, we have systematically analyzed the ability of the scanning mutants to inhibit the formation of complexes B1, B2 and B3 in competition EMSAs. If mutations are introduced at critical positions, then a decrease in the ability of the unlabeled mutant oligonucleotide to compete for factor binding will be detected by the increase in band intensity as compared to the competition with the wild-type site B. Mutations at non-essential bases are expected to demonstrate binding competition similar to the wild-type. The competition assays were performed using site B wild-type as probe, nuclear extracts from the A3.01 cell line and unlabeled wild-type and substitution mutant site B double-stranded oligonucleotides as competitors (Figure 2B). Phosphorimaging analyses were used to quantify the percentage of competition for formation of complexes B1 and B3 by each of the unlabeled oligonucleotides. The competition activities of mutant sites B m1–m9 were expressed relative to the competition activity of the wild-type site B, which was arbitrarily assigned a value of 100% of competition. Figure 2B shows a graphic representation of the quantification results for complex B1. Site B mutants m6, m7 and m8 exhibited the lowest percentages of competition (Figure 2B, 44.4, 34.1 and 32.7%, respectively), indicating that the base pairs which were substituted in these three double-stranded oligonucleotides, were the most important for complex B1 formation. Quantification of the bands corresponding to complex B3 indicated the importance of the same base pair (data not shown). Similar results were observed for complexes B1 and B2 when identical experiments were conducted with Raji nuclear extracts (Figure 2C). These results demonstrate that nucleotides between 4534 and 4542 in site B are critical for formation of all three complexes B1, B2 and B3, suggesting the presence of overlapping binding sites in site B or competition among factors for the same site.
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We next prepared seven scanning mutants of site C (named m1–m7), which sequentially replace 3 bp at a time of the entire site C sequence with a CCC sequence (Figure 2D). We tested the ability of this set of substitution mutants to inhibit formation of complexes C1, C2 and C3 in competition EMSAs using wild-type site C as probe, A3.01 nuclear extracts and the unlabeled wild-type and mutant double-stranded oligonucleotides as competitors (Figure 2E). Phosphorimaging analyses were used to quantify the percentage of competition for the formation of complexes C1 and C3 by each of the unlabeled oligonucleotides. The competition activities of mutant sites C m1–m7 were expressed relative to the competition activity of the wild-type site C, which was arbitrarily assigned to a value of 100% of competition. Figure 2E shows a graphic representation of the quantification results for complex C1. Site C mutants m3, m4 and m5 exhibited significantly reduced competition activities (Figure 2B, 55.4, 20.3 and 25.2%, respectively), indicating that the base pairs which were substituted in these three oligonucleotides were the most important for complex C1 formation. Quantification of the bands corresponding to complex C3 indicated the importance of the same base pair (data not shown). Similar results were observed for complexes C1 and C2 when identical experiments were conducted with Raji nuclear extracts (Figure 2F). These results demonstrate that nucleotides between 4687 and 4695 in site C are critical for the formation of all three complexes C1, C2 and C3, suggesting the presence of overlapping binding sites in site C or competition among factors for the same site.
In addition, we confirmed these results by complementary experiments in which a labeled probe containing site B was used and complex formation was competed by unlabeled oligonucleotides containing wild-type and mutant sites C, as well as by reverse experiments in which a labeled probe containing site C was used and complex formation was competed by unlabeled oligonucleotides containing wild-type and mutant sites B (data not shown), confirming that factors binding to site C have a DNA-binding specificity similar to that of factors binding to site B, except for PU.1. These data are in agreement with our previous work (7).
In conclusion, our results indicate that the DNA sequences identified here as critical for factors binding to sites B and C are located between nt 4534 and 4542 and between nt 4687 and 4695, respectively.
The transcription factors Oct-1 and Oct-2 bind to sites B and C
Based on the above systematic mutational analysis, we performed a new computer search for potential transcription factor binding sites, which was focused on the critical nucleotide sequences of sites B and C. This search revealed the presence of a putative octamer regulatory sequence both in site B and in site C (Figure 3A, nt 4532–4539 in site B and nt 4689–4696 in site C). These two putative octamer sequences presented three mismatches with respect to the Oct consensus. Oct-1 and Oct-2 along with Pit-1 and Unc-86 are founding members of the POU family of transcription factors. These members show homology in the domain responsible for specific DNA-binding, the so-called POU domain. While Oct-1 is ubiquitous, Oct-2 is predominantly expressed in B-cells, as well as in activated T-cells and in nervous system. However, B-cells appear to contain the highest amounts of Oct-2 (22,23).
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To determine whether transcription factors binding to sites B and C are related to Oct proteins, unlabeled double-stranded oligonucleotides were prepared and used as competitors in EMSAs. Site C probe was incubated with nuclear extracts from Jurkat cells in the absence or in the presence of different competitor double-stranded oligonucleotides: the unlabeled homologous oligonucleotide, an unlabeled heterologous oligonucleotide corresponding to site D and an unlabeled oligonucleotide corresponding to the octamer consensus sequence (Figure 3B). In the absence of competitor, two retarded complexes were detected: the ubiquitous complex C1 and the T-cell specific complex C3 (Figure 3B, lane 1). Formation of complex C1 was inhibited by a molar excess of unlabeled homologous oligonucleotide (Figure 3B, lanes 2–7), by the Oct consensus oligonucleotide (Figure 3B, lanes 14–19), but not by an oligonucleotide with an unrelated sequence containing site D (Figure 3B, lanes 8–13). Complex C3 was also out-competed by the homologous site C oligonucleotide and by the Oct consensus oligonucleotide (although to a lesser extent than complex C1) but not by the site D oligonucleotide. Similar competition experiments using site C as probe and nuclear extracts from Raji cells confirmed the above results for complex C1 and indicated that complex C2 was inhibited by an excess of unlabeled homologous oligonucleotide and by the Oct consensus, but not by the unrelated site D oligonucleotide (data not shown). We conclude from these experiments that the ubiquitous complex C1, the B-cell-specific complex C2 and the T-cell-specific complex C3 could contain a member(s) of the Oct proteins. Similar competition experiments performed with site B as probe supported the notion that formation of complexes B1, B2 and to a lesser extent B3 also resulted from binding of proteins belonging to the Oct family (data not shown).
To identify directly the factors present in the specific complexes B1/C1, B2/C2 and B3/C3, we performed supershift assays using antibodies directed against two individual members of the Oct family of transcription factors: the ubiquitously expressed Oct-1 protein and the lymphoid-specific Oct-2 protein. Labeled site C oligonucleotide probe was incubated with nuclear extracts from Jurkat or Raji cells in the presence of either an anti-Oct-1 antibody or an anti-Oct-2 antibody or purified rabbit IgG as control (Figure 3C). With Jurkat nuclear extracts, we observed, in the presence of the IgG control, the formation of the ubiquitous complex C1 and of the T-cell-specific complex C3 (Figure 3C, lane 1). Addition of the anti-Oct-1 antibody resulted in the complete disappearance of complex C1 (Figure 3C, lane 3), whereas the addition of the anti-Oct-2 antibody did not modify the migration profile of complexes C1 and C3 (Figure 3C, lane 5). With Raji nuclear extracts, we observed, in the presence of the IgG control, the formation of the ubiquitous complex C1 and the B-cell-specific complex C2 (Figure 3B, lane 2). The anti-Oct-1 antibody completely eliminated complex C1 (Figure 3C, lane 4) and complex C2 was eliminated by the anti-Oct-2 antibody (Figure 3C, lane 6). Similar results were obtained for complexes B1, B2 and B3 when identical experiments were conducted with site B as probe (data not shown).
In order to identify the protein(s) present in complexes B3/C3, we first tested, by supershift experiments, a series of protein candidates for potential transcription factor binding sites revealed by our in silico analysis: among others, the members of the MEF-2 transcription factor family. However, the latter assays did not provide any clue about the identity of the complexes B3/C3. Of note, the validity and/or the specificity of the different 
-MEF-2 (-MEF-2A, -2B, -2C and -2D) antibodies were established in separate experiments (data not shown). We also tried to purify complexes B3/C3 by affinity chromatography. However, despite many attempts, we did not succeed in the identification of these B3/C3 complexes. Additional purification and microsequencing studies are currently underway in our laboratory to identify the nature of the protein(s) present in complexes B3/C3.
From these data, we conclude that the ubiquitous complexes B1/C1 contain the Oct-1 transcription factor and that the B-cell-specific complexes B2/C2 contain the Oct-2 transcription factor. The transcription factor(s) present in the T-cell-specific complexes B3/C3 still remains (remain) to be identified.
The transcription factor PU.1 specifically interacts with site B of the HS7 region to form complex B4
Complex B4 was previously demonstrated by our laboratory to contain a protein related to the transcription factor PU.1 (7). Indeed, our laboratory has previously reported that a SV40 PU.1 consensus oligonucleotide inhibits specifically the formation of complex B4, whereas no competition of binding in complexes B1 and B2 was observed. We have also previously shown that the PU.1 protein, derived from lysates of Cos-1 cells transfected with a PU.1 expression vector, binds to site B in a PU box-dependent manner. Nevertheless, because proteins that bind DNA by means of an Ets domain share DNA recognition properties, we could not rule out the possibility that other Ets family members also bind to the PU box present in the HS7 site B (Figure 4A).
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To determine directly the presence of PU.1 in complex B4, we performed supershift assays. Preimmune serum or antiserum specific for PU.1 was incubated with nuclear extracts from U937 cells before the addition of the labeled probe corresponding to site B (Figure 4B). We observed that the addition of the anti-PU.1 antibody interfered with the formation of complex B4, leading to its disappearance (Figure 4B, lane 2). In contrast, the preimmune serum did not affect complex B4 formation (Figure 4B, lane 1), indicating the specificity of the protein–antibody interaction. These results indicated that the B4 complex contained PU.1. In addition, we also demonstrated that PU.1 protein produced in baculovirus bound to site B and induced the formation of a complex comigrating with complex B4 (Figure 4B, compare lanes 3 and 4).
In conclusion, the macrophage- and B-cell-specific transcription factor PU.1 specifically interacts with site B in the HIV-1 HS7 region. Moreover, the comigration of complex B4 with purified PU.1 protein and the complete disappearance of complex B4 when using the anti-PU.1 antiserum suggest that this complex does not contain other Ets proteins.
The transcription factors Sp1 and Sp3 specifically interact with the GC-box located in the HIV-1 HS7 region
We have previously identified a GC-box with close homology to the Sp1 consensus sequence at position nt 4623–4631 in the HS7 region and we have shown, by in vitro DNase I footprinting analysis, that this site binds affinity-purified human Sp1 protein (7). In order to further assess the presence of Sp1 and/or other Sp family members on this GC-box, we designed a double-stranded oligonucleotide, and designed site Sp wt (nt 4616–4638) encompassing the potential Sp site (Figure 5A). This oligonucleotide was radiolabeled and tested in EMSAs for DNA–protein interactions with nuclear extracts from Jurkat cells (Figure 5B). Two retarded protein–DNA complexes were observed. Similar results were obtained with nuclear extracts from the U937, Raji and A30.1 cell lines (data not shown). To evaluate the sequence specificity of the binding to the Sp wt probe, we performed competition EMSAs using increasing concentrations of different unlabeled double-stranded competitor oligonucleotides (Figure 5B). The specificity of the protein binding was demonstrated because their formation was inhibited by competition with molar excesses of the unlabeled homologous Sp wt oligonucleotide (Figure 5B, lanes 2–5), but not by the same molar excesses of a heterologous oligonucleotide corresponding to site B (Figure 5B, lanes 14–17). A Sp1 consensus oligonucleotide (named Sp1 cons) inhibited the formation of the retarded complexes even at lower concentrations than the homologous oligonucleotide (Figure 5B, lanes 6–9). In contrast, these complexes were not competed by a mutated version of the Sp1 consensus oligonucleotide (named Sp1 cons mut) containing a GG to TT substitution, thereby demonstrating the specificity of the retarded complexes to the HS7 Sp motif (Figure 5B, lanes 10–13). These results support the hypothesis that both complexes contain Sp family members.
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To identify directly the Sp family members within the two retarded complexes observed with the HS7 Sp wt probe, we performed supershift assays using specific antibodies directed against individual members of the Sp family of transcription factors (Figure 5C). The HS7 Sp wt probe was incubated with nuclear extracts from Jurkat cells and polyclonal antibodies directed against Sp1, Sp2, Sp3 or Sp4 were added to the binding reaction mixture (Figure 5C and data not shown). The
-Sp1 antibody selectively supershifted the major slower migrating complex (Figure 5C, lane 3) and the -Sp3 antibody resulted in the appearance of a supershifted complex and the corresponding disappearance of the faster migrating complex (Figure 5C, lane 4). We confirmed these results when both anti-Sp1 and anti-Sp3 antibodies were included in the same binding reaction (Figure 5C, lane 5). Similar relative mobilities of Sp1 and Sp3 EMSA complexes were reported in previous studies (24–27). In contrast, the binding pattern was not affected by the addition of the antibodies directed against the other Sp proteins (Sp2 and Sp4) (data not shown), showing that the two complexes did not seem to involve these other proteins. Moreover, the binding pattern was not affected by the addition of an unrelated antibody against MEF-2, used as a negative control (Figure 5C, lane 2). Overall, these results demonstrate that Sp1 and Sp3 transcription factors interact with the GC-box (renamed Sp site hereafter in the manuscript) located in the intragenic HS7 region of HIV-1.
Transcription factor(s) binding to site D seems (seem) to contain a zinc finger DNA-binding domain
Our laboratory has previously reported that an ubiquitously expressed factor(s) interacts (interact) specifically with site D (7). This site is well conserved among many HIV-1 isolates, but computer analysis has revealed no relevant homology between this binding site and the recognition sequences for known transcription factors (7). Methylation interference analysis of site D has identified the guanine residues that are important for binding: methylation of two guanine residues at positions 4830 and 4833 on the coding strand and of one guanine residue at position 4835 on the non-coding strand has been shown to strongly interfere with binding to site D (7).
In this study, we further characterized site D by EMSAs to identify the protein(s) present in complex D. Since many transcription factors are zinc finger proteins, the probe corresponding to site D was incubated with nuclear extracts from Jurkat cells in the absence or in the presence of increasing concentrations of a chelator of zinc, the 1,10-phenanthroline. Incubation of this site D probe resulted in one major ubiquitous retarded complex (labeled D) (Figure 6, lane 1) as previously reported by our laboratory (7). We observed that complex D disappeared in the presence of phenanthroline (Figure 6, lanes 2–5). The presence of increasing concentrations of ZnCl2 caused the restoration of complex D (Figure 6, lanes 6–9), whereas the presence of increasing concentrations of another ion (MgCl2) did not allow the reappearance of complex D (Figure 6, lanes 10–13).
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We conclude that the binding of the ubiquitously expressed protein(s) to site D is Zn2+-dependent. This observation suggests that this (these) protein(s) could contain zinc finger domain(s). However, despite many purification and protein sequencing attempts, we did not succeed to identify this (these) protein(s).
Tat expression does not affect the binding of the nuclear factors to the HS7 binding sites
To determine whether Tat can influence the binding of factors to the HS7 binding sites, we performed EMSAs, using nuclear extracts prepared from clonal Jurkat cell lines that stably expressed either the one-exon form of Tat [72 amino acids (Tat72)] (Figure 7, lane 2) or the two-exon form of Tat [101 amino acids (Tat101)] (Figure 7, lane 3) (8). Nuclear extracts from a clone transfected with the empty expression cassette vector were used as controls (Figure 7, lane 1). Both forms of Tat were used because Tat101 is expressed both early and late in the virus life cycle, while Tat72 is expressed solely in the late phase (28). Probes corresponding to HS7 site B, Sp site, site C and site D were incubated with these nuclear extracts. No difference in binding activity between the Tat72 and the Tat101 clones and the control clone was noted when the four probes were used (Figure 7, compare lanes 2 and 3 with lane 1).
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We conclude from these in vitro experiments that Tat has no effect on nuclear factor binding to the HS7 intragenic region.
Sp1, Sp3, Oct-1 and PU.1 are recruited to the HS7 regulatory region in vivo
To demonstrate in vivo in the context of chromatin the relevance of our in vitro binding studies, we performed chromatin ChIP assays using both the lymphoid HIV-infected ACH2 cell line and the monocytic HIV-infected U1 cell line. Formaldehyde cross-linked chromatin from these cell lines was used for immunoprecipitation with antibodies directed against Sp1, Sp3, Oct-1 or PU.1 or a purified rabbit IgG as negative control. Following reverse of the cross-link, the purified DNA was subjected to PCR analysis using a set of primers flanking the HS7 region (Figure 8). In ACH2 cells, Oct-1, Sp1 and Sp3 binding to the HS7 region was detected (Figure 8A, lanes 1, 2 and 3, respectively), whereas, as expected, immunoprecipitations of the cross-linked chromatin with the anti-PU.1 antibody or with the purified rabbit IgG gave no signal (Figure 8A, lanes 4 or 5, respectively). In U1 cells, analysis of PCR products from immunoprecipitated DNA showed significant enrichment of the HS7 region when immunoprecipitation was carried out with the anti-Oct-1, anti-Sp1, anti-Sp3 or anti-PU.1 antibodies (Figure 8B, lanes 1, 2, 3 or 4, respectively). In contrast, no such enrichment was observed following immunoprecipitation of the cross-linked chromatin with the purified rabbit IgG (Figure 8B, lane 5). As a control, we used another set of primers flanking a region of the pol gene located 2 kb upstream of the HS7 region that has not been reported so far as binding any of these transcription factors. Immunoprecipitations with all the antibodies did not enrich eluates with DNA from this control region in both ACH2 and U1 cell chromatin, demonstrating the specificity of the HS7 interactions (data not shown).
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These data, thus, demonstrate the occupancy in vivo of the HS7 region by Oct-1, Sp1 and Sp3 in both ACH2 and U1 cell lines as well as, consistently, the binding of PU.1 to the HS7 only in the U1 cells.
Identification of point mutations abolishing factor binding to the DNA motifs in the HIV-1 HS7 region
To further characterize physically the DNA motifs located in the HS7 region of the pol gene, we studied by using EMSA the effect of selected mutations on binding affinity. Point mutations were designed to abolish binding of factors to their respective sites without modifying the underlying amino acid sequence of the integrase.
- Site B: We demonstrated that Oct-1, Oct-2 and PU.1 bound to site B. To abolish binding of Oct-1 and Oct-2, three adenine residues at positions 4535, 4538 and 4541 were substituted with guanine residues (Figure 9A). The oligonucleotide corresponding to this mutation was designated site B mutOct. To abolish PU.1 binding to site B, two cytosine residues at positions 4526 and 4529 were substituted with thymine residues in the site B PU box (Figure 9A). The oligonucleotide corresponding to this mutation was designated site B mutPU.1. The effect of these selected mutations was analyzed on binding affinity by using EMSAs. The wild-type and the two mutated site B oligonucleotides were used as probes and were incubated with nuclear extracts from U937 (Figure 9A), Raji and Jurkat (data not shown) cells. Figure 9A demonstrated the lack of Oct-1 binding to the site B mutOct oligonucleotide probe (lane 2) and the lack of PU.1 binding to the site B mutPU.1 oligonucleotide probe (lane 3). Similar experiments using Raji nuclear extracts showed the lack of Oct-2 binding to the site B mutOct oligonucleotide and confirmed the lack of PU.1 binding to site B mutPU.1 oligonucleotide probe (data not shown). Similar EMSAs using Jurkat nuclear extracts confirmed the lack of Oct-1 binding and showed the lack of T-cell-specific binding to the site B mutOct oligonucleotide probe (data not shown). Taken together, our results identify an octamer motif and a PU box in the HS7 site B. The octamer motif specifically binds the octamer proteins Oct-1 and Oct-2 in vitro. The PU box specifically binds the Ets protein PU.1. We report a 3 bp mutation, referred to as site B mutOct, that abrogates the binding of Oct-1 and Oct-2 and binding of the T-cell-specific factor to site B and a 2 bp point mutation, referred to as site B mutPU.1, that abrogates PU.1 binding to site B.
- Sp site: We showed that Sp1 and Sp3 bound to the HS7 GC-box. Two guanine residues at positions 4629 and 4632 were substituted with two adenine residues in this motif (Figure 9B). The wild-type and the mutated oligonucleotide (referred to as site Sp wt and site Sp mut, respectively) were used as probes and incubated with nuclear extracts from U937 cells. Figure 9B showed the lack of Sp1 and Sp3 binding to the site Sp mut oligonucleotide probe, thereby demonstrating that the selected 2 bp mutation abolished Sp binding to the HS7 Sp site.
- Site C: We demonstrated that Oct-1, Oct-2 and a T-cell factor(s) bound to the HS7 site C. To abolish factor binding to this site, two thymine residues at positions 4688 and 4694 were substituted with cytosine residues in the octamer sequence of site C (Figure 9C). The effect of this 2 bp mutation on binding affinity was analyzed by EMSAs. The wild-type and the mutated oligonucleotides (referred to as site C wt and site C mutOct, respectively) were used as probes and incubated with Raji (Figure 9C) or Jurkat nuclear extracts (data not shown). Figure 9C showed the lack of Oct-1 and Oct-2 binding to the site C mutOct oligonucleotide probe. Similar EMSAs using Jurkat nuclear extracts confirmed the lack of Oct-1 binding and showed the lack of T-cell-specific binding to the same mutated probe (data not shown). These results demonstrated that the selected point mutations introduced in site C abolished both the binding of Oct-1 and Oct-2 and the binding of the T-cell-specific factor.
- Site D: To abolish binding of nuclear factors to site D, we substituted one adenine residue at position 4832 for a cytosine residue and one cytosine at position 4835 for a thymine residue (Figure 9D). The adenine residue (nt 4832) is located between two guanine residues on the coding strand (nt 4830 and 4833), which were previously demonstrated by our laboratory to be critical for complex D formation by methylation interference (7). The cytosine residue (nt 4835) corresponds to the guanine residue previously identified on the non-coding strand by the same technique as critical for complex D formation. The wild-type and the mutated oligonucleotides (referred to as site D mut) were used as probes in EMSAs with U937 nuclear extracts. Figure 9D demonstrated that the selected 2 bp mutation abolished formation of complex D.
The HS7 binding sites are involved in the transcriptional activity of the pol gene region
Our laboratory has previously shown using chloramphenicol acetyltransferase (CAT) transient expression plasmids that a 500 bp fragment encompassing the HS7 region (nt 4481–4982) exhibited transcription enhancing activity (2-fold) when it was cloned in its natural position with respect to the HIV-1 promoter in U937 and CEM cells in the presence of Tat. However, in the absence of Tat, the CAT values were too low to detect any enhancing activity of the HS7 region.
In order to address the potential functional role of the HS7 region in the basal (Tat-independent) activity of the HIV-1 promoter, we used the more sensitive luciferase transient expression system. We subcloned the 500 bp HS7 fragment into the construct pLTR in the sense orientation with respect to the transcriptional unit, downstream of the luciferase reporter gene, thereby generating the pLTR-HS7wt. The pLTR construct contains the complete HIV-1 LAI 5'-LTR (plus the leader sequence up to the ATG of gag) driving the expression of the luciferase gene. The constructs pLTR and pLTR-HS7wt were transiently transfected into Jurkat cells (Figure 10A) or U937 cells (Figure 10B). The reporter constructs were cotransfected into the cells with pRL-TK, and used as an internal control to measure the transfection efficiency. At 44 h post-transfection, cells were lysed and assayed for luciferase activity. In Jurkat cells, transfection of plasmid pLTR-HS7wt caused a 1.58-fold increase in luciferase activity compared with that of the control plasmid pLTR tested under the same conditions (Figure 10A). In U937 cells, transfection of pLTR-HS7wt caused a 1.98-fold increase in luciferase activity compared with that of pLTR (Figure 10B). These results indicate that, in the absence of Tat, the region associated with the intragenic HS7 shows a weak transcriptional enhancing activity when it is cloned in a position similar to that observed within the viral genome (downstream and in sense orientation relative to the 5'-LTR-luc transcriptional unit).
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In order to determine the relative contribution of each factor binding sites (site B, site C, Sp site and site D) to the HS7 enhancing activity, the point mutations identified in EMSAs (Figure 9) were introduced individually or in combination in the context of the pLTR-HS7wt plasmid. The mutated plasmids were designed pLTR-HS7totmut, pLTR-HS7mutSp, pLTR-HS7mutD, pLTR-HS7mutB/PU, pLTR-HS7mutB/Oct, pLTR-HS7mutC and pLTR-HS7mutCmutB/Oct. These plasmids were assayed for luciferase activity after transient transfection in Jurkat or U937 cells (Figure 10A or B, respectively). Remarkably, in both cell lines, transfection of plasmid pLTR-HS7totmut exhibited luciferase activity similar to that obtained with the control vector pLTR, thereby demonstrating that the enhancing effect observed with pLTR-HS7wt required intact site B, site C, Sp site and site D motifs. Moreover, in both cell lines, transfection of the plasmids containing the individual mutations presented luciferase activities similar to that obtained with the wild-type pLTR-HS7wt construct. In conclusion, mutations in the HS7 binding sites in combination abolish the transcription-enhancing activity of the pol HS7 region, suggesting that these sites are responsible for most of this activity. However, we could not demonstrate the functional role played by one individual site in the HS7 enhancing activity (Figure 10A and B).
We next studied the effect of mutating the HS7 binding sites on the response of the HIV-1 promoter to Tat. We observed that the pol HS7 region exhibited transcriptional enhancing activity in the presence of Tat, in agreement with our previous results (7), and that mutations in all the HS7 binding sites in combination abolished this activity (data not shown).
Our functional results, thus, demonstrate a positive regulatory role of the pol HS7 region in both Tat-independent and Tat-dependent HIV-1 promoter-driven gene expression. These data indicate that the loss of transcriptional enhancing activity caused by mutations in the HS7 binding sites correlated with the loss in factor binding to these sites. Moreover, mutation of each HS7 binding site individually did not affect the enhancing activity of the intragenic positive regulatory region.
Functional analysis of the individual HS7 region binding sites by overexpression assays
We next wanted to further characterize the functionality of each HS7 site individually (sites for which we were able to identify by the above experiments the bound transcription factors) and to examine whether these transcription factors (PU.1, Sp1, Sp3, Oct-1 and Oct-2) act through the HS7 binding sites. Because these transcription factors present distinct cell-specific expression, we decided to study each site separately by overexpression assays using for each individual factor a cell line which does not express the factor considered. To this end, we produced a series of artificial luciferase reporter constructs in which multimerized copies of each wild-type and mutated HS7 binding sites were inserted upstream of the HSV TK minimal promoter into the pTK construct. The resulting constructs were cotransfected with expression vectors for the various transcription factors (PU.1, Sp1, Sp3, Oct-1 and Oct-2) and assayed for luciferase activity. The minimal TK promoter was used instead of the HIV-1 promoter (LTR) because the latter is known to contain binding sites for Oct, Ets and Sp family members (29) and could have therefore complicated the interpretation of the transfection results. In these experiments, we used multimerized HS7 binding sites because multimerization of a transcription factor binding site allows the amplification of its transcriptional effect.
Ectopic expression of the Oct-1 and Oct-2 transcription factors down-regulates TK promoter activity through multimerized sites B and C
To determine whether the HS7 sites B and C can act as independent regulators of transcription, we constructed four luciferase reporter plasmids driven by the HSV TK minimal promoter with three tandem repeats of the wild-type and Oct-mutated site B or site C inserted upstream of the TK promoter. These four plasmids were referred to as p(Bwt)3TK, p(BmutOct)3TK, p(Cwt)3TK and p(CmutOct)3TK, respectively. To examine the response of these reporter constructs to the POU-homeodomain transcription factors Oct-1 and Oct-2, murine NIH3T3 fibroblats were transiently cotransfected with each of them and increasing amounts of either the Oct-1 or the Oct-2 expression vector (Figure 11A and B, respectively) and then assayed for luciferase activity. The murine NIH3T3 cell line was used in these experiments because our EMSAs performed with NIH3T3 nuclear extracts and with probes corresponding to site B or site C revealed no binding activity corresponding to complexes B1/C1, B2/C2, B3/C3 and even no binding at all (data not shown).
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As shown in Figure 11, the control pTK construct was repressed by Oct-1 up to 2.22-fold (i.e. by 55%) and by Oct-2 up to 2.12-fold (i.e. by 53%). Cotransfection of the reporter construct p(Bwt)3TK with the Oct-1 or Oct-2 expression vectors resulted in a dose-dependent decrease in luciferase activity by ectopically expressed Oct-1 (up to 5.00-fold) and Oct-2 (up to 5.88-fold); thus, representing a 2.3- and 2.8-fold down-regulation when compared with the Oct-1 and Oct-2 responses of the control pTK construct devoid of upstream HS7 sites B. This effect required intact Oct motifs in site B, because mutations in these motifs [p(BmutOct)3TK] resulted in levels of Oct-1- and Oct-2-mediated repression similar to those obtained with the control pTK (Figure 11A and B, respectively). Similar results were obtained when examining site C. Indeed, addition of three copies of the HS7 site C upstream of the TK promoter [p(Cwt)3-TK] resulted in a dose-dependent decrease in luciferase activity by ectopically expressed Oct-1 (up to 9.09-fold) and Oct-2 (up to 5.26-fold), thus representing a 4.1-fold and 2.5-fold down-regulation when compared with the Oct-1 and Oct-2 responses of the control pTK devoid of upstream HS7 sites C (Figure 11A and B, respectively). This effect also required intact Oct motifs in site C [see p(CmutOct)3-TK in Figure 11A and B].
We conclude from these experiments that ectopic Oct-1 and Oct-2 proteins have a site B- or site C-dependent inhibitory effect on the heterologous TK promoter containing multiple upstream site B or site C, respectively. These results, thus, demonstrate that the HS7 sites B and C function as negative regulatory elements in response to ectopic Oct-1 or Oct-2 proteins in a heterologous context, suggesting that Oct-1 and Oct-2 might be negatively involved in the HS7 transcriptional activity.
Ectopic expression of the PU.1 transcription factor up-regulates TK promoter activity through multimerized site B
In order to define the potential role of PU.1 through the PU box of the HS7 site B, we generated a mutant derivative of the p(Bwt)3TK construct with the upstream insertion of three tandem repeats of a site B mutated in the PU box sequence. This derivative was designated p(BmutPU.1)3TK. The constructs pTK, p(Bwt)3TK and p(BmutPU.1)3TK were cotransfected with increasing amounts of the PU.1 expression vector pJ6-PU.1 into PU.1-negative NIH3T3 cells (Figure 12). Transfected cells were assayed for luciferase activity. R