Characterization of a human immunodeficiency virus type 1 pre-integration complex in which the majority of the cDNA is resistant to DNase I digestion

Dheeraj K. Khiytani1 and Nigel J. Dimmock1

Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, UK1

Author for correspondence: Nigel Dimmock. Fax +44 2476 523568. e-mail ndimmock{at}bio.warwick.ac.uk


   Abstract
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Abstract
Introduction
Methods
Results
Discussion
References
 
The human immunodeficiency virus type 1 (HIV-1) pre-integration complex (PIC) is a cytoplasmic nucleoprotein structure derived from the core of the virion and is responsible for reverse transcription of viral RNA to cDNA, transport to the nucleus and integration of the cDNA into the genome of the infected target cell. Others have shown by Mu phage-mediated PCR footprinting that only the LTRs of the cDNA of PICs isolated early in infection are protected by bound protein, while the rest of the genome is susceptible to nuclease attack. Here, using DNase I footprinting, we confirmed that the majority of the cDNA of PICs isolated at 8·5 h after infection with cell-free virus was sensitive to digestion with DNase I and that only part of the LTRs (approximately 6% of the total cDNA) was protected. However, PICs isolated 90 min later (at 10 h post-infection) were very different in that the majority (approximately 90%) of cDNA was protected from nuclease degradation. These late PICs were integration active in vitro. We conclude that HIV-1 has at least two types of PIC, an early PIC characterized by protein bound only at the LTRs, and a late, and possibly more mature form, in which protein is bound along the length of the cDNA.


   Introduction
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Abstract
Introduction
Methods
Results
Discussion
References
 
The human immunodeficiency virus type 1 (HIV-1) virion is bounded by a lipid bilayer into which is inserted the envelope protein, a homotrimer of gp120–gp41, and a variety of host proteins. Internal to this is the matrix (MA) protein, which surrounds the capsid (CA) protein core structure. In turn this encloses two identical single-stranded, plus-sense viral RNA strands, associated with nucleocapsid (NC), reverse transcriptase (RT) and integrase (IN) proteins. Closely associated with the core are the Vif and Nef proteins. The viral accessory protein, viral protein R (Vpr), is also found within the virion but is most probably located outside the core. Tat may also be located inside virions. Infection is initiated by gp120 attaching to specific target cells via a primary receptor, the CD4 protein and a co-receptor, usually CCR5 or CXCR4 (for a review, see Levy, 1998 ). As a result of these interactions the viral lipid bilayer fuses with that of the plasma membrane. Following fusion, it is hypothesized that the virus core develops into an immature reverse transcription complex (Zennou et al., 2000 ), and in turn this matures into a better defined nucleoprotein structure, termed the pre-integration complex (PIC) (Farnet & Haseltine, 1990 ; Karageorgos et al., 1993 ). These are present in the cytoplasm. PICs were first observed in experiments with murine leukaemia virus (MLV) (Bowerman et al., 1989 ; Farnet & Haseltine, 1991 ; Fujiwara & Mizuuchi, 1988 ) when detection of viral cDNA was combined with a functional assay for integration. HIV-1 PICs are derived from the virion core structures and contain IN, RT, Vpr, the cellular high mobility group protein HMG 1(Y) and a reduced amount of MA (Bukrinsky et al., 1993b ; Ellison & Brown, 1994 ; Farnet & Bushman, 1997 ; Farnet & Haseltine, 1991 ; Gallay et al., 1995b ; Karageorgos et al., 1993 ; Miller et al., 1997 ). All these proteins are associated with the viral RNA/cDNA. It is not clear if other internal virion proteins are present. PICs sediment on sucrose velocity gradients between 160S and 640S after treatment with RNase (Bowerman et al., 1989 ; Farnet & Haseltine, 1990 ; Karageorgos et al., 1993 ; Miller et al., 1997).

The RT protein associated with the PIC reverse transcribes the virion RNA (for a review, see Brown, 1997 ). PICs are also responsible for transport of newly synthesized viral cDNA into the nucleus and integration of the cDNA into the target cell genome (Bukrinsky et al., 1992 ; Bushman et al., 1990 ). Vpr has been implicated in the targeting of PICs to the host cell nucleus but has not been shown directly to be part of the complex (Gallay et al., 1996 ; Heinzinger et al., 1994 ; Jenkins et al., 1998 ; Karni et al., 1998 ; Popov et al., 1998a , b ; Vodicka et al., 1998 ; Zhang et al., 1998 ). MA is also thought to contribute to nuclear targeting of the viral cDNA (Bukrinsky et al., 1993a ; Gallay et al., 1995b , 1996 ; Heinzinger et al., 1994 ; Popov et al., 1998b ; von Swedler et al., 1994 ), although there is also evidence that this may not be the case (Fouchier et al., 1997 ). Phosphorylation of MA may be important for its karyophilic properties (Bukrinskaya et al., 1996 ; Gallay et al., 1995a , b ). The viral IN also appears to have a role in nuclear targeting (Gallay et al., 1997 ; Pluymers et al., 1999 ) and possibly reverse transcription (Wu et al., 1999 ), in addition to its major activity of integrating viral cDNA into the host cell genome (Bushman et al., 1990; Ellison & Brown, 1994 ; Farnet & Haseltine, 1990 ). Recently, it has been shown that a central DNA flap (a 99 nucleotide plus-strand overlap created during HIV-1 reverse transcription at the boundary at which left- and right-hand segments of nascent plus-strand cDNA merge), acts as a cis-determinant of HIV-1 DNA nuclear import (Zennou et al., 2000 ).

Much has been done to determine the protein composition and in vitro function of PICs through the use of immunoprecipitation, integration assays, nuclear import assays and, to a certain extent, nuclease protection assays. However, less is known about the formation and maturation of PICs especially in regard to nuclear targeting, translocation and integration of viral cDNA with the host genome. Although the exact conformation of PICs is not well understood, experiments have shown that the ends of the cDNA may be joined through dimerization by the viral IN protein or the cell HMG 1(Y) protein (Ellison & Brown, 1994 ; Farnet & Bushman, 1997 ; Miller et al., 1997). Recently, the protein–DNA structure of PICs partially purified from HIV-1-infected cells at 8·5 h after infection has been analysed by Mu-mediated PCR footprinting (Chen et al., 1999). The HIV-1 cDNA termini (LTRs), but not the rest of the genome, were protected from nuclease attack by bound protein. The termini form a unique structure, resembling the ends of MLV PIC DNA (Wei et al., 1997 ), which suggests that this may be a common feature of retrovirus PIC DNAs.

The consensus thus far is that HIV-1 PICs contain condensed cDNA in a complex resembling a partially dissociated viral core in which proteins are tightly associated with the ends of the cDNA but only loosely associated, if at all, with the intervening sequence (Chen et al., 1999 ; Farnet & Bushman, 1997 ). The work described in this report found an almost identical structure at 8·5 h post-infection using the different technique of DNase I footprinting (protease digestion followed by DNase digestion). However, PICs isolated in exactly the same way at 10 h post-infection were almost completely resistant to DNase I digestion. These late PICs were active and integrated HIV-1 cDNA into phage DNA in vitro. Thus late PICs have a complex protein–DNA structure, are functionally integrative and may represent a more mature structure than PICs present at an earlier time.


   Methods
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Abstract
Introduction
Methods
Results
Discussion
References
 
{blacksquare} Cells, viruses and infection.
C8166 (Salahuddin et al., 1983 ) and H9 (Popovic et al., 1984 ) CD4+ T cell lines were obtained via the Centralized Facility for AIDS Reagents, NIBSC, Potters Bar, UK, and maintained in RPMI 1640 medium (Gibco BRL Life Technologies) containing 10% foetal calf serum (FCS; Helena BioSciences) and 2 mM L-glutamine (Gibco). Virus was prepared by co-cultivating H9 cells chronically infected with HIV-1 IIIB with non-infected H9 cells, at a ratio of 1:4. After 2 days the tissue culture fluid was harvested, clarified and virus concentrated by pelleting through 20% sucrose. Infectivity was determined by syncytium formation on C8166 cells (McLain & Dimmock, 1994 ).

{blacksquare} Antibodies.
We used the following antibodies: mAb H12-5C to the HIV-1 CA protein (B. Chesebro and K. Wherly); rabbit antiserum to Vpr (V. Ayyavoo and D. Weiner) – both obtained from the NIH AIDS Research and Reference Reagent Program, Rockville, USA; mAb 4H2B1 to MA (R. B. Ferns and R. B. Tedder); mAb IIG10E6 to RT (D. Helland and A. M. Szilvay); rabbit antiserum to IN (S. Ranjbar) – all obtained from the Centralized Facility for AIDS Reagents.

{blacksquare} Preparation of PICs.
PICs were prepared essentially as described previously (Farnet & Haseltine, 1990 ). Infection was initiated by mixing 108 syncytium-forming units of cell-free virus with 5x107 C8166 cells in 3 ml medium at 37 °C. After 5 h, 5 ml of fresh medium was added. Cells harvested at the required time were washed twice in buffer K (20 mM HEPES, pH 7·4, 5 mM MgCl2, 150 mM KCl, 1 mM dithiothreitol and 20 µg/ml aprotinin) and lysed in buffer K containing 1% Triton X-100 for 30 min at 20 °C. The Triton X-100 concentration was optimized by cell fractionation and monitored by phase-contrast microscopy. Nuclei and cell debris were removed by successive centrifugations at 1000 g for 3 min and 8000 g for 10 min. The resulting supernatant (cytoplasmic extract) was treated with 20 µg/ml RNase A for 30 min at 20 °C, and centrifuged on a 5 ml, 15–30% sucrose gradient for 105 min at 149000 g (Farnet & Haseltine, 1991 ). Sixteen fractions were collected from the top of the gradient. Putative PICs containing viral cDNA as judged by PCR were located in fractions 8–10. These co-sedimented with a 160S cowpea mosaic virus marker. To investigate viral cDNA in the infected cell nuclei, the 1000 g nuclear pellet (see above) was washed in buffer solution and Dounce homogenized in 300 µl buffer solution. Disruption of nuclei was monitored by phase contrast microscopy. Lysed nuclear extracts were proteinase K treated (1 mg/ml; Roche Molecular Biochemicals, PCR grade) for 1 h at 55 °C. DNA was phenol–chloroform extracted and recovered by precipitation with ethanol overnight at -20 °C. PCR involved preheating in a Touchdown PCR instrument (Hybaid) at 94 °C for 3 min, denaturing at 94 °C for 45 s, annealing at 56 °C for 45 s and an elongation step at 72 °C for 1 min. A final elongation step at 94 °C for 10 min was performed. DNA was subjected to PCR for 30 cycles. PCR products were analysed by electrophoresis on 3% agarose gels. Experiments were carried out in duplicate on different lots of infected cells.

{blacksquare} Immunoprecipitation of PICs from sucrose velocity gradient fractions.
Fifty µl of sucrose gradient fractions containing detectable 160S cDNA were incubated overnight at 4 °C with 10 µl of monoclonal antibodies against RT, MA, CA at 10 µg/ml or 10 µl of polyclonal antibodies (1/300) against IN or Vpr. Next antibody–antigen complexes were collected for 2 h at room temperature using either protein A or G Sepharose beads (Sigma–Aldrich) in their respective binding buffer (50 mM Tris, 150 mM NaCl, pH 8·0 or 0·01 M NaH2PO4, 0·15 M NaCl, 0·01 M EDTA, pH 7·0). Beads with bound complexes were then washed three times in wash buffer (10 mM Tris–HCl, pH 7·4, 150 mM NaCl and 1% Triton X-100). These were then digested with proteinase K at 1 mg/ml for 1 h at 55 °C to release immunoprecipitated complexes from the Sepharose beads and the beads were removed by centrifugation. The supernatant containing any viral cDNA was phenol–chloroform extracted, ethanol precipitated and amplified by PCR as described above using the GAG2 primer pair (see below and Table 1) and 30 cycles of amplification.


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Table 1. Summary of the 23 regions of viral cDNA amplified and sizes of PCR products

 
{blacksquare} Integration assay.
Cytoplasmic extract (300 µl), purified PICs or deproteinized PICs were incubated with 500 ng of linearized {phi}X174 phage DNA (Sigma) for 45 min at 37 °C. Reactions were stopped by treatment with proteinase K (1 mg/ml) for 1 h at 55 °C, and DNA extracted and recovered by precipitation with ethanol. Integration was assessed by PCR (Pryciak & Varmus, 1992 ) with an HIV-1 cDNA-specific forward primer and a reverse primer that recognized phage DNA. The advantage of this assay is that there can be no PCR product until HIV-1 cDNA has integrated. Various primer pairs were tried with 30 cycles of PCR, and one using a forward primer from HxB2, a molecular clone of HIV-1 (8953 GCTGCTTGTGCCTGGCTAGA 8972) and a reverse primer from {phi}X174 (144 AGCTGCGCAAGGATAGGTCG 163), gave a product of approximately 900 bp. This does not argue for a single integration site, but rather that there is a bias to certain integration sites, as others have shown (Bo et al., 1996 ). In the integration reaction the viral integrase cuts the host DNA in an exchange reaction that covalently links each viral strand to phage DNA at one end, but not the other, where there is a gap (Whitcomb & Hughes, 1992 ). Thus repair of the gap is not needed for a successful PCR reaction. PCR products were analysed as above and partially sequenced. Experiments were done in duplicate with different lots of infected cells.

{blacksquare} DNase I footprinting.
PICs were treated with DNase I (RNase free; Sigma–Aldrich) at 0·1, 1, 10 and 30 µg/ml for 30 min at 37 °C, and then reacted with proteinase K (1 mg/ml) at 55 °C for 1 h to remove protein. DNA was phenol–chloroform extracted and precipitated with ethanol overnight at -20 °C. PCR was then carried out for 24, 26, 28 and 30 cycles using 23 primer pairs designed by Primer Designer for Windows (Version 3.0; Scientific and Educational Software) from the sequence of HxB2. These covered the entire HIV-1 genome in approximately 500 bp overlapping fragments (Table 1). PCR conditions were optimized for each primer pair so that each had specific MgCl2 and primer concentrations in the PCR mix. Positive controls were amplified following proteinase K treatment (1 mg/ml) but in the absence of DNase I and negative controls (-) were amplified following proteinase K and then DNase I treatment at 0·1 µg/ml. Footprinting was carried out twice on PIC preparations from different batches of infected cells.


   Results
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Abstract
Introduction
Methods
Results
Discussion
References
 
Time-course of synthesis of 160S HIV-1 PIC cDNA, accumulation of viral cDNA in the cell nucleus and virion proteins associated with PIC cDNA
Viral cDNA was not detected in 160S PIC cDNA from cytoplasmic extracts at 2·5 and 5 h post-infection but was clearly present at 7·5, 8·5 and 10 h post-infection (Fig. 1a). However, the 12·5 h sample contained little amplifiable material. In nuclei, cDNA was first detected as a weak band at 10 h post-infection and there was a very strong band present at 12·5 h (Fig. 1b). The experiment was repeated with a new batch of infected cells with very similar results (data not shown). Absence of cDNA in the nucleus at 8·5 h suggests that there was no significant contamination of nuclei with cytoplasm and the absence of cytoplasmic cDNA at 12·5 h suggested there was little nuclear contamination of cytoplasm. No cDNA was detected in any of the nuclear washes (data not shown). The co-precipitation of cDNA by antibodies to various virion proteins incubated with 160S PICs isolated at 10 h post-infection is shown in Fig. 1(c). There were PCR products after immunoprecipitation with antibodies specific for RT, IN, MA and Vpr proteins but not with antibodies specific for CA or the influenza NP protein or in the absence of antibody. Similar data were obtained also with 8·5 h PICs (data not shown). Although there are slight differences in kinetics, the above data indicate that our PICs have properties similar to those described in the literature by others (Li & Burrell, 1992 ).



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Fig. 1. Time-course and cellular location of HIV-1 cDNA isolated from infected C8166 cells and the detection of PIC-associated virion proteins. (a) Detection of cDNA present in gradient-purified 160S PICs. (b) Detection of cDNA present in nuclear extracts from the same cells. Samples were harvested at the times shown post-infection (h). (c) Detection of cDNA complexed with virion proteins in 160S gradient-purified PICs from 10 h cytoplasmic extracts after immunoprecipitation of PICs with antibodies to RT, IN, MA, Vpr and CA. In all panels cDNA was detected using PCR, the GAG2 primer pair, and 30 cycles of amplification. PCR products were analysed by agarose gel electrophoresis and visualized by ethidium bromide staining. The arrow represents a DNA marker of 506 bp. In panel (c) immunoprecipitated complexes were captured and then eluted from protein G– or protein A–Sepharose beads. DNA was then extracted with phenol–chloroform, ethanol precipitated, and amplified by PCR. C8+, C8-, unfractionated cytoplasmic extracts from HIV-1-infected and uninfected C8166 cells respectively; Mo, Ra, immunoprecipitation mix with normal mouse or rabbit antiserum replacing virus-specific mouse or rabbit antibody respectively; IgG1, immunoprecipitation mix with influenza NP-specific IgG1; PG, PA, immunoprecipitation mix with protein G– or protein A–Sepharose respectively but no antibody; PCR, PCR mix only. Each experiment was repeated at least twice.

 
Integration activity of 160S PICs
We next determined the integration activity of 160S purified PICs isolated at 2·5, 5, 7·5, 8·5 and 10 h post-infection into {phi}X174 phage DNA. Preliminary experiments suggested that there were multiple integration sites, but that one particular site was favoured (see Methods). Thus we could use PCR with specific primers for HIV-1 DNA and for {phi}X174 phage DNA to demonstrate that integration had taken place, as there could be no product until the cDNA had integrated. This approach takes less time than Southern blotting and was consistently reproducible. Fig. 2 shows that there was no PCR product at 2·5 and 5 h post-infection, and that a single band, approximately 900 bp, was first detected at 7·5 h post-infection. Much stronger PCR bands were evident at 8·5 and 10 h post-infection (Fig. 2). There was no PCR product using deproteinized PICs, or when PICs and phage DNA were held together at 4 °C. The 900 bp PCR product was excised from the gel and partially sequenced using the HIV-1 and {phi}X174 PCR primers. These gave approximately 250 bp of sequence with 93% similarity with the HxB2 sequence and a similar length of sequence with 97% phage similarity respectively (data not shown). This confirmed that HIV-1 cDNA had integrated into the phage DNA and thus that purified PICs isolated between 7·5 and 10 h post-infection were integration active. This activity corresponded well with the appearance of PIC cDNA described in Fig. 1(a).



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Fig. 2. In vitro integration activity of PICs. PICs were purified on sucrose velocity gradients from cytoplasmic extracts of HIV-1-infected C8166 cells at the times shown post-infection (h). Purified 160S PICs or 160S PICs deproteinized with proteinase K were incubated with 500 ng of linearized {phi}X174 phage DNA for 45 min at 37 °C. After DNA extraction, samples were subjected to PCR for 30 cycles using a primer that recognized HIV-1 and another that recognized {phi}X174 DNA, so that no product was obtained without integration. The arrow indicates a DNA marker of 1018 bp. PCR products were analysed by electrophoresis on agarose gels. Similar data were obtained from a second experiment using a different batch of infected cells.

 
DNase footprinting of PICs harvested at 8·5 h post-infection shows that only part of the LTR is protected
The extent of protein–DNA interactions within PICs was determined by DNase footprinting across the whole HIV-1 genome, using PCR to indicate which regions were protected. Since PCR can be variable, we minimized primer-to-primer variation and stringency by optimizing PCR conditions with viral cDNA, so that all 23 primer pairs gave an amplicon of approximately equal band intensity after 30 cycles (data not shown). In addition, carrying out a complete set of PCRs on the same infected cell preparation further minimized variation.

PICs were harvested from C8166 cells infected with cell-free virus and harvested at 8·5 h post-infection. Cytoplasms were prepared and fractionated by sucrose velocity gradient centrifugation. PICs sedimenting at 160S were subjected to DNase footprinting using four concentrations of DNase I (0·1, 1, 10 and 30 µg/ml). Digestion with proteinase K was then carried out and protected DNA amplified for 24, 26, 28 and 30 cycles with 23 primer pairs that covered the entire HIV-1 genome. After analysis on agarose gels it was clear that PCR products were only seen with primer pair LTR1 and with 10, 1 and 0·1 µg/ml DNase I (Fig. 3). No PCR product was seen using any of the other 22 primer pairs, even with 30 cycles of PCR, or at any DNase concentration. The negative result shown with GAG4 primers was representative of data with the other 21 primer pairs (Fig. 3). PCR was continued up to 36 cycles in combination with the lowest DNase I concentration, but still no product was seen (data not shown). Positive controls (+) were successfully amplified after protease treatment (1 mg/ml) but in the absence of DNase I (Fig. 3), and no amplicon was seen in negative controls (-) following proteinase K and DNase I treatment (0·1 µg/ml). These data showing protection of only part of the LTR(s) are entirely consistent with Mu-mediated PCR footprinting on PICs harvested at a similar time (Chen et al., 1999 ).



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Fig. 3. Nuclease protection assay of cDNA from HIV-1 PICs harvested at 8·5 h post-infection using 23 primers pairs to cDNA regions 1–23 (Table 1). PICs were purified by sucrose gradient centrifugation and the 160S PIC cDNA digested with DNase I at 30, 10, 1 and 0·1 µg/ml as indicated by the triangle. The positive control (+) was treated with proteinase K but not DNase I. After proteinase K treatment for 1 h at 55 °C, DNA was extracted with phenol–chloroform, and precipitated with ethanol overnight. PCR was done for 24, 26, 28 and 30 cycles. Products were electrophoresed on 3% agarose gels. The negative control (-) was treated with proteinase K and then 0·1 µg/ml DNase I before PCR. PCR products were obtained using LTR1 primers (region 1; top panel), but with none of the other primers. Only the 30 cycle PCR data are shown. The lower panel shows the negative result with the GAG4 (region 7) primer pair that is representative of all primer pairs other than LTR1. The arrows indicate a DNA marker of 506 bp. This experiment is representative of two independent experiments each carried out on different batches of infected cells.

 
DNase footprinting of PICs harvested at 10 h post-infection shows that the majority of cDNA is protected
In the next part of this study we carried out DNase I footprinting exactly as described above, except that 160S PICs were harvested at 10 h post-infection. A DNA dilution series using three different primer pairs showed that approximately the same amount of DNA was present in both 8·5 and 10 h samples (Fig. 4).



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Fig. 4. Titration and PCR of cDNA from 160S HIV-1 PICs harvested at 8·5 and 10 h post-infection, as described in Fig. 1, using the primers pairs shown and 30 cycles of PCR. The arrows indicate a DNA marker of 506 bp.

 
DNase I protection with primer pairs 1–23 amplifying the entire genome from PICs isolated at 10 h post-infection is shown in Fig. 5. Data from two independent experiments are shown. PCR products were analysed after 24, 26, 28 and 30 cycles and with four concentrations of DNase I. Only the 30 cycle PCR data are shown. Positive controls, amplified in the absence of DNase I, were uniform in appearance and intensity for the most part, suggesting that the PICs amplified were representative of the 10 h PIC population as a whole. No PCR product was obtained in any of the negative controls. The first line of Fig. 5 shows protection of the genome from the LTR to the gagpol junction. All three LTR regions (detected with primer pairs LTR1, LTR2 and LTR/GAG) were well protected. This is in marked contrast with 8·5 h PICs where only region 1 was protected. Regions of gag were also well protected. For example in region 4, a PCR product was detected with primer pair GAG1 after 26 cycles and 0·1 µg/ml DNase (not shown). However, further along the genome, region 7 (GAG4) gave PCR products only after 30 cycles and 1 µg/ml DNase. Nonetheless the adjacent region 8 (GAG/POL) was well amplified after 26 cycles and 1 µg/ml DNase (not shown).



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Fig. 5. Nuclease protection assay of the entire cDNA genome isolated from 160S HIV-1 PICs harvested at 10 h post-infection. Two repeat experiments using different batches of infected cells are shown to demonstrate reproducibility. Twenty-three primer pairs (numbered 1–23 and described according to the relevant region of the genome – see also Table 1) were used. The first lane (-) is the negative control, the triangle indicates digestion with DNase I at concentrations of 30, 10, 1, 0·1 µg/ml, and (+) is the positive control (all as described in Fig. 3). PCRs were done using 24, 26, 28 and 30 cycles but only the 30 cycle data are shown. The arrowheads indicate a DNA marker of 506 bp.

 
The second line of Fig. 5 shows amplification of regions 9–16 from pol to the vprvpu junction. Regions 9–11 and 13–16 were quite well protected, with clear PCR products after 26 or 28 cycles and 0·1 or 1 µg/ml DNase (not shown). Region 12 (in the centre of pol) showed weak protection (upper data set) or no protection (lower data set). However, this was a relatively poor primer set, with positive control bands apparent only after 28 or 30 cycles of PCR. The third line of Fig. 5 shows amplification of regions 17–23 from vpu to nef. Overall, there was less amplification than with the upstream regions, although there was clear protection of regions 17–19, 21 and 22. However, none of the regions gave a product with 24 and 26 cycles of PCR, even at the lowest DNase concentration (not shown). PCR products were, however, obtained at 28 cycles for region 17 (VPU/ENV) and region 21 (ENV4), but only at DNase concentrations of 0·1 and 1 µg/ml. A PCR product was seen for region 18 (ENV1) after 30 cycles and 0·1 µg/ml DNase and for region 19 (ENV2) and 22 (ENV/NEF) only after 28 cycles at 0·1 µg/ml DNase. There was no protection of region 20 (ENV3) even though the positive control gave products after 26 cycles of PCR in both experiments. There was a weak product in region 23 (NEF/LTR) after 30 cycles of PCR.


   Discussion
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Abstract
Introduction
Methods
Results
Discussion
References
 
Recent work has provided insights into the composition, structure and function of HIV-1 PICs (see Introduction for references). The aim here was to elucidate PIC structure in more detail in order to begin to understand how PICs function in vivo. We initially characterized the time-course of appearance of viral cDNA in gradient-purified PICs in the cytoplasm and the accumulation of viral cDNA in the cell nucleus. 160S PIC cDNA was detected in the cell cytoplasm at 7·5, 8·5 and 10 h post-infection, but not at 12·5 h post-infection (Fig. 1a). Viral cDNA was first detected in the nucleus as a weak band at 10 h post-infection and then as a strong band at 12·5 h post-infection (Fig. 1b). These observations are consistent with synchronous transport of PICs to the nucleus between 10–12·5 h post-infection. In agreement with others (see Introduction for references) we found virion proteins RT, IN, MA and Vpr, but not CA, associated with PIC cDNA (Fig. 1c). The appearance of integration activity in 160S PICs (Fig. 2) closely followed their detection shown in Fig. 1(a), suggesting that the majority of the cDNA-containing PIC population was integration-competent (Fig. 2).

The main thrust of this report is the investigation of DNA–protein interactions over the entire HIV-1 genome using DNase I footprinting combined with PCR. In total, 23 overlapping sequences from the 5' LTR to the 3' LTR were probed and we examined PICs isolated at 8·5 and 10 h after infection of C8166 cells with cell-free virus. In the 8·5 h PICs, we found that only part of the LTR (region 1, amplified with primer pair LTR1) was protected from DNase I (Fig. 3). This amounted to approximately 6% of the cDNA, and is entirely consistent with earlier data (Chen et al., 1999 ). Together these data suggest that very limited amounts of protein are tightly bound to the cDNA of 8·5 h PICs. However, when we analysed PICs isolated at 10 h post-infection in exactly the same way, a completely different picture emerged. Here nearly all (approximately 90%) of the viral cDNA genome was protected by bound protein from digestion by DNase I. The fact that the one protected region 1 (LTR1) in 8·5 h PICs (Fig. 3) was amplified to a similar extent in the 10 h sample (Fig. 5) underlines the significance of areas of higher nuclease resistance seen elsewhere in the genome. However, not all the 10 h PIC cDNA genome was protected from DNase digestion and region 20 was completely sensitive (Fig. 5). Regions of high protection did not correspond with either of the two genomic polypurine tracts (Charneau et al., 1994 ).

As HIV-1 infection of cells in culture comprises a series of multiple non-synchronous infection events, we cannot fully discount the fact that the 10 h PICs may comprise mixtures of structures at various states of maturity that have been derived from virions entering the cell asynchronously. However, data in Fig. 1(a, b) suggest that there is synchronous movement of PIC cDNA from cytoplasm to nucleus. In addition, the 8·5 and 10 h post-infection patterns of nuclease protection differ radically as shown above, and in two separate experiments the nuclease protection findings were reproducible even to the extent of nuclease concentration sensitivity and number of PCR cycles required. Such data suggest that the characteristics of infection did not vary significantly from infection to infection. The positive controls shown in Fig. 5 suggest that the PICs isolated at 10 h post-infection represent the PIC population as a whole, and thus they may be an intermediate on the pathway into the nucleus. Further, PICs described here are unlikely to be proviral contaminants, as in our system, as discussed above, there was no detectable cross-contamination of cytoplasm and nucleus. In addition, our 10 h PICs were well characterized as originating from a cytoplasmic fraction that sedimented at 160S (Bowerman et al., 1989 ; Farnet & Haseltine, 1990 ; Karageorgos et al., 1993 ; Miller et al., 1997 ) and being associated with the viral proteins RT, IN, Vpr and MA, but not CA, as others have also found (see Introduction for references). It is interesting that there was no difference in the types of virion protein detected in our 8·5 and 10 h PICs. Clearly, the increased nuclease resistance at 10 h could only be explained if the distribution or association of PIC-associated proteins had changed as hypothesized in Fig. 6(a–c), or more (possibly cell) proteins had been recruited (Fig. 6d).



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Fig. 6. Diagram showing two conventional views of the structure of 8·5 h PICs (Chen et al., 1999 ; Farnet & Bushman, 1997 ) and possible structures of 10 h PICs. All are drawn with exactly the same types and amounts of virion proteins. (a, b) 8·5 h PICs where all proteins are tightly bound to and protect only the LTRs from nuclease digestion (a), or some proteins are tightly bound to and protect the LTRs and the others are loosely associated with and do not protect the rest of the viral genome (b). (c, d) 10 h PICs where all associated proteins are tightly bound to and protect most of the genome (c), or which resemble model (a), but have recruited cellular proteins which are tightly bound and protect from nuclease digestion (d).

 
It is postulated that a number of cytoplasmic viral intermediates are formed following HIV-1–cell fusion. These are (a) an immature reverse transcription complex (Zennou et al., 2000), (b) an early (8·5 h) PIC described by (Chen et al., 1999 ) and by us above, in which only the LTRs are tightly associated with protein, (c) the late (10 h) PIC described above in which approximately 90% of the genome is tightly associated with protein and (d) a PIC structure with a central DNA flap, a 99 nucleotide sequence, located approximately halfway along the HIV (Zennou et al., 2000 ) and feline immunodeficiency virus genomes (Whitwam & Poeschla, 2001 ). The central DNA flap is hypothesized to act as a determinant in translocating HIV-1 DNA through the nuclear pore (Zennou et al., 2000 ). Interestingly, the position of the DNA flap corresponds with regions 13 and 14 of our 10 h PICs where there was reasonably good protection (Fig. 5). It remains to be determined how late PICs harvested 10 h post-infection and the PICs with the central DNA flap are related structurally and in terms of the sequence of events on the PIC pathway. However, our 10 h PICs were integration-competent (Fig. 2) and appear to be nuclear translocation-competent (Fig. 1a, b). However, because all virus preparations have a high particle:infectivity ratio, it cannot be proved that late PICs are destined to become integrated provirus.

In conclusion, we have described a new form of nuclease-resistant PIC present at 10 h post-infection that appears late in the cytoplasm. This may be a mature and integration-competent intermediate that appears just prior to nuclear membrane docking and translocation. More work is needed to support this view, and we are currently engaged in identifying proteins that are bound near the central DNA flap of the late PIC, and investigating if and how they might aid nuclear translocation.


   Acknowledgments
 
Cells were kindly provided by Dr H. Holmes at the Centralized Facility for AIDS Reagents (NIBSC, Potters Bar, UK). Antibodies were also provided by the Centralized Facility for AIDS Reagents: Dr S. Ranjbar (IN antibodies), Drs D. Helland and A. M. Szilvay (RT antibodies), Drs R. B. Ferns and R. S. Tedder (MA antibodies); and by the NIH AIDS Research and Reference Reagent Program (Rockville, USA): Dr J. Kopp (Vpr antibodies) and Drs B. Chesebro and K. Wehrly (CA antibodies). We also thank Axis Genetics plc for purified cowpea mosaic virus and Dr Steve Busby (University of Birmingham, UK) for helpful discussions on nuclease footprinting. Work in the NJD laboratory was supported by grants from the National Heart, Lung, and Blood Institute, NIH (5R01HL59726) and The WPH Charitable Trust.


   References
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Abstract
Introduction
Methods
Results
Discussion
References
 
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Received 18 March 2002; accepted 25 June 2002.