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INTRODUCTION |
A pivotal step in the retroviral life cycle is forming the
provirus, an integrated cDNA copy of the viral RNA genome. The key
viral players in integration are the trans-acting integrase (IN)1 protein and the
cis-acting DNA attachment site. Integration proceeds through
three steps, the first two of which are known to require IN function.
The linear ends of the cDNA are initially processed adjacent to
phylogenetically conserved CA dinucleotides, resulting in a pair of
recessed 3' ends. After nuclear localization, the exposed 3'-hydroxyls
are joined to the 5'-phosphates of a double-stranded staggered cut in
chromosomal DNA. The final step is DNA repair, wherein the
single-stranded gaps at the sites of joining are sealed, resulting in
the sequence duplication of the double-stranded cut flanking the
integrated provirus (for a review, see Ref. 1).
In infected cells, integration is mediated by large nucleoprotein
complexes known as preintegration complexes (PICs), which are derived
from the cores of infecting virions (2). PICs isolated from infected
cells can integrate their endogenous cDNA into an exogenously added
target DNA in vitro (2-7). Additionally, recombinant IN
proteins purified after expression in bacteria can cut and join
oligonucleotide attachment site DNA substrates (reviewed in Ref. 1).
These latter in vitro assays have been invaluable for
deciphering the structure and function of retroviral IN proteins. IN
can be divided into three distinct functional domains: the amino-terminal, catalytic core, and carboxyl-terminal domains (reviewed
in Ref. 8). The central domain contains the highly conserved D,D(35)E
amino acid motif that comprises the IN active site (9, 10).
Although simplified in vitro assays using purified IN
proteins have provided essential knowledge toward understanding the overall integration process, these systems only partially mimic integration in vivo, because the predominant recombination
products result from the insertion of only one viral DNA end into just one strand of target DNA. Virus replication requires integration of
both DNA ends into both strands of target DNA; the single-ended activity typical of some in vitro systems would not yield a
productive viral infection. Although altering the source of purified IN
protein from bacterial to viral and/or modifying reaction conditions
can increase the frequency of two-ended integration products (11-13), these systems still do not recapitulate the efficiency of two-ended integration activity displayed by PICs isolated from infected cells
(14).
The discrepancy in reaction products catalyzed by PICs as compared with
purified IN proteins suggests that efficient two-ended integration
activity might require higher-order protein-protein and/or protein-DNA
interactions specific to nucleoprotein complexes derived from infected
cells. To begin to address this, we have used in vitro
Mu-mediated polymerase chain reaction (MM-PCR) footprinting to analyze
the protein-DNA structure of human immunodeficiency virus type I
(HIV-I) PICs partially purified from infected cells. We have
established an efficient HIV-I infection system initiated from
transfected cell supernatant that yields active PICs. This has allowed
us to analyze the structure and function of HIV-I PICs derived from a
number of IN mutant viruses. Our results indicate that multiple IN
functions are required to form the native protein-DNA structure of the
HIV-I intasome.
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MATERIALS AND METHODS |
Cells and Viruses--
MOLT IIIB (5), SupT1 (15), C8166 (16),
MT-4 (17), Jurkat (18), and CEM-12D7 (19) T-cell lines were maintained in RPMI 1640 medium containing 10% fetal calf serum. 293T (20) and
HeLa-CD4 (21) cells were grown in Dulbecco's modified Eagle's medium
containing 10% fetal calf serum. Two different infection systems were
used to isolate HIV-I PICs. In one system, chronically infected MOLT
IIIB cells were cocultured with uninfected SupT1 cells, essentially as
described previously (5). HIV-I production (HTLV-IIIB strain) was
stimulated 2- to 3-fold by pretreating MOLT IIIB cells (4 × 105 cells/ml) for 24 h with phorbol 12-myristate
13-acetate (10 µg/ml). In the other infection system, cell-free HIV-I
(NL4-3 strain) produced from transfected 293T cells was used to infect
C8166 cells (see below).
Plasmids and Oligonucleotides--
Wild-type pNL4-3 (22) and IN
mutants D116N (23) and K156E/K159E (24) have been described previously.
HIV-I (NL4-3) carrying the substitution of Lys for Gln-62 (Q62K) in IN
displays a replication-defective phenotype similar to that of
K156E/K159E.2 The W235E IN
change (25) was incorporated into pNL4-3 using overlapping PCR
(23).
Oligonucleotides were gel-purified before MM-PCR. Mu50 (5'-
GTTTTCGCATTTATCGTGAAACGCTTTCGCGTTTTTCGTGCGCCGCTTCA)
and Mu54
(5'-CTGCTGAAGCGGCGCACGAAAAACGCGAAAGCGTTTCACGATAAATGCGAAAAC) were annealed to generate Mu right-end DNA; Mu25
(5'-GCATTTATCGTGAAACGCTTTCGCG) was the Mu-specific PCR primer. HIV-I
primers were chosen to match both the NL4-3 and HXBc2 (26) strains. The
primers and their nucleotide positions in HXBc2 are as follows. AE330
(referred to as LA9 in Ref. 27; 805 to 787; 5'-GACGCTCTCGCACCCATCTC)
and AE529 (8690-8708; 5'-GCTGAGGGGACAGATAGGG) were used in the first round to amplify the 5' and 3' regions, respectively, of HIV-I. Minus-strand primers AE604 (454 to 432; 5'-CAGTACAGGCAAAAAGCAGCTGC) and
AE584 (215 to 196; 5'-CACAGGGTGTAACAAGCTGG) were used in second-round PCRs to analyze the 5'-long terminal repeat (LTR). Plus-strand primers
AE461 (9332-9356; 5'-AGTGTTAGAGTGGAGGTTTGACAGC) and AE459 (9522-9546;
5'-GCTTTTTGCCTGTACTGGGTCTCTC) were used in the second rounds to analyze
the 3'-LTR. The plus-strand primer AE525 (3220-3240; 5'-ACCTCCATTCCTTTGGATGGG) was used in the first round to analyze a
region of HIV-I internal to the LTR ends; AE524 (3264-3281; 5'-GGACAGTACAGCCTATAG) was used in this second-round PCR.
Infections, PIC Isolation, and in Vitro Integration
Assay--
SupT1 cells (4 × 107) were cocultivated
with phorbol 12-myristate 13-acetate-treated MOLT IIIB cells in 20 ml
of MOLT IIIB-conditioned medium (24-h supernatant of unstimulated cells
seeded at 1 × 106 cells/ml) at a ratio of 10:1. After
5 h, cells were washed twice in buffer K (20 mM HEPES,
pH 7.5, 5 mM MgCl2, 150 mM KCl, 1 mM dithiothreitol, and 20 µg/ml aprotinin) and lysed in 1 ml of buffer K-0.025% (w/v) digitonin. Crude cytoplasmic extract
cleared by brief centrifugation was treated with RNase A (0.1 mg/ml)
for 30 min at room temperature. To partially purify PICs, 1.5 ml of lysate was passed through a bovine serum albumin-coated Sepharose CL-4B
(Amersham Pharmacia Biotech) spin column (12 ml) equilibrated in buffer
K-0.025% digitonin. The eluate was further purified on a 10-ml,
10-50% (w/v) Nycodenz gradient prepared in buffer K. Centrifugation
was for 16 h at 274,000 × g in a Beckman SW 41 Ti
rotor at 4 °C. Twelve fractions were collected from the top of the gradient.
293T cells were seeded at 5.8 × 104
cells/cm2 in a 14-cm plate 24 h before transfection.
Cells were transfected with plasmid DNA (50 µg) using calcium
phosphate coprecipitation (28). Transfected cell supernatant was passed
through 0.45 µm filters and treated with DNase I (Promega; 2 units/ml) in the presence of 10 mM MgCl2 for 45 min at 37 °C. This step reduced the level of plasmid carry-over from
transfection to the limit of detection by Southern blotting. For
analyzing reverse transcription in cell lines, T cells (2 × 107 cells) and HeLa-CD4 cells (4 × 106
cells plated in a 10-cm plate 20 h before infection) were infected for 8 h with 10 ml of cell supernatant. Crude cytoplasmic extracts (0.5 ml) treated with RNase A were deproteinized by adding SDS, EDTA,
and proteinase K to final concentrations of 0.5% (w/v), 6 mM, and 0.6 mg/ml, respectively. After incubating at
56 °C for 1 h, DNA was recovered by phenol/chloroform
extraction and precipitation with ethanol. To isolate PICs, C8166 cells
(3 × 107) were infected for 8 h with 20 ml of
transfected cell supernatant. RNase A-treated cytoplasmic extract was
loaded directly onto Nycodenz gradients.
PIC activity was assayed essentially as described previously (5).
Briefly, 200 µl of crude cytoplasmic extract, spin column eluate, or
gradient fraction was incubated with 600 ng of linearized
X174 DNA
for 45 min at 37 °C. Reactions were deproteinized, and DNA was
recovered by precipitation with ethanol. DNA was electrophoresed through 0.6% agarose gels in Tris acetate-EDTA (28), transferred to
GeneScreen Plus membrane (NEN Life Science Products), and probed with a
LTR-specific riboprobe (HXBc2 nucleotides 8896-9615). Integration activity was quantified as the percentage of cDNA substrate
converted into product using either PhosphorImager analysis (Molecular
Dynamics) or densitometry (IS-1000 Digital Imaging System).
Western Blotting--
Gradient-purified C8166 cell extracts
adjusted to 15% (w/v) glycerol were frozen in liquid N2
and stored at
80 °C. SDS was added to a final concentration of
0.25% to thawed samples (200 µl), and proteins were recovered by
methanol-chloroform-H2O extraction essentially as described
previously (29). Briefly, samples were mixed with 4 volumes of
methanol, followed by 2 volumes of chloroform. Two phases were
separated after mixing 3 volumes of H2O and centrifuging at
9,000 × g for 1 min. Proteins at the interface were
precipitated using 3 volumes of methanol and spinning at 9,000 × g for 2 min. Pelleted proteins were resuspended in 1×
sample buffer (12 mM Tris-HCl, pH 6.8, 5% glycerol, 0.4%
SDS, 2.88 mM 2-mercaptoethanol, and 0.02% bromphenol
blue), boiled for 10 min, and electrophoresed through 10%
SDS-polyacrylamide gels. Proteins were transferred to Hybond-C extra
membrane (Amersham Pharmacia Biotech), and IN was detected using
monoclonal antibody 8E5 (30) with an ECL Western blotting kit (Amersham
Pharmacia Biotech). Recombinant HIV-I IN protein was purified after
expression in Escherichia coli as described previously
(31).
MM-PCR Footprinting--
Mu A protein was kindly provided by Dr.
Michiyo Mizuuchi (National Institute of Diabetes and Digestive and
Kidney Diseases). Two different naked DNA footprinting controls were
generally used. In one, plasmid DNA was added to buffer K to a final
concentration of 1.25 ng/ml. The other control was deproteinized PICs
resuspended in buffer K. Mu transpososomes were assembled by mixing the
annealed Mu end (40 nM) with Mu A transposase (232 nM) in 40 mM Tris-HCl, pH 8.0, 0.005% (w/v)
bovine serum albumin, 30% (w/v) glycerol, 30% (v/v) dimethyl
sulfoxide, 0.2% (v/v) Triton X-100, and 150 mM KCl. After
a 10-min incubation at 30 °C, the mixture was diluted 1:3 in the
same buffer. Assembled Mu transpososomes (200 µl) were mixed with an
equal volume of gradient-purified PICs or naked DNA controls, and
transposition was initiated by adding CaCl2 to a final
concentration of 10 mM. After 30 min at 30 °C, reactions were stopped by adding SDS, EDTA, EGTA, and proteinase K to final concentrations of 0.5%, 6 mM, 10 mM, and 0.6 mg/ml, respectively. Deproteinized Mu transposition products were
recovered by precipitation with ethanol.
DNA was dissolved in 50 µl of H2O, of which 3 µl was
used in nested PCR. The first round (50 µl) contained 20 mM Tris-HCl, pH 8.8, 10 mM KCl, 10 mM (NH4)SO4, 4 mM
MgSO4, 0.1% Triton X-100, 0.4 µM each of
Mu25 and a viral-specific primer, 0.4 mM of each dNTP, 0.1 mg/ml bovine serum albumin, and 1 unit of Vent DNA polymerase (New
England Biolabs). Twenty-five cycles were performed on a Perkin-Elmer
GeneAmp PCR system with the following PCR profile: preheating at
96 °C for 4 min, denaturation at 96 °C for 15 s, annealing
at 58 °C for 1 min, and elongation at 74 °C for 1 min. A final
10-min elongation at 74 °C was then performed. An aliquot (2 µl)
was transferred for the second round (25 µl; 25 cycles), which
contained 20 mM Tris-HCl, pH 8.8, 10 mM KCl, 10 mM (NH4)SO4, 2 mM
MgSO4, 0.1% Triton X-100, 0.2 µM each of
Mu25 and a nested viral primer, 5 × 105 cpm of 5' end
32P-labeled viral primer, 0.2 mM of each dNTP,
0.1 mg/ml bovine serum albumin, and 1 unit of Vent (exo
)
DNA polymerase (New England Biolabs). PCR was carried out essentially as described above, except that the elongation time was adjusted from
10 to 45 s, based on the length of the fragment being amplified. Aliquots of second-round PCRs were analyzed on 5% denaturing
sequencing gels.
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RESULTS |
Active HIV-I PICs from Molecularly Cloned DNA--
In this study,
HIV-I PICs were isolated from acutely infected cells using one of two
different tissue culture infection systems. In one system, HIV-I
(HTLV-IIIB strain) infection was initiated by mixing uninfected SupT1
cells with chronically infected MOLT IIIB cells, essentially as
described previously (5). Cells were lysed 5 h after infection,
and cytoplasmic extract containing HIV-I PICs was purified by spin
column chromatography. This step removed 80-90% of the total protein
present in the cell extract, yielding PICs that reproducibly supported
a higher level of integration activity than those in the starting
extract (Fig. 1A, lanes
1-4; data not shown). The column eluate was further purified on a
Nycodenz gradient, which was fractionated and analyzed for HIV-I DNA
content and PIC activity. DNA and integration activity co-sedimented to fractions 6, 7, and 8 (Fig. 1A, lanes 5-10). The
integration activity of the crude cell extract varied from 10% to
50%, depending on the individual preparation.

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Fig. 1.
Integration activity of HIV-I PICs.
A, Southern blot of PICs isolated from infected SupT1 cells.
X174 target DNA was included in integration reactions as indicated.
Lanes 1 and 2, crude cytoplasmic extract (about
24% of the substrate was converted to product in lane 2);
lanes 3 and 4, spin column eluate (about 37% of
the substrate was converted to product in lane 4.)
Lanes 5-10, activity of gradient-purified PICs; about 37%
of the substrate was converted to product in lane
6, about 32% of the substrate was converted to product in
lane 8, and about 26% of the substrate was converted to
product in lane 10. B, cDNA
synthesis in various CD4-positive cell lines. Lane 1, SupT1
cells; lane 2, HeLa-CD4 cells; lane 3, C8166
cells; lane 4, MT-4 cells; lane 5, Jurkat cells;
lane 6, CEM-12D7 cells. C, activity of PICs
isolated from C8166 cells. Lane 1, activity of crude
cytoplasmic extract. Lanes 2-4, activities of gradient
fractions 6-8, respectively. About 60% of HIV-I cDNA was
converted to product in each of these reactions. cDNA,
9.7-kilobase pair HIV-I substrate; IP, 15.1-kilobase pair
integration product.
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One goal of this study was to analyze the structure and function of
HIV-I PICs derived from IN mutant viruses. Thus, we set out to
establish an infection technique using transfected cell supernatant as
the source of cell-free virus. Preliminary experiments tested the
ability of six different CD4-positive cell lines to support reverse
transcription after infection with molecularly cloned HIV-I (NL4-3
strain). Of these six cell lines, C8166 T cells (16) supported the
highest level of cDNA synthesis (Fig. 1B). A kinetic
analysis revealed maximum levels of reverse transcription approximately
8 h after infecting C8166 cells (data not shown). Because HIV-I
cDNA in the crude cytoplasmic extract of C8166 cells was lost
during spin column chromatography (data not shown), these extracts were
directly purified by Nycodenz gradient centrifugation. The gradients
were fractionated and analyzed for HIV-I DNA content and PIC activity.
Both cDNA and integration activity colocalized to fractions 6-8
(Fig. 1C, lanes 2-4). PICs from C8166 cells
routinely integrated more of their cDNA substrate into the target
(30-80% of substrate converted to product) as compared with PICs
isolated from the coculture infection system.
MM-PCR Analysis of Wild-type HIV-I PICs--
MM-PCR footprinting
was developed to analyze the native protein-DNA structure of Moloney
murine leukemia virus (MoMLV) PICs (32). In this footprinting
technique, preassembled Mu transpososomes are the DNA cleavage reagent.
Mu A transposase inserts Mu-end DNA at the site of cleavage, and, as in
any protein-DNA footprinting technique, bound protein interferes with
the ability of the cleavage reagent to cut the nucleic acid. The
advantage of this coupled cutting and DNA joining technique over other
footprinting methods is that the DNA cleavage reagent itself becomes a
substrate for subsequent PCR amplification (Fig.
2). We thus applied this technique to
analyze the native protein-DNA structure of HIV-I PICs isolated from
cells using two different tissue culture infection systems.

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Fig. 2.
MM-PCR footprinting. Transposition
results in the insertion of Mu end DNA at the site of nuclease attack
(step 1). The frequencies and distribution of Mu cutting are detected
using two rounds of PCR (steps 2 and 3). Regions of protein binding
that prevent Mu from cutting the DNA (indicated by X) are
revealed after denaturing polyacrylamide gel electrophoresis (step 4).
Lane 1, MM-PCR pattern of deproteinized control; lane
2, pattern of native nucleoprotein complex.
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Three different substrates, naked plasmid, deproteinized PICs, and
native PICs, were generally analyzed. PICs partially purified after the
coculture infection were initially tested. As predicted, deproteinized
PICs supported a pattern of Mu transposition similar to the pattern
detected using naked plasmid DNA (Fig. 3,
A and C, compare lanes 2 to
lanes 1). Native PICs, however, revealed dramatically different transpositional patterns. Bound proteins protected more than 100 base pairs from each LTR end (Fig. 3, A and C, lanes 3). In addition to
these large footprinted regions, transpositional enhancements were
detected near the very ends of HIV-I (Fig. 3, A and
C, lanes 3). Both the footprinted and enhancement
regions were characteristic of the viral DNA ends; internal regions of
deproteinized and native PICs supported similar distributions of Mu
transposition (Fig. 3B). However, subtle differences were
detected (Fig. 3B), suggesting that proteins may loosely associate with internal regions of HIV-I. PICs partially purified from
infected C8166 cells yielded the same overall results. These PICs
generally displayed clearer footprinted regions (Fig. 3, D
and E), perhaps due to their higher integration
activity.

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Fig. 3.
MM-PCR footprinting of wild-type HIV-I
PICs. Mu transposition reactions were deproteinized and analyzed
by sequencing gels after two rounds of PCR. The top panel
shows the relative positions of HIV-I primers used in second-round
PCRs. A, native structure of the U3 end of HTLV-IIIB.
Lane 1, naked plasmid DNA; lane 2, deproteinized
PICs; lane 3, native PICs. B, internal HIV-I
region. Lanes 1 and 2, deproteinized and native
samples, respectively. C, native structure of the U5 end of
HTLV-IIIB. Lanes 1-3 were the same as in A.
D and E, native structure of the U3 and U5 ends,
respectively, of NL4-3. Lanes 1-3 in each panel were the
same as in A. Nycodenz gradient fraction 7 was analyzed in
A-E; MM-PCR of fraction 6 from infected SupT1 cells
revealed footprinting and enhancement patterns indistinguishable from
those of fraction 7. Numbers to the left of the
panels refer to the nucleotide position in HIV-I. The HXBc2 molecular
clone of HTLV-IIIB terminates at nucleotide 9718; NL4-3 U5 ends at
nucleotide 9709. E, regions of transpositional enhancement;
F, footprinted regions.
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The internal boundaries of the footprints were next identified. Whereas
the U3 footprint extended approximately 250 base pairs from the end of
HIV-I, the U5 footprint extended about 200 base pairs (Fig.
4). PICs isolated from infected C8166
cells displayed sharper footprint boundaries than those isolated from
the coculture infection (Fig. 4; data not shown).

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Fig. 4.
Extent of protein-DNA footprint. The
top panel shows the relative positions of viral primers used
in the second-round PCRs. A, structure of the U3 end of
NL4-3. B, the U5 end of NL4-3. Other labeling is as
described in the legend to Fig. 3.
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Multiple IN Functions Are Required to Form the HIV-I
Intasome--
Because IN is essential for retroviral DNA integration,
we tested the effects of different IN mutations on the structure and function of HIV-I PICs. HIV-I IN mutants can be divided into two classes, class I and II, based on their effects on the virus
replication cycle (33). Whereas class I mutants are blocked
specifically at the integration step, class II mutants display reverse
transcription and/or virus assembly defects. Thus, measurable
quantities of PICs can only be derived from class I IN mutant viral
infections. Most HIV-I deletions, as well as a number of point changes,
unfortunately fall into class II (33).
HIV-I IN contains three functional domains, the amino-terminal,
catalytic core, and carboxyl-terminal domains (8), and each domain
contains at least one amino acid residue that is conserved among all
retroviruses (33). The amino-terminal domain contains two conserved His
residues and two conserved Cys residues. Viral mutants carrying
substitutions of any one of these residues display the class II
replication phenotype (33), therefore measurable quantities of these
PICs were not recoverable (data not shown). The catalytic domain
contains the conserved active site Asp and Glu residues of the D,D(35)E
motif as well as residues such as Lys-159 (24, 34) and Gln-62 (31, 34,
35) implicated in viral DNA end binding. Substituting any of the three
active site residues yields the class I mutant viral phenotype, as do certain substitutions of Lys-159 (the double mutant K156E/K159E was
used here) and Gln-62 (Q62K). The substitution of Glu for the conserved
carboxyl-terminal domain residue Trp-235 (W235E) also yields the class
I mutant viral phenotype (25). At present, the exact defect of the
W235E mutant virus is unknown. We thus analyzed four different class I
mutant viruses that affected three different IN functions: D116N, which
was defective for catalysis; Q62K and K156E/K159E, each of which was
defective for viral DNA end binding; and W235E, which was defective for
an unknown carboxyl-terminal domain function.
Cytoplasmic extracts of C8166 cells were prepared after infection with
either wild-type or class I IN mutant viruses. Whereas wild-type PICs
converted about 50% of the cDNA substrate to the integration
product, none of the mutant viral PICs supported detectable integration
activity (Fig. 5). The preparations were
then centrifuged into Nycodenz gradients. Each mutant sedimented in
Nycodenz to the same position as did wild-type, indicating that the
nucleoprotein structure of each PIC was largely intact (data not
shown). The gradient-purified samples were then subjected to MM-PCR
footprinting. The results of this analysis showed that none of the
mutant PICs displayed the pattern of protein-DNA footprinting and
transpositional enhancements indicative of the wild-type HIV-I intasome
(Fig. 6). To ensure that the lack of
intasome structure for each of the mutants was not simply due to the
absence of the IN protein, gradient-purified samples were analyzed by
Western blotting using an anti-IN monoclonal antibody. The results of
this experiment showed that each of the mutant PICs contained IN
protein at a level comparable to that of wild-type (Fig.
7).

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Fig. 5.
Integration activity of HIV-I IN mutant
PICs. PICs isolated after infection with the indicated IN mutant
viruses were assayed for in vitro integration activity.
WT, wild-type NL4-3; N, D116N IN mutant;
E/E, K156E/K159E; E, W235E; K, Q62K.
The samples in the even-numbered lanes were reacted with
X174 target DNA. Other labeling is the same as that described in the
legend to Fig. 1.
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Fig. 6.
MM-PCR analysis of HIV-I IN mutant PICs.
IN mutant PICs partially purified by Nycodenz gradient centrifugation
were analyzed by MM-PCR footprinting. A, structure of the U3
end of HIV-I. B, structure of the U5 end. The samples in the
odd-numbered lanes were deproteinized before Mu
transposition; even-numbered lanes contained the
matched native samples. Other labeling is the same as that described in
the legends to Figs. 3 and 5.
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Fig. 7.
IN protein content of HIV-I mutant PICs.
Gradient-purified samples were analyzed by Western blotting.
Lanes 1-3 contained 50, 25, and 12.5 ng, respectively, of
recombinant HIV-I IN protein. The samples in lanes 4-8 were
isolated after infection with wild-type, D116N, K156E/K159E, W235E, and
Q62K HIV-I, respectively. The sample in lane 9 contained a
gradient-purified extract from mock-infected cells. The positions of
molecular mass standards in kilodaltons are indicated on the
left.
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DISCUSSION |
In this report, we describe the native protein-DNA structure of
preintegrative HIV-I as detected by MM-PCR footprinting. The structure
has two distinguishable characteristics. Firstly, large regions
(200-250 base pairs) of protein binding were detected at the ends of
HIV-I. Secondly, enhanced regions of nuclease attack were detected near
the very ends of the virus. The overall structure is quite similar to
the one previously reported for MoMLV (32). Thus, similar intasome
structures may be common to the preintegrative DNAs of a variety of retroviruses.
Numerous results point to the functional relevance of the retroviral
intasome. Firstly, functionally intact IN was required to form both the
MoMLV (32) and HIV-I (Fig. 6) structures. Also, both the MoMLV (32) and
HIV-I (36) structures, as well as PIC function, were undetectable after
stripping bound proteins with high concentrations of salt, and both
structure and function were restored in parallel by adding back
extracts from uninfected cells. Finally, Mu insertion into the enhanced
regions destroyed the ability of MoMLV PICs to undergo subsequent
in vitro integration (32). This interference highlights the
functional relevance of the retroviral intasome as detected by MM-PCR footprinting.
The basis for the transpositional enhancements at the ends of MoMLV and
HIV-I DNA is unknown. Retroviral integration and Mu transposition share
numerous features, notably their mechanisms of polynucleotidyl transfer
(reviewed in Ref. 37). Recombinant HIV-I IN protein preferentially
integrates oligonucleotide attachment DNA substrates into regions of
target DNA distortion (38), suggesting that protein binding in native
PICs may distort viral end regions and create hot spots for Mu
insertion. Alternatively, it is possible that proteins bound near the
ends of retroviral DNA physically interact with the Mu nuclease.
Bacterially expressed MoMLV IN protein bound to recombinant viral DNA
displayed the viral end footprinting and enhancement patterns
associated with the endogenous intasome isolated from infected cells
(39), suggesting that IN is solely responsible for the MoMLV structure
as detected by MM-PCR footprinting. However, we have been unable to
detect evidence for the HIV-I intasome structure using purified
recombinant IN with either recombinant viral DNA or deproteinized
cDNA substrates purified from infected cells (data not shown).
To study the role of IN in the formation of the HIV-I intasome, we
established a highly efficient infection system initiated with
molecularly cloned virus. Numerous CD4-positive cell lines were
screened for their ability to support reverse transcription after
infection (Fig. 1B), and one T-cell line, C8166 (16), was
not only found to support efficient DNA synthesis, but the resulting
HIV-I PICs displayed efficient in vitro integration activity
(Fig. 1C). As predicted, PICs isolated after infection with
class I viral mutants defective for either IN catalysis (D116N), viral
DNA end binding (Q62K and K156E/K159E), or an unknown carboxyl-terminal function (W235E) did not display detectable levels of in
vitro integration activity (Fig. 5). Despite containing their
mutant IN proteins (Fig. 7), none of the mutant PICs displayed the
protein-DNA footprints and viral end enhancements indicative of the
wild-type HIV-I intasome (Fig. 6). Although this was somewhat expected
for IN core domain mutants K156E/K159E and Q62K, whose defects are predicted to disrupt HIV-I attachment site DNA binding (24, 34, 35),
this was unanticipated for both the carboxyl-terminal domain mutant
(see below) and the D116N active site mutant. One IN active site
residue, Glu-152, has been implicated in interacting with the HIV-I
attachment site (34). In contrast, recombinant D116N mutant proteins
have not revealed evidence of DNA binding defects using either
functional (34) or physical (40) in vitro assay systems.
Thus, we speculate that IN catalysis may be closely linked to HIV-I
intasome formation in infected cells. There is a precedent in other DNA
recombination systems, notably Mu transposition, that the
recombinationally active Mu A protein-Mu DNA transpososome becomes more
stable with each successive step along the recombination pathway (37).
Thus, 3' processing of the viral ends by IN may stabilize the HIV-I
nucleoprotein complex. A kinetic analysis revealed that the MoMLV
intasome formed relatively slowly (32), consistent with the
interpretation that IN catalysis may be required for intasome formation
in infected cells.
The W235E defect in infected cells is unknown, in part because
recombinant W235E mutant IN protein displays wild-type levels of 3'
processing and DNA strand transfer activities in in vitro integration assays (41). A region of the carboxyl terminus between residues 247 and 270 has been implicated in binding to the HIV-I attachment site (35, 42). In contrast, the region between residues 213 and 246, where Trp-235 resides, cross-linked to the target DNA portion
of an in vitro disintegration DNA substrate (42). It is
therefore possible that the W235E mutant virus is defective for
interacting with chromosomal DNA in infected cells. However, because
the structure of W235E PICs purified from cytoplasmic extracts was
indistinguishable from the structure of either the D116N or attachment
site DNA binding mutants, we propose that the W235E virus is blocked at
a step that precedes target DNA interaction in the nuclei of infected
cells. Determining the precise defect of the W235E mutant may reveal
higher-order protein-protein and/or protein-DNA interactions important
for PIC activity that are dispensable for recombinant IN function in
more simplified in vitro integration assays. We also plan to
continue to analyze the structure and function of HIV-I IN and
attachment site mutant PICs isolated from infected cells.