Department of Cancer Immunology and AIDS,
Dana-Farber Cancer Institute, and Department of Pathology, Harvard
Medical School, Boston, Massachusetts 02115
Functional retroviral integrase protein is thought to be essential
for productive viral replication. Yet, previous studies differed on the
extent to which integrase mutant viruses expressed human
immunodeficiency virus type 1 (HIV-1) genes from unintegrated DNA.
Although one reason for this difference was that class II integrase
mutations pleiotropically affected the viral life cycle, another reason
apparently depended on the identity of the infected cell. Here, we
analyzed integrase mutant viral infectivities in a variety of cell
types. Single-round infectivity of class I integration-specific mutant
HIV-1 ranged from <0.03 to 0.3% of that of the wild type (WT) across
four different T-cell lines. Based on this approximately 10-fold
influence of cell type on mutant gene expression, we examined class I
and class II mutant replication kinetics in seven different cell lines
and two primary cell types. Unexpectedly, some cell lines supported
productive class I mutant viral replication under conditions that
restricted class II mutant growth. Cells were defined as permissive,
semipermissive, or nonpermissive based on their ability to support the
continual passage of class I integration-defective HIV-1. Mutant
infectivity in semipermissive and permissive cells as quantified by
50% tissue culture infectious doses, however, was only 0.0006 to
0.005% of that of WT. Since the frequencies of mutant DNA
recombination in these lines ranged from 0.023 to <0.093% of the WT,
we conclude that productive replication in the absence of integrase
function most likely required the illegitimate integration of HIV-1
into host chromosomes by cellular DNA recombination enzymes.
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INTRODUCTION |
Retroviruses carry two enzymes,
reverse transcriptase (RT) and integrase (IN), which function early in
the viral life cycle. Soon after infection, RT converts genomic RNA
into linear double-stranded cDNA. This DNA, which contains a copy of
the viral long terminal repeat (LTR) at each end, is the substrate for
IN-mediated DNA recombination. IN initially processes the 3' ends of
the cDNA adjacent to phylogenetically conserved CA dinucleotides and
then inserts these cleaved ends into a target DNA site in a cell
chromosome. The cis-acting end regions important for
integration define the viral attachment (att) sites, which
are comprised of U3 and U5 sequences in the upstream and downstream
LTRs, respectively. (For a recent review of retroviral integration, see
reference 6.)
In addition to the linear DNA product of reverse transcription, various
types of circular DNA form in retroviral-infected cells. Whereas some
of these result from IN-mediated autointegration of the viral cDNA into
itself, others result from host-mediated enzyme activities. One type of
host-mediated circle, which contains a single copy of the LTR, most
likely forms by homologous recombination between the LTRs. A second
type contains two tandem LTRs and probably forms by ligation of the two
cDNA ends. Host-mediated DNA circles are considered to be dead end
products of reverse transcription (6).
Although IN function is believed to be essential for retroviral growth
(6), IN and att site mutations can disrupt the
viral life cycle at distinct steps. Whereas some mutations (class I) block replication specifically at the integration step, others (class
II) cause pleiotropic defects. Cells infected with integration-specific class I mutants contain higher levels of unintegrated DNA circles than
do wild-type (WT)-infected cells (reviewed in reference
14), and class I human immunodeficiency virus type 1 (HIV-1) IN mutants display 12 to 19% of WT activity in the
multinuclear activation of galactosidase indicator (MAGI) assay
(2, 15, 60). This single-round infection assay requires de
novo synthesis of the viral Tat protein and its subsequent
trans-activation of an integrated 
-galactosidase
(-Gal) gene in HeLa-derived CD4-LTR/-Gal cells (15, 27,
31). Class I IN mutants recombine with host cell chromosomes
about 10
4 as frequently as the WT, but this low
level of integration is apparently mediated by host enzyme activities
and not viral IN (19, 30). Thus, although IN mutant
viruses have not been shown to support productive retroviral
infections, they can apparently support some transient level of gene
expression in HIV-1-infected cells (2, 8, 15, 53, 60).
Single-round class I IN mutants carrying the gene for firefly
luciferase (Luc) in the viral nef position, however,
displayed only about 0.2% of WT activity in infected human
rhabdomyosarcoma (RD) cells (34). Thus, the level of
transient gene expression from unintegrated HIV-1 DNA might depend on
the identity of the infected cell (8) and/or the
particular indicator assay.
To further examine the influence of target cell type, we determined
levels of WT and class I IN mutant gene expression in four different
T-cell lines by using single-round HIV-1 carrying the gene for
chloramphenicol acetyltransferase (CAT). Mutant CAT activity ranged
from <0.03 to 0.3% of WT activity, indicating an approximately
10-fold influence of cell type on gene expression from unintegrated DNA
relative to WT-integrated proviral DNA. Based on this, we reexamined
replication capacities of class I and class II mutant HIV-1 in a
variety of transformed and primary CD4-positive human cells.
Unexpectedly, certain T-cell lines supported productive class I mutant
replication under conditions that restricted class II mutant growth.
Remarkably, class I mutant HIV-1 could be continually passaged in
permissive cells without reversion to an integration-competent
genotype. Our results highlight contributions of cell type to gene
expression in the absence of IN function, suggesting that unintegrated
DNA may contribute to HIV-1 pathogenesis and the development of AIDS in
certain settings.
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MATERIALS AND METHODS |
Plasmids.
Single-round CAT expression vectors carrying
either WT (20) or IN mutant D116A (41) were
previously described. Plasmid pHI.Luc was built by amplifying the Luc
gene from pGL3-Basic (Promega Corp., Madison, Wis.) and ligating
digested DNA to BamHI-NotI-digested pHI.libGFP, a
single-round vector similar to previously described pHIvec2.GFP
(22). HIV-1 vectors carrying the gene for puromycin acetyltransferase (pac) were built by amplifying
pac from pPur (Clontech, Palo Alto, Calif.) and ligating
digested DNA to either BamHI-XhoI-digested
pHVSL3P (42) or BamHI-NotI-digested
pHI.libGFP. Plasmid pRL--actin, which expresses the gene for
Renilla luciferase (R-Luc) from the -actin promoter
(55), was kindly provided by Marianne Sweetser (University
of Washington, Seattle).
Gag-Pol was expressed from a separate plasmid when using pac
expression vectors. For this, WT IN was expressed from
CMV
P1envpAvpu/vpr (38), and a D116A derivative was
built by ligating the pol-containing PflMI
fragment from pHXBH10envCAT(D116A) (41) with
PflMI-digested CMVP1envpAvpu/vpr. Plasmids pSVIII-env
(20) and pHCMV-G (62), which express the
HIV-1 and vesicular stomatitis virus G (VSV-G) glycoproteins,
respectively, were previously described. Rev was expressed from a
separate plasmid as previously described (22, 42) when
using VSV-G.
WT pNL4-3 (1) and mutants D116N (15),
K156E/K159E (24), and 0A/0B (5) were
previously described. Overlapping PCR (15) was used to
incorporate the codon for D64N into pNL43(D116N), yielding double
mutant pNL43(N/N). Mutants E152Q and 1-212 were similarly built by PCR.
To facilitate the combination of IN mutations, pUC(A/P) was built by
introducing unique AgeI and PflMI sites into the
pUC19 polylinker. The AgeI-PflMI fragments from
N/N and E152Q were subcloned into pUC(A/P), and then the
BsmI-PflMI fragment from pUC(E152Q) was swapped
for the corresponding fragment in pUC(N/N), yielding pUC(NNQ). The
AgeI-PflMI fragment from pUC(NNQ) was
reintroduced into WT pNL43/XmaI (5), yielding
pNLX(NNQ). The AgeI-PflMI fragment from pNLX(NNQ)
was placed into pNLX(0A/0B), yielding pNLX(NNQ.LTR). CCR5-tropic N/N
was made by placing the EcoRI-BamHI fragment from
pNL(AD8) (17) into pNLX(N/N). The presence of IN and
att site mutations, as well as the absence of off-site
changes, was confirmed by sequencing PCR-generated DNAs.
Cells, viruses, and infections.
293T cells (39)
were grown in Dulbecco's modified Eagle medium (DMEM) containing 10%
fetal calf serum. HeLa-CD4 cells (27) were grown in DMEM
containing 0.1 mg of G418 sulfate per ml, and CD4-LTR/-Gal cells
(27) were grown in DMEM containing 0.1 mg of G418 sulfate
and 0.05 mg of hygromycin B per ml. T-cell lines were grown in RPMI
1640 medium containing 10% fetal calf serum. Peripheral blood
mononuclear cells (PBMC) and monocyte-derived macrophages (MDM) from
HIV-1-seronegative donors were purified by Ficoll-Paque density
gradient centrifugation. PBMC grown in RPMI were stimulated for 3 days
with 1 µg of phytohemagglutinin (Sigma, St. Louis, Mo.)/ml. One day
prior to infection, interleukin 2 (Chiron Corp., St. Louis, Mo.) was
added to the final concentration of 100 U/ml. Enriched preparations of
MDM cultured in macrophage-driving media were purified by adherence to
plastic essentially as previously described (10).
Virus stocks were prepared by transfecting 293T cells with DNA in the
presence of calcium phosphate (47). Whereas CAT viruses were prepared as previously described using two plasmids
(20), pac viruses required three or four
different plasmids depending on the identity of the viral envelope
(22, 42). Transfected cell supernatants were tested for
Mg2+-dependent 32P-RT
activity as previously described (15) and for p24 levels by enzyme-linked immunosorbent assay as recommended by the manufacturer (NEN Life Science Products, Boston, Mass.). Virus stocks were filtered
through 0.45-µm-diameter pores prior to infection. Viruses were
pretreated with DNase I as previously described (11) to degrade residual plasmid when infected cells were lysed for DNA extraction.
T cells (2 × 106) infected with
107 cpm of RT activity (107
RT-cpm) of HIV-1 CAT in 0.5 ml for 16 h were washed twice with
serum-free RPMI and cultured for an additional 44 h in 5 ml of
RPMI. Cells were counted, lysed in 20 µl of 0.25 M Tris-HCl (pH 7.6)
per 106 cells by freeze-thaw, and processed for
in vitro CAT assays essentially as previously described
(20). Twofold dilutions of lysate were tested in CAT
assays, and the percent of acetylated chloramphenicol was determined by
scintillation counter or phosphorimager scanning (Molecular Dynamics,
Sunnyvale, Calif.). Nonspecific CAT activities from cells mock infected
with envelope-lacking viruses were subtracted from WT and D116A values.
Percent acetylation was graphed against lysate volume, and standard
curves were generated to determine the volume of lysate required for
50% acetylation of added substrate. This volume was then converted to
the corresponding level of input virus (in RT-cpm).
Titers of WT and D116A CAT were determined using the MAGI assay as
previously described (15, 27). For CAT activity, 1.1 × 106 CD4-LTR/-Gal or HeLa-CD4 cells seeded
in 10-cm-diameter plates were infected the following day with 1.2 × 108 RT-cpm (4 ml) for 16 h. Cells washed
twice in serum-free DMEM were then cultured for 32 h in 10 ml of
DMEM. Cells were trypsinized, lysed in 100 µl of 0.25 M Tris-HCl (pH
7.6) per 106 cells by freeze-thaw, and processed
for in vitro CAT assays as described above.
Unless otherwise specified, T cells (2 × 106) infected with 2 × 106 RT-cpm of WT or mutant NL4-3 in 0.5 ml for
18 h were washed three times with serum-free RPMI and resuspended
in 5 ml of RPMI. Cultures were split at regular intervals, and aliquots
of supernatants were saved for RT assays. For determination of 50%
tissue culture infectious dose (TCID50), cells
infected with serial 10- or 3-fold dilutions of virus were plated in
24-well plates at 105 cells/well and split at
regular intervals for 3 to 4 weeks, and aliquots of the media were
saved for RT assays. Whereas MT-4- and C8166-containing plates were
scored by cytopathic effect, 174xCEM, CEM-12D7, and Jurkat cells were
scored positive when RT activity was threefold over mock-infected
values. TCID50 was calculated according to the
method of Spearman and Karber (29). Each determination was
performed at least twice.
Frequency of IN mutant DNA recombination.
Toxic
concentrations of puromycin dihydrochloride (Sigma) were determined for
the following cell lines as previously described (12):
Jurkat, 10 µg/ml; CEM-12D7 and MT-4, 0.5 µg/ml; C8166, 0.4375 µg/ml; 174xCEM, 0.375 µg/ml. Cells (2 × 106) infected with 2 × 106 RT-cpm of WT or D116A pac in 0.5 ml for 18 h were washed three times with serum-free RPMI, drug was
added 3 days postinfection (dpi), and 6 dpi, cells were serially
diluted into 96-well plates in the presence of 20% conditioned media.
Cells were fed at regular intervals, and single-cell colonies were
counted 2 to 3 weeks postseeding. Mutant DNA recombination frequency
was calculated by dividing the number of mutant puromycin-resistant
colonies by the number of WT colonies. Each determination was performed at least twice.
Cell fractionation, Southern blotting, and PCR.
For cloning
passed virus, cells were fractionated by the Hirt method as previously
described (18). Supernatant DNA (4 to 10 µl) was
amplified by Pfu polymerase (Stratagene, La Jolla, Calif.)
using AgeI- and PflMI-tagged (15) or
LTR-specific primers (5, 11) in buffer supplied by the
manufacturer. Reaction mixtures heated at 96°C for 1 min were cycled
20 to 25 times by denaturing at 96°C for 20 s, annealing at
58°C for 45 s, and extending at 72°C for 4 min, followed by a
final 10-min extension at 72°C. Fragments purified from
polyacrylamide gels (47) were either sequenced directly or
digested with AgeI and PflMI and ligated to
AgeI-PflMI-digested pNL4-3 prior to sequencing.
For Southern blotting and inverse PCR, cytoplasmic DNA was recovered as
previously described (11) after lysing 3.0 × 107 MT-4 cells in 1 ml of buffer K (20 mM HEPES
[pH 7.5], 5 mM MgCl2, 150 mM KCl, 1 mM
dithiothreitol, 20 µg of aprotinin/ml, 0.025% [wt/vol] digitonin).
Nuclei resuspended in 2 ml of buffer P1 (Qiagen Inc., Valencia, Calif.)
were lysed as recommended by the manufacturer. The supernatant obtained
after centrifugation was applied to an equilibrated Qiagen column, and
DNA was recovered as recommended by the manufacturer. The genomic DNA
pellet was resuspended in 6 ml of buffer G (6 M guanidine-HCl, 0.1 M
sodium acetate [pH 5.5], 5% Tween 20, 0.5% Triton X-100) by heating
at 56°C for several hours. Ethanol was layered onto the suspension,
and DNA was recovered by spooling onto glass essentially as previously
described (4).
Southern blotting was performed as previously described
(11), and levels of unintegrated DNA were quantified by a
phosphorimager (Molecular Dynamics). Integration was detected by
inverse PCR essentially as previously described (31). For
this, genomic DNA (2 to 15 µg) digested with HindIII
was recovered following extraction with phenol-chloroform and
precipitation with ethanol. DNA (500 ng) was ligated overnight in 50 µl at 16°C with 1 U of T4 DNA ligase (New England Biolabs, Beverly,
Mass.). Following heat inactivation (65°C for 10 min), DNA was
recovered by precipitation with ethanol and was resuspended in 10 µl
of H2O, and 2 µl was amplified using nested
PCR. The first round (50 µl) contained 0.5 µM (each) primers AE459
(11) and AE452 (5) and 2 U of AmpliTaq DNA
polymerase (Perkin-Elmer, Foster City, Calif.) in buffer recommended by
the manufacturer. Following heating at 95°C for 2 min, reactions were
cycled 20 times by denaturing (95°C for 15 s), annealing (58°C
for 1 min), and extending (72°C for 45 s), followed by a 7-min
extension at 72°C. DNA (2 µl) transferred to a second PCR (25 µl)
was amplified as in the first round by using 1 U of DNA polymerase,
primers AE322 and AE347, and 200,000 cpm of 5'-end-labeled AE347
(5). Reaction products were detected by autoradiography
following polyacrylamide gel electrophoresis.
Determination of HIV-1 promoter strength by transfection.
T
cells (3 × 106) were transfected with 1 µg each of pEGFP-C1 (Clontech), pRL--actin (55), and
pHI.Luc by using LipofectAMINE 2000 (Life Technologies, Inc.,
Rockville, Md.). For this, DNA in 100 µl of serum-free RPMI was mixed
with 100 µl of serum-free RPMI containing 10.3 µl of Lipofectamine,
and 45 min later, this mixture was added to cells in a final volume of
1 ml serum-free RPMI. Following 4 h at 37°C, RPMI (4 ml) was
added. Cells were counted, washed, and lysed in 20 µl of Passive
Lysis Buffer (Promega Corp.) per 106 cells
48 h posttransfection. Lysed cells were centrifuged at 20,630 × g for 15 min at 4°C, and 5 µl of supernatant was
analyzed using the DualLuciferase Reporter Assay System (Promega Corp.) as instructed by the manufacturer. Luminescence was detected using a
EG&G Berthold Microplate LB 96V Luminometer. Relative HIV-1 promoter
strength was calculated by normalizing the level of pHI.Luc-driven Luc
activity to the level of pRL--actin-driven R-Luc activity from a
total of six independent transfections.
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RESULTS |
Cell type-dependent HIV-1 gene expression in single-round infection
assays.
Previous studies reported that levels of HIV-1 class I IN
mutant gene expression varied over a wide range, from approximately 0.2 to 19% of WT expression, using two different infection assays (2, 15, 34, 60). Because of this, we evaluated levels of
WT and class I mutant gene expression across numerous cell lines using
a single infection strategy. For this, single-round HIV-1 carrying the
bacterial CAT gene in the viral nef position and either WT
or class I IN mutant D116A (2, 7, 14) was pseudotyped with
the HIV-1 envelope as previously described (20, 42). Four
different CD4-positive T-cell lines, C8166 (45), MT-4
(35), Jurkat (59), and CEM-12D7
(43), each of which was previously used to study HIV-1 IN
mutant replication (7, 15, 16, 28, 50, 53), were infected
with equal RT-cpm of virus. Cells were lysed 60 h postinfection,
and the level of virus inoculum that yielded 50% acetylation in in
vitro CAT assays was determined. By this method, a lower RT-cpm value
equated to a more efficient infection.
C8166 was the most permissive cell line (Table
1). Compared to C8166, about 7-, 30-, and
40-fold more WT virus was required to achieve 50% acetylation using
MT-4, Jurkat, and CEM-12D7 cells, respectively (Table 1). A similar
hierarchy of cell line activity was observed in the absence of IN
function. C8166 was still the most permissive cell line, but CEM-12D7
cells were now more permissive than Jurkat cells: compared to C8166,
about 2-, 24-, and >97-fold more D116A virus was required to achieve
equal CAT activities using MT-4, CEM-12D7, and Jurkat cells,
respectively (Table 1). Although D116A was more active in C8166 cells
than in the other lines, the level of D116A activity relative to WT
activity was greatest in MT-4 cells (Table 1). Compared to this
relative level of CAT activity, D116A was about 2-, 3-, and >10-fold
less active in CEM-12D7, C8166, and Jurkat cells, respectively (Table
1). Based on these differences in both absolute and relative levels of
CAT activity in these four cell lines, we decided to reevaluate the
replication capacity of integration-defective HIV-1 in a variety of
CD4-positive cell types.
Cell type-dependent replication in absence of IN function.
We
used four different HIV-1 mutants, three class I and one class II, in
preliminary assays for viral spread. In order to suppress the frequency
of reversion back to WT during productive replication, we analyzed
mutants that differed from the WT by more than 1 nucleotide. Since our
previously described class I IN active site mutant D116N
(15) carried just a single base change, we incorporated a
second active-site substitution, D64N, yielding double mutant N/N (Fig.
1). The other two class I mutants were
previously described. One, E/E, disrupted two lysines important for
IN-att DNA binding (24), and the other, 0A/0B
(5), was an att site mutant that disrupted the
invariant CA dinucleotides in both U3 and U5 (Fig. 1). Since deletion
mutants of IN fall into the class II category (14), we
engineered three stop codons after Glu-212 in IN, yielding C-terminal
deletion mutant 1-212 (Fig. 1).
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FIG. 1.
IN and att site mutations. Shown beneath
the genetic map of HIV-1 are the plus-strand termini of proviral U3 and
U5 and a diagram of IN highlighting active-site (Asp-64, Asp-116, and
Glu-152) and att DNA binding (Lys-156 and Lys-159)
residues. The nucleotide changes in mutants 0A/0B, N/N, NNQ, and E/E
are shown beneath the corresponding WT sequence. The size of the 1-212 deletion mutant is shown relative to the 288-residue WT protein. LTR,
long terminal repeat.
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Viruses produced from transfected 293T cells were normalized for RT
activity, and cells (2 × 106 cells) were
infected with 107 RT-cpm (about 1.5 µg of p24)
of WT or mutant HIV-1. Jurkat (Fig. 2A)
and CEM-12D7 (not shown) cells supported peak WT replication 5 and 7 dpi, respectively, under these conditions. Neither of these cell lines
supported detectable levels of mutant replication over prolonged
observation periods: Jurkat and CEM-12D7 cells infected with 0A/0B were
monitored for 1 month, and Jurkat cells infected with N/N and 1-212 were monitored for 2 months (Fig. 2A and data not shown). C8166 cells
supported peak WT replication 4 dpi (Fig. 2B). Unexpectedly, C8166
cells also supported the replication of N/N, E/E, and 0A/0B (Fig. 2B).
Replication of 1-212, however, was not detected under these conditions.
MT-4 cells also supported replication of N/N and E/E but not that of
1-212 (Fig. 2C). We note that MT-4 cells infected with as much as
3.0 × 107 RT-cpm did not support detectable
1-212 replication over 1 month of observation. Thus, MT-4 and C8166
cells supported class I mutant replication under conditions that
restricted class II mutant growth. In contrast, both classes of IN
mutants were restricted from replicating in Jurkat and CEM-12D7 cells.
Indistinguishable from the effects of WT, we note that virtually all
MT-4 and C8166 cells were lysed as a result of class I mutant viral
replication. Since class I mutant virus treated with DNase I prior to
infection replicated identically to untreated samples (data not shown),
we conclude that plasmid carryover from transfection played no role in
the observed mutant viral replication phenotypes.
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FIG. 2.
Replication kinetics of WT and mutant HIV-1 in various
T-cell lines. Cells infected with the indicated viruses were monitored
for RT activity at the indicated time points. Jurkat (A) and C8166 (B)
cells were infected with WT ( ), N/N ( ), E/E ( ), 1-212 (×),
0A/0B ( ), or mock-treated supernatant (+). (C) MT-4 cells were
infected with WT (), N/N (), E/E (), 1-212 (×), or virus-free
supernatant (+). (D) 174xCEM cells were infected with WT (), N/N
(), or mock-treated supernatant (+).
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Since both C8166 and MT-4 cells were originally transformed with human
T-cell leukemia virus type 1 (HTLV-1), we considered the possibility
that preexisting HTLV-1 IN might complement the class I HIV-1
integration defects in these cells. This seemed unlikely because 0A/0B,
which grew similarly to IN mutants N/N and E/E (Fig. 2B), contained
changes in the parts of the att sites that are conserved
among all retroviruses (5, 6, 14) (Fig. 1). That is, it
seemed unlikely that 0A/0B cDNA could be a viable substrate for any
retroviral IN protein. Consistent with this interpretation, 0A/0B
preintegration complexes isolated from infected C8166 cells did not
show evidence for in vivo 3' processing by IN or detectable levels of
DNA strand transfer activity in in vitro integration reaction mixtures
(H. Chen and A. Engelman, unpublished data). However, to further rule
out the possibility of IN complementation in MT-4 and C8166 cells,
other HTLV-negative human T-cell lines and primary cell types were
screened for their ability to support mutant virus replication. Whereas
neither H9 (40), Molt-4 Clone 8 (26), PBMC,
nor MDM supported a detectable level of class I mutant growth (data not
shown), N/N replicated in 174xCEM cells (Fig. 2D), a T-B cell hybrid
(46) that to the best of our knowledge does not contain a
preexisting retroviral IN protein.
To investigate mutant replication further, supernatant collected from
MT-4 cells at the peak of N/N growth (9 dpi with
106 RT-cpm) was passed onto fresh MT-4 and Jurkat
cells. Whereas Jurkat cells infected with 107
RT-cpm of this virus did not support a detectable level of replication, MT-4 cells infected with either 106 or
107 RT-cpm supported efficient HIV-1 growth (data
not shown). MT-4 cells undergoing this second round of infection were
lysed by Hirt extraction (21), and HIV-1 in the cell
supernatant was molecularly cloned and sequenced. All of the clones
analyzed (six total) retained both substitutions in the active site of
the IN enzyme. Two of these clones were sequenced across their entire IN regions, and each sequence was identical to that of N/N (one had an
A
G transition that did not alter the encoded amino acid). Thus, both
the starting genotype and cell type-dependent replication phenotype of
N/N produced in 293T cells were maintained after infection and HIV-1
replication in MT-4 cells.
Definition of permissive, semipermissive, and nonpermissive T-cell
lines.
Although the previous results showed that N/N could be
passed onto fresh MT-4 cells, a past study reported self-limiting
replication of integration-defective HIV-1 (8). To test if
the replication described here might also be self-limiting, we
repeatedly passaged class I mutant HIV-1 in MT-4 and 174xCEM cells. To
further limit the possibility of reversion during extensive tissue
culture, the remaining IN active site residue in N/N, Glu-152, was
replaced with Gln. The resulting NNQ mutant was then combined with the four att sites changes in 0A/0B, yielding the
multiple-mutatant NNQ.LTR. NNQ.LTR would minimally require 7 nucleotide changes to revert back to WT (Fig. 1).
As expected, NNQ.LTR replicated in C8166 (data not shown), MT-4 (Fig.
3A) and 174xCEM (Fig. 3G) cells, but not
in either CEM-12D7 or Jurkat cells. Also as expected, the supernatant
from infected MT-4 cells initiated a second round of replication when
passed onto fresh MT-4 cells (Fig. 3B) but not Jurkat cells (not
shown). NNQ.LTR growth peaked 12 days after passage, which was similar to its replication profile in the initial round of infection (Fig. 3A
and B). At 12 dpi, the 5-ml culture contained approximately 1.4 × 108 total RT-cpm, or about 70-fold more NNQ.LTR
than was used to initiate the second round of infection (Fig. 3B).
Thus, NNQ.LTR initiated a productive HIV-1 infection, and the yield of
progeny virus was similar to that of the WT under these conditions
(Fig. 3B). NNQ.LTR maintained this approximately 8-day delay in WT
levels of productivity following five serial passages in MT-4 cells
(Fig. 3A to F), and as was the case for virus collected after the first passage, Jurkat cells infected with fifth-round NNQ.LTR did not support
a detectable level of mutant viral replication. MT-4 cells undergoing
the sixth round of infection were lysed by Hirt extraction, and HIV-1
DNA in the Hirt supernatant was amplified by PCR using primers specific
for IN and the U3 and U5 att sites. DNA sequence analysis
revealed that all of the IN and att site mutations were maintained, proving that class I integration-defective HIV-1 replicated over several months of serial passage in MT-4 cells without acquiring an integration-competent phenotype (Fig. 3A to F) or genotype.
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FIG. 3.
Serial passage (pssg) of WT and NNQ.LTR in MT-4 and
174xCEM cells. (A) MT-4 cells infected with WT (), NNQ.LTR ( ), or
mock supernatant (+) were monitored for RT activity at the indicated
time points. (B) Virus recovered at the days of peak WT and NNQ.LTR
replication in panel A were used to inoculate fresh MT-4 cells (passage
1). (C to F) WT and NNQ.LTR replication upon passages 2 through 5, respectively. (G) 174xCEM cells were infected as described for panel A. (H and I) Passages 1 and 2, respectively, of WT and NNQ.LTR onto fresh
174xCEM cells.
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In contrast to the results with MT-4, NNQ.LTR recovered from infected
174xCEM cells replicated for only one serial passage (Fig. 3G to I). We
therefore conclude that NNQ.LTR replication in 174xCEM cells was
self-limiting. Based on the ability of class I integration-defective
HIV-1 to continuously replicate upon repeated passage, we propose the
following classification of CD4-positive human T-cell lines and primary
cells: permissive, semipermissive, and nonpermissive. Whereas MT-4
cells are permissive, 174xCEM cells are semipermissive and Jurkat
cells, CEM-12D7 cells, H9 cells, Molt-4 Clone 8 cells, PBMC, and MDM
are nonpermissive (Table 2). Although
C8166 cells were not analyzed by serial passage, we speculate that they
are permissive based on WT and class I mutant CAT activities (Table 1)
and TCID50 values (see below). We note that
Jurkat cells infected with class I mutant viruses derived from
transfected 293T cells occasionally yielded detectable levels of RT
activity 20 to 30 dpi. However, fresh Jurkat cells infected with these
supernatants did not support detectable RT production over prolonged
observation periods. Thus, although Jurkat cells can support some
transient gene expression under these conditions, we characterize them
as nonpermissive because the resulting virus was noninfectious in fresh
Jurkat cells.
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TABLE 2.
Summary of permissive, semipermissive, and nonpermissive
T-cell lines and primary cells as defined in text
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|
WT and class I mutant infectivities by TCID50.
Although we have shown that HIV-1 can replicate in certain cell lines
in the absence of IN function, our experiments until now were performed
using relatively high levels of input virus. To further investigate
class I mutant replication, we next quantitated WT and NNQ.LTR
infectivities under conditions that required viral spread. For this, WT
and NNQ.LTR TCID50 values were determined by
end-point dilution on five different T-cell lines. Each cell type was
infected with dilutions of WT or NNQ.LTR, cells were then extensively
washed prior to seeding in 24-well plates, and 2 to 4 weeks
postinfection, the viral dilution that yielded replication in 50% of
the wells was calculated as previously described (29). Under these conditions, most infections were initiated by one virus particle.
Whereas about 0.12, 0.11, 4.7, 1.5, and 1.2 pg of WT p24 were
sufficient to infect 50% of C8166, MT-4, 174xCEM, CEM-12D7, and Jurkat
cells, respectively, about 16, 6.4, 95, >1,500, and 
1,500 ng of
NNQ.LTR p24, respectively, were required (Table
3). Two important conclusions can be
gleaned from these results. First, NNQ.LTR initiated bona fide
spreading and productive HIV-1 replication in permissive and
semipermissive T-cell lines. Second, spreading infection assays are
extremely insensitive to large differences in titer and infectivity.
For example, WT was 4 to 5 orders of magnitude more infectious than
NNQ.LTR in permissive and semipermissive cells (Table 3). Yet, these
large differences in TCID50 equated to only about
a 1-week delay in peak viral growth (Fig. 3).
Class I mutant replication occurs in absence of normal
integration.
Our results until now demonstrated that class I IN
mutants initiated productive, spreading infections in certain T-cell
lines. We next directed our attention to the mechanism of mutant virus replication. Class I mutants recombine with chromosomes about 0.01% as
frequently as WT HIV-1, but integration in this case is apparently
promoted by host enzyme activities and not IN (19, 30).
Thus, two possibilities we considered for mutant viral replication were
gene expression from unintegrated DNA templates and/or transcription
from illegitimately integrated proviruses. To begin to distinguish
between these possibilities, WT and N/N-infected MT-4 cells were
fractionated and analyzed for HIV-1 DNA content. Cytoplasmic DNA was
recovered following cell lysis in isotonic buffer, and nuclei were
lysed using alkali. Whereas unintegrated nuclear DNA was recovered
using Qiagen columns, the genomic DNA pellet was recovered essentially
as previously described (4). Unintegrated DNA was detected
by Southern blotting, and integration was analyzed by inverse PCR
(31) as follows.
Although local chromosomal hotspots exist for retroviral integration
(61), HIV-1 for the most part integrates randomly
throughout the human genome (9). Inverse PCR uses
restriction enzyme digestion and intramolecular ligation to detect the
frequency and distribution of genomic HindIII sites
adjacent to a population of integrated proviruses (Fig.
4A)
(31). In addition to a ladder of DNA products reflecting
the randomness of genomic integration, two different internal HIV-1
fragments are detected using this strategy. All DNA forms yield a
1,102-bp fragment, but only host-mediated two LTR-containing circles
yield a 261-bp product (Fig. 4A).
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FIG. 4.
N/N replicates in MT-4 cells in the absence of normal
proviral formation. (A) Inverse PCR strategy. Shown are unintegrated
linear cDNA, integrated provirus, and host-mediated DNA circles. The
HindIII sites at positions 531, 8131, and 9606 in the
NL4-3 strain of HIV-1 are marked by vertical arrows, and the locations
of LTR-specific PCR primers are indicated by horizontal arrows. The
sizes of inverse PCR products (bold) are indicated in base pairs. VAR,
the variable inverse PCR product whose length is determined by
the location of an adjacent genomic HindIII site. A
variable-length ladder of DNA products reflects relative frequency and
distribution of retroviral integration (31). Dashed line,
host genomic DNA. (B) WT and N/N replication in MT-4 cells
(108) infected with 109 RT-cpm for 2 h at
37°C. RT activity was measured at the indicated times. (C)
Cytoplasmic (above) and nuclear (below) unintegrated DNA from WT (W)-,
N/N (N)-, 1-212 (2)-, and mock (m)-infected cells lysed at the
indicated times. The migration positions of circular DNA containing
either one LTR (1-LTR) or two LTRs (2-LTR) are indicated. Linear DNA
was most likely absent from nuclear extracts due to alkaline lysis
(3). Unintegrated DNA levels were quantified by adding the
signals from both panels. (D) Inverse PCR. Lane 1, genomic DNA from
mock-infected cells; lane 2, 1 ng of plasmid pNL4-3 was mixed with 10 µg of mock-infected DNA prior to HindIII digestion;
lanes 3 to 8, genomic DNA prepared from the indicated infections; lane
9, 10 µg of mock-infected DNA was mixed with Hirt supernatant from
approximately 5 × 104 N/N-infected MT-4 cells. The
migration positions of the internal 1,102- and 261-bp products are
indicated to the right of the gel. Some of the inverse PCR products
detected in infected-cell lysates resulted from either nonspecific
amplification of plasmid-related sequences (approximately 530-bp
product in lane 2) or amplification of 2-LTR circles in the genomic DNA
fractions (lane 9). Each of the four intense bands that migrated
between 261 and approximately 400 bp in lanes 5 through 9 were
attributed to 2-LTR circles (lane 9 and data not shown). The migration
positions of molecular mass standards are indicated on the left (in
base pairs). LTR, long terminal repeat.
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|
Consistent with previous reports (2, 15, 30, 60), cells
infected with N/N contained a higher level of host-mediated circles
than WT-infected cells at 1 dpi and DNA synthesis in class II-infected
cells was undetectable (Fig. 4C). In the WT infection, the level of
unintegrated DNA increased dramatically from 1 to 2 dpi, in parallel
with HIV-1 production (Fig. 4B and C). Replication was initially
detected in N/N-infected cells 2 dpi, and HIV-1 levels increased over
time roughly in parallel with the level of unintegrated DNA (Fig. 4B
and C). At their replication peaks (7 dpi for N/N, 2 dpi for WT),
N/N-infected cells contained approximately 70% as much unintegrated
DNA (Fig. 4C) and yielded about half as much virus (Fig. 4B) as
WT-infected cells. As expected, a large population of integrated
proviruses was detected in the genomic DNA of WT-infected cells (Fig.
4D, lanes 3 and 4). In contrast, this DNA ladder was virtually absent
from the N/N infection: the only readily detectable
HindIII fragments in this case were attributed to
amplification of either plasmid-related sequences (lane 2) or
cofractionating two LTR circles (lanes 5 to 9). We therefore conclude
that class I IN mutant viral replication produced near-normal levels of
unintegrated HIV-1 DNA and progeny virions without yielding the normal
level of integrated provirus. Although these results are consistent
with the model that N/N replicated from unintegrated DNA, we note that
a minor population of novel bands was detected in N/N-infected genomic
DNA upon long autoradiographic exposure. Although difficult to
precisely quantitate by inverse PCR, we speculate that this low level
of integration represents the approximate 104
illegitimate recombination frequency previously reported for class I IN
mutant viruses (19, 30). We therefore considered the
possibility that the frequency of class I mutant DNA recombination within a given cell line might be an important determinant of its
permissivity. To investigate this hypothesis, we quantitated the level
of D116A integration in MT-4, C8166, 174xCEM, CEM-12D7, and Jurkat cells.
Permissive and nonpermissive cell lines support similar mutation
recombination frequencies.
Single-round HIV-1 carrying the
pac gene in place of nef and either WT or D116A
IN were pseudotyped with either the HIV-1 or VSV-G glycoprotein
essentially as previously described (42). Cells infected
with equal RT-cpm of envelope-matched viruses were extensively washed,
incubated with predetermined levels of puromycin for 3 days, and then
serially diluted into 96-well plates in the presence of drug. After 2 to 3 weeks, puromycin-resistant colonies were counted and mutant DNA
recombination frequency was calculated as colony number per RT-cpm of
input D116A versus WT.
Unexpectedly, nonpermissive CEM-12D7 and Jurkat cells supported more WT
pac colonies than either MT-4 or 174xCEM cells (Table 4). This was most likely due to
differences in plating efficiencies among cell lines. For example, in
the absence of drug, about 20 MT-4 cells and 150 174xCEM cells were
required per well for cell growth. Jurkat cells, in contrast, tolerated
densities as low as one cell per well without drug. Despite this
limitation, precise DNA recombination frequencies were determined for
four of the five cell lines. MT-4 was the only cell line that did not
support a detectable level of D116A pac-resistant colonies
(Table 4).
The level at which a particular cell line supported D116A integration
did not correlate with its permissivity. For example, nonpermissive
CEM-12D7 and permissive C8166 cells supported the highest and lowest
recombination frequencies, respectively (Table 4). We note that we did
not investigate the mechanism of D116A integration here. However, a
previous sequence analysis of 11 proviruses derived from three
different class I IN mutant viruses revealed little evidence for
IN-mediated DNA recombination (19). Integration normally
yields a 5-bp target site duplication flanking the HIV-1 provirus
(57, 58), and 10 of 11 sequenced mutants lacked this
characteristic. Although one mutant displayed a 5-bp duplication, it
contained thymidine substituted for the invariant adenine residue at
the U3 att terminus that normally joins to target DNA
(6), suggesting that recombination enzymes other than IN
most likely played a critical role in its integration (19). Based on those results, we speculate that host
enzymes were responsible for most if not all of the D116A
pac recombination events that occurred in C8166, 174xCEM,
CEM-12D7, and Jurkat cells.
| |
DISCUSSION |
Although it is widely believed that IN function is absolutely
required for productive retroviral replication, previous reports differed on the extent to which unintegrated DNA could serve as a
template for gene expression and synthesis of HIV-1 proteins (2,
8, 15, 44, 53, 60). We previously noted that this was in part
influenced by the nature of the viral mutation: whereas class I IN
mutations blocked HIV-1 replication specifically at the integration
step, class II mutations caused more pleiotropic damage, including
defects in virion morphogenesis and reverse transcription
(14). Because of this pleiotropy, class II mutants displayed only 0.1 to 0.8% of WT titers in the MAGI assay under conditions where class I mutants yielded 12 to 19% (2, 14, 15). In addition to this mutational influence, the identity of
the target cell also appeared to affect the level of gene expression in
the absence of IN function. In this case, self-limiting replication of
integration-defective HIV-1 was detected in primary MDM but not PBMC
(8). In the present study, we focused primarily on this
influence of target cell type on gene expression. Unexpectedly, we
found that a subset of transformed T-cell lines supported productive HIV-1 replication in the absence of IN-mediated DNA recombination.
Cell type and assay type-dependent gene expression in the absence
of IN function.
Although 12 to 19% of WT titers were previously
observed using the MAGI assay (2, 15, 60), class I IN
mutants carrying the Luc gene in the place of nef displayed
only about 0.2% of WT activity in RD cells
(34). To investigate this apparent cell type difference,
we determined levels of WT and class I mutant HIV-1 required for 50%
CAT activity in four different T-cell lines. Large differences in HIV-1
activity were detected regardless of IN function. That is, about 40-, 30-, and 7-fold more WT virus was required for CEM-12D7, Jurkat, and
MT-4 cells, respectively, than for C8166 cells (Table 1). These results
are consistent with previous reports that certain T-cell lines are more
permissive than others in their ability to support HIV-1 replication
(43, 52). In addition to these differences, however, we
detected an approximately 10-fold effect of cell type on levels of
D116A CAT activity relative to WT activity (Table 1). These levels, which ranged from <0.03% for Jurkat cells to 0.3% for MT-4 cells, are in-line with the previous 0.2% level reported for single-round Luc
viruses (34). Not surprisingly, we conclude that the
identity of the indicator gene in the viral nef position
does not influence the level of gene expression from class I IN mutant
viruses in transient-infection assays. Thus, we were perplexed as to
why class I mutant titers were as high as 19% of the WT titers in the
MAGI assay (2).
Since the MAGI assay uses HeLa-derived CD4-LTR/-Gal cells
(27), we tested whether HeLa cells might present an
especially favorable environment for gene expression from unintegrated
DNA. For this, we determined levels of WT and D116A required for 50% CAT activity using two related cell lines, HeLa-CD4 and CD4-LTR/-Gal (27), and compared these values to the relative MAGI
titers of the viruses. Whereas approximately 2.4 × 105 and 3.0 × 105
RT-cpm of WT CAT were required in HeLa-CD4 and CD4-LTR/-Gal cells,
respectively, about 1.2 × 108 and 1.1 × 108 RT-cpm of D116A CAT were required,
respectively. Thus, D116A supported about 0.2 and 0.27% of WT CAT
activity in HeLa-CD4 and CD4-LTR/-Gal cells, respectively. We
therefore conclude that the relative level of class I IN mutant gene
expression in HeLa cells was similar to those of other cell types when
the indicator gene was in the viral nef position (Table 1).
Yet, as expected, D116A CAT yielded about 10% as many blue-staining
cells as WT CAT in the MAGI assay (data not shown). Thus, the indicator
assay itself can apparently influence the outcome of class I mutant gene expression relative to WT as much as 37-fold in a single cell
type. Since the Tat protein responsible for indicator gene function in
both assays was expressed from the incoming mutant viral genome
(15, 31), we conclude that the activation of integrated
-Gal is more sensitive than the activation of viral CAT or Luc for
detecting gene expression from unintegrated HIV-1 DNA.
Cell-type dependent HIV-1 replication without IN function.
Based on our result that cell type affected the relative level of D116A
gene expression as much as 10-fold (Table 1), we reevaluated class I
and class II mutant replication kinetics in a variety of CD4-positive
cells (Table 2). Our results for the first time show that IN function
is not required for bona fide spreading and productive retroviral
replication (Fig. 2 and 3). Similar to previous observations with the
MAGI assay (2, 15, 60), we found that only class I
mutants, and not class II, functioned in viral replication assays (Fig.
2). It is important, however, to highlight two limitations of class I
mutant viral replication. First, mutant infectivity in permissive and
semipermissive cell lines as quantified by TCID50
was 20,000- to 170,000-fold less efficient than that of the WT (Table
3). Second, mutant replication was not detected in either primary PBMC
or MDM. Thus, despite finding that as little as 6.4 ng of class I
mutant p24 was sufficient to initiate replication in 50% of MT-4 cells
(Table 3), we stress that IN remains an important target for the
development of antiviral drugs.
This work highlights the importance of paying attention to target cell
type when determining mutant viral replication phenotypes. Permissive
MT-4 (28, 53) and C8166 (7) cells were
previously used to study HIV-1 IN mutant viral replication. Stevenson
et al. (53) studied deletions of IN, which are predicted
to behave as class II mutants (14). Thus, the lack of
mutant viral replication in that study was probably due to class II
pleiotropy. Whereas the IN analyzed by Lafemina et al. contained the
E152Q class I mutation, it also contained a second change, V151D
(28). The effects of V151D alone on MT-4-dependent
replication were not addressed in that study, so it is possible that
the double mutant behaved differently than a true class I mutant.
Cannon et al. (7) used 50 ng of class I D116A to infect
5 × 105 C8166 cells, which, based on our
data, should have replicated (Fig. 2 and Table 3). Since infectivity
was scored by syncytium formation 3 to 4 dpi, we speculate that those
cell cultures were terminated prior to mutant replication (Fig. 2).
D116A replication also went undetected in H9 cells despite multiple
weeks of observation (7). Consistent with that result, we
determined here that H9 was a nonpermissive T-cell line (Table 2).
Cara et al. detected self-limiting replication of integration-defective
HIV-1 in primary MDM (8). The mutation analyzed in that
study was an IN deletion which, as mentioned, is predicted to behave as
class II. Yet, we were unable to detect class I mutant HIV-1
replication in MDM despite using relatively high levels of virus and
cells derived from multiple blood donors (Table 2). It is possible that
viral strain differences or cell culture techniques contributed to
these different results. We note that our results are consistent with
previous reports that MDM do not support detectable levels of class I
IN mutant replication (17, 60). We propose that future
investigations aimed at determining deleterious effects of IN and
att site mutations on HIV-1 replication should avoid permissive and semipermissive target cell lines. On the other hand,
infecting MT-4 cells with 50 ng of p24, for example, should help
determine whether mutants identified as replication-defective in
nonpermissive cells are of the class I or class II phenotype (Fig. 2
and Table 3).
Mechanism of class I IN mutant replication in permissive cell
lines.
Since MT-4 and C8166 cells were originally transformed with
HTLV-1, we considered the possibility that preexisting HTLV-1 IN might
complement class I HIV-1 integration defects in these cells and in
doing so form the basis for cell line permissivity. Our finding that
174xCEM cells also supported mutant viral replication (Fig. 2D),
however, ruled out the necessity for preexisting retroviral IN protein.
However, 174xCEM cells were notably less permissive than either MT-4 or
C8166 cells (Fig. 2 and 3). Thus, might the preexisting HTLV-1 genome
somehow contribute to class I IN mutant viral replication? We feel that
the HTLV-1 Tax protein, which is expressed in both MT-4 and C8166 cells
(23) and is a potent trans-activator of the
HIV-1 promoter (51), plays a role in permissivity (see
below). However, our finding that N/N replicated in MT-4 cells without
forming the normal level of integrated provirus (Fig. 4) together with
the results that MT-4 and C8166 cells supported frequencies of D116A
pac recombination that did not differ significantly from
nonpermissive Jurkat and CEM-12D7 cells (Table 4), demonstrates that
replication occurred in permissive cell lines independent of an HTLV-1
or cellular function(s) that restored functional integration to class I
IN mutant HIV-1.
Other parameters of cell line permissivity. (i) Viral entry.
We therefore considered other parameters that might contribute to cell
line permissivity. Since we observed as much as a 40-fold difference in
WT HIV-1 CAT activity across cell lines (Table 1), we focused our
attention on functions that influenced permissivity independent of
integration. Srivastava et al. (52) previously highlighted
rate and efficiency of virus entry as major determinants of
permissiveness: whereas 30 min was sufficient to allow 50% of WT HIV-1
to enter C8166 cells, about 4 h was required for Jurkat cells
(52). To minimize the effects of these different entry rates, we allowed viral infections to proceed for 16 to 18 h. Under these conditions, we infer that virtually all and between 90 and
95% of potentially infectious virus entered C8166 and Jurkat cells,
respectively. We therefore speculate that virus entry did not
significantly limit the first round of HIV-1 replication. Although
CEM-12D7 cells were not directly tested, we note that, similar to
Jurkat cells, 4 h was required for 50% entry into A3.01 cells
(52). Since CEM-12D7 cells were derived from A3.01 cells (43), we speculate that entry did not limit the first
round of replication in CEM-12D7 cells. Consistent with this
interpretation, we determined by fluorescence-activated cell sorting
that the levels of HIV-1 receptors CD4 and CXCR4 on the surface of
CEM-12D7 cells were equal to or greater than the levels of these
proteins on Jurkat, 174xCEM, C8166, and MT-4 cells (data not shown).
Despite controlling for entry in first-round infections, however, we
speculate that limited entry most likely played an important role in
restricting the spread of class I mutant HIV-1 throughout nonpermissive
cell cultures.
(ii) Transcription from the HIV-1 promoter.
Both MT-4 and
C8166 cells express the HTLV-1 Tax protein (23). Since Tax
is a strong trans-activator of the HIV-1 promoter (51), we investigated promoter activity in the different
cell lines. For this, cells were cotransfected with two different
reporter gene constructs. Whereas one drove Tat-dependent Luc
expression from the HIV-1 promoter, the other construct expressed the
gene for R-Luc under the control of the -actin promoter
(55). Since -actin is a constitutively expressed gene,
its expression is used to normalize levels of heterologous gene
expression in a variety of cell types and experimental conditions
(reviewed in reference 54). Thus, to assess the relative
strength of the HIV-1 promoter across cell lines, HIV-1-driven Luc
activity was normalized to -actin-driven R-Luc activity in different
transfected cell extracts.
The results show that nonpermissive CEM-12D7 and Jurkat cells supported
similar low levels of HIV-1 gene expression (Fig. 5). In comparison, MT-4, 174xCEM, and
C8166 cells supported about 2.2-, 7.6-, and 23-fold more activity,
respectively (Fig. 5). We note that HIV-1 transcription levels in
CEM-12D7, Jurkat, MT-4, and C8166 cells (Fig. 5) correlated roughly
with the levels of WT virus required for 50% CAT activity with these
cells (Table 1). This result is consistent with our conclusion that
entry did not play a major role in restricting the initial round of mutant viral replication. The results are also consistent with the
interpretation that efficient transcription from the HIV-1 promoter
played a role in permissivity. We note that the B-lymphoblastoid parent
of 174xCEM, 721.174 (46), was transformed with
Epstein-Barr virus (13), and that various Epstein-Barr
virus proteins can up-regulate the HIV-1 promoter (25, 33,
48). Thus, we propose that a discerning feature of permissive
and semipermissive cells is their expression of heterologous viral
proteins that enhance HIV-1 transcription.
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FIG. 5.
Cell type-dependent HIV-1 promoter activity. The
activity of the HIV-1 promoter relative to -actin is shown for the
indicated cell lines. The average activity following six independent
transfections is noted beneath each bar. Error bars represent standard
deviation from the mean.
|
|
C8166 cells supported about 10-fold higher levels of HIV-1
transcription than did MT-4 cells (Fig. 5). Similarly, about sevenfold more WT CAT virus was required for MT-4 cells than for C8166 cells (Table 1). Yet, both cell types were equally infected with the WT as
measured by TCID50s (Table 3). Thus, we conclude
that MT-4 cells are more efficient than C8166 cells at fulfilling a
step(s) in the HIV-1 cycle that occurs after transcription and
translation but prior to the next round of virus entry. Based on this,
we speculate that MT-4 cells are particularly efficient at assembling and releasing HIV-1 particles. Similar to the results reported here, we
note that other studies reported that MT-4 cells supported the
replication of HIV-1 mutants that failed to replicate in
less-permissive Jurkat (41) and CEM-12D7 (36) cells.
(iii) Gene expression from unintegrated DNA versus integrated
proviruses.
What function(s) might contribute to productive class
I IN mutant replication? To address this, we compiled in Table
5 a comparison of mutant and WT
activities as measured by CAT activity (Table 1),
TCID50 (Table 3), and DNA recombination frequency (Table 4). In C8166 cells, for example, D116A integrated about 0.023%
as frequently as WT, and D116A CAT was about 0.1% as active as WT CAT
(Table 5). We therefore conclude that integration could have accounted
for as much as 23% of the total CAT activity in D116A-infected C8166
cells. Similarly, we conclude that D116A integration accounted for as
much as 31 and 62% of the CAT activity in MT-4 and CEM-12D7 cells,
respectively. Based on our results with CD4-LTR/-Gal cells, however,
we note that the activity of D116A relative to that of the WT varied as
much as 37-fold in the same cell type. It would therefore appear that
unintegrated DNA can express the majority of the Tat protein in cells
transiently infected with class I mutants.
In contrast to concluding that unintegrated DNA contributed at least in
part to class I mutant gene expression during transient infections, we
note that the relative TCID50s of NNQ.LTR and WT in 174xCEM, C8166, MT-4, Jurkat, and CEM-12D7 cells were about 8-, 38-, 55-, 267-, and 1,250-fold lower, respectively, than the frequencies of
D116A integration in these cell lines (Table 5). Based on this, we
conclude that DNA recombination most likely played an important role in
class I IN mutant viral replication. In other words, although class I
IN mutants supported productive HIV-1 replication in the absence of IN
function, they most likely required the integration of their genomes
into cell chromosomes to achieve this result. Although integration may
be necessary, we note that it alone is not sufficient for mutant viral
replication (Table 5). We therefore conclude that the ability of
certain T-cell lines to support productive class I IN mutant viral
replication depends on a number of permissivity factors, most of which,
including virus entry, assembly/release, and transcription, would
appear to influence the efficiency of HIV-1 replication independent of integration.
Unintegrated HIV-1 DNA and AIDS dementia.
Large levels of
unintegrated HIV-1 DNA have been detected in the brains of some
patients with AIDS dementia (37, 56). Although the extent
to which this DNA might serve as a template for gene expression and
subsequent protein production has not been directly tested, our finding
that different T-cell lines affected the level of gene expression from
unintegrated DNA as much as 10-fold suggests that certain in vivo cell
types may also have the ability to preferentially express unintegrated
forms of HIV-1. Since secreted Tat protein has been implicated in the development of AIDS dementia (32, 49) and our results are consistent with the interpretation that Tat can be expressed from unintegrated DNA, we propose that it will be worthwhile to assess the
activity of class I IN mutant HIV-1 relative to WT in infected human
neuronal cell cultures in vitro. The results of these experiments may
shed light on whether gene expression from unintegrated DNA plays a
role in HIV-1 pathogenesis and the development of AIDS dementia in vivo.
We thank H. Göttlinger, W. Marasco, J. Sodroski, and
M. T. Sweetser for plasmid DNA, H. Chen for help with Southern
blotting, K.-T. Jeang for stimulating discussion in the early phase of
this project, H. Göttlinger, J. Sodroski, and Å. Ohagen for
valuable discussion, and Å. Ohagen for critical review of the manuscript.
This work was supported by NIH grant AI45313 (A.E.), the G. Harold and
Leila Y. Mathers Foundation (A.E.), and the Japanese Foundation for
AIDS Prevention (N.N.).
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Abstract
Introduction
Present address: Department of Pathology, National Institute of
Infectious Diseases, Shinjuku-ku, Tokyo, 162-8640, Japan.
