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*
Department of Molecular Microbiology and Immunology, School of Hygiene and Public Health, Johns Hopkins University, Baltimore, MD 21205; and
Department of Gynecology and Obstetrics, Johns Hopkins School of Medicine, Baltimore, MD 21205
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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75% of children born to HIV+ mothers
escape infection. The proportion of first and second trimester fetuses
harboring HIV DNA exceeds the percentage of ultimately infected
neonates (10, 11), suggesting that in vivo antiviral activity protects
the fetus from infection or results in the in utero clearance of HIV.
Data from the Women-Infants Transmission Study (1) indicate rapidly
increasing viremia within weeks of birth in nearly all infected
children, suggesting the presence of maternal or placental protective
mechanisms that inhibit viral replication before delivery and hold in
utero infection in check until the postpartum period, when it is
released from antiviral influences.
A discussion of the maternal
fetal transmission of HIV must consider
various cytokines that have been invoked as important components of the
host defense system. These include C-C chemokines (RANTES,
MIP-1
,7 MIP-1ß), the
C-X-C chemokine SDF-1, IL-16, IFN-, TNF-, and CD8+ T
cell-derived antiviral factor, or CAF (12, 13, 14, 15, 16, 17). The HIV-suppressive
effect of chemokines is mediated via competitive binding to chemokine
receptors that serve as coreceptors for HIV-1 isolates, thereby
inhibiting fusion mediated by the corresponding viral envelope proteins
(18, 19). The C-C chemokines suppress infection of lymphocytes by
macrophage-tropic CCR5-dependent HIV-1 strains, while the CXC chemokine
SDF-1 is the ligand for CXCR4 (Fusin), a coreceptor that mediates
fusion and entry of T cell-tropic HIV-1. In contrast, CAF from
HIV-infected and uninfected individuals inhibits HIV
replication in vitro by a post-entry, tropism-independent
mechanism (15).
Attempts to diagnose placental infection at the time of delivery have been negative or nonpredictive of transmission (6, 8, 10, 11, 20). Anatomic, physiologic, and immunologic barriers presented to maternal virus and cells coming in contact with placental tissues may play defensive roles (21). Transformed cell lines derived from placenta are resistant to infection by cell-free virions, but susceptible to infection by cell-associated HIV (22, 23). In contrast, studies from our laboratory have shown that fetal placental chorionic villus-derived, nontransformed stromal cells are not infectible with cell-free or cell-associated HIV. Heavily irradiated HIV-1-infected lymphocytes, when incubated with placental stromal cells, survived for up to 1 mo without detectable virus production, but virus could be rescued when fresh PHA blasts were added (24).
Preliminary unpublished studies revealed that the supernatant from placental stromal cells was sufficient for the protection of HIV-infected irradiated lymphocytes and suppression of virus production. We therefore examined the supernatant derived from placental stromal cells for antiviral activity and attempted to determine the mechanism of suppression of HIV-1. While cognizant of the fact that the observed activities may be the product of several cosegregating factors, we use the convention of calling this net activity placental factor (PF) throughout.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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First trimester (10–12 wk) chorionic villus biopsy samples from
HIV-1-seronegative subjects, obtained for metaphase chromosome
analysis, were used as a source of placental stromal cells. The method
for obtaining the cells has been described (24). Briefly,
microscopically dissected villi were digested with trypsin and
collagenase. The resultant cell suspension was cultured in Chang medium
(Irvine Scientific Santa Ana, CA) with 5% FBS for the first 2
wk, and thereafter maintained in RPMI 1640 or MEM (Life
Technologies, Gaithersburg, MD) supplemented with 10% heat-inactivated
FBS, 2 mM L-glutamine, penicillin (200 U/ml), and
streptomycin (200 µg/ml).
Preparation and storage of culture supernatant containing PF
Stromal cells were maintained in RPMI 1640 medium for 2–3 wk. Culture supernatants were collected every 3 to 4 days, and the cultures were refed with fresh medium. The supernatants were centrifuged at 1800 rpm for 10 min. The clear supernatant (PF) was pooled and stored at 4°C or frozen in aliquots at -20°C. Culture supernatant was also obtained from a fibroblast cell line CCD18 (American Type Culture Collection, Manassas, VA).
Reagents
HIV-1 strains MN and IIIB were grown and titered in H9 cells,
while HIV-1BaL was grown and titered in macrophages
(Advanced Biotechnologies, Columbia, MD). RANTES, MIP-1, and
MIP-1ß were obtained from R&D Systems (Minneapolis, MN). HeLa
CD4-LTR/ß-gal cells were obtained from AIDS Research and Reference
Reagent Program (Rockville, MD). Polyclonal anti-cyclophilin A
(CyPA) Abs were obtained from Affinity Bioreagents (Golden, CO).
Assay of antiviral activity of PF
PBMC from HIV-1-negative individuals were isolated by
Ficoll-Hypaque gradient centrifugation. The cells were washed with PBS
and stimulated with PHA-M (2.5 µg/ml) in RPMI 1640 medium (RPMI 1640
with 10% heat-inactivated FBS, 2 mM L-glutamine, 200 U/ml
penicillin, and 200 µg/ml streptomycin). Cell viability was
measured by trypan blue exclusion. PBMC stimulated with PHA for 48–72
h (PHA-PBMC) were infected with HIV-1MN,
HIV-1IIIB, or HIV-1BaL using 100
TCID50 of commercially titered stock virus per 2 x
106 PHA-PBMC. After 60 min of incubation at 37°C, cells
were washed free of unabsorbed virus and resuspended in RPMI 1640
medium plus 2 U/ml IL-2 with or without C-C chemokines, RANTES (200
ng/ml), MIP-1 (400 ng/ml) and MIP-1ß (400 ng/ml), AZT (10 µM),
fibroblast culture supernatant, or PF. The cells were cultured at
2 x 106/ml in a 24-well tissue culture plate. The
cultures were refed at 2- to 3-day intervals, when an aliquot of
supernatant was taken for measuring HIV-1 in the cultures. Cultures
were maintained up to 2 wk postinfection and monitored for HIV-1 growth
by measuring p24 Ag (Organon Teknika, Durham, NC). Cells were also
monitored for cell viability using trypan blue exclusion at 2- to 3-day
intervals. Simultaneously, 1 x 103 cells from each
culture were immunostained with FITC-CD3/phycoerythrin-CD19,
FITC-CD4/phycoerythrin-CD8 Abs (Becton Dickinson, Mountain View, CA)
and analyzed on a Coulter (Hialeah, FL) flow cytometer. Proliferation
of PBMC maintained in PF was measured by [3H]thymidine
incorporation 72 h poststimulation with PHA or 7 days
poststimulation with Candida Ag (20 µg/ml), tetanus toxoid
(2.7 U/ml), and anti-CD3 (5 ng/ml). All of the experiments were set
up as duplicate cultures and repeated three to five times.
Multinuclear activation of galactosidase indicator (MAGI) assay
HeLa CD4-LTR/ß-gal indicator cells (25) were placed in 12-well
plates at 8 x 104 to 1 x 105
cells/well and incubated at 37°C for 24 h in DMEM with 10% FBS
or PF diluted 1/1 with DMEM/10% FBS. Cell-free HIV-1MN or
HIV-1lllB at 103 TCID50 or
104 TCID50 was added and allowed to infect the
monolayer over a 2-h period. For experiments using cell-associated
virus, 1 x 106 PHA-stimulated PBMC infected with
HIV-1MN or 1 x 106 H9-IIIB cells were
cocultured with the indicator cell line for 48 h. IL-2 (1 IU/ml)
was added to wells containing stimulated PBMC. The cultures were
maintained in PF diluted 1/1 throughout the time course of the
experiment. In addition, for experiments using cell-associated virus,
all of the cultures were maintained with 10 µM AZT. As a control,
SDF-1 (100 ng/ml) was added to HeLa indicator cells 30 min before
the addition of infected PHA blasts. After 48 h, the medium in
each well was removed and the indicator cells were fixed with 2 ml of
1% formaldehyde/0.02% glutaraldehyde, at room temperature for 5 min.
The cells were then washed three times with PBS and stained with 500
µl of 4 mM potassium ferrocyanide, 4 mM potassium ferricyanide, 2 mM
magnesium chloride, and 0.4 mg/ml X-Gal for 50 min. The staining
solution was removed and the cells were washed twice with PBS. Six
x100 microscopic fields per culture were scored and averaged for blue
cells.
Measurement of infectivity of cell-free HIV-1 after direct treatment with PF
HIV-1MN or HIV-1BaL (1 x 106 TCID50) was incubated with PF (diluted 1/1 with IL-2-containing medium) for 1 h at 37°C. The treated virus diluted 1/1,000 (1 x 103 TCID50) and 1/10,000 (1 x 102 TCID50) was used to infect PHA-PBMC, as described above. HIV-1 treated with RPMI 1640 + IL-2 medium under similar conditions was used as control. Culture supernatants harvested at day 7 or day 10 postinfection were assayed for p24 content.
Measurement of infectivity of virus from PF-treated PBMC
PHA-PBMC infected with HIV-1MN for 72 h in the absence of PF were incubated with RPMI 1640 + IL-2 medium and 10 µM AZT for 3 days to insure that all subsequently recovered virus was derived from PF-treated cells. The cells were washed three times with RPMI 1640 medium and transferred to PF diluted 1/1 with RPMI 1640 + IL-2 medium. Culture supernatant was harvested after 7 days of continuous treatment with PF, filtered through 0.45-um syringe filters (Nalge Rochester, NY), and stored at -80°C or used immediately. An aliquot of culture supernatant was assayed for p24 content. Infectivity assays were performed by using viral stocks from PF-treated and untreated cultures normalized for p24 content, diluted and added at serial fourfold dilutions from 100 pg to 1.25 pg p24 per 2 x 106 PHA-PBMC. Each concentration of viral stock was used to infect six replicate cultures of fresh PHA-stimulated PBMC. Cultures were refed with RPMI 1640 + IL-2 medium at 3-day intervals. Culture supernatants from day 7 or day 10 were assayed for p24 production. Similar experiments were also conducted using HIV-1IIIB and HIV-1BaL derived from PF-treated and untreated chronically infected PM1 cells.
Polymerase chain reaction
Cellular proviral DNA PCR amplification and detection were performed on cryopreserved and fresh samples using a sensitive nested PCR protocol for amplification of a region of viral env, as described previously (26). Briefly, a two-step nested PCR amplification was performed using the following primers corresponding to HXB2, accession number K03455. Outer primers, PND1 5'-CAGCACAGTACAATGTACACATGGAAT (6949–6975 sense) and PND2 5'-ATTACAGTAGAAAAATTCCCCTCCAC (7317–7338 antisense); inner primers, PND3 5'-TGGCAGTCTAGCAGAAGAAG (7009–7028 sense) and PND4 5'-ACAATTTCTGGGTCCCCTCCT (7355–7381 antisense), generating a final product of approximately 330 bp. Products from the outer primer reaction were diluted 1/100 in water, and 5 µl of the diluted product was used for the second (inner) PCR, which was performed under identical conditions as the first reaction. PCR products were electrophoresed in a 2% agarose gel and stained with ethidium bromide.
Reverse-transcriptase assay
Cell culture supernatants were harvested and assayed for RT activity (27). Briefly, supernatants were cleared of any particulate debris by centrifugation in a Sorvall (Newtown, CT) MC 12V centrifuge at 14,000 rpm for 2 min. For each sample, 250 µl of culture supernatant was mixed with 125 µl of 30% polyethylene glycol-8000, 0.5 M NaCl, at 4°C overnight. The samples were centrifuged at 2500 rpm for 30 min (Sorvall RT 6000B), viral pellets were dissolved in 25 µl of RT lysis buffer (1% Triton X-100, 20 mM Tris-HCl, pH 7.5, 60 mM KCl, 1 mM DTT, 30% glycerol), and 10 µl of each sample was used for the RT reaction. Viral lysates were combined with 90 µl of RT reaction mixture (40 mM Tris-HCl, pH 7.8, 8 mM DTT, 10 mM MgCl2, 0.05 A260 U poly(rA)-(dT)15 (Boehringer Mannheim, Indianapolis, IN), and 2.5 µCi [3H]dTTP) at 37°C for 2 h. The reaction product was precipitated with 3 ml of chilled 10% (w/v) TCA using tRNA as a carrier. Incorporation of [3H]dTTP was determined by material bound to GF/C glass microfiber filters following five washes with chilled 5% TCA, and quantified by a Beckman LS 6500 scintillation counter.
Western blot analysis
Virion-associated viral proteins were prepared from cell culture supernatants by centrifugation at 3000 rpm for 30 min in a Sorvall 6000B centrifuge to remove cellular debris and then filtered through a 0.22-um-pore-size membrane (27). Supernatants containing virus particles were concentrated by centrifugation through a 20% sucrose cushion at 100,000 x g for 2 h in a Sorvall Ultra80. Viral pellets were resuspended in a loading dye (0.08 M Tris, pH 6.8, 2% SDS, 10% glycerol, 0.1 M DTT, and 0.2% bromophenol blue) and analyzed on 15% polyacrylamide gels. Proteins were then transferred by passive diffusion for 48 h or by electrophoretic transfer for 1 h to nitrocellulose and probed with polyclonal serum from a HIV-1-positive patient (diluted 1/200) and polyclonal rabbit anti-CyPA Ab (diluted 1/1000). Alkaline phosphatase-conjugated second Ab was used. Gel images were digitized with an Eagle Eye system (Stratagene, La Jolla, CA). Densitometric analysis was performed on a Macintosh Centris 650 computer using the public domain National Institutes of Health Image program (National Institute of Health, Bethesda, MD).
Physicochemical and immunochemical characterization of PF
Filtration and centrifugation techniques using Centriprep 10,
30, and 50 concentrators (Amicon, Beverly, MA) were used to obtain
crude m.w. fractions of PF. Different fractions of PF were tested for
the antiviral activity. Also, PF was heated at 100°C for 1 h,
followed by assay for antiviral activity, as described earlier.
Alternatively, PF was treated with 1 M HCL or 1 M NaOH for 30 min and
neutralized before use in viral growth assays. PF was also assayed
after digestion with proteinase K (100 g/ml) or Pronase-coated beads
(Sigma, St. Louis, MO) at 37°C overnight. IL-4, IL-10, IL-12,
IFN-, IFN-ß, IFN-
, TNF-, TNF-ß, granulocyte-macrophage
CSF, DHEA, DHEA-S, TGF-ß1, TGF-ß2, vascular endothelial growth
factor, platelet-derived growth factor, RANTES, MIP-1, and MIP-1ß
were measured with commercial enzyme immunoassay kits (R&D Systems).
Single test Limulus amebocyte lysate assay (BioWhittaker,
Gaithersburg, MD) was used to exclude the presence of endotoxin in PF.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PF showed activity against BaL, MN, and IIIB strains of HIV-1,
reducing the levels of p24 to less than 2% of those obtained from
untreated cultures or cultures exposed to supernatants from a
fibroblast line, CCD18. This effect was comparable with suppression
with 10 µM AZT (Table I
). Inhibition of
viral growth was less pronounced with the C-C chemokines RANTES (200
ng/ml), MIP-1 (400 ng/ml), and MIP-1ß (400 ng/ml) added alone or
in combination. Moreover, although MIP-1 alone had some effect on
HIV-1MN, inhibitory activity of the C-C chemokines was
restricted primarily to the macrophage-tropic strain
HIV-1BaL. Partial loss of antiviral activity was observed
when PF was used at 1/10 dilution. Occasional batches of PF
demonstrated antiviral activity even at 1/100 dilution (Fig. 1), but activity was always lost at
1/1000. Flow-cytometric analysis showed that PF-treated cultures
maintained significantly higher CD4+ cell proportions as
compared with the infected cells maintained in the absence of PF
(Fig. 2).
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).
PHA-PBMC infected with HIV-1 and maintained in PF demonstrated enhanced
survival as compared with parallel control cultures (Fig. 1
). This was
further confirmed by the absence of DNA fragmentation when
HIV-1-infected cells maintained in the continued presence of PF were
analyzed by agarose gel electrophoresis (data not shown). Thus,
reduction in measured p24 per viable cell was even greater than the
reduction in supernatant concentration.
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No direct effect on the infectivity of cell-free viruses.
Pretreatment of cell-free HIV-1MN or HIV-1BaL
(1 x 106 TCID50) with PF for 1 h at
37°C did not have any direct effect on the infectivity of virus
compared with IL-2-containing medium under similar conditions (Fig. 3). These results were in contrast to the
antiviral effect observed when PHA-PBMC infected with cell-free HIV-1
were maintained in the continuous presence of PF (Fig. 1).
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. In the presence of PF, there was a
10-fold or greater decrease in the number of infected indicator cells
following 48-h exposure to cell-free HIV-1MN and
HIV-1IIIB. This reflects decreased first and/or second
round productive infection. However, PF treatment did not decrease the
number of blue indicator cells when PHA-PBMC infected with
HIV-1MN were cocultured with the Hela-CD4-LTR-ß-gal cells
for 48 h in the presence of AZT (i.e., in the absence of de novo
reverse transcription, Table III). Therefore, fusion of infected PBMC,
permitting preformed Tat to enter HeLa cells and induce ß-gal (Fig. 4), was unimpaired. In contrast, there
was a 20-fold reduction in the number of blue cells scored after
exposure to infected PHA-PBMC in the presence of 100 ng/ml of SDF-1,
the natural ligand for CXCR4. Viral DNA could be amplified from HeLa
indicator cells that had been exposed to cell-free HIV-1MN
(103 TCID50/ml) for 1 h in the presence of
PF, washed free of nonadherent virions, and washed again at 24 h
after trypsinization and DNase treatment to remove any contaminating
input DNA or surface-bound but unfused virions. This indicated that
virus entry and initial reverse transcription were not blocked (Fig. 5).
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To test the effect of PF on viral production from infected cells,
PHA-PBMC infected with HIV-1 (IIIB, MN, or BaL) were grown in IL-2
medium for 3 days, followed by addition of 10 µM AZT for an
additional 72 h. Thereafter, the cells were washed free of AZT and
then treated with PF diluted 1/1 with IL-2-containing RPMI 1640 medium.
Culture supernatants from day 7 were assayed for p24 and RT and
normalized for p24 content. The difference in p24 and RT levels between
supernatant from the PF-treated and untreated cultures was
proportionally related (p < 0.001,
r2 = 0.85). Infectivity was evaluated by adding
the supernatant fluid to fresh PHA-PBMC. Supernatants were diluted at
least 1000-fold to eliminate any antiviral effect of residual PF before
being added at serial fourfold dilutions (100 to 1.25 pg) to the
PHA-PBMC. Measurement of supernatant p24 on days 3, 7, and 10
postinfection showed that PF-treated HIV-1 was 10–100-fold less
infectious than untreated control viruses analyzed by the method of
Reed and Muench (28). Day 7 p24 values gave the earliest detectable
time point for limit dilution inocula analysis (Fig. 6). Similar results were obtained when
HIV-1IIIB or HIV-1BaL obtained from PF-treated
PM1 cells were used to infect PHA-PBMC (not shown).
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Sucrose gradient-purified virions produced by PF-treated PBMC were
subjected to Western blot analysis with polyclonal sera containing Abs
to all the major virus-encoded proteins. Compared with untreated
virions, no qualitative differences were observed in HIV-1-encoded
structural proteins. Viral protease activity was intact, as evidenced
by complete processing of the Gag and Gag-Pol polyproteins to cleavage
products p24, p51, and p66. Preliminary data from Western-blotted
proteins associated with purified HIV-1 virions, probed with
anti-HIV-1 and anti-CyPA Ab (Fig. 7), revealed that virions produced by
PBMC maintained in the continuous presence of PF incorporated two- to
threefold less CyPA per unit p24 than did untreated virions (Fig. 8).
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PF was found to be relatively resistant to heat and acid
treatments. Cultures treated with PF that had been heated to 100°C
for 1 h (Fig. 9) or treated with 1 M
HCl to a pH of 4 for 30 min and neutralized before use (data not shown)
showed minimal loss of antiviral activity as compared with the
untreated PF. However, treatment with 1 M NaOH to attain a transient pH
of 9.5 for 30 min before normalization abrogated the antiviral effect
of PF. Antiviral activity was found to be restricted to fractions less
than 50 kDa.
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, IFN-ß, IFN-,
TNF-, TNF-ß, granulocyte-macrophage CSF, DHEA, DHEA-S, TGF-ß1,
TGF-ß2, vascular endothelial growth factor, platelet-derived growth
factor, RANTES, MIP-1, and MIP-1ß, as determined by enzyme
immunoassays (R&D Systems). Trace amounts of IL-8 (8.65 ng/ml) detected
in PF are far below the 100 ng/ml concentration reported to inhibit HIV
replication in naturally infected cells. Also, IL-8 reportedly has no
effect on HIV replication in acutely infected CD4+ cells
(29). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In contrast to earlier reports demonstrating increased virus production
when virus-induced apoptosis was inhibited by transfecting cells with
the bcl-2 gene (33), we found decreased virus production and
delayed cell death when lymphoblasts were treated with conditioned
medium from placental stromal cells. The observed antiviral activity of
PF may be independent of, and mechanistically distinct from, any
postulated antiapoptotic effect. On the other hand, our results are
consistent with more recent findings of Strack et al. showing
transfected HIV-1 protease-depleted cells of bcl-2, and
promoted apoptosis and oxidative stress-dependent activation of nuclear
factor-
B. Overexpression of bcl-2 decreased both
HIV-induced cell death and production of virus, while factors that
blocked the intracellular cleavage of bcl-2 prevented cell
death following HIV infection of lymphocytes (34). In our experiments,
PF treatment had no apparent effect on protease function, as shown by
patterns of viral proteins in supernatants from PF-treated cells.
However, it is still possible that PF acts by suppressing intracellular
production of reactive oxygen intermediates.
PF lacks identity with the C-C chemokines and other cytokines reported to have antiviral activity. The stage of HIV replication at which PF acts has not been precisely defined. DNA PCR amplification suggests that cell-free virus can enter and at least initiate reverse transcription up to the env gene in normal quantities, but MAGI assay results indicate a block before high Tat production. Further evidence of unimpeded HIV envelope-mediated membrane fusion comes from the experiments using HeLa CD4 ß-gal cells and infected PBMC. At the other end of the life cycle, 10–100-fold decreased infectivity of released material was seen. Together, decreased susceptibility of PF-treated cells, and decreased infectivity of released material from HIV-infected PF-treated cells, can account for the decrease in virus production observed over 1–2 wk in culture.
Previous studies have shown that a cellular peptidyl-prolyl isomerase, CyPA, is incorporated into HIV-1 virions via direct interaction with the HIV-1 Gag protein. Disruption of the Gag-CyPA interaction results in noninfectious HIV-1 particles (35). Data from the present study indicate a two- to threefold decrease in the ratio of CyPA:p24 in virion particles purified from culture supernatants of PF-treated cells, which may partly account for their decreased infectivity. These findings must be viewed carefully, in light of recent reports showing the presence of host cell-derived vesicles copurifying with preparations of HIV-1 virions (36). It is possible that the true CyPA:Gag ratio is even more skewed in virions from PF-treated cells, since supernatants from uninfected cultures may contain large amounts of CyPA that would minimize net measured differences.
PF is a small, heat- and pH-stable molecule with broad activity against different strains of HIV-1, and does not share identity with any other known cytokines. The CD8+ T cell-derived antiviral factor reported by Levy and others to inhibit HIV replication without target cell lysis or suppression of target CD4+ T cell activation is also reported to be heat and pH stable and act against a broad range of HIV isolates with different tropisms (14, 16, 37). CAF has been reported to suppress HIV transcription (15, 38), an activity we have not observed clearly with PF, although inhibition of Tat expression as seen in experiments with HeLa-CD4 cells may suggest a block at the transcriptional level. More recently, a CXC chemokine derived from T cell lines, MDC, has been reported to broadly inhibit HIV in vitro (39). MDC is thus a candidate for the elusive Levy Factor, but has important differences from PF, notably, lack of effect on PM1 cells, and susceptibility to heat denaturation.
Our results suggest that PF could play an important role in protecting the fetus and newborn. Further studies to purify and molecularly characterize the factor(s) responsible for PF activity are in progress. Identification of a naturally produced molecule of human origin with anti-HIV activity holds therapeutic potential.
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Acknowledgments |
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Footnotes |
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2 Current address: Department of Microbiology and Immunology, School of Medicine, University of Maryland, Baltimore, MD 21201.
3 Current address: Division of Clinical Immunology, Johns Hopkins Bay View Medical Center, 5501 Hopkins Bayview, Baltimore, MD 21224.
4 Current address: Department of Infectious Diseases, Johns Hopkins School of Medicine, Baltimore, MD 21287.
5 Current address: Department of Clinical Virology, Christian Medical College, Vellore, India 632004.
6 Address correspondence and reprint requests to Dr. David H. Schwartz, Department of Molecular Microbiology and Immunology, School of Hygiene and Public Health, Johns Hopkins University, 615 N. Wolfe Street, Baltimore, MD 21205.
7 Abbreviations used in this paper: MIP, macrophage-inflammatory protein; AZT, 3'-azido-3-deoxythymidine; ß-gal, ß-galactosidase; CAF, CD8+ cell antiviral factor; CyPA, cyclophilin A; DHEA, dehydroepiandrosterone; LTR, long terminal repeat; MAGI, multinuclear activation of galactosidase indicator; PF, placental factor; RT, reverse transcription; SDF, stromal derived factor; TCID50, tissue culture 50% infective dose.
Received for publication September 12, 1997. Accepted for publication July 24, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, MIP-1ß as the major HIV-suppressive factors produced by CD8+ T cells. Science 270:1811. inhibits entry of human immunodeficiency virus type 1 into primary human macrophages: a selective role for the 75-kilodalton receptor. J. Virol. 70:7388.[Abstract]
, and MIP-1ß suppress HIV-1 replication in monocytes/macrophages. Proc. Natl. Acad. Sci. USA 93:15341.fetal transmission. Proc. Natl. Acad. Sci. USA 92:978.This article has been cited by other articles:
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  J. M. Fakruddin, R. A. Lempicki, R. J. Gorelick, J. Yang, J. W. Adelsberger, A. J. Garcia-Pineres, L. A. Pinto, H. C. Lane, and T. Imamichi Noninfectious papilloma virus-like particles inhibit HIV-1 replication: implications for immune control of HIV-1 infection by IL-27 Blood, March 1, 2007; 109(5): 1841 - 1849. [Abstract] [Full Text] [PDF]
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 M. Derrien, A. Faye, G. Dolcini, G. Chaouat, F. Barre-Sinoussi, and E. Menu Impact of the Placental Cytokine-Chemokine Balance on Regulation of Cell-Cell Contact-Induced Human Immunodeficiency Virus Type 1 Translocation across a Trophoblastic Barrier In Vitro J. Virol., October 1, 2005; 79(19): 12304 - 12310. [Abstract] [Full Text] [PDF]
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