Arora, Reetakshi PhD*; Bull, Lara PhD†; Siwak, Edward B PhD*; Thippeshappa, Rajesh MS*; Arduino, Roberto C MD†; Kimata, Jason T PhD*
Viral load has a strong predictive value for HIV-1 disease progression.1-3 Studies suggest that viral set points are the result of a complex and dynamic interplay between the infecting virus and the host.4,5 Both host genetic factors (eg, HLA Class I; chemokine receptors, CCR2, CCR5; chemokines, CCL3L1, CCL4, CCL2) and viral genetic factors have been implicated in influencing the level of plasma viral load and disease progression.6-10 Absent from these earlier studies is the evaluation of whether the efficiency of cell-mediated virus transmission may also influence viral load in the host.
Dendritic cells (DCs) have been hypothesized to play a key role in HIV-1 infection and replication.11 Notably, DCs are among the first cell types, along with CD4+ T cells and macrophages, that encounter HIV-1 after mucosal transmission.12 HIV-1 exploits the biological function of DCs to facilitate its transport from the site of entry at mucosal tissue to lymphoid tissues where robust replication occurs in CD4+ T cells. Additionally, DCs efficiently capture and transmit HIV-1 to T cells via infectious synapses that concentrate HIV and viral receptors.13 Moreover, it has also been shown that a cell-contact-dependent mechanism of HIV-1 infection is more efficient than cell-free infection, suggesting that cell-cell interactions may be critical for transmission and replication and that contact between DCs and T cell may play a key role in driving HIV-1 replication.14,15 Productive infection of DCs by HIV-1 is not required for enhanced infection as DCs express a number of viral attachment factors including DC-specific ICAM3-grabbing nonintegrin (DC-SIGN), syndecans, CD4, and glycosphingolipids that can bind, concentrate, and retain infectious virions for up to 48 hours.11 Together, these data suggest that adaptation of HIV-1 to antigen-presenting cell-T cell interactions may influence replication and persistence. However, although significant progress has been made toward understanding the mechanisms underlying DC-mediated transmission of HIV-1, whether these interactions contribute to the level of viral replication in the host remains largely unexplored.
In the present study, we sought to determine whether replicative capacity of HIV-1 contributes to the level of plasma viral load. Using HIV-1 variants isolated from chronically infected antiretroviral treatment (ART)-naive individuals, we investigated the relationship between ex vivo infectivity and replication capacity of CCR5-tropic HIV-1 and plasma viral RNA levels and CD4+ cell counts in the host. In particular, as viral replication mainly occurs in lymphoid tissues of the host and is likely promoted by antigen-presenting cell-T cell interactions,14,16-18 we examined whether the level of HIV-1 infection of T cells mediated by DCs in trans would correlate with plasma viral RNA measurements from the infected individuals. Our data suggest a significant role for viral replicative capacity in determining the level of plasma viral load and further suggest a role for DCs in driving viral replication in T cells.
A cohort of individuals infected with HIV-1 were recruited from patients seeking treatment at the Thomas Street Health Center, Houston, TX. The subjects had broad range of plasma viral RNA levels (ranging <1000 to >100,000 RNA copies/mL) and CD4+ cell counts (ranging from 26 to 1027 cell/mm3) (Table 1). All the individuals participating in the study were ART naive at the time of blood draw. Informed consent was obtained from all the participants in this study in accordance with University of Texas Health Science Center (UTHSC)--Houston Committee for the Protection of Human Subjects. Buffy coats from healthy anonymous donors were obtained from Gulf Coast Regional Blood Center, Houston, Texas.
Quantification of Plasma Viral Load
Plasma HIV-1 RNA was quantified by using the Amplicor HIV-1 Monitor test, version 1.5 (Roche Diagnostics, Indianapolis, IN) with a detection limit of 400 copies per milliliter.
Analysis of CD4+ Cell Count
Immunophenotyping for CD4+ cells was done by flow cytometry. Peripheral blood mononuclear cells (PBMCs) isolated from HIV-1-infected individuals were stained with CD4 monoclonal antibody according to standard procedures for cytometric analysis (BD Biosciences, San Jose, CA).
Genomic DNA was isolated from the PBMCs of HIV-1-infected individuals or healthy donors using QIAamp DNA Blood Mini kit (Qiagen, Germantown, MD). CCR5 was amplified by PCR, and the DNA products were analyzed on ethidium bromide-stained 3% agarose gels as previously described.19 Homozygous wild-type and homozygous mutant for delta 32 basepairs deletion were run in parallel as control.
HIV-1 Isolation and Titer Determination
HIV-1 was isolated from the infected PBMCs by coculturing with PBMCs from anonymous healthy donors according to standard techniques.20 Donor PBMCs used for HIV-1 isolation and titer determination were first screened for the CCR5 delta 32 mutant allele. Donor samples positive for the mutation were excluded from being used for virus isolation or titer determination. Briefly, PBMCs from HIV-1-infected individuals were diluted at a concentration of 1 × 106 in RPMI 1640 medium supplemented with 20% heat inactivated fetal bovine serum, 2mM L-glutamine, 100 U/mL of penicillin, 0.1 μg/mL of streptomycin constituting complete RPMI and Interleukin-2 (IL-2) (Roche Diagnostics) (50 U/mL). PBMCs from infected individuals were then cocultured with equal number of Phytohemagglutinin (PHA) and IL-2 stimulated healthy donor PBMCs (1 × 106/mL) in T-25 flask. Every week, culture supernatants were tested for p24 HIV-1 antigen by enzyme-linked immunosorbent assay (ELISA) kit (Beckman Coulter, Fullerton, CA). At the same time, half of the culture volume was replaced with fresh PHA and IL-2 stimulated donor PBMCs and propagated for 3 weeks. From these cultures, the cell-free supernatant was stored at −70°C until use. Viral stock titers were determined using healthy donors' PBMCs that were stimulated with PHA and IL-2 (PHA-lymphoblasts) for 3 days as previously described.20 The 50% tissue culture infective dose was calculated by the Spearman-Karber method.
Characterization of Coreceptor Usage
GHOST indicator cells were used to determine coreceptor usage of each viral isolate. GHOST-X4 and GHOST-Hi5 cell lines were obtained from AIDS Research and Reference Reagent Program. The cell lines were maintained in Dulbecco's modified Eagle's medium (DMEM) containing 10% heat inactivated fetal bovine serum, 2 mM L-glutamine, 100 U of penicillin per mL, and 100 μg of Streptomycin per mL (DMEM). GHOST assays were performed as described earlier.21
In Vitro HIV Replication Capacity
Human monocyte-derived DCs were generated from PBMCs of HIV-negative anonymous blood donors as previously described.22 Briefly, CD14+ monocytes were isolated using anti-CD14+ microbeads and miniMACS systems according to manufacturer instructions (Miltenyi Biotec, Auburn, CA). The CD14+ monocytes were then washed with complete RPMI media and cultured with 1000 U/mL of granulocyte-macrophage colony stimulating factor (GM-CSF) and Interleukin-4 (IL-4) (R&D Systems, Minnneapolis, MN) in complete RPMI media for 7 days to generate DCs. By fluorescence-activated cell sorting (FACS) analysis, high expression (>95%) of DC-SIGN (CD209) on DCs confirmed the purity of the population.
DCs (1 × 105) were incubated with 2000 50% tissue culture infections dose (TCID50) of different viral isolates in triplicates in a total volume of 200 μL in U-bottom 96-well plate and incubated for 3 hours. The cells were then washed 3 times with RPMI media and cocultured with 2 × 105 peripheral blood lymphocytes from the same donor. The cells were cocultured for 14 days in presence of IL-2 (50 U/mL) in complete RPMI. Supernatants were collected every second day and replenished with fresh media and IL-2. Viral replication was assessed by measuring the amount p24 HIV-1 antigen in culture supernatants by ELISA (Beckman Coulter, Fullerton, CA). Replication rates were determined by calculating the slope (p24 pg/mL/day) of the exponential phase p24gag accumulation. All the donor PBMCs were genotyped for delta 32 basepairs deletion in CCR5 gene. Only donors who were homozygous wild-type at the CCR5 locus were used to measure replication rates.
Nef Amplification, Sequencing, and Infection Assays
Genomic DNA was isolated from HIV-1-infected PBMCs as described above. Variant nef alleles from each patient were amplified by nested Polymerase Chain Reaction using primers as described earlier23,24 and cloned into TOPO vector pCR 2.1 (Invitrogen) for sequencing. Sequences were aligned using MacVector (version 10.0.2) and compared with HIV-1NL4-3 as a standard sequence. Nef alleles were further cloned into the bicistronic expression vector, pIRES2-EGFP (BD Biosciences Clonetech, Palo Alto, CA). For expression analyses, cell lysates, prepared after the transfection of Nef-IRES-EGFP variants in 293 T cells, were run on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Bio Rad, Hercules, CA) and immunoblotted against Nef antisera24 or an anti-green fluorescent protein (GFP) monoclonal antibody (Invitrogen). Furthermore, to assess viral infectivity, single-cycle infection assays were also carried out in DC capture-transfer assays using HIV-luciferase (HIV-luc) pseudotyped with a R5-tropic envelope (Q168env225) and trans complemented with Nef variants. Infectivity was measured by luciferase assay. Incorporation of Nef into virions was also assessed by recovering virions from cotransfected cell culture supernatants by centrifugation at 23,600g for 1 hour at 4°C and analyzing viral lysates after SDS-PAGE by immunoblotting using Nef antisera and p24 specific antibodies.
Real-Time PCR for the Determination of Gene Copy Number of CCL3L1
Taqman assay for CCL3L1 was similar to that used earlier.6 Briefly, real-time PCR was done using an ABI 7500 (Applied BioSystems Inc.) with the following universal cycling conditions: 2minutes at 50°C, 10 minutes at 95°C, and 40 cycles of 15 seconds at 95°C, and 1 minute at 60°C. All the samples were tested in triplicate. Calculations were based on the estimates of template quantities, after comparing the threshold cycle (Ct) numbers at which positive (over background) fluorescence was detected.
Correlations between viral replicative capacity and plasma viral load were examined by linear regression. The relationships between viral replicative capacity or plasma viral RNA loads and CD4+ T-cell counts were analyzed by Poisson regression. Pearson correlation was used to test for correlation between relative Nef-mediated enhancement of infection activity and plasma viral load. Correlation between plasma viral load and copy number of CCL3L1 was tested using Spearman correlation. Statistical analyses were performed using STATA 10 software.
HIV-1 isolation was attempted from PBMC specimens collected from 20 chronically HIV-1-infected individuals before ART. Characteristics of the HIV-1-infected individuals are presented in Table 1. Subjects were selected, irrespective of disease status, and included individuals with a wide range of plasma viral RNA levels. The group included 5 females and 15 males of mean age 37.8 ± 4.9 (range, 33-46) and 38.13 ± 4.8 (range, 19-50), respectively. We were not able to recover infectious virus from PBMCs of 2 patients (#7 and #365). The median plasma HIV-1 RNA level was 84,252.5 copies per milliliter (range, 428-750,000 copies/mL) and, the median CD4+ cell count was 316 cells per cubic millimeter (range, 26-1027 cells/mm3). The HIV-1-infected individuals included in this study showed significant inverse relationship between the plasma viral load and CD4+ cell counts (P = 0.02; 95% CI: −0.512 to −0.044) (Fig. 1).
Coreceptor Specificity of Viral Isolates
To determine the coreceptor usage of the viral isolates, we used the GHOST indicator cell lines expressing either the CCR5 or CXCR4 coreceptors. HIV-1 clones, Yu2 and NL4-3, were used as controls for R5 and X4-dependent infections, respectively. GFP expression was detected in GHOST-Hi5 cells in case of infection by all the viral isolates, indicating that all the viral isolates were R5 tropic except for isolate JA, which was R5X4 dual-tropic by this assay (Table 1).
Relationship Between HIV-1 Replication Capacity and Plasma Viral RNA Concentration
Initially, we examined the replicative capacity of the viral isolates in PHA lymphoblasts. However, several of the viral isolates replicated poorly or inconsistently, prompting us to consider an alternative cell culture system to evaluate viral replicative capacity (data not shown). We, therefore, measured the replication rate of viral isolates by performing p24 HIV-1 antigen ELISA on supernatants from DC capture-transfer assays. Importantly, viral replication has been shown to be most robust in these culture conditions, and DCs may enhance infection and replication in the host.11,14,15,26-28 Viral replication was compared using cells derived from the same donor. The replication rates were determined from the slope of the exponential phase of p24 production. Of 18 viral isolates, 3 did not show any detectable p24 antigen by ELISA. The median of replication rate was 0.2763 log10 pg/mL/day (range, 0.0653-0.7389 log10 pg/mL/day). There was a significant linear relationship (r2 = 0.283, P = 0.04) between the plasma viral load and HIV-1 replication capacity (Fig. 2A), indicating that replication capacity of the virus may significantly influence plasma viral load in the host. Additionally, HIV-1 replication of selected viral isolates was determined in monocyte-derived dendritic cells-peripheral blood lymphocytes (DC-PBL) cocultures using cells from a different donor to compare the interindividual variability. Comparable replication rates were observed with the second donor (Table 2).
HIV-1 Replication Capacity and CD4+ T-Cell Count
To determine if there was also a relationship between replication capacity (RC) values and CD4+ T-cell counts. Correlation between replication rates of the viral isolates and CD4+ T-cell count in the host was statistically determined by Poisson regression analysis. The data demonstrate a decreasing trend for CD4+ T-cell counts with an increase in replication rate (Fig. 2B), although it was not statistically significant (P = 0.09; 95% CI: −3.02 to 0.23).
Plasma HIV-1 Viral Load and Viral Genetic Determinant
Because Nef function significantly affects viral replication, we examined whether mutations in Nef affecting viral infectivity accounted for the differences in viral load or replicative capacity in DC-mediated in trans infection of T cells. We amplified and sequenced the predominant nef variants from the HIV-1-infected patients and confirmed their expression in 293 T cells by immunoblot (Fig. 3A). No null nef mutants were identified and Nef protein was detected from each allele. Furthermore, to examine the effect of Nef on viral infectivity in DC capture-transfer assays, a nef deleted HIV-1 clone containing a luciferase reporter was complemented with each of the Nef variants in trans and a CCR5-tropic Env. HIV-1NL4-3 nef and a construct with no nef gene served as positive and negative controls, respectively, and comparisons were made with the activity of the HIV-1NL4-3 nef allele to calculate a relative viral infectivity value. Interestingly, incorporation of Nef in virions was only observed in 6 of the 15 viruses (Fig. 3B). Furthermore, there was no significant difference (P = 0.89) between the average infectivity of the Nef-negative viruses [1.45 ± 0.26 (mean + standard error of the mean)] and Nef-positive viruses (1.51 ± 0.40), suggesting that virion association of Nef does not enhance viral infectivity. However, we found a significant linear association (r = 0.5362; P = 0.039) between relative Nef activity and viral load (Fig. 3C), indicating a significant influence of Nef enhancement of infectivity on viral load in the host. There was also a weak, but not significant, association observed between Nef activity and in vitro replicative capacity of viral isolates (data not shown). These results were confirmed in cells derived from 2 additional donors.
Plasma HIV-1 Viral Load and Host Genetic Determinants
Because all of the viral isolates tested were CCR5-tropic, we examine the virus donors for the 32 basepairs deletion in CCR5 gene and copy number of CCL3L1 genes to exclude the possibility that genetic determinants in these genes accounted for the differences in viral loads. By PCR analysis, we did not observe any CCR5-delta32 homozygous mutant. All the HIV-1-infected individuals were homozygous wild-type for CCR5 locus except subject #JA (Table 1). Next, we examined the gene copy number of CCL3L1 in HIV-1-infected individuals by quantitative real-time PCR (Table 1). No correlation between the plasma viral RNA levels and copy number of CCL3L1 gene in HIV-1-infected individuals was observed (P > 0.05, nonparametric Spearman correlation).
HIV-1 disease progression has been correlated with plasma viral RNA levels,1-3 although the viral and host determinants contributing to the level of virus are incompletely understood. Early studies demonstrated that phenotypic changes in the infecting virus were associated with progression to AIDS, particularly the appearance of cytopathic variants that replicated efficiently in CD4+ T cells.9 Indeed, our studies with the simian immunodeficiency virus (SIV)-macaque model demonstrated that the appearance of variants with greater replicative capacity in CD4+ T cells influenced plasma viral load and the rate of disease progression.29 Importantly, highly pathogenic variants also show greater replicative capacity in DC T-cell cocultures compared with the minimally pathogenic parental virus,26,30 suggesting that SIV, and by extension HIV-1, evolve during infection to exploit DC-T cell interactions for viral replication. The data presented here support this hypothesis and show a direct relationship between plasma viral RNA levels in the host and infectivity and replication capacity of the infecting viruses in DC T-cell cocultures.
We conducted a study with 20 HIV-1-infected individuals who were antiretroviral naive at the time of virus isolation to evaluate the replication rate of HIV-1 viral isolates in vitro from the patients with low or high viral load irrespective of disease status. Several approaches have been used to measure in vitro HIV-1 replication capacity.31 Most commonly, replication rate has been studied by measuring the p24 HIV-1 antigen in supernatants after infecting the PHA lymphoblasts from healthy donors. Because several of the viruses we isolated replicate poorly in PHA-lymphoblasts, we performed virion capture and transfer assays with DCs and T cells to study the replication rate of R5-tropic viral isolates. In particular, a DC-mediated capture-transfer assay was used because HIV-1 infection of CD4+ T cells has been shown to be most efficient when virions are presented in association with macrophages or DCs.11,14,15,26-28 We observed a significant linear relationship between HIV-1 replication rate in DC-PBL cocultures and plasma HIV-1 RNA concentration in the host. Thus, these data suggest that replication capacity of the virus is a major determinant of plasma viral load, and hence, disease progression. Furthermore, variants that efficiently utilize antigen-presenting cell-T cell interactions may have a replicative advantage in the host. Variation in transmission efficiency of different isolates may also reflect upon replication capacity, but transmission differences could be in part due to the levels of expression of DC-SIGN,32 DC-SIGN-mediated infectious synapse formation,33 or level of CD4 coexpression.34
Because our analysis remained limited to R5-tropic HIV-1, we evaluated the role of genetic determinants of the virus contributing to plasma viral load other than those in envelope that alter coreceptor usage. Viruses carrying deleterious mutations in the nef gene have been shown to be less fit than the wild-type viruses.35 Furthermore, the slow replicating viruses associated with a low viral load in case of long-term survivors has been reported to harbor defect in the nef gene. Thus, we PCR amplified and sequenced Nef variants and performed single-cycle infection assays in DC-PBL cocultures to assess Nef-mediated viral infectivity. There was significant linear association observed between the viral infectivity enhancement by the predominant Nef variants in DC-mediated in trans infection of T cells and viral RNA levels in the plasma of the hosts from which the nef alleles were cloned, further demonstrating the importance of Nef-mediated enhancement of infectivity on viral replication in vivo.35-37 Interestingly, although Nef protein was expressed from all nef alleles, only a third of the Nef proteins were found to be virion associated. Moreover, there was no correlation between virion incorporation of Nef and enhanced viral infectivity. These data are consistent with recent studies indicating that Nef affects virion infectivity during virion biogenesis in the infected cell, and not because of incorporation into virions.38-40 Finally, although our study group showed strong correlation between replicative capacity and viral RNA levels in plasma, we observed little association between Nef activity and replicative capacity, indicating that there may be other viral factors influencing RC either independent or in conjunction with Nef-mediated enhancement of infectivity.
Because all the viral isolates were R5-tropic, we examined whether CCR5 gene deletion (CCR5Δ32) or gene copy number of CCL3L1 was influencing plasma viral load. Both CCR5 and CCL3L1 have been shown to have a strong influence on the disease progression.6,10 All the individuals studied in our study were wild-type homozygous for CCR5 gene, except for 1 who was heterozygous. There was no significant association between plasma viral load and CCL3L1 copy number. Although recent findings implicated that the individuals with lower than the average number of CCL3L1 genes for their ethnic group were found to be more susceptible to HIV-1 infection in contrast to the ones having more than the average number of CCL3L1 copies, large cohorts were required to observe this effect.6,41 Contrary to this and in support of our observation, Shao et al42 recently reported that copy number variations in CCL3L1 failed to exert major impact on the outcome after HIV-1 infection in smaller adolescent population. In view of these observations, our data suggest that replication capacity of HIV-1 is a significant determinant of viral load. What remains unclear are the cellular factors affecting replication capacity of the virus.
In conclusion, our study shows a linear correlation between replication capacity of HIV-1 variants with plasma viral load. Because this was observed in the context of DC-PBL cocultures, it supports our earlier observations with the SIV-macaque model that the efficiency of replication in response to DC-T cell interactions influences plasma viral load. Thus, adaptation of HIV-1 to antigen-presenting cell-T cell interactions during the course of infection may drive viral replication and disease progression.
We thank Claudia Kozinetz and Xiaoying Yu of the Design and Analysis Core of the Baylor-UT Houston Center for AIDS Research (CFAR) for assistance with statistical analyses. The following reagents were obtained through the National Institutes of Health AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health: IL-2 from Roche Diagnostics; GHOST indicator cell lines from Dr Vineet N. KewalRamani and Dr Dan R. Littman; pYU-2 from Dr Beatrice Hahn and Dr George Shaw; pNL4-3 from Dr Malcolm Martin.
1. Mellors JW, Kingsley LA, Rinaldo CR Jr, et al. Quantitation of HIV-1 RNA in plasma predicts outcome after seroconversion. Ann Intern Med. 1995;122:573-579.
2. Mellors JW, Rinaldo CR Jr, Gupta P, et al. Prognosis in HIV-1 infection predicted by the quantity of virus in plasma. Science. 1996;272:1167-1170.
3. O'Brien WA, Hartigan PM, Martin D, et al. Changes in plasma HIV-1 RNA and CD4+ lymphocyte counts and the risk of progression to AIDS. Veterans Affairs Cooperative Study Group on AIDS. N Engl J Med. 1996;334:426-431.
4. Desrosiers RC. Strategies used by human immunodeficiency virus that allow persistent viral replication. Nat Med. 1999;5:723-725.
5. Piatak M Jr, Saag MS, Yang LC, et al. High levels of HIV-1 in plasma during all stages of infection determined by competitive PCR. Science. 1993;259:1749-1754.
6. Gonzalez E, Kulkarni H, Bolivar H, et al. The influence of CCL3L1 gene-containing segmental duplications on HIV-1/AIDS susceptibility. Science. 2005;307:1434-1440.
7. Hogan CM, Hammer SM. Host determinants in HIV infection and disease. Part 2: genetic factors and implications for antiretroviral therapeutics. Ann Intern Med. 2001;134:978-996.
8. Hogan CM, Hammer SM. Host determinants in HIV infection and disease. Part 1: cellular and humoral immune responses. Ann Intern Med. 2001;134(9 Pt 1):761-776.
9. Kimata JT. HIV-1 fitness and disease progression: insights from the SIV-macaque model. Curr HIV Res. 2006;4:65-77.
10. Liu R, Paxton WA, Choe S, et al. Homozygous defect in HIV-1 coreceptor accounts for resistance of some multiply-exposed individuals to HIV-1 infection. Cell. 1996;86:367-377.
11. Wu L, KewalRamani VN. Dendritic-cell interactions with HIV: infection and viral dissemination. Nat Rev Immunol. 2006;6:859-868.
12. Hladik F, McElrath MJ. Setting the stage: host invasion by HIV. Nat Rev Immunol. 2008;8:447-457.
13. McDonald D, Wu L, Bohks SM, et al. Recruitment of HIV and its receptors to dendritic cell-T cell junctions. Science. 2003;300:1295-1297.
14. Cameron PU, Freudenthal PS, Barker JM, et al. Dendritic cells exposed to human immunodeficiency virus type-1 transmit a vigorous cytopathic infection to CD4+ T cells. Science. 1992;257:383-387.
15. Gummuluru S, KewalRamani VN, Emerman M. Dendritic cell-mediated viral transfer to T cells is required for human immunodeficiency virus type 1 persistence in the face of rapid cell turnover. J Virol. 2002;76:10692-10701.
16. Brenchley JM, Schacker TW, Ruff LE, et al. CD4+ T cell depletion during all stages of HIV disease occurs predominantly in the gastrointestinal tract. J Exp Med. 2004;200:749-759.
17. Haase AT. Perils at mucosal front lines for HIV and SIV and their hosts. Nat Rev Immunol. 2005;5:783-792.
18. Pantaleo G, Graziosi C, Demarest JF, et al. HIV infection is active and progressive in lymphoid tissue during the clinically latent stage of disease. Nature. 1993;362:355-358.
19. Huang Y, Paxton WA, Wolinsky SM, et al. The role of a mutant CCR5 allele in HIV-1 transmission and disease progression. Nat Med. 1996;2:1240-1243.
20. Hollinger FB, Bremer JW, Myers LE, et al. Standardization of sensitive human immunodeficiency virus coculture procedures and establishment of a multicenter quality assurance program for the AIDS Clinical Trials Group. The NIH/NIAID/DAIDS/ACTG Virology Laboratories. J Clin Microbiol. 1992;30:1787-1794.
21. Kimata JT, Gosink JJ, KewalRamani VN, et al. Coreceptor specificity of temporal variants of simian immunodeficiency virus Mne. J Virol. 1999;73:1655-1660.
22. Yu Kimata MT, Cella M, Biggins JE, et al. Capture and transfer of simian immunodeficiency virus by macaque dendritic cells is enhanced by DC-SIGN. J Virol. 2002;76:11827-11836.
23. Patel PG, Yu Kimata MT, Biggins JE, et al. Highly pathogenic simian immunodeficiency virus mne variants that emerge during the course of infection evolve enhanced infectivity and the ability to downregulate CD4 but not class I major histocompatibility complex antigens. J Virol. 2002;76:6425-6434.
24. Shugars DC, Smith MS, Glueck DH, et al. Analysis of human immunodeficiency virus type 1 nef gene sequences present in vivo. J Virol. 1993;67:4639-4650.
25. Long EM, Rainwater SM, Lavreys L, et al. HIV type 1 variants transmitted to women in Kenya require the CCR5 coreceptor for entry, regardless of the genetic complexity of the infecting virus. AIDS Res Hum Retroviruses. 2002;18:567-576.
26. Kimata JT, Wilson JM, Patel PG. The increased replicative capacity of a late-stage simian immunodeficiency virus mne variant is evident in macrophage- or dendritic cell-T-cell cocultures. Virology. 2004;327:307-317.
27. Pope M, Betjes MG, Romani N, et al. Conjugates of dendritic cells and memory T lymphocytes from skin facilitate productive infection with HIV-1. Cell. 1994;78:389-398.
28. Weissman D, Barker TD, Fauci AS. The efficiency of acute infection of CD4+ T cells is markedly enhanced in the setting of antigen-specific immune activation. J Exp Med. 1996;183:687-692.
29. Kimata JT, Kuller L, Anderson DB, et al. Emerging cytopathic and antigenic simian immunodeficiency virus variants influence AIDS progression. Nat Med. 1999;5:535-541.
30. Biesinger T, Yu Kimata MT, Kimata JT. Changes in simian immunodeficiency virus reverse transcriptase alleles that appear during infection of macaques enhance infectivity and replication in CD4+ T cells. Virology. 2008;370:184-193.
31. Bates M, Wrin T, Huang W, et al. Practical applications of viral fitness in clinical practice. Curr Opin Infect Dis. 2003;16:11-18.
32. Pohlmann S, Baribaud F, Lee B, et al. DC-SIGN interactions with human immunodeficiency virus type 1 and 2 and simian immunodeficiency virus. J Virol. 2001;75:4664-4672.
33. Arrighi JF, Pion M, Garcia E, et al. DC-SIGN-mediated infectious synapse formation enhances X4 HIV-1 transmission from dendritic cells to T cells. J Exp Med. 2004;200:1279-1288.
34. Wang JH, Janas AM, Olson WJ, et al. CD4 coexpression regulates DC-SIGN-mediated transmission of human immunodeficiency virus type 1. J Virol. 2007;81:2497-2507.
35. Blaak H, Brouwer M, Ran LJ, et al. In vitro replication kinetics of human immunodeficiency virus type 1 (HIV-1) variants in relation to virus load in long-term survivors of HIV-1 infection. J Infect Dis. 1998;177:600-610.
36. Mariani R, Kirchhoff F, Greenough TC, et al. High frequency of defective nef alleles in a long-term survivor with nonprogressive human immunodeficiency virus type 1 infection. J Virol. 1996;70:7752-7764.
37. Premkumar DR, Ma XZ, Maitra RK, et al. The nef gene from a long-term HIV type 1 nonprogressor. AIDS Res Hum Retroviruses. 1996;12:337-345.
38. Fackler OT, Moris A, Tibroni N, et al. Functional characterization of HIV-1 Nef mutants in the context of viral infection. Virology. 2006;351:322-339.
39. Laguette N, Benichou S, Basmaciogullari S. Human immunodeficiency virus type 1 Nef incorporation into virions does not increase infectivity. J Virol. 2009;83:1093-1104.
40. Qi M, Aiken C. Selective restriction of Nef-defective human immunodeficiency virus type 1 by a proteasome-dependent mechanism. J Virol. 2007;81:1534-1536.
41. Julg B, Goebel FD. Susceptibility to HIV/AIDS: an individual characteristic we can measure? Infection. 2005;33:160-162.
42. Shao W, Tang J, Song W, et al. CCL3L1 and CCL4L1: variable gene copy number in adolescents with and without human immunodeficiency virus type 1 (HIV-1) infection. Genes Immun. 2007;8:224-231.
© 2010 Lippincott Williams & Wilkins, Inc.