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Stable engraftment of human lymphoid tissue in NOD/scid-IL-2Rγcnull mice after CD34+ hematopoietic stem cell reconstitution permits the evaluation of ongoing HIV-1 infection for weeks to months. We demonstrate that HIV-1 infected rodents developed virus-specific cellular immune responses. CD8+ cell depletion, 2 or 5-7 weeks after viral infection, resulted in significant increase of HIV-1 load, robust immune cell activation and cytopathology in lymphoid tissues but preserved CD4/CD8 double-positive thymic T cell pools. Human CD8+ cells reappeared in circulation as early as 2-3 weeks. These data support the role of CD8+ cells in viral surveillance and the relevance of this humanized mouse model for the studies of HIV-1 pathobiology and virus-specific immunity.
In the past two decades, our laboratories have developed and characterized small animal models for the studies of HIV-1 infection and human disease (1-4). Recently, NOD/scid-γcnull (NOD/Shi-scid, NOG, or NOD/LtSz-scid, NSG) mice transplanted with human CD34+ hematopoietic stem cells (HSC) alone or in combination with fetal liver/thymus implant (BTL mice) have become promising models to study HIV-1 infection due to graft longevity and the establishment of chronic viral infection (5-8). HSC, when transplanted into immunodeficient mice, on either NSG/NOG or Balb/c-Rag2-/-γc-/- (BRG) backgrounds, developed a functional human immune system. These chimeric mice are susceptible to HIV-1 infection and demonstrate natural human disease progression (4, 9-16). However, neither BRG nor NSG/NOG humanized mice transplanted with HSC alone have shown temporal control of viral replication, robust humoral and cellular virus-specific adaptive responses [as was found in BLT animals (8)], or establishment of a stable virologic set point, as might be attributed to cytotoxic T lymphocyte (CTL)-mediated control of viral replication.
Control of HIV-1 replication is dependent on human and viral genetics, innate and adaptive (humoral and cellular) immune responses [reviewed in (17, 18)]. Levels of HIV-1 replication in an infected human host markedly decline after an initial viremia and establish a stable set-point. The temporal relationship between the decrease in viral load and the appearance of HIV-specific CD8+ CTL responses, suggests that the latter may regulate virus levels (19). CD8+ CTL control in treatment naïve patients was determined by limited dilution functional cytotoxic assay (20) or tetramer staining (21) combined with intracellular cytokine profiles of CD8+ cells (22, 23). The administration of CD8-specific antibodies to macaques that had been infected with simian immunodeficiency virus (SIV) or SIV/HIV(SHIV) has been shown to abrogate the decline in viremia from its peak level, result in increased peripheral viral load, accelerate CD4+ cell destruction and disease progression (24-35). The most efficient depletion of CD8+ cells in monkeys (lasting up to 6 weeks, with near total depletion of CD8+ cells from blood and lymph nodes) was achieved by using cM-T807 chimeric antibodies in which the heavy and light chain variable region genes were isolated from the murine M-T807 hybridoma and ligated to the human γ1 heavy chain and κ light chain genes, respectively. Complement-independent mechanisms have been shown to be primarily responsible for cM-T807-induced CD8+ lymphocyte depletion in vivo, although long-term use of these antibodies resulted in the development of humoral immune responses in macaques, due to xenoreactivity (27).
We now posit that further manipulations of the human immune system can be achieved in the small animal model (NSG/hCD34) of HIV-1 infection, affecting the course of disease. Herein, we demonstrate that NSG/hCD34 mice mount an HIV-specific cellular immune response following virus infection. This was shown by detecting IFN-γ and IL-2 cytokine production in response to HIV-1-derived peptide pools by human CD8 and CD4 T cells collected at five weeks after infection. CD8+ cell depletion strategies in virus-infected chimeric mice were then applied. Acceleration of HIV-1 replication was observed when CD8+ cell depletion was done two weeks after viral infection. The viral load was also increased, but at a lesser extent, when depletion was conducted at 5-7 weeks after viral infection. Following the CD8+ cell removal, preservation of T cell development in the thymus with the presence of CD4/CD8 double-positive cells was observed, and re-appearance of human CD8+ cell in circulation was seen as early as 2 to 3 weeks after depletion. Our findings underscore the importance of CD8+ T cell-mediated control of HIV-1 infection, are reflective of viral and CD4+ T cell dynamics seen previously for SIV-infected monkeys, and support the importance of this rodent model for the studies of HIV-1 immunobiology.
NOD/scid-IL-2Rγcnull mice were obtained from the Jackson Laboratories (Bar Harbor, ME) and bred under specific-pathogen-free conditions in accordance with ethical guidelines for care of laboratory animals at the University of Nebraska Medical Center (UNMC), as set forth by the National Institutes of Health.
Human cord blood was obtained, with parental written informed consent, from healthy full-term newborns (Department of Gynecology and Obstetrics, UNMC). After density gradient centrifugation, CD34+ cells were enriched using immunomagnetic beads according to the manufacturer's instructions (CD34+ selection kit; Miltenyi Biotec Inc., Auburn, CA). Purity of CD34+ cells isolated was evaluated by flow cytometry and was >90%. Cells were either frozen or immediately transplanted into newborn mice at 105 cells/mouse intrahepatically (i.h.) in 20 μl phosphate-buffered saline (PBS) using a 30-gauge needle. Before cell transplantation, newborn pups were irradiated at 1 Gy using a C9 cobalt 60 source (Picker Corporation, Cleveland, OH). Cells were transplanted between 4-12 hr after irradiation. Ranging from two to seven, littermates were reconstituted with one cord blood sample derived from one donor. The number of animals reconstituted was dependent on the number of CD34+ cells isolated from cord blood. Mice were weaned at 3 weeks of age and randomly distributed between different experimental groups.
The CCR5 coreceptor-utilizing HIV-1ADA strain was propagated in human monocyte-derived macrophages (MDM) (36). Viral preparations were screened and found to be negative for endotoxin (<10 pg/ml) (Associates of Cape Cod, Woods Hole, MA) and Mycoplasma (Gen-Probe II; Gen-Probe, San Diego, CA). The viral titers were assayed on MDM and determined to be 105 tissue culture infectious dose50 (TCID50)/ml.
HIV-1ADA was injected intraperitoneally (i.p.) at 104 TCID50. The levels of viral RNA copies/ml in plasma were analyzed using automated COBAS Amplicor System (Roche Molecular Diagnostics, Basel, Switzerland); detection limit of 50 viral RNA copies/ml. Mouse plasma samples (20 μl each) were diluted up to the volume 700 μl with normal human serum for assay use, which made the detection limit to 1750 copies/ml. HIV-1 infection was confirmed by virologic and histologic examinations. Reconstituted not infected animals of similar age served as controls.
The chimeric cM-T807 mAb were obtained from the National Institutes of Health National Center for Research Resources (Dr. Keith Reinmann). Mice were administered 10 mg of mAb/kg of antibody subcutaneously (s.c.) and 5 mg/kg i.p. in three day interval. The doses of cM-T807 were selected to create a sufficient and sustained concentration of antibody for elimination of CD8+ cells due to the C5a component of complement deficiency and to prevent rapid clearance of chimeric antibodies in immune deficient mice (37). Two different time points were used for depletion: 2 and 5-7 weeks after infection. Information of all the animals used in the study (age, period of infection, time after depletion, levels of IgM/IgG) is shown on Table 1 and in the Supplemental Table.
Peripheral blood samples were collected into EDTA-coated tubes from the facial vein by using lancets (MEDIpoint, Inc., Mineola, NY) or by cardiocentesis at the end of observation. Four-seven color flow cytometric analyses of whole blood samples were performed to monitor changes in T cell populations. In brief, 100 μl aliquots of whole blood were incubated with respective antibodies for 30 min at 4°C, and red blood cells were first lysed with FACS Lysing Solution® (Becton Dickinson, San Jose, CA) and then washed twice with PBS containing 2% fetal bovine serum. Spleen was divided into two halves; one was used for flow cytometry and the other for immunohistochemistry. Blood leukocytes and splenocytes were tested for human pan-CD45, CD3, CD4, CD8, CD11c, CD14, CD19 and HLA-DR markers as four or seven-color combinations. Antibodies and isotype controls were obtained from BD PharMingen (San Diego, CA) and staining was analyzed with a FACS DIVA (BD Immunocytometry Systems, Mountain View, CA). To reproducibly resolve CD8+ cells in the presence of cM-T807, anti-CD8-PE clone DK25 (Dako, Carpinteria, CA) was used. Depletion was confirmed by showing an absence of CD3+CD8+ and CD3-CD8+ lymphocytes. Results are expressed as percentages of total number of gated lymphocytes. The gating strategy was human CD45→CD3→CD4/CD8, CD45→CD19, and CD45→CD14.
Splenocytes were collected and enriched for human CD45 positive cells using magnetic bead separation (Miltenyi Biotec, Bergisch Gladbach, Germany). Enriched cells were cultured in RPMI-1640 supplemented with 10% fetal calf serum and penicillin/streptomycin, at 2 million cells per ml, in the presence of either HIV-1 gag or envelope peptide pools (AIDS Research and Reference Reagent Program, Division of AIDS, NIH). Peptide pools were used at 1μg/ml each. Cells were incubated at 37°C for 18hr with Brefaldin A (100ng/ml) added during last 6 hr. Cells incubated in the presence of peptide diluents served as non-stimulated control. At the end of incubation, cells were collected and stained for surface antigens- human CD45, 3, 4, 8- followed by intracellular staining with antibodies to human IFN-γ and IL-2. All antibodies and reagents for intracellular staining were obtained from eBioscience, Inc. (San Diego, CA). Stained cells were analyzed using BD LSR II and gated for human CD45→CD3→CD4 or CD8→ IFN-γ or IL-2.
Mouse plasma samples were diluted five times. The concentration of human cytokines: IFNγ, IL-2, IL-4, IL-6, IL-10, and TNFα were determined by FACSArray according to manufacturer's instruction (BD Bioscience).
Spleen and lymph node samples were fixed with 4% paraformaldehyde overnight and embedded in paraffin. Five-micron-thick sections were stained with mouse HLA-DR (clone CR3/43, 1:100), CD8 (clone 144, 1:50), CD20 (clone L26, 1:50), CD68 (clone KP-1, 1:50), or HIV-1 p24 (clone Kal-1, 1:10); all obtained from Dako (Carpinteria, CA). Mouse monoclonal antibodies to Ki-67 (clone DVB-2, 1:50), CD14 (clone 7, 1:50) and CD4 (clone 1F6, 1:40) were purchased from Biocare Medical, LLC (Concord, CA) and Novocastra (Norwell, MA), respectively. The polymer-based HRP-conjugated anti-mouse Dako EnVision systems are used as secondary detection reagents, and 3,3'-diaminobenzidine (DAB, Dako) was used as the chromogen. All paraffin-embedded sections were counterstained with Mayer's hematoxylin. Deletion of primary Ab or mouse IgG served as controls. Images were obtained by Optronics digital camera fixed to Nikon Eclipse E800 (Nikon Instruments, Melville, NY) using MagnaFire 2.0 software.
The plasma levels of IgM and IgG were determined by ELISA (Bethyl Laboratories, Montgomery, TX).
Plasma collected at different time points were analyzed for HIV-1 specific human immunoglobulins by using concanavalin A (Con A) based ELISA, as previously described (38). HIV-1ADA envelope protein was expressed in a baculoviral system from a plasmid obtained from Dr. Lee Ratner (39). Purified envelope protein was used to coat the ELISA plate at 0.5mg/well. Peroxidase conjugated secondary antibodies to either human IgM or IgG were used (Bethyl Laboratories) and finally developed with 3,3′,5,5′-tetramethylbenzidine (TMB) substrate. Plasma samples from uninfected humanized mice were used as negative controls, and HIV-1 seropositive human serum was used at 1:10,000 dilutions as a positive control.
Data were analyzed using GraphPad Prizm and Excel softwares; statistical tests employed were non-parametric Mann-Whitney U-test, Wilcoxon test and one-way ANOVA for comparisons of multiple groups. A P value of <0.05 was considered statistically significant. For statistical analyses, viral load values of <1750 and >35000000 were set to 1750 and 35000000 (which represent the lower and upper thresholds for the assay). Since a Wilcoxon test was used to compare the distributions, setting these interval values to their lower and upper limits respectively had no effect on the results.
Newborn NOD/scid-IL-2Rγcnull mice (NSG) were reconstituted with human (hCD34+) HSC isolated from umbilical cord blood. Phenotypic development of human lymphoid tissue was evaluated by flow cytometry analyses of human cells in peripheral blood (CD45, CD3, CD4, CD8, CD19 and CD14) to determine the relative abundance of immune cell groups. Presence of human antibodies in peripheral blood was analyzed by anti-human IgG/IgM ELISA (Table 1, Supplemental Material). At the age of ~5 months, chimeric mice showed partial maturation of a reconstituted human immune system. The number of human cells in circulation became stable at 5-7 months of age, with the majority being CD3+ cells (Fig. 1). At this stage humanized mice were infected with HIV-1ADA (a macrophage-tropic CCR5 utilizing virus) by administering 104 TCID50 i.p. Progression of viral infection was determined by measuring plasma viral load at different time points. Virus was rarely detected in the serum as early as 2 weeks post-infection, but by 5-6 weeks all tested animals were infected. Figure 1 demonstrates the percentages of various phenotypes of human immune cells in spleens from uninfected and HIV-1 infected animals, as analyzed by flow cytometry. The total number of human cells (CD45+) in the spleen was unaffected by virus infection. However, in HIV-1 infected animals, CD4+ T cell numbers declined and a parallel reduction of CD4:CD8 ratio was noted when compared to uninfected animals (n = 10 per group; P =0.032 and P =0.026, respectively). Peripheral viral load, number of CD4+ and CD8+ human cells in circulation were analyzed in three animals bi-weekly (Fig. 2) and for the rest of animals at weeks 4 – 8 or at time of sacrifice (weeks 8 – 13).
In prospectively studied animals, the peak of infection occurred at 5-7 weeks following viral inoculation, when it reached 5.5 (#149) and 6.5 log10 (#159) copies of viral RNA, and the viral load declined by 1.5 (#149) and 0.5 (#159) log10 at 9-10 weeks. With the selected infectious dose of 104 TCID50, which provides successful infection of ~50% of animals, the peak of viral load was observed at 5-6 weeks post-infection and median reached 5.31 log10 with 4.72 – 6.00 range (n = 6). By 8-10 weeks post-infection, viral load declined to 4.80 log10 with a range of 4.12 – 5.39 (P = 0.023 by Mann-Whitney U test, n = 6).
To confirm the involvement of CD8+ cells in control of HIV-1 replication in this model and to characterize the cellular immune responses, we infected and sacrificed 5 additional humanized animals at 5 weeks post-infection. To detect cellular immune responses, splenocytes were enriched for human CD45+ cells by magnetic bead positive selection for in vitro stimulation with HIV-1 antigens. Three uninfected animals similar in the levels of reconstitution served as controls. To evaluate the presence of cellular responses and their functional properties, we used intracellular cytokine staining (ICS). The number of IFN-γ, IL-2 or dual cytokine producing cells was analyzed following stimulation with HIV-1 gag and env peptide pools. Representative FACS plots for three infected and one uninfected mice are shown in Fig 3. Four of five infected animals efficiently responded to HIV gag peptide pool stimulation compared to HIV env peptide pool and had detectable levels of CD8+IFN-γ+ cells, and 30 – 87% of these cells were double positive for IFNγ+ and IL-2+. All 5 mice showed a virus-specific CD4+ T cell response, as evidenced by CD4+IL-2+ cells; a subset of these CD4+ cells was double positive for both IFN-γ and IL-2. These data confirm that HIV-1 infection elicited antigen-specific human cellular immune responses.
To understand what would happen if we depleted the CTL control of viral replication in this humanized mouse model of HIV-1 infection and to provide a proof-of-concept that this animal model can be used for such immune manipulations, we used CD8 depletion strategies. CD8+ cell depletion was achieved by using two sequential injections of cM-T807 antibodies (s.c. and i.p. within 3 days). Two schemes of depletion were used: 1) 5-7 weeks post-infection (w.p.i., as a model of established HIV-1 infection, and 2) 2 weeks post-infection to prevent the development of CD8+ cell-mediated anti-viral responses and to accelerate the development of neuropathology as discussed in detail in our companion paper (Gorantla et al., submitted).
Dynamics of CD4+ and CD8+ cells and viral load in blood from two representative uninfected and HIV-infected/CD8 cell depleted animals engrafted with the same donor cells are shown in Fig. 4A. CD8 cell depletion at 5 w.p.i increased viral load in circulation and reduced temporarily CD4+ cell number. In spleens of these animals, the percentage of CD4+ cells was 8.7 ± 3.9% and was not different from the number of CD4+ cells in the spleens of HIV-1-infected animals analyzed at 8-10 weeks post-infection (11.2 ± 3.2%, n = 5). CD8+ cell-depleted/uninfected animals of similar age and similar total number of human CD45+ cells in spleen (25.2 ± 2.6%, n = 4) analyzed at 1-3 weeks post-depletion did not experience loss of CD3+CD4+ cells, but a number of CD8+ cells were significantly reduced to 0.38 ± 2.6% (range 0 – 1.4%). CD8+ cells re-appeared in circulation by 3 weeks post-depletion (Fig. 4B) with appearance of a significant number of CD4+CD8+.
Changes in CD4+ and CD8+ cell numbers and the viral load in peripheral blood of four HIV-infected/CD8-depleted (at 5-7 w.p.i and 2 weeks post-CD8 depletion) and five HIV-1 infected/non-depleted control mice analyzed at the first peak of viremia 5-6 weeks and 2 weeks later were compared (Table 2). When CD8+ cells were depleted in animals with established infection, an increase in viral load was detected in all mice with Δlog10 ranging from 0.57-2.3 (P = 0.008, Mann-Whitney U-test). In non-depleted mice, declines could be associated with considerable decrease in CD4+ T cell count in blood. However, in depleted animals, even with lowered CD4+ T cell numbers, a substantial increase in viral load was observed.
In contrast to the small changes in viral load that were caused by depleting CD8+ cells in mice with established HIV-infection (5-7 w.p.i), depletion of CD8+ cells at 2 weeks after infection had a more profound effect on viral load (Fig 5B). Only one out of five mice showed significant levels of virus at 2 weeks post-infection (m341 had 6.50 Log10 viral copies/ml andwas excluded from statistical analysis). Following CD8 depletion, all animals exhibited substantial levels of virus in circulation (geometrical mean 5.13 vs 3.70 for all non-depleted animals, P = 0.006). The summary of the peripheral viral load values in non-depleted, versus CD8+ cell depleted animals is shown in Fig.5.
In order to investigate the other possible mechanisms in addition to CTL loss for the increase of viral replication, we evaluated the levels of human cytokines (IL-2, IL-4, IFN-γ, TNF-α, IL-6 and IL-10) in plasma after i.p. injection of antibodies by CBA. Increased viral replication in HIV-1 infected/CD8+ cell depleted animals could also be associated with the homeostatic proliferation of CD4+ cells and increased production of cytokines in the lymphoid tissue, which are capable of stimulating HIV-1 replication. We collected plasma from 7 animals at 3 days before the first s.c. injection and 3 days after the second i.p. injection of anti-CD8 antibodies. Only human IL-2 levels were increased from 67.2 ± 25.9 pg/ml to 158.2 ± 28.1 pg/ml (P = 0.019, with exception of one mouse with the drop of IL-2 concentration below the level of detection limit 10 pg/ml). Three of these seven animals were infected. We noted that in two infected animals human cells produced detectable levels of IL-4 (514 and 309 pg/ml) after CD8+ cell depletion.
Effect of CD8 depleting antibodies on lymph node pathologies and double-positive CD4/CD8 thymocyte differentiation in thymus was studied. Two weeks after depletion the formation of syncytia in lymph nodes was noted (Fig. 6, HIV-1p24 staining). CD8+ cell depletion resulted in significant increase of HIV-1 p24+ cells, changes in B cell distribution (CD79α staining), and increased activation (HLA-DR) in the lymph nodes. To test the ability of chimeric mice to restore the CD8+ cell pool after depletion, we analyzed thymic tissues by immunohistochemistry and flow cytometry. Thymic tissues of uninfected/non-depleted animals were populated by human CD45RO+CD4+CD8+ thymocytes (Fig. 7A). Cortical areas had single positive human CD4+ cells and fewer CD8+ cells. Murine epithelial claudin-3-positive cells and a small number of human HLA-DR+ cells with dendritic cell morphology were also seen in thymi (not shown). Depletion of CD8+ cells significantly reduced the number of CD4/CD8 double-positive cells and increased the density of Ki-67+ human cells. Figure 7A shows the immunohistology of thymi obtained from animal infected for 8 weeks with CD8+ cell depletion for one week before euthanasia. The reduction of CD4/CD8 double-positive cell numbers was also demonstrated by flow cytometry performed at two weeks after depletion (Fig. 7B). These data confirm that CD8+ cell depletion did not abolish the generation of CD4/CD8 double positive progenitor cells in thymus.
We previously demonstrated that Balb/c-Rag2-/-γc-/- mice could be repopulated with human cells within the lymphoid tissue structures following human CD34+ cell transplantation. These mice developed a functional human immune system that was susceptible to HIV-1 infection (4). In the current report, NSG mice that are more readily engrafted with human HSC were used. Animals with an established human immune system were infected with a macrophage-tropic HIV-1 virus, resulting in a chronic virus infection and CD4+ T cell depletion. Viremia reached significant levels by 5-6 weeks after infection. We observed the presence of HIV Gag and Env specific, IFN-γ– and IL-2-producing human CD8+ and CD4+ cells in the spleens of these mice, using intracellular cytokine staining. Most of the HIV-specific CD8+ cells reacted with Gag, and the number of IFN-γ/IL2 double producers exceeded the number of single IFN-γ-producing cells. Most of the virus-specific CD4+ cells produced only IL-2 although some CD4+ cells that produced both IFN-γ and IL-2 were also detected. These results support the notion that humanized NSG mice can generate polyfunctional cellular immune responses to HIV-1, as was shown for human subjects (40-42). We did not find strong humoral immune responses in evaluated animals. In 10 evaluated animals 8 had detectable levels of HIV-1-specific IgM that declined by 10-11 weeks after infection without the development of IgG responses (data not shown).
The best evidence for the role of CD8+ CTLs in the control of immunodeficiency virus infection was obtained in SIV-infected rhesus macaques that were subjected to the immunodepletion of CD8+ cells. In the current model, depletion of CD8+ cells 2 weeks after infection led to the acceleration in viral replication during the acute phase of infection. CD8+ cell depletion at 5-7 weeks post-infection also enhanced the viral replication, and prevented small decline observed in non-depleted mice. CD8+ cell depletion was associated with a significant reduction in the number of human CD4/CD8 double-positive cells in thymus. However, the pool of CD8+ T cells started to return at 2-3 weeks post-depletion. Overall, observations made with CD8+ T cell depletion using cM-T807 antibody in our humanized mice partially were reminiscent of previously published data from non-human primate SIV/SHIV model systems. The durable levels of depletion was achieved and a strong correlation between disease outcome and viral-specific CTL restoration was observed by use of the mouse-human chimeric monoclonal antibody (cM-T807) during early stages SIV infection in rhesus macaques (26). In humanized NSG mice, earlier depletion of CD8+ cells accelerated viral replication. A significant reduction in CD4+ cell number, observed in our study in blood was also reported in SIV model (33). Our data are also concordant with the recently published observations that CD8+ lymphocyte depletion can induce a transient CD4+ T cell increase, and thus providing increased number of targets for SIV in animals during acute SIV infection without influence on the disease outcomes (35).
The generation of HIV-1-specific human CTL in “humanized” mice is influenced by issues of T cell maturation, receptor repertoire, interactions with murine stromal cells and more. Nonetheless, virus-specific human CD8+ T cells have been observed in previous studies using immune deficient mice reconstituted with human peripheral blood lymphocytes (3, 43-49). A more sophisticated bone marrow-liver-thymus (BLT) mouse model, where NOD/scid or NSG mice were transplanted with fetal thymus/liver tissue combined with intravenous transplantation of CD34+ stem cells (8, 50-52) also showed the generation of antigen-specific human immune responses. Our data show that in NSG mice reconstituted at birth with HSC the thymus is highly populated with human CD45RO+ thymocytes, and we also observed that the T cell repertoire, analyzed by staining for variable ß chain of T cell receptor, is preserved (data not shown). The presence of double-positive T cells in peripheral blood of HIV-1-infected/CD8+ cell depleted mouse and reduction of viral load suggests that these cells may play a role in the restoration of the lymphocyte pool following CD8+ cell depletion and contribute to control the viral replication (53). We cannot exclude that this is an example of immunopathology induced by combination of significant levels of HIV-1 replication and CD8 depletion. This observation needs further investigation.
We have direct evidence that our chimeric mice developed cellular immune response against HIV-1. However, we cannot over-look the possibility that this increase in viral replication following CD8+ T cell depletion may be due to, at least in part, to increased production of IL-2/IL-4 and homeostatic expansion of CD4+ T cells (54). Nonetheless, our results clearly show that the NSG/hCD34 mouse model can be effectively employed to study the progression and pathogenesis of HIV-1 infection in a live animal host and to dissect out the role of specific immune subsets in the control of infection. This is expected to enable future investigations of viral diversity, developmental therapeutics and antiretroviral immunity.
We would like to thank Dr. John G. Sharp (UNMC) for active discussions and critical insights into the immune reconstitution strategies; Dr. Keith A. Reimann for helpful suggestions regarding anti-CD8 antibodies application, Dr. Lee Ratner (Washington University, St. Louis, MO) for providing plasmids for the HIV-1ADA envelope gp120 protein, Mrs. Victoria Smith and Dr. Charles Kuszynski for FACS analyses (UNMC), Ms. Valerie K. Shostrom, Statistical Coordinator, College of Public Health Department of Biostatistics (UNMC), and Dr. Dawn Eggert for animal breeding (UNMC).
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1This work was supported by NIH grant R21NS060642-01 (to L.P.), and P20 RR15635 and 1 P01 NS043985-01 (to L.P. and H.E.G.) and 2R37 NS36126 and 5 P01 MH64570-03 (to H. E. G.).