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Journal of Virology, April 2005, p. 4965-4976, Vol. 79, No. 8
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.8.4965-4976.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Preferential Feline Immunodeficiency Virus (FIV) Infection of CD4+ CD25+ T-Regulatory Cells Correlates both with Surface Expression of CXCR4 and Activation of FIV Long Terminal Repeat Binding Cellular Transcriptional Factors

Anjali Joshi, Himanshu Garg,{dagger} Mary B. Tompkins, and Wayne A. Tompkins*

Immunology Program, College of Veterinary Medicine, North Carolina State University, Raleigh, North Carolina

Received 11 August 2004/ Accepted 19 November 2004


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ABSTRACT
 
Previously, we have characterized feline CD4+ CD25+ T-regulatory (Treg) cells with regard to their immune regulatory properties and ability to support feline immunodeficiency virus (FIV) replication in vitro and in vivo. Our studies showed that while CD4+ CD25+ cells were capable of replicating FIV in the presence of interleukin-2 (IL-2) alone, CD4+ CD25– cells harbored a latent infection that required a strong mitogenic stimulus to activate virus replication. In the present study, we investigated the mechanisms governing the preferential replication of FIV in highly purified CD4+ CD25+ Treg cells compared to their CD4+ CD25– counterparts. Studies aimed at elucidating mechanisms regulating infection of these cells revealed that CD4+ CD25– cells were less susceptible to FIV binding and entry than CD4+ CD25+ cells, which correlated with increased surface expression of FIV coreceptor CXCR4. In addition, the number of CD4+ CD25+ cells that expressed the primary receptor CD134 was greater than for CD4+ CD25– cells. Although increased permissiveness to FIV infection of CD4+ CD25– cells following mitogenic stimulation correlated strongly with upregulation of surface CXCR4, it did not correlate with CD134 expression. Further, study of intracellular factors regulating FIV replication revealed that CD4+ CD25+ but not CD4+ CD25– T cells showed constitutive and IL-2-responsive transactivation of activating transcription factor, CAAT enhancer binding protein, and activating protein 1 transcription factors that are important for FIV replication. These factors were upregulated in CD4+ CD25– T cells following ConA stimulation, which correlated with FIV replication. This is the first report elucidating the mechanisms that allow for productive lentiviral infection of CD4+ CD25+ Treg cells.


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INTRODUCTION
 
Feline immunodeficiency virus (FIV), a lentivirus belonging to the family Retroviridae, is remarkably similar to the human lentivirus human immunodeficiency virus type 1 (HIV-1) both in terms of genomic organization and disease pathogenesis (4). These similarities in the pathogenesis of the two viruses make FIV an excellent small animal model for HIV-1-induced immunodeficiency in humans. Despite the marked similarities between HIV and FIV, the two viruses also have apparent differences. While HIV-1 utilizes CD4 as the primary receptor and shows a major tropism for cells of the CD4 lineage (28), FIV does not utilize the CD4 molecule as a receptor and its tropism is more generalized for cells of the CD4, CD8, as well as B lineage (12, 17). In this regard, the primary receptor for FIV has only recently been identified as CD134, a CD4+ T-cell activation marker belonging to the tumor necrosis factor receptor superfamily (40), while the utilization of CXCR4 as the coreceptor for FIV entry is well established (18, 53).

Studies have established that in HIV-infected individuals, the virus persists mainly as a latent reservoir in memory CD4+ T (CD45RO+) cells, which possess the inherent ability to survive for a prolonged period of time (10, 37). While HIV infection has also been demonstrated in naive T cells in vivo (33), establishment of a stable HIV infection in resting T cells in vitro remains controversial. Some studies suggest that HIV can efficiently bind and enter resting T cells (42, 46), whereas others have shown that naive T cells are resistant to de novo HIV infection (9, 19). In this regard, the data suggest that susceptibility of different lymphocyte subsets to HIV infection is regulated, to some extent, by differential expression of viral coreceptors. Pertaining to HIV binding and entry, studies have shown that the HIV coreceptors CXCR4 (X4) and CCR5 (R5) are differentially expressed on activated versus naive or resting T cells. While CXCR4 is expressed on both naive and memory T cells, CCR5 is a marker of activated and replicating T cells that are infectible with R5-tropic viruses (16, 38). In support of this, studies have demonstrated that while resting CD4+ T cells can be infected by X4-tropic viruses, they are resistant to infection by R5-tropic viruses (38).

In addition to cell surface receptor expression, the activation-proliferative state of T cells also determines the extent of virus replication (43). Further, it has been demonstrated that HIV and FIV replicate efficiently in CD4+ CD25+ T cells that are partially activated but not in naive CD4+ CD25– resting T cells (7, 9, 24, 43). To date there have been limited reports regarding FIV replication in activated versus resting T cells. We have recently reported that FIV establishes a productive infection in feline CD4+ CD25+ T cells but not in naive CD4+ CD25– T cells (24). We subsequently phenotypically and functionally characterized these CD4+ CD25+ T cells as T-regulatory (Treg) cells (51). These cells have a partial activation phenotype, are anergic, and demonstrate immunosuppressive function in the presence of interleukin-2 (IL-2) (51). While little is known of the factors regulating anergy in Treg cells, the activation-proliferative state of lymphocytes is largely controlled by a group of cellular transcription factors that regulate IL-2 gene expression, such as NF-{kappa}B and AP-1 (1), which also regulate transcription of HIV and FIV genes (5, 11).

HIV and FIV depend on both cellular and viral factors for efficient transcription of their genomes, and the activity of their promoters within the long terminal repeat (LTR) regions are highly dependent on the level of host cell activation (5, 45). HIV and FIV LTRs encompass binding sites for several cellular transcription factors, including activating protein 1 (AP-1), activating transcription factor (AP-4), ATF CAAT enhancer binding protein (CEBP), NF-{kappa}B, and SP-1 (5, 11, 36). Although the role of NF-{kappa}B in HIV mRNA transcription is well established (11, 36), the role of a putative NF-{kappa}B binding site in FIV LTR transactivation remains unclear. However, several studies have established that the binding sites for AP-1, ATF, and CEBP in the FIV LTR are critical for FIV replication in infected cells. Kawaguchi et al. (25) demonstrated by site-specific mutations that the CEBP site in the FIV LTR is necessary for efficient viral replication in both feline fibroblasts and a T-lymphoblastoid cell line. A number of studies demonstrated that mutations or deletions of both ATF and AP-1 sites resulted in severely reduced basal promoter activity from the FIV LTR and impaired ability of viruses to replicate both in feline lymphocytes and in macrophages (5, 21, 23).

In the present study, we extended our previous observations to elucidate the mechanisms governing the differential replication of FIV in CD4+ CD25+ and CD4+ CD25– T cells. Our results demonstrate that CD4+ CD25– T cells exhibit restrictions to FIV infection both at the level of viral entry and transcriptional activation. While viral entry correlates more significantly with surface CXCR4 than CD134 expression, FIV transcriptional activation correlates significantly with AP-1, CEBP, and ATF activation but not that of NF-{kappa}B.


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MATERIALS AND METHODS
 
Cats. Specific-pathogen-free cats were obtained from Liberty Labs (Liberty Corners, N.J.) or Cedar River Laboratory (Mason City, Iowa) and housed at the Laboratory Animal Resource Facility at the College of Veterinary Medicine, North Carolina State University. At the time of the study, cats ranged in age between 6 to 7 years and were seronegative for FIV.

Blood and LN cell collection. Lymphocytes were obtained either from lymph nodes (LN) or whole blood. LN cells were obtained by peripheral LN biopsies as described previously (48). Peripheral blood mononuclear cells (PBMC) were isolated from whole blood by Percoll (Sigma, St. Louis, Mo.) density gradient centrifugation as described by Tompkins et al. (50). Preliminary studies using cells from both sources gave equivalent results.

Antibodies and flow cytometry analysis. PBMC or purified cell populations were stained for surface expression of various markers using phycoerythrin (PE)-, fluorescein isothiocyanate (FITC)-, or allophycocyanin-conjugated anti-CD4 (49) and FITC-conjugated anti-CD25 antibodies (31). Cell surface CXCR4 expression was determined using the cross-reacting human anti-CXCR4 antibody (44717) followed by PE-conjugated goat anti-mouse immunoglobulin G (IgG) (Jackson ImmunoResearch Laboratories, West Grove, Pa.). Surface CD134 expression was determined using cross-reacting (40) PE-conjugated BerACT35 monoclonal antibody (Ancell Corporation, Bayport, Minn.) by following the manufacturer's instructions. Samples were analyzed using a FACSCalibur flow cytometer (Becton Dickinson, Los Angeles, Calif.). Generally, 3 x 105 to 5 x 105 cells were used for staining and 15,000 events were acquired using Becton Dickinson Cell Quest software.

Purification of T cells. To investigate single lymphocyte subsets, CD4+ CD25+ and CD4+ CD25– cells were stained with anti-CD4 and anti-CD25 antibodies, as described above, and sorted into pure populations using a Cytomation MoFlo fluorescence-activated cell sorter (FACS). The purity of FACS-sorted cell populations was ≥97.0%. For some experiments, CD4+ subsets were enriched by negative selection using goat anti-mouse IgG-coated magnetic beads (Dynabeads M-450; Dynal, Oslo, Norway) as described previously (24, 51). Briefly, PBMC were depleted of B cells, CD8 cells, and monocytes by using anti-CD21, anti-CD8, and anti-CD14 monoclonal antibodies, respectively. The remaining cells were further treated with 5 mM leucine methyl ester for 1 h at 25°C (Sigma) to deplete natural killer cells. The CD25-expressing CD4+ cells were then enriched by positive selection using anti-CD25 monoclonal antibody-coated beads. The purity of enriched CD4+ CD25+ or CD4+ CD25– cells was verified by flow cytometric analysis and varied from 90 to 97% for the CD4+ cell population and >98.0% for CD25+ and CD25– cells (Fig. 1).



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  		FIG. 1. Representative dot plots indicating the purity of CD4+ CD25+ and CD4+ CD25– cells used in the study. LN-derived cells were sorted into CD4+ CD25+ and CD4+ CD25– populations by using antibody-coated magnetic beads as described previously. (A) Total LN cells, (B) purified CD4+ CD25+ cells, and (C) purified CD4+ CD25– cells were stained with PE-conjugated anti-CD4 and FITC-conjugated anti-CD25 antibody and analyzed by flow cytometry. Numbers represent the percentages of cells within each quadrant.

Cell culture and stimulation. PBMC, LN cells, or purified T-cell subsets were cultured at 106 cells/ml in growth medium (RPMI-1640 containing 10% heat-inactivated fetal bovine serum, 1% penicillin-streptomycin, 1% sodium bicarbonate, 1% sodium pyruvate, 1% L-glutamine, and 1 mM HEPES buffer) in the presence or absence of 2-µg/ml ConA or 100-U/ml recombinant human IL-2 (NIH AIDS Research and Reagent Program). Feline CD4E (FCD4E) cells were established through long-term culture of PBMC from a specific-pathogen-free cat in the presence of IL-2 and were cultured in RPMI-1640 medium. These cells are 100% positive for the feline pan-T-cell marker 1.572 and 60 to 65% positive for the feline CD4 homolog (17). HeLa cells were maintained in Dulbecco's modified Eagle's medium (Mediatech) supplemented with 10% fetal bovine serum, penicillin (100 U/ml), and streptomycin (100 µg/ml). Medium for HeLa cells selected to stably express feline CD134 (HeLa-CD134) was supplemented with G-418 (Gibco-BRL, Gaithersberg, Md.) at 700 µg/ml. Viability for different cell populations was determined by trypan blue dye exclusion assay and was routinely greater than 90%.

FIV-NCSU1 virus stock generation and in vitro infection. FIV-NCSU1 virus stock was prepared from PBMC derived from FIV-positive cats as described previously (24). No virus-associated DNA was present in the stock as analyzed by PCR using FIV-specific primers. In vitro infection assays were performed using a multiplicity of infection (MOI) of 0.1 or as indicated. Before infections, the virus stock was treated with DNase I (300 U/ml) for 30 min at room temperature. Cells were exposed to the virus for 1 to 2 h at 37°C. Following virus adsorption, the cells were washed three times with culture medium and plated at a concentration of 106 cells/ml of culture medium.

FIV p24 antigen capture ELISA. FIV gag-p24 antigen in culture supernatants of infected cells was detected using an antigen capture enzyme-linked immunosorbent assay (ELISA) as described previously (24).

PCR and RT-PCR. DNA was extracted from equal numbers of infected cells using a QIAamp DNA Blood Mini kit (QIAGEN, Valencia, Calif.) and quantified by measuring the A260 using a UV-spectrophotometer (Bio-Rad, Hercules, Calif.). Generally, 0.5 µg of DNA was used for PCRs unless otherwise indicated. For some experiments, DNA was harvested by direct cell lysis in PCR tubes to minimize loss during extraction. Briefly, 105 infected cells were pelleted in a PCR tube, lysed in 50 µl of lysis buffer (10 mM Tris-Cl [pH 8.3], 0.45% Nonidet P-40, 0.45% Tween 20, 50 µg of proteinase K/ml), and incubated for 3 h at 56°C followed by denaturation for 10 min at 100°C. Ten microliters of this lysate was used for PCR. FIV-specific products in infected cells were detected using primers LTR-sense and LTR-antisense, and a GAPDH (glyceraldehyde-3-phosphate dehydrogenase) primer pair was used as a control to ensure the use of equal amounts of DNA template in identical experiments as described previously (24). For reverse transcription (RT)-PCR studies, RNA was isolated using a RNeasy Micro kit (QIAGEN). During extraction the samples were treated with RNase-free DNase as per the manufacturer's protocol to get rid of contaminating DNA.

Real-time PCR and RT-PCR. RNA or DNA was isolated from equal numbers of infected cell populations as described above. Purified RNA (0.5 µg) was reverse transcribed and amplified using a TaqMan One Step RT-PCR kit (Applied Biosystems, Foster City, Calif.) in a total reaction volume of 50 µl with cycling conditions recommended by the manufacturer. In order to calculate viral copy numbers, a standard curve was generated using serial 10-fold dilutions of gag-RNA standard of known copy number. The gag-RNA was prepared by in vitro transcription (Ambion, Austin, Tex.) from a plasmid containing a portion of the NCSU1 gag gene under the control of the T7 promoter. The gag primers and probe derived from the FIV-NCSU1 sequence have been described previously (8) and are as follows: forward primer, 5'-GAT TAG GAG GTG AGG AAG TTC AGC T-3'; reverse primer, 5'-CTT TCA TCC AAT ATT TCT TTA TCT GCA-3'; probe, 5'-6FAM-CAT GGC CAC ATT AAT AAT GGC CGC A-TAMRA-3'. Samples and standards were amplified in 96-well plates (Axygen, Union City, Calif.) using the Bio-Rad iCyler. In parallel, RT-PCRs for the GAPDH gene were used as a housekeeping control. Quantitative real-time PCR for cell-associated proviral DNA was also performed with the above-mentioned primer pairs in a total reaction volume of 25 µl by using a TaqMan universal PCR kit (Applied Biosystems). To calculate proviral copy numbers, a standard curve was generated using serial 10-fold dilutions of the NCSU1 plasmid with known copy numbers. Equal amounts (0.5 µg) of DNA were used for each reaction and amplified in the iCycler (Bio-Rad) under the following conditions: 95°C for 10 min, 45 cycles of 95°C for 15 s, and a final extension of 60°C for 1 min. PCR for GAPDH DNA was used as a housekeeping control for normalizing the DNA quantity.

Viral binding and entry assays. FIV binding and entry assays were performed as described previously (27). Briefly, for viral binding, 106 purified CD4+ CD25+ and CD4+ CD25– cells were chilled on ice for 20 min followed by incubation with FIV-NCSU1 for 1 h on ice. Subsequently, the cells were washed four times with chilled phosphate-buffered saline (PBS) and lysed immediately to isolate total RNA. For entry assays, 106 cells were incubated with FIV at 37°C for 1 h and treated with trypsin-EDTA (0.25%) for 7 min at 37°C to remove cell surface-attached virions, followed by four washes with PBS. Cells were then lysed, total RNA isolated and used for real time RT-PCR. The efficiency of trypsin-EDTA treatment in removing virus attached to the cell surface was determined by incubating cells with FIV on ice for 1 h followed by trypsinization as indicated above and by subsequently performing RT-PCR using FIV-specific primers.

Blocking studies using the CXCR4 antagonist AMD-3100. CD4+ CD25+ and CD4+ CD25– cells (2 x 105 per treatment) were incubated for 30 min with serial 10-fold dilutions of AMD-3100 (a kind gift of Edward Hoover, Colorado State University), starting with a maximum concentration of 10 µg/ml. Cells were subsequently infected with FIV-NCSU1 in either the presence or absence of inhibitor. Following infection, cells were washed with PBS and cultured in RPMI medium supplemented with IL-2 for CD4+ CD25+ cells and with ConA for CD4+ CD25– cells. Culture medium for both the cell types was replenished with the respective above-mentioned concentrations of AMD-3100 to minimize second-round infections. Infected cells were cultured for 12 days and culture supernatants harvested every 3 days postinfection for FIV-p24 antigen capture ELISA.

Transfection of cells. The feline CD134 expression vectors were a kind gift form Brian J. Willet (University of Glasgow). HeLa cells were transiently transfected using the Effectene transfection reagent (QIAGEN) as previously described (18). Forty-eight hours posttransfection, cells were analyzed for surface CD134 expression by flow cytometry. Selection of HeLA cell clones stably expressing feline CD134 was done by single-cell cloning followed by culturing in media supplemented with G-418 at 700 µg/ml.

Electrophoretic mobility shift assay (EMSA). Nuclear extracts were prepared from 106 cells as per the protocol of Dignam and coworkers (15) either immediately after isolation or after culture for 5 days in the presence of IL-2 (100 U/ml) or ConA (2 µg/ml). Protein concentrations in the nuclear extracts were estimated using a colorimetric Coomassie blue protein assay reagent kit (Pierce Biotechnology, Rockford, Ill.) following the manufacturer's protocol. Oligonucleotides were designed using the predicted binding sites for AP-1, ATF, CEBP, and NF-{kappa}B in the FIV-Petaluma and FIV-PPR genomes and incorporating the changes in the FIV-NCSU1 genome. The sequences for the sense strand are as follows: AP-1, 5'-AGC ATG ACT CAT AGT TAA AG-3'; CEBP, 5'-TGC TTA ACC GCA AAA CCA CAT CCT A-3'; NF{kappa}B, 5'-GGA CTA GTG ACT GTT TAC GA-3'; and ATF, 5'-GCT GAT GAC GTA TAA TTT GC-3'. In addition, a consensus oligonucleotide for NF-{kappa}B having the sequence 5'-AGT TGA GGG GAC TTT CCC AGG C-3' was also used in the study. The double-stranded oligonucleotides were labeled at the 5' end with radiolabeled [{gamma}-32P]ATP (3,000 Ci/mmol at 10 mCi/ml; Amersham Biosciences) using T4 polynucleotide kinase (10 U/µl; Invitrogen). Nuclear extracts (2.5 µg for AP-1 and CEBP, 5 µg for ATF and consensus NF-{kappa}B, and 10 µg for LTR NF-{kappa}B) were incubated with 1 µl of radiolabeled oligonucleotides (0.2 ng, 5 x 104 to 8 x 104 cpm) in binding buffer (10 mM Tris-HCl [pH 7.5], 50 mM NaCl, 1 mM MgCl2, 0.5 mM dithiothreitol, 0.5 mM EDTA, 4% glycerol, 50 µg of poly(I):poly(C)/ml) for 15 min at room temperature. Specific competition was accomplished by using a 50-fold molar excess of unlabeled probe. In addition, nonspecific competitors that did not contain the AP-1, ATF, CEBP, or NF-{kappa}B binding sequence were used to demonstrate specificity and have been described elsewhere (21, 25, 30). Nuclear extracts for specific and nonspecific competition were derived from 1.5 x 106 T cells activated with 2-µg/ml ConA for 5 days. Following binding, the complexes were resolved in a 6% nondenaturing acrylamide-bisacrylamide (30:1) gel in a 22.5 mM Tris borate and 0.5 mM EDTA buffer. The gels were subsequently dried and autoradiographed.

Statistical analysis. The Student's t test was used to compare differences in p24 antigen for infected CD4+ CD25+ and CD4+ CD25– cells. Differences in viral copy numbers and cell surface markers for CD4+ CD25+ and CD4+ CD25– cells were determined using the Mann-Whitney's test (t test-like for nonparametric data). Differences were considered significant at a P of <0.05. Correlation analysis between variables was performed using the Pearson's correlation coefficient.


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RESULTS
 
CD4+ CD25– cells show a compromised ability for FIV binding and entry compared to CD4+ CD25+ cells. Previously we have shown that CD4+ CD25+ cells replicated FIV in vitro in the presence of IL-2 alone, whereas CD4+ CD25– cells established a latent infection and could be activated to produce infectious virus only via a strong mitogenic stimulus such as ConA (24). However, ConA-stimulated CD4+ CD25– cells produced significantly less virus than IL-2-cultured CD4+ CD25+ cells, suggesting that other factors in addition to those controlling virus replication may govern permissiveness of CD4+ CD25– cells to FIV infection. To determine if these cell types differed with respect to FIV permissiveness at the level of viral entry, virus binding and entry assays were performed on highly purified CD4+ CD25+ and CD4+ CD25– T-cell subsets purified by antibody-coated magnetic beads or FACS sorting as demonstrated in Fig. 1. PCR amplification of proviral DNA was performed 8 h postinfection on purified CD4+ CD25+ and CD4+ CD25– T-cell subsets infected with serial dilutions of FIV. As shown in Fig. 2A, proviral DNA was detectable at 16-fold-lower virus dilutions in CD4+ CD25+ cells compared to those in CD4+ CD25– cells, suggesting that CD4+ CD25+ cells are more permissive to FIV infection. Stimulation of CD4+ CD25– cells with ConA prior to infection enhanced their permissiveness to FIV, which was comparable to that seen in CD4+ CD25+ cells (Fig. 2A).



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  		FIG. 2. Differential permissiveness to FIV infection in CD4+ CD25+ versus CD4+ CD25– cells. (A) LN-derived cells were sorted into highly purified CD4+ CD25+ and CD4+ CD25– populations by using FACS. CD4+ CD25+, CD4+ CD25–, or CD4+ CD25– cells (105 each) stimulated with ConA for 72 h were infected in vitro with serial twofold dilutions of FIV-NCSU1 starting with a MOI of 0.1 (lane 1). Cells were harvested 8 h postinfection and DNA was isolated by direct cell lysis for PCR of proviral DNA by using FIV LTR-specific primers. Water control reaction did not yield any visible amplification product. For GAPDH controls, three samples were picked up randomly from each group of CD4+ CD25+ (lanes 1-3), CD4+ CD25– (lanes 4-6), and CD4+ CD25– (lanes 7-9) cells stimulated with ConA. The experiment was repeated four times with similar results. (B) Estimation of viral copy numbers for binding and entry assays in CD4+ CD25+ and CD4+ CD25– cells using real-time RT-PCR. LN-derived CD4+ CD25+ and CD4+ CD25– cells of high purity (>97%) were obtained using a combination of immunomagnetic beads and flow cytometric sorting. Sorted cells (106) were used for viral binding and viral entry experiments as described in Materials and Methods. RNA (0.5 µg) from each cell type was used for real-time RT-PCR using FIV-NCSU1 gag-specific primers. Viral copy numbers were determined using in vitro-transcribed gag RNA standard. To determine if CD4+ CD25– cells show an increase in viral receptiveness following mitogenic stimulation, cells were stimulated for 72 h with 2-µg/ml ConA prior to binding and entry assays. The experiment was repeated thrice withsimilar results. Error bars represent means ± standard deviations of triplicate wells. The P values represent significant increases in viral binding or entry in CD4+ CD25+ or ConA-stimulated CD4+ CD25– cells versus unstimulated CD4+ CD25– cells. RT-PCR using GAPDH-specific primers was used to demonstrate equal RNA loading per reaction (data not shown). Water control reaction lacked any detectable amplification products (data not shown).

Having seen a difference with respect to proviral DNA in in vitro-infected CD4+ CD25+ versus CD4+ CD25– cells, viral binding and entry assays were performed in the above cell types by determining viral RNA copy numbers by real-time RT-PCR. For binding assays, purified cell populations were incubated with FIV on ice for 1 h, and total RNA was isolated immediately following incubation and washing. For detecting viral entry, cells were incubated with FIV at 37°C for 1 h and trypsinized to remove cell surface-attached virions prior to RNA isolation. While CD4+ CD25+ cells bound FIV twice as efficiently as did CD4+ CD25– cells, the difference was approximately 50-fold in terms of viral entry (Fig. 2B). CD4+ CD25– cells, on the other hand, when stimulated with a mitogen prior to virus incubation were as efficient as CD4+ CD25+ cells with regard to both FIV binding and entry (Fig. 2B). Trypsinization control experiments suggested that trypsin-EDTA treatment was capable of stripping most of the virus attached to the cell surface (data not shown). The control reaction in which the RT step was omitted did not yield any visible amplification product, indicating the absence of contaminating DNA (data not shown). The above findings suggest that CD4+ CD25– cells are compromised with respect to FIV binding, and an additional restriction occurs at the level of viral entry into cells.

Differential expression of FIV coreceptor CXCR4 on CD4+ CD25+ and CD4+ CD25– cells. Having seen a difference in CD4+ CD25+ and CD4+ CD25– cells at the level of viral binding and entry, we investigated the expression of FIV coreceptor CXCR4 on the two cell types. As seen in Fig. 3A, two-color flow cytometry staining of feline LN cells with CD4 and CXCR4 antibodies revealed that >95% of the CD4+ cells were CXCR4 positive. As CD4+ CD25+ cells represent 20 to 30% of the total LN CD4+ T cells (51), these data indicate that the majority of CD4+ CD25+ as well as CD4+ CD25– cells express CXCR4. However, comparison of the mean fluorescence intensity (MFI) of CXCR4 expression on CD4+ CD25+ and CD4+ CD25– cells revealed that CD4+ CD25+ cells inherently expressed higher levels of CXCR4 on a per cell basis (MFI of 401.58 versus 116.26) when assayed immediately postisolation (Fig. 3B). IL-2 treatment for 3 days led to only moderate increases in CXCR4 expression levels on both CD4+ cell subsets (Fig. 3B; 401.58 versus 425.65 for CD4+ CD25+ and 116.26 versus 157.47 for CD4+ CD25– cells). Interestingly, treatment with ConA for 3 days led to a significant increase in CXCR4 expression in CD4+ CD25– cells (Fig. 3B; 116.26 versus 385.19) but downregulated CXCR4 expression in CD4+ CD25+ cells (Fig. 3B; 401.58 versus 219.48). These findings suggest that the reduced viral permissiveness exhibited by CD4+ CD25– cells when compared to CD4+ CD25+ cells could be due to a lower surface CXCR4 expression in these cells.



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  		FIG. 3. Expression of CXCR4 in CD4+, CD4+ CD25+, and CD4+ CD25– cells. (A) LN-derived cells (5 x 105) were incubated with cross-reacting anti-CXCR4 antibody (44717) followed by PE-conjugated goat anti-mouse IgG. The cells were then washed, stained with FITC-labeled feline anti-CD4 antibody (30A), and analyzed by flow cytometry. Numbers represent the percentages of cells within each quadrant. Ab, antibody. (B) LN-derived cells were sorted into CD4+ CD25+ and CD4+ CD25– populations. Purified cells (3 x 105) were stained for surface CXCR4 expression as described above either immediately postisolation or after stimulation for 3 days with 100-U/ml IL-2 or 2-µg/ml ConA and analyzed by flow cytometry. One representative of four independent experiments is shown. Shaded region, secondary antibody (Ab) alone.

Having determined that CD4+ CD25+ and CD4+ CD25– cells differ with respect to levels of CXCR4 expression, we asked whether the specific CXCR4 antagonist AMD-3100 could block infection in a concentration-dependent manner. CD4+ CD25+ and CD4+ CD25– cells were infected with FIV-NCSU1 in the presence of various concentrations of AMD-3100 and p24 antigen in culture supernatants detected by an ELISA. As seen in Fig. 4, concentrations of AMD-3100 below 0.01 µg/ml had a minimal effect in blocking FIV infection of both of the cell types. However, concentrations of AMD-3100 in the range of 0.01 to 1 µg/ml were significantly more efficacious in blocking FIV infection of CD4+ CD25– (50% inhibitory concentration, 0.0295) compared to CD4+ CD25+ (50% inhibitory concentration, 0.2063) cells, again suggesting a higher density of CXCR4 on the Treg cells.



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  		FIG. 4. Blocking FIV infection in CD4+ CD25+ and CD4+ CD25– cells by using the CXCR4 antagonist AMD-3100. LN-derived CD4+ CD25+ and CD4+ CD25– cells (2 x 105) were infected with FIV-NCSU1 at a MOI of 0.1 either in the absence of AMD-3100 or in the presence of serial 10-fold dilutions of AMD-3100 starting with a maximum concentration of 10 µg/ml. Following infection, CD4+ CD25+ and CD4+ CD25– cells were cultured in the presence of 100-U/ml IL-2 or 2-µg/ml ConA, respectively, for 12 days. Culture media was replenished with AMD-3100 to minimize second-round infections. Supernatants were harvested every 3 days for detection of p24 antigen by ELISA. Only results from 9-day harvests are shown. Data points represent means of duplicate cultures. The experiment was repeated thrice with similar results. The P values represent significantly higher levels of p24 antigen in culture supernatants of CD4+ CD25+ versus CD4+ CD25– cells at corresponding concentrations of AMD-3100. {diamondsuit}, CD4+ CD25+; {square}, CD4+ CD25–.

To further establish a correlation between CXCR4 expression levels and susceptibility to FIV infection, a time study was performed to determine kinetics of CXCR4 upregulation in parallel with viral entry into CD4+ CD25– cells. CD4+ CD25– cells were either left unstimulated or were stimulated with ConA for different time periods prior to infection with FIV as indicated in Fig. 5. Following stimulation, cells were monitored for viral entry by real-time PCR and CXCR4 expression by flow cytometry. The MFI of CXCR4 expression started to increase after 4 h of stimulation and continued to increase thereafter up to 72 h (Fig. 5B). In parallel, viral entry in CD4+ CD25– cells increased in an identical pattern (Fig. 5A). These results indicate a strong correlation (R = 0.9845; Fig. 5C) between increase in viral entry in CD4+ CD25– cells following ConA stimulation and an upregulation of cell surface CXCR4 expression on a per cell basis.



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  		FIG. 5. Correlation of the increase in viral entry in CD4+ CD25– cells following ConA stimulation with the increase in MFI of CXCR4 expression. Magnetic bead-purified CD4+ CD25– cells were stimulated with 2-µg/ml ConA for 4, 8, 24, 48, and 72 h or were left unstimulated. (A) Stimulated and unstimulated cells (106) were then infected with FIV, trypsinized to get rid of cell surface-attached virions, and cultured for 8 h. Subsequently, DNA was isolated from infected cells and 0.5 µg was used for real-time PCR using NCSU1 gag-specific primers. Error bars represent means ± standard deviations of triplicates. The experiment was repeated twice. One representative result is depicted. (B) Stimulated or unstimulated CD4+ CD25– cells (3 x 105) were stained for surface CXCR4 expression as described in the legend for Fig. 3A and analyzed by flow cytometry. Data represent means of duplicates. Real-time PCR data and CXCR4 staining from the same experiment are shown. (C) Linear correlation between MFIs of CXCR4 expression and viral copy numbers in CD4+ CD25– cells following ConA stimulation. Increase in viral entry in CD4+ CD25– cells following ConA stimulation is plotted against CXCR4 expression levels. The line represents linear correlation and the Pearson correlation coefficient (R).

FIV receptor CD134 (OX-40) is differentially expressed on CD4+ CD25+ and CD4+ CD25– T-cell populations. Recently it has been reported that FIV uses CD134 as the primary receptor for entry into cells (40). Hence we investigated whether CD4+ CD25+ and CD4+ CD25– cells differed with respect to the cell surface expression of CD134. Flow cytometry analysis using the human CD134 antibody clone previously described to cross-react with the feline CD134 homolog (40) revealed that within the PBMC or LN population the number of CD134+ cells ranged from 6 to 14% in eight normal cats analyzed. Figure 6A represents a typical flow cytometry analysis of CD134 expression on feline LN cells. Similar analysis on sorted cell populations revealed that approximately 25 to 30% of CD4+ CD25+ and 10 to 15% of CD4+ CD25– cells expressed CD134 (Fig. 6B). Surprisingly, stimulation with ConA at a concentration of 2 µg/ml did not upregulate surface CD134 expression on either CD4+ T-cell subset (Fig. 6C). On the contrary, there seemed to be a downregulation of surface CD134 expression on CD4+ CD25+ and CD4+ CD25– cells following ConA stimulation (Fig. 6B versus C). At a similar concentration of ConA, 90 to 95% of CD4+ cells became positive for the activation marker CD25 (Fig. 6C, top) compared to 5 to 10% of the CD4+ CD25+ population present prior to stimulation (Fig. 1A). Treatment with 100-U/ml IL-2 did not have any effect on the surface expression of either CD25 or CD134 (data not shown). In order to rule out the possibility that the low levels of CD134 expression on feline CD4+ CD25+ and CD4+ CD25– cells were due to nonspecific antibody binding, HeLA cells were either transiently transfected or stably selected to express feline CD134, and surface CD134 expression was detected by flow cytometry. As depicted in Fig. 6D, the anti-human BerACT35 antibody did bind specifically to feline CD134, as detected by an increase in CD134 expression on transiently transfected cells as opposed to untransfected controls. Expression of CD134 was even higher in clones that were selected to stably express the feline CD134 homolog. From these findings we can conclude that although CD4+ CD25+ cells express higher levels of FIV receptor and coreceptor than do CD4+ CD25– cells, the increase in viral entry in CD4+ CD25– cells following ConA stimulation correlates with increased CXCR4 but not CD134 expression.





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  		FIG. 6. Expression of FIV receptor CD134 (OX40) on total LN and CD4+ CD25+ versus CD4+ CD25– cells. (A) LN (5 x 105) cells were stained with PE-conjugated anti-CD134 antibody and analyzed by flow cytometry. (B) LN-derived cells were sorted into CD4+ CD25+ and CD4+ CD25– populations by using FACS. Surface CD134 expression in the above cell types was determined immediately after sorting by staining with PE-conjugated anti-CD134 antibody (BerACT35) followed by flow cytometry. Numbers represent the percentages of CD4+ CD25+ or CD4+ CD25– cells expressing CD134. One representative of four independent experiments is depicted. (C) Total unsorted LN cells were stimulated with 2-µg/ml ConA for 72 h. Cells (5 x 105) were then stained with allophycocyanin-conjugated anti-CD4, FITC-conjugated anti-CD25, and PE-conjugated anti-CD134 antibodies and analyzed by flow cytometry. (Top) CD25 expression on CD4+ cells stimulated with ConA for 72 h. (Bottom) Gated R2:CD4+ CD25+ cells and R3:CD4+ CD25– cell populations were analyzed for surface CD134 expression. Numbers represent the percentages of ConA-stimulated CD4+ CD25+ or CD4+ CD25– cells coexpressing CD134. (D) HeLA cells either untransfected, transiently transfected, or stably selected to express feline CD134 were stained for surface CD134 expression followed by flow cytometry analysis. Numbers represent the percentages of CD134-positive cells. Dotted line, untransfected HeLa cells; solid line, HeLa cells expressing feline CD134.

CD4+ CD25+ cells constitutively express high levels of cellular transcription factors known to activate FIV gene expression. Previously we have shown that while CD4+ CD25+ cells replicate FIV in vitro in the presence of IL-2 alone, CD4+ CD25– cells require a strong mitogenic stimulus like ConA for a productive FIV infection (24). An important block to retroviral replication in resting and naive T cells is at the level of cellular transcriptional activity (45). Hence, we analyzed the activation status of the transcription factors ATF, AP-1, and CEBP, which are known to play an important role in FIV replication (5, 25). Figure 7A depicts the nucleotide sequence of the FIV-NCSU1 LTR from position 40 to 200 and the predicted binding sites for the transcription factors. The shift assay revealed constitutive transactivation of CEBP and ATF in CD4+ CD25+ T cells when analyzed immediately after isolation (Fig. 7B, lane 4), which was further enhanced with IL-2 treatment (Fig. 7B, lane 5). CD4+ CD25+ cells also expressed low constitutive activation of transcription factor AP-1 (Fig. 7B, lane 4), which was significantly upregulated by IL-2 treatment (Fig. 7B, lane 5). In contrast, CD4+ CD25– cells expressed low or no constitutive activity of these transcription factors (Fig. 7B, lane 7), all of which were highly upregulated by ConA (Fig. 7B, lane 9) but not IL-2 (Fig. 7B, lane 8) treatment. Hence, while CD4+ CD25+ cells demonstrated constitutive or IL-2-dependent activation of transcription factors known to be important in FIV replication, CD4+ CD25– cells expressed these transcription factors only following treatment with ConA. This correlates with FIV replication in CD4+ CD25+ cells in the presence of IL-2 alone but in CD4+ CD25– cells only when stimulated with a strong mitogenic stimulus.



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  		FIG. 7. (A) Sequence of the FIV-NCSU1 LTR (sense strand only) from nucleotide positions 40 to 200. The predicted binding sites for the transcription factors NF-{kappa}B, AP-1, CEBP, and ATF are underlined. The oligonucleotides used for the detection of NF-{kappa}B, CEBP, AP-1, and ATF are indicated by bold letters. (B) EMSA of nuclear extracts derived from CD4+ CD25+ and CD4+ CD25– cells. LN-derived cells were sorted into CD4+ CD25+ and CD4+ CD25– populations using immunomagnetic beads. Nuclear extracts were prepared from 106 cells either immediately following isolation or after stimulation for 5 days with 100-U/ml IL-2 or 2-µg/ml ConA as indicated (lanes 4-9). Equal amounts of extracts from different cell types or treatments were incubated with 5' [{gamma}-32P]ATP-labeled ATF, AP-1, or CEBP oligonucleotides for 15 min at room temperature. Fifty-fold molar excesses of specific competitor (SC, lane 2) or nonspecific competitor (NSC, lane 3) were used to demonstrate specific protein binding of each probe. Following binding, the complexes were resolved on a nondenaturing polyacrylamide gel, dried, and autoradiographed. The experiments were repeated thrice with similar results.

NF-{kappa}B binding to the FIV LTR is not important for a productive infection. While a putative NF-{kappa}B motif is variably expressed on isolates of FIV, specific protein binding to this sequence has not been established (41, 47). We also observed that the putative NF-{kappa}B oligonucleotide derived from the FIV-NCSU1 LTR did not bind efficiently to NF-{kappa}B by EMSA (data not shown). Moreover, the binding was not enhanced in mitogen-treated cells, suggesting that this FIV-NCSU1 sequence does not provide a good fit for NF-{kappa}B. To further explore if NF-{kappa}B is activated in CD4+ CD25+ Treg cells, we developed consensus oligonucleotides for the NF-{kappa}B binding site (Fig. 8A). With this probe, CD4+ CD25+ cells showed constitutive high expression of NF-{kappa}B, which was further upregulated by IL-2 treatment (Fig. 8A, lane 4 versus lane 5). CD4+ CD25– cells revealed no constitutive NF-{kappa}B activity (Fig. 8A, lane 7), but activation was observed following ConA and to a lesser extent IL-2 treatment (Fig. 8A, lane 8 versus lane 9). In order to elucidate the discrepancy seen with the NF-{kappa}B probe derived from the FIV LTR versus the consensus NF-{kappa}B oligonucleotide, cross-competition experiments were performed using nuclear extracts derived from FCD4E cells. Incubation of FCD4E nuclear extracts with an excess of unlabeled consensus NF-{kappa}B probe prior to addition of 32P-labeled FIV LTR NF-{kappa}B probe abolished weak binding of the FIV LTR probe (Fig. 8B, lane 3 versus lane 7). In contrast, incubation with an excess of unlabeled LTR-derived NF-{kappa}B oligonucleotide prior to addition of radiolabeled consensus NF-{kappa}B oligonucleotide did not have any effect on NF-{kappa}B binding to consensus oligonucleotide (Fig. 8B, lane 4 versus lane 8). These results suggest that, while NF-{kappa}B is upregulated in CD4+ CD25+ Treg cells, the putative NF-{kappa}B binding site in the FIV LTR is not a perfect match for binding to NF-{kappa}B and probably does not play an important role in FIV replication.



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  		FIG. 8. NF-{kappa}B binding to the FIV LTR is not required for a productive FIV infection. (A) EMSA of nuclear extracts from CD4+ CD25+ and CD4+ CD25– cells using consensus NF-{kappa}B oligonucleotide. Nuclear extracts were prepared from CD4+ CD25+ and CD4+ CD25– cells after different treatments (lanes 4-9) as described in the legend for Fig. 7B. Nuclear extracts were then incubated with consensus NF-{kappa}B oligonucleotide for 15 min at room temperature in the presence or absence of specific competitor (SC, lane 2) or nonspecific competitor (NSC, lane 3). Complexes were resolved on a polyacrylamide gel, dried, and autoradiographed. One representative of three independent experiments is depicted. (B) Cross-competition assays using radiolabeled NF-{kappa}B oligonucleotide derived from NCSU1 LTR sequence or consensus NF-{kappa}B sequence. Nuclear extracts were prepared form 1.5 x 106 FCD4E cells, an IL-2-dependent CD4+ cell line highly permissive for FIV-NCSU1 infection. Five micrograms of extracts was incubated either with end-labeled NCSU1 LTR-derived (lane 3) or consensus NF-{kappa}B (lane 4) oligonucleotides. Complexes were subsequently resolved on a 6% polyacrylamide gel. For competition (lanes 5-6) and cross-competition (lanes 7-8) assays, nuclear extracts were incubated with a 50-fold molar excess of the indicated unlabeled probes prior to incubation with the radiolabeled probe. One representative of four independent experiments is depicted.


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DISCUSSION
 
Previously we have characterized feline CD4+ CD25+ T-regulatory cells both with regard to their immune regulatory properties and ability to support FIV-NCSU1 replication (24, 51). Our studies showed that feline CD4+ CD25+ cells possess the key characteristics of Treg cells with regard to their anergic state and ability to suppress proliferation of CD4+ CD25– cells via inhibition of IL-2 production. Further, we also demonstrated that these cells are arrested in the G0/G1 state of the cell cycle and express on their surface CTLA4, a marker associated with Treg cells (51). Virus replication studies revealed that while CD4+ CD25+ cells harbored a productive FIV infection in the presence of IL-2 alone, CD4+ CD25– cells established a latent infection and required a strong mitogenic stimulus such as ConA to produce infectious virus. While CD4+ CD25+ cells efficiently replicated FIV, they themselves remained anergic to mitogenic stimulus and activation-induced cell death, suggesting their potential to act as a long-term reservoir of productive FIV infection (24). The present study was aimed at investigating the mechanisms governing infection and replication of FIV in CD4+ CD25+ cells.

Ramilo et al. (39) demonstrated that PBMC depleted of CD25+ cells by using anti-CD25 immunotoxin were severely restricted in their ability to replicate HIV in vitro. Further, Chou et al. (9) demonstrated that, whereas CD4+ CD25+ T cells could be productively infected with HIV-1 in vitro, highly purified CD4+ CD25– cells were incapable of being infected. However, they did not elucidate the mechanism(s) restricting HIV infection of CD4+ CD25– cells. In the present study, we found that CD4+ CD25– cells were less susceptible than CD4+ CD25+ T cells to FIV infection in vitro which correlated with reduced surface expression of the FIV coreceptor CXCR4 on their surface. In agreement with increased density of CXCR4 on CD4+ CD25+ T cells compared to CD4+ CD25– T cells is the observation that CXCR4 antagonist AMD-3100 was able to block FIV infection of CD4+ CD25– cells at significantly lower concentrations than that required to block infection of CD4+ CD25+ Treg cells. Further, CD4+ CD25– cells express significantly less CXCR4 on their surface, which is consistent with the finding that they bind twofold less virus on their surface. However, the level of viral entry seen in the CD4+ CD25– cells was approximately 50-fold less than that seen in CD4+ CD25+ cells, which is most likely explained by the fact that cell surface components other than the viral receptor and coreceptor like heparan sulfate proteoglycans serve as facilitators in nonspecific cellular attachment of HIV and FIV (13, 32, 35). Our findings of CXCR4 expression on feline cells corroborate the findings of others reporting that essentially all CD4+ cells express CXCR4 (16, 38). Increased expression of CXCR4 on CD4+ CD25+ Treg cells as opposed to resting CD4+ CD25– T cells may be due to the fact that the Treg cells are partially activated (29, 51). This is well established, as evidenced by studies that CXCR4 surface expression is increased following cell activation (6). The importance of CXCR4 expression levels on a per cell basis in HIV infection has also been emphasized by others. Harouse et al. (20) found that although comparable percentages of CD3+ CD4+ CXCR4+ cells are present in the gut-associated lymphoid tissue and periphery, the level of CXCR4 expression is significantly lower in CD3+ lamina propria lymphocytes than the LN or peripheral blood. This could explain why X4-tropic viruses cause a more profound loss of circulating CD4+ T cells than intestinal CD4+ cells.

Shimojima et al. (40) recently showed that FIV utilizes CD134 (OX-40), a CD4+ T-cell activation marker belonging to the tumor necrosis factor receptor superfamily, as the primary receptor for entry into cells. In the present study, we found that the percentage of CD4+ CD25+ Treg cells that expressed surface CD134 was significantly higher than for CD4+ CD25– cells, suggesting a relationship between increased susceptibility of Treg cells to FIV infection and CD134 expression. However, ConA stimulation did not upregulate surface CD134 expression on CD4+ CD25– T cells, although it markedly increased their susceptibility to FIV-NCSU1 infection in vitro. Barten et al. (3) also found only a moderate increase in surface CD134 expression in rat whole blood stimulated with high concentrations of ConA. However, stimulation of feline CD4+ CD25– cells with ConA resulted in an increase in CXCR4 expression, which correlates significantly with viral entry. Moreover, the timeline of increased susceptibility of ConA-stimulated CD4+ CD25– cells showed a linear correlation with upregulation of CXCR4 on their surface. Thus, while the NCSU1 isolate of FIV utilizes the CD134 receptor (B. J. Willett, personal communication), our findings suggest that CXCR4 expression levels and not CD134 may be the limiting factor for FIV entry into CD4+ CD25+ and CD4+ CD25– T cells. In support of this, De Parseval et al. (14) recently showed that overexpression of CXCR4 in Crandell feline kidney cells renders them permissive to infection and to fusion of a primary isolate of FIV independent of CD134 expression.

To further explore potential differences in CD4+ CD25+ and CD4+ CD25– cells that could account for a productive infection in Treg cells but a latent replication-competent infection in the naive T-helper cells, we analyzed expression of a number of cellular transcription factors known to regulate the FIV promoter. EMSA of several cellular transcription factors revealed a striking difference in their activation status in CD4+ CD25+ and CD4+ CD25– T cells. Consistent with other reports (22, 26, 45, 54), our studies show that resting CD4+ CD25– cells have a low basal activity of AP-1, ATF, CEBP, and NF-{kappa}B that is upregulated strongly following ConA but not IL-2 stimulation. Activation of these transcription factors in CD4+ CD25– T cells by ConA stimulation but not IL-2 is consistent with the observation that ConA but not IL-2 converts a latent infection to a productive infection in these cells (24). In contrast to CD4+ CD25– cells, CD4+ CD25+ Treg cells showed constitutive high basal activity of ATF and CEBP and a strong AP-1 transactivation in response to IL-2. As it is well established that these transcription factors are required for FIV replication, these data support the argument that the ability of CD4+ CD25+ Treg cells to support FIV replication lies in the high basal and IL-2-responsive activity of these cellular transcription factors.

It is well established that NF-{kappa}B is an important transcription factor in regulating HIV replication (11, 45). However, we found that the putative NF-{kappa}B binding sequence in the NCSU1-FIV LTR did not bind NF-{kappa}B or compete with a consensus NF-{kappa}B probe, suggesting that NF-{kappa}B does not play a role in FIV replication in Treg cells. In support of this, a number of studies have reported that mutations in the putative NF-{kappa}B binding site in the FIV LTR does not affect FIV replication and probably does not play an important role in viral replication (41, 47). Interestingly, however, we did find by using the NF-{kappa}B consensus sequence that NF-{kappa}B was activated in unstimulated and IL-2-treated CD4+ CD25+ Treg cells but not CD4+ CD25– T cells.

Constitutive and IL-2-responsive activity of AP-1 and NF-{kappa}B in CD4+ CD25+ Treg cells as opposed to CD4+ CD25– cells is also of interest from the point of view that these transcription factors regulate IL-2 gene expression, and survival of anergic CD4+ CD25+ Treg cells is dependent upon IL-2 (34, 52). Thus it is possible that constitutive and/or IL-2-induced activation of these transcription factors not only maintains a constant basal level activation of the IL-2 promoter necessary for Treg cell survival but also provides necessary signals for FIV replication.

Overall, our results implicate an important role for CD4+ CD25+ cells as a reservoir for FIV replication both in vitro and in vivo. Although IL-2 is the limiting factor for viral replication in these cells in vitro, we believe that in the lymphoid milieu or during instances of immune activation the cytokine levels in vivo would be sufficient for these cells to support a productive FIV infection. Moreover, as IL-2 is necessary for the survival of CD4+ CD25+ Treg cells, activation of these transcription factors may be a component of their cell survival program and their ability to resist apoptosis (2, 24, 34, 44, 51). Thus, the partial activation status of CD4+ CD25+ Treg cells, as manifested in part by constitutive transactivation of transcription factors capable of promoting FIV replication, may be critical to maintaining a long-term stable reservoir of productive FIV infection. The CD4+ CD25– cells, on the other hand, could serve as a latent viral reservoir capable of competent reactivation when stimulated.

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ACKNOWLEDGMENTS
 
We thank Janet Dow and Debra Anderson for their excellent technical assistance. We also thank Koishi Ohno (University of Tokyo) for providing the anti-CD25 antibody, Edward Hoover (Colorado State University) for providing the CXCR4 antagonist AMD-3100, and Brian J. Willett (University of Glasgow) for providing the feline CD134 expression vectors.

The present study was supported in part by National Institute of Health grants AI38177, AI43858, and AI058691.


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FOOTNOTES
 
* Corresponding author. Mailing address: College of Veterinary Medicine, North Carolina State University, 4700 Hillsborough St., Raleigh, NC 27606. Phone: (919) 515-7394. Fax: (919) 515-4237. E-mail: Wayne_Tompkins{at}ncsu.edu. Back

{dagger} Present address: National Cancer Institute, Frederick, MD 21702. Back


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Journal of Virology, April 2005, p. 4965-4976, Vol. 79, No. 8
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.8.4965-4976.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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  • Holmes, D., Knudsen, G., Mackey-Cushman, S., Su, L. (2007). FoxP3 Enhances HIV-1 Gene Expression by Modulating NF{kappa}B Occupancy at the Long Terminal Repeat in Human T Cells. J. Biol. Chem. 282: 15973-15980 [Abstract] [Full Text]  
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