Little is known about what effector populations are associated with the control of human herpesvirus 8 (HHV-8) infection in vivo. We compared T lymphocyte subsets among HIVHHV-8+ and HIVHHV-8 infected human individuals. αβ+ T cells from HHV-8-infected individuals displayed a significantly higher percentage of differentiated effector cells among both CD4+ and CD8+ T cell subsets. HHV-8 infection was associated with significant expansion of γδ+ Vδ1 T cells expressing a differentiated effector cell phenotype in peripheral blood. In vitro stimulation of PBMC from HHV-8-infected individuals with either infectious viral particles or different HHV-8 viral proteins resulted in γδ Vδ1 T cell activation. In addition, γδ Vδ1 T cells displayed a strong reactivity against HHV-8-infected cell lines and prevented the release of infectious viral particles following the induction of lyric replication. These data indicate that γδ T cells play a role in both innate and adaptive T cell responses against HHV-8 in immunocompetent individuals.

Human herpesvirus 8 (HHV-8),3 also known as Kaposi’s sarcoma-associated herpesvirus (KSHV) is the etiological agent of several infectious diseases including Kaposi’s sarcoma, primary effusion lymphoma (PEL), and multicentric Castleman’s disease (1). Like other herpesviruses, HHV-8 is able to establish a predominantly latent, life-long infection in its host. The increased incidence of these diseases in immunocompromised individuals suggests that host immune control may be essential in preventing HHV-8-associated diseases (2). Several studies have demonstrated the anti-HHV-8 specificity of TCR αβ+CD8+ CTL (3, 4, 5, 6, 7, 8, 9, 10). HHV-8 specific CTL responses to latent and lytic viral proteins have been detected during primary HHV-8 infection (11). The contribution of these T cells in the resolution of HHV-8 infection has yet to be documented, because several immune evasion mechanisms targeting infected cell recognition by CTL have been described including blocking Ag presentation, inhibiting costimulatory molecule surface expression, and deregulating T cell activation signaling (12).

The large majority of T cells present in the peripheral blood of healthy individuals express αβ TCR, with T cells expressing the complementary γδ TCR typically accounting for <5% of the circulating T cells (13). Two main subsets of human γδ T cells have been described. One, expressing the TCR variable region Vδ2, represents the majority of peripheral blood γδ lymphocytes. γδ T cells of this subset play a role in the defense against intracellular pathogens and hematological malignancies (14, 15). By contrast, the second subset of Vδ1 T cells is resident mainly in the oral and intestinal epithelia, where these cells might provide a first line of defense against viral infections or malignancies (16). γδ T cells have been implicated in antiviral immune responses on the basis of their selective expansion in the peripheral blood of patients infected with HIV, CMV, EBV, and HSV (17, 18, 19, 20, 21).

In the present study, we examined the relative frequency of differentiated effector T cells in peripheral blood samples from HHV-8-infected and uninfected individuals. We report that HHV-8-infected immunocompetent individuals have a significant expansion of T lymphocytes expressing the γδ Vδ1 TCR and that these cells exhibit anti-HHV-8 specificity and antiviral activity.

Participants for this study were recruited from the Seattle, WA area for participation in studies of the natural shedding of HHV-8 infection (22). All HHV-8 positive subjects we studied were men who have sex with men, were HIV-1-negative as shown by ELISA, had Abs to HHV-8 in a combined whole virus ELISA plus a confirmatory immunofluorescence assay (IFA), and were also observed to shed HHV-8 DNA in saliva on ≥2 days of observation. HHV-8 seronegative controls were enrolled from cohorts followed with known low rates of HHV-8 and who demonstrated persistent seronegativity to HHV-8 over time. Both HHV-8 seropositive and seronegative populations were age matched. The study protocol was approved by the University of Washington Institutional Review Board, Seattle, WA.

PBMC were isolated from 50 ml of heparinized blood by Ficoll-Hypaque centrifugation and cryopreserved. Cells were stained with fluorescently labeled Abs as described previously. The staining combinations used were: CD3-Qdot 605, CD4-Alexa Fluor 405 (Caltag Laboratories/Invitrogen Life Technologies), CD8-allophycocyanin-Alexa Fluor 750 (Caltag Laboratories/Invitrogen Life Technologies), CD27-Qdot 655, Vδ1-FITC (δTCS1 clone; Pierce/Endogen), Vδ2-PE (BD/Pharmingen), CD45R0-Texas Red-PE (Beckman/Coulter), CD11a-allophycocyanin (BD/Pharmingen), and CD57-Alexa Fluors680. All Abs (mAb) were purchased labeled except CD3, CD27, and CD57, which were purchased purified from BD Pharmingen and conjugated in the laboratory using standard protocols (www.drmr.com/abcon). Qdot and Alexa dyes were purchased from Invitrogen Life Technologies. Dead cells were excluded following staining with propidium iodide (Sigma-Aldrich). Samples were analyzed on a LSR II flow cytometer (BD Biosciences). Data analysis, including postacquisition compensation, was performed using FlowJo software (Tree Star).

JMP software produced by the SAS Institute was used for all statistical analyses. Significant values for comparisons between groups were determined by the nonparametric Wilcoxon’s rank sum analysis. Data are shown as box plots in which the ends of the box are the 25th and 75th percentiles, and the line across the middle indicates the median. The lines above and below the box extend to the outermost data that falls within 1.5× interquartile range.

All cell lines were maintained in complete medium consisting of DMEM (Invitrogen Life Technologies) supplemented with antibiotics (100 IU/ml penicillin and 0.1 mg/ml streptomycin) and 10% heat-inactivated bovine serum (Gemini Bio-Products). The cell lines BCBL-1, JSC-1, Raji, Vero, and Ramos were obtained from the American Type Culture Collection. The lymphoblastoid cell line TM was obtained from Dr. S. R. Ridell (Fred Hutchinson Cancer Research Center, Seattle, WA). The Bjab cell line was obtained from M. Lagunoff (University of Washington, Seattle, WA). C1R, a human lymphoblastoid cell line, was provided by V. Groh (Fred Hutchinson Cancer Research Center). The rKSHV.219 infected JSC-1 cell line (JSC/rKSHV.219) has been described elsewhere (23).

Bjab infection by rKSHV.219 was achieved using cell to cell virus spread (24). Briefly, rKSHV.219 lytic replication was induced in latently infected Vero cells (105 cells/well in a 6-well plate) using sodium butyrate (0.1 μM, Sigma) and a recombinant baculovirus expressing KSHV ORF50 as described elsewhere (23). After overnight incubation, culture medium was removed and cells were washed with PBS. Induced Vero cells were cocultured with Bjab (106 cells/well) for 4 days, after which puromycin selection (0.5 mg/ml; Calbiochem) began. After 2 wk KSHV-infected Bjab were tested for GFP expression and Vero cells’ residual presence by flow cytometry using anti-CD19 Ab (BD Pharmingen).

PBMC (2 × 106 cells/ml, 5 ml/well in 6-well plate) were stimulated with infectious viral particles (5 × 104 infectious U/ml) in the presence of rhIL-2 (20 U/ml; Hemagen Diagnostics) and rhIL-7 (100 U/ml; R&D Systems). After 2 wk, γδ T cells were purified using magnetic cell separation according to manufacturer’s protocol (Miltenyi Biotec). The derived cell lines were further expanded under polyclonal expansion using PHA as previously described (25). Briefly, purified γδ T cells were expanded using PHA stimulation (0.8 μg/ml; Murex Diagnostics) in the presence of irradiated heterologous feeders (2 × 106 cell/ml, 5,000 rad).

HVS (strain C4488) was kindly provided by R. Desrosiers (New England Regional Primate Research Center, MA). Owl monkey kidney (OMK) cells were used for propagation of HVS. γδ T cell immortalization by HVS infection was performed as previously described (26). Briefly, γδ T cells were simultaneously stimulated with PHA (0.8 μg/ml; Murex Diagnostics) and irradiated heterologous PBMC (5,000 rad) with recombinant human (rh)IL-2 (20 U/ml) and rhIL-7 (100 U/ml) addition on the following day. Sample wells were infected over a 7-day period with 10% (v/v) prepared HVS virus supernatant. Immortalization was assessed based on the γδ T cell line’s ability to proliferate extensively in vitro independently of any external stimulation. T cell clones established from an immortalized γδ T cell line were generated as previously described (27). γδ T cells from an uninfected individual were isolated by FACS using a Vδ1-specific Ab (Pierce/Endogen). Cells were then immortalized by HVS as described above.

PBMC (2 × 106 cells/ml) were stimulated in the presence of the purified HHV-8 recombinant proteins gB, K8.1, open reading frame (ORF)65, and ORF73 (0.5 μg/ml) or infectious viral particles (105 infectious U/ml) in complete medium supplemented with rhIL-2 and rhIL-7. γδ T cells (5 × 105 cells/ml) were cocultured in a 96-well round-bottom plate with different cell lines (5 × 104 cells/ml). When specified, commercially available viral lysates from HHV-8 or CMV (Applied Biosystems) were added to the culture at a final concentration of 1 μg/ml. In some experiments cells from different PEL lines were mixed with γδ T cells (2 × 106 cells/ml) at various dilutions. When indicated, purified OKT3 or isotype control were added at a final concentration of 5 μg/ml. For purified viral protein stimulation, HVS-immortalized γδ T cells (5 × 105 cells/ml) were mixed with irradiated (5,000 rad) heterologous PBMC (105 cells/ml) in the presence of the purified HHV-8 recombinant proteins gB, K8.1, ORF65, and ORF73 (0.5 μg/ml). After 48 h, culture supernatants from triplicate wells were pooled and tested for the presence of cytokines. Measurements of human IFN-γ and TNF-α were analyzed using a sandwich ELISA. Samples were tested in duplicate. The coefficient of variation was always <10%. Matched pair mAbs were purchased from Endogen. The lowest detection limit for the IFN-γ assay is 0.3 pg/ml and 1 pg/ml for the TNF-α assay.

Highly purified γδ Vδ1 T cells from different cell lines were isolated using flow cytometry. Following an automated process, the positive cells were distributed in 24 wells at 10 cells/well into a 96-well PCR plate preloaded with 5 μl of lysis buffer (1× recombinant Thermus thermophilus buffer (Applied Biosystems), 0.005% Triton X-100, 0.5% 2-ME, and 3.3% proteinase K). Cell lysis was achieved by incubating the plate at 55°C for 15 min followed by a 5-min incubation at 95°C. Samples were then subjected to a nested PCR using γδ Vδ1-specific primers as previously described (28). Following the first PCR (recombinant Thermus thermophilus DNA polymerase; profile: 60°C for 30 min, 95°C for 5 min, and then at 95°C for 20 s/52°C for 20 s/60°C for 1 min for 45 cycles), 10 μl of the reaction mix was transferred into 90 μl of fresh PCR buffer before running the second PCR (AmpliTaq DNA polymerase; profile: 95°C for 2 min and then at 95°C for 30 s/54° for 30 s/72°C for 30 s for 35 cycles and then at 72°C for 7 min). Positive amplifications were visualized on a 2% agarose gel and cloned into the PCR2.1 TOPO vector (Invitrogen Life Technologies). Positive colonies were expanded overnight in 2 ml of Luria-Bertani medium. Plasmid DNA extraction was performed using a Qiagen kit according to the manufacturer’s instructions. M13 forward and reverse primers were used for the sequencing reactions. Nucleotide sequences were assigned to TCR-δ gene segments based on identities of V and J germline sequences published in GenBank using Blastn.

The HHV-8-GST fusion proteins gB, ORF73, ORF65, and K8.1A were a gift from Dr. B. Chandran (University of Kansas Medical Center, Kansas City, KS) and were expressed from recombinant baculoviruses (29). Protein expression was achieved by infecting SF9 cells with recombinant baculovirus. Supernatants were harvested after 5 days. Recombinant proteins were isolated by ammonium sulfate precipitation followed by dialysis against PBS. The proteins were further purified using a metal affinity resin (BD Biosciences) according to manufacturer protocol. Proteins concentration was assessed by a Bradford protein assay (Pierce).

Cells from the JSC/rKSHV.219 cell line were incubated with pooled culture supernatants from HVS-immortalized Vδ1 T cells previously stimulated for 48 h with JSC/rKSHV.219 as described above. Cells were cultured for 24 h in the presence of a blocking Ab specific for the IFN-γ receptor α-chain (10 μg/ml; BD Pharmingen) or an isotype control before the induction of lytic replication with sodium butyrate (0.1 μM). Cell-free supernatants were recovered 48 h later. Virus titrations were performed as described elsewhere (23). Briefly, virus yield was determined using a susceptible cell line by evaluating the number of GFP-expressing cells. Because this is not a plaque assay, viral yields were reported as infectious units per milliliter instead of PFU per milliliter. Infectious units of rKSHV.219 were determined by titering virus present in cell-free supernatants on 293 cells. The number of GFP-positive cells was determined 2 days postinfection by visually counting cells using an inverted Nikon fluorescence microscope. No GFP-positive cells were present on the same day as inoculation. Treatment of the 293 cell line with recombinant human IFN-γ did not affect its susceptibility to HHV-8 infection (data not shown).

We assessed T cell phenotypes by using memory and differentiation markers to determine whether there were any significant differences between HHV-8-infected and seronegative men. As shown in Table I, no significant difference in the relative percentage of CD4+ or CD8+ T cell subpopulations was noted between HHV-8-infected individuals and controls. However, we noted a significant increase in γδ lymphocytes expressing the Vδ1 receptor chain and a relative decrease in γδ lymphocytes expressing Vδ2 receptor chain in PBMC from HHV-8-infected individuals as compared with seronegative controls. Although the decrease in the percentage of Vδ2 T cells was not statistically significant, the Vδ1 T cell subset expansion and the overall reduction of the Vδ2 T cell subset led to a significant alteration in the peripheral Vδ1/Vδ2 T cell ratio in HHV-8-infected individuals (Table I).

Table I.

Phenotypic characteristic of αβ and γδ T lymphocytes in HHV-8 seropositive versus HHV-8 seronegative individualsa

HHV-8+ (n = 7)HHV-8 (n = 5)p Valueb
Age 45c 42  
Gender Male Male  
HIV status Negative Negative  
HHV-8 immunofluorescence assay Positive Negative  
HHV-8 PCR (saliva) Positive NAd  
Percentage CD4 among CD3 cells 57.9 63.8 0.1 
Percentage CD8 among CD3 cells 37.7 31.3 0.1 
Percentage Vδ1 among CD3 cells 4.2 (1.8–6.5)e 0.5 (0.3–1.2) 0.01 
Percentage Vδ2 among CD3 cells 1.3 4.1 0.3 
Vδ1/Vδ2 ratio 3.4 (0.8–4.8) 0.30 (0.08–0.6) 0.01 
HHV-8+ (n = 7)HHV-8 (n = 5)p Valueb
Age 45c 42  
Gender Male Male  
HIV status Negative Negative  
HHV-8 immunofluorescence assay Positive Negative  
HHV-8 PCR (saliva) Positive NAd  
Percentage CD4 among CD3 cells 57.9 63.8 0.1 
Percentage CD8 among CD3 cells 37.7 31.3 0.1 
Percentage Vδ1 among CD3 cells 4.2 (1.8–6.5)e 0.5 (0.3–1.2) 0.01 
Percentage Vδ2 among CD3 cells 1.3 4.1 0.3 
Vδ1/Vδ2 ratio 3.4 (0.8–4.8) 0.30 (0.08–0.6) 0.01 
a

All patients were clinically normal.

b

Determined by nonparametric Wilcoxon test.

c

Data are median.

d

NA, not available.

e

Interquantile range.

Previous studies have shown that CD57 surface expression on both CD4+ and CD8+ T lymphocyte subsets is characteristic of terminally differentiated cells in individuals with persistent viral infection (30, 31, 32). Using this specific marker, we determined the relative percentage of effector cells among αβ+ T lymphocytes. As shown in Fig. 1, A and B, we observed a significant increase in the relative percentage of CD57+ effector cells in both CD4+ (median 4.8/infected vs 0.9/uninfected) and CD8+ (median 43/infected vs 13.7/uninfected) peripheral T cell populations from HHV-8-infected individuals. We have previously shown that specific surface markers can be used to determine an effector T cell population not only among αβ but also for γδ T cells (30, 33). Consequently, we also used CD57 as a marker to determine the relative percentage of effector memory cells in both Vδ1 and Vδ2 γδ T cell subpopulations. As seen in Fig. 1, among the HHV-8-infected individuals only the Vδ1 T cell subset had a significant increase in the relative percentage of CD57+ effector cells similar to what we observed for the αβ+ T cells (median 64.8/infected vs 24.3/uninfected). In contrast, the frequency of effector cells among the Vδ2 T cell subpopulation remained unchanged between HHV-8-infected individuals and seronegative controls. For three infected individuals, PBMC from sequential blood draws were available. The observed Vδ1 T cell frequency always remained above the Vδ2 T cell frequency in all samples for a period of up to 28 mo (Fig. 1 C).

FIGURE 1.

αβ+ and γδ+ T cell phenotypes in HHV-8-infected individuals and uninfected controls. A, PBMC (107 cells) were stained using a panel of Abs as described in Material and Methods. Cells were scatter-gated on lymphocytes and then gated on CD3 to identify T cells. T cells not expressing Vδ1 or Vδ2 were then gated on CD4+ or CD8+. Relative percentages of cells expressing CD57+ within each T cell subset for representative HHV-8 seropositive and seronegative individuals are shown. B, Statistical analyses of the relative percentage of CD57+ cells in each T cell subset for both populations. C, Relative percentages of cells expressing Vδ1 and Vδ2 in sequential blood samples from a representative HHV-8-infected subject.

FIGURE 1.

αβ+ and γδ+ T cell phenotypes in HHV-8-infected individuals and uninfected controls. A, PBMC (107 cells) were stained using a panel of Abs as described in Material and Methods. Cells were scatter-gated on lymphocytes and then gated on CD3 to identify T cells. T cells not expressing Vδ1 or Vδ2 were then gated on CD4+ or CD8+. Relative percentages of cells expressing CD57+ within each T cell subset for representative HHV-8 seropositive and seronegative individuals are shown. B, Statistical analyses of the relative percentage of CD57+ cells in each T cell subset for both populations. C, Relative percentages of cells expressing Vδ1 and Vδ2 in sequential blood samples from a representative HHV-8-infected subject.

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We next investigated whether γδ Vδ1 T cells were able to respond to HHV-8 stimulation in vitro. PBMC from HHV-8-infected patients or uninfected controls were stimulated in the presence of IL-2 and IL-7 with or without infectious viral particles. Infectious viral particles were obtained from HHV-8-infected Vero cells in which lytic viral replication was artificially triggered as previously described (23). As shown in Table II, stimulation with HHV-8 infectious viral particles demonstrated no significant expansion among either αβ+ CD4+CD8+ or γδ+ Vδ1+Vδ2+ T cell subsets in PBMC from uninfected individuals. In contrast, stimulation of PBMC from all HHV-8-infected individuals resulted in a 3- to 6-fold expansion of the γδ Vδ1 T cell population.

Table II.

γδ T cell expansion following stimulation with HHV-8 viral particles and purified viral recombinant proteinsa

SubjectsCD4+CD8+Vδ1+Vδ2+
HHV-8 seropositive     
 Serpos I     
  Vector 46.3 36.0 2.4 3.3 
  gB 36.1 47.1 2.3 4.3 
  K8.1 34.6 47.9 2.0 2.2 
  ORF65 38.3 46.8 2.5 2.3 
  ORF73 38.2 43.2 2.2 2.6 
  Infectious particles 46.3 37.6 10.2 2.4 
 Seropos II     
  Vector 49.9 34.1 2.0 0.8 
  gB 51.4 32.4 4.1 1.6 
  K8.1 50.3 30.6 5.9 1.4 
  ORF65 53.3 31.4 3.3 1.9 
  ORF73 43.1 40.8 3.2 1.6 
  Infectious particles 50.6 30.3 5.9 2.4 
 Seropos III     
  Vector 54.0 34.2 1.2 1.0 
  gB 55.2 33.1 2.5 1.1 
  K8.1 50.1 37.4 2.5 1.0 
  ORF65 45.1 44.6 3.3 1.1 
  ORF73 47.5 41.1 3.0 2.2 
  Infectious particles 58.2 30.9 7.9 0.8 
 Seropos IV     
  Medium 36.6 49.0 4.4 0.9 
  Infectious particles 14.9 68.0 17.9 6.3 
 Seropos V     
  Medium 39.0 53.1 3.4 0.5 
  Infectious particles 46.0 36.0 17.0 1.3 
 Seropos VI     
  Medium 49.9 31.4 7.3 0.8 
  Infectious particles 10.3 35.5 42.2 4.0 
 Seropos VII     
  Medium 40.2 55.7 0.6 0.4 
  Infectious particles 39.0 53.1 3.4 0.5 
HHV-8 seronegative     
 Seroneg I     
  Vector 33.5 24.4 3.0 33.6 
  gB 32.8 26.5 3.1 27.5 
  K8.1 28.2 28.9 4.1 29.0 
  ORF65 26.1 30.3 3.3 27.0 
  ORF73 31.2 27.4 3.1 27.5 
  Infectious particles 27.2 33.3 3.4 27.0 
 Seroneg II     
  Vector 66.3 17.1 1.7 0.9 
  gB 54.8 25.5 1.9 0.7 
  K8.1 55.3 26.8 2.3 1.4 
  ORF65 69.6 19.6 1.8 0.6 
  ORF73 76.3 15.2 1.4 0.7 
  Infectious particles 69.4 20.2 0.7 0.9 
 Seroneg III     
  Vector 72.6 10.2 1.3 7.5 
  gB 66.2 14.6 1.7 9.7 
 K8.1 71.7 11.7 1.6 7.0 
  ORF65 72.9 10.8 1.7 6.7 
  ORF73 74.7 10.3 1.4 5.8 
  Infectious particles 74.4 9.7 1.0 6.0 
 Seroneg IV     
  Medium 44.0 32.0 8.0 2.3 
  Infectious particles 49.0 30.0 7.6 1.6 
 Seroneg V     
  Medium 46.9 19.7 2.4 11.7 
  Infectious particles 42.2 19.0 2.5 12.6 
SubjectsCD4+CD8+Vδ1+Vδ2+
HHV-8 seropositive     
 Serpos I     
  Vector 46.3 36.0 2.4 3.3 
  gB 36.1 47.1 2.3 4.3 
  K8.1 34.6 47.9 2.0 2.2 
  ORF65 38.3 46.8 2.5 2.3 
  ORF73 38.2 43.2 2.2 2.6 
  Infectious particles 46.3 37.6 10.2 2.4 
 Seropos II     
  Vector 49.9 34.1 2.0 0.8 
  gB 51.4 32.4 4.1 1.6 
  K8.1 50.3 30.6 5.9 1.4 
  ORF65 53.3 31.4 3.3 1.9 
  ORF73 43.1 40.8 3.2 1.6 
  Infectious particles 50.6 30.3 5.9 2.4 
 Seropos III     
  Vector 54.0 34.2 1.2 1.0 
  gB 55.2 33.1 2.5 1.1 
  K8.1 50.1 37.4 2.5 1.0 
  ORF65 45.1 44.6 3.3 1.1 
  ORF73 47.5 41.1 3.0 2.2 
  Infectious particles 58.2 30.9 7.9 0.8 
 Seropos IV     
  Medium 36.6 49.0 4.4 0.9 
  Infectious particles 14.9 68.0 17.9 6.3 
 Seropos V     
  Medium 39.0 53.1 3.4 0.5 
  Infectious particles 46.0 36.0 17.0 1.3 
 Seropos VI     
  Medium 49.9 31.4 7.3 0.8 
  Infectious particles 10.3 35.5 42.2 4.0 
 Seropos VII     
  Medium 40.2 55.7 0.6 0.4 
  Infectious particles 39.0 53.1 3.4 0.5 
HHV-8 seronegative     
 Seroneg I     
  Vector 33.5 24.4 3.0 33.6 
  gB 32.8 26.5 3.1 27.5 
  K8.1 28.2 28.9 4.1 29.0 
  ORF65 26.1 30.3 3.3 27.0 
  ORF73 31.2 27.4 3.1 27.5 
  Infectious particles 27.2 33.3 3.4 27.0 
 Seroneg II     
  Vector 66.3 17.1 1.7 0.9 
  gB 54.8 25.5 1.9 0.7 
  K8.1 55.3 26.8 2.3 1.4 
  ORF65 69.6 19.6 1.8 0.6 
  ORF73 76.3 15.2 1.4 0.7 
  Infectious particles 69.4 20.2 0.7 0.9 
 Seroneg III     
  Vector 72.6 10.2 1.3 7.5 
  gB 66.2 14.6 1.7 9.7 
 K8.1 71.7 11.7 1.6 7.0 
  ORF65 72.9 10.8 1.7 6.7 
  ORF73 74.7 10.3 1.4 5.8 
  Infectious particles 74.4 9.7 1.0 6.0 
 Seroneg IV     
  Medium 44.0 32.0 8.0 2.3 
  Infectious particles 49.0 30.0 7.6 1.6 
 Seroneg V     
  Medium 46.9 19.7 2.4 11.7 
  Infectious particles 42.2 19.0 2.5 12.6 
a

PBMC (2 × 106 cells/ml) from HHV-8 seropositive (Seropos I–VII) and seronegative (Seroneg I–V) subjects were stimulated with the purified recombinant viral proteins gB, K8.1, ORF65, and ORF73 (0.5 μg/ml) or infectious viral particles (105 infectious units/ml) in the presence of rhIL-2 and rhIL-7. PBMC cultures in the presence of GST (vector) or cytokines alone (medium) were used as controls. After 2 wk, cells were recovered and analyzed by flow cytometry using a panel of Abs as described in Material and Methods. Cells were scatter-gated on lymphocytes. Relative percentages of positive cells for each marker 2 wk poststimulation are shown.

We next determined whether the observed expansion of the γδ Vδ1 T cell population could also be triggered in response to viral protein stimulation. To do so, PBMC from seropositive (n = 3) and seronegative (n = 3) individuals were cultured in the presence of the purified HHV-8 GST-fusion-proteins gB, ORF65, ORF73, and K8.1. As seen in Table II, PBMC stimulation with several purified viral proteins (gB, K8.1, and ORF65) resulted in a 2- to 3-fold expansion of the γδ Vδ1 T cell population in two of three HHV-8-infected individuals. In contrast, following similar stimulation no γδ Vδ1 expansion was observed in PBMC from seronegative individuals. No consistent expansion in the αβ+ CD4+ or CD8+ T cell populations were seen in the HHV-8+ persons.

We did, however, note some expansion in the Vδ2 T cell population in three of seven HHV-8 seropositive individuals (Seropos II, IV, and VI in Table II), an effect not observed ex vivo in PBMC from HHV-8 seropositive individuals. It is possible that this could reflect a bystander activation effect from a previous exposure to other related herpesviruses, especially EBV, which are known to stimulate this particular γδ T cell subset (34, 35). The serological status of the individuals enrolled in our study for these viruses was not available.

Following in vitro stimulation with infectious viral particles γδ T cells were purified and the derived cell lines were further expanded with PHA. Flow cytometry analysis revealed that the resulting cell lines were exclusively expressing the Vδ1 TCR chain (data not shown). We then mixed the expanded γδ Vδ1 T cells with two HHV-8+ infected cell lines: 1) JSC-1, a PEL cell line coinfected with EBV and HHV-8; and 2) BCBL-1, a PEL cell line infected with just HHV-8. We used the Burkitt’s lymphoma Raji (HHV-8EBV+) as the negative control in these experiments because there is no HHV-8 uninfected PEL line. Following coculture, large amounts of IFN-γ were detected for two γδ Vδ1 cell lines when stimulated with either BCBL-1 or JSC-1 (Fig. 2,A). In addition, significant levels of TNF-α secretion were also detected with one of the γδ Vδ1 cell lines in response to both HHV-8-infected PEL lines. In all experiments only minimal amounts of IFN-γ and TNF-α were measured following stimulation with Raji. Similar stimulation of two Vδ1 T cell lines established from seronegative individuals did not result in any significant IFN-γ secretion (Fig. 2 B). These data suggested that the γδ Vδ1 T cell reactivity we observed was directed against the HHV-8-infected tumor cell lines independently of EBV presence.

FIGURE 2.

γδ T cell stimulation with HHV-8-infected PEL lines. A, γδ T cell lines (2 × 106 cells/ml) established from two HHV-8 seropositive individuals were stimulated with cells from different HHV-8-infected PEL lines (JSC-1 and BCBL-1) or uninfected control (Raji) at various dilutions. B, γδ T cell lines (2 × 106 cells/ml) established from two seronegative individuals were cocultured with the JSC-1, BCBL-1, or Raji cell lines at a responder/stimulator ratio of 25:1. Cytokines secretion was measured at 48 h in pooled supernatants from triplicate cultures. Cytokines titers were determined by ELISA. The coefficient of variation for cytokine titration was <10%. Data are representative of three independent experiments.

FIGURE 2.

γδ T cell stimulation with HHV-8-infected PEL lines. A, γδ T cell lines (2 × 106 cells/ml) established from two HHV-8 seropositive individuals were stimulated with cells from different HHV-8-infected PEL lines (JSC-1 and BCBL-1) or uninfected control (Raji) at various dilutions. B, γδ T cell lines (2 × 106 cells/ml) established from two seronegative individuals were cocultured with the JSC-1, BCBL-1, or Raji cell lines at a responder/stimulator ratio of 25:1. Cytokines secretion was measured at 48 h in pooled supernatants from triplicate cultures. Cytokines titers were determined by ELISA. The coefficient of variation for cytokine titration was <10%. Data are representative of three independent experiments.

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Because the reactivity of the Vδ1 cell lines against HHV-8-infected PEL lines did not preclude a possible specific recognition of nonviral Ags, we established a new HHV-8-infected cell line from a Burkitt’s lymphoma line called Bjab using a previously described HHV-8 recombinant virus, rKSHV.219 (23). This recombinant carried the puromycin gene resistance that allowed a quick selection of a long-term infected cell population. It also provided an easy identification of latent or lytic replication in living cells thanks to the selective expression of the green (GFP) or red fluorescent protein (RFP), respectively. Bjab cells were cocultured with rKSHV.219-infected Vero cells in which lytic viral replication was artificially induced. Successfully infected cells expressed GFP and could be expanded under drug selection. As shown in Fig. 3,A, substantial amounts of IFN-γ were detected when Vδ1 T cells were stimulated with HHV-8-infected Bjab/rKSHV.219 or PEL lines. A similar level of IFN-γ secretion was measured when a commercially available viral lysate from HHV-8 but not from CMV was used to stimulate Vδ1 T cells in the presence of uninfected Bjab cells. Uninfected cell lines including Bjab did not induce any significant IFN-γ secretion. Stimulation of a Vδ1 T cell line established from an HHV-8-seronegative individual did not result in any significant IFN-γ secretion. These data indicated that the specificity of Vδ1 T cell lines from seropositive subjects is directed against HHV-8 viral Ags. To assess whether the activation of Vδ1 T cells by HHV-8-infected cells was mediated through engagement of the γδ TCR, cocultures of T cells with the JSC-1 PEL line were performed in the presence of OKT3. As shown in Fig. 3 B, the addition of OKT3 resulted in a strong inhibition of IFN-γ secretion. These data indicate that the HHV-8-infected PEL line triggered the production of IFN-γ by Vδ1 T cells through a mechanism involving TCR engagement.

FIGURE 3.

γδ T cells stimulation with HHV-8-infected cell lines. A, γδ T cell lines (5 × 105 cells/ml) established from a HHV-8 seropositive and a seronegative individual were stimulated with cells from different HHV-8-infected cell lines or uninfected controls at a responder/stimulator ratio of 10:1.Viral lysates from HHV-8 or CMV were added to a coculture of uninfected Bjab and γδ T cells at a final concentration of 1 μg/ml. B, γδ T cells (2 × 106 cells/ml) from an HHV-8-infected individual were stimulated with JSC-1 cells (responder/stimulator ration of 20:1) in the presence of an OKT3 Ab (5 μg/ml) or an isotype control. IFN-γ secretion was measured after 48 h by ELISA in pooled supernatants from triplicate cultures. The coefficient of variation for cytokine titration was <10%. Data are representative of three independent experiments.

FIGURE 3.

γδ T cells stimulation with HHV-8-infected cell lines. A, γδ T cell lines (5 × 105 cells/ml) established from a HHV-8 seropositive and a seronegative individual were stimulated with cells from different HHV-8-infected cell lines or uninfected controls at a responder/stimulator ratio of 10:1.Viral lysates from HHV-8 or CMV were added to a coculture of uninfected Bjab and γδ T cells at a final concentration of 1 μg/ml. B, γδ T cells (2 × 106 cells/ml) from an HHV-8-infected individual were stimulated with JSC-1 cells (responder/stimulator ration of 20:1) in the presence of an OKT3 Ab (5 μg/ml) or an isotype control. IFN-γ secretion was measured after 48 h by ELISA in pooled supernatants from triplicate cultures. The coefficient of variation for cytokine titration was <10%. Data are representative of three independent experiments.

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To further examine whether Vδ1 T cell expansion was Ag driven, we compared the γδ Vδ1 T cell repertoire diversity in PBMC from HHV-8-infected or uninfected individuals. We determined the CDR3 sequences expressed by γδ Vδ1 T cells following in vitro stimulation with or without HHV-8 as described in Table II. To do so, we developed a new method to determine the sequence of the third hypervariable CDR (CDR3) of the Vδ1 gene segment. Our method combines the use of flow cytometry and highly sensitive PCR. It allows us to gather CDR3 sequences from an extremely low number of T cells. The CDR3 repertoire expressed by each Vδ1 T cell population was analyzed based on how many different sequences are obtained (diversity) and how many times the same sequence is detected (complexity/oligoclonality). As shown in Fig. 4, the γδ TCR repertoire in the unstimulated PBMC from an HHV-8-seronegative individual contained a restricted number of Vδ1 clonotypes (2/10 distinct Vδ1 sequences). Following HHV-8 stimulation, the Vδ1 T cell repertoire became more diverse (7/9 distinct Vδ1 sequences) and no clonal amplification was detected. In contrast, the γδ Vδ1 repertoire in unstimulated PBMC from an HHV-8-infected patient exhibited a large junctional diversity (12/19 distinct Vδ1 sequences). The Vδ1 transcripts were oligoclonal and a dominant clonotype was detected. In vitro stimulation with HHV-8 resulted in a restricted Vδ1 repertoire (5/14 distinct Vδ1 sequences) characterized by the clonal expansion of the same dominant transcript detected in unstimulated PBMC. This particular clonotype was detected in 6 of 19 independent PCR in unstimulated PBMC and in 9 of 14 independent PCR in PBMC stimulated with HHV-8. The sequence of the dominant Vδ1 transcript was identical with the TCR sequence of a HHV-8-specific γδ T cell clone isolated from the same individual. These data provided direct evidence supporting enrichment for HHV-8-reactive γδ Vδ1 T cells within PBMC from infected patients.

FIGURE 4.

Junctional diversity of the Vδ1 T cell subpopulation. PBMC from a seropositive and a seronegative individual were stimulated with or without HHV-8 in the presence of rhIL-2 and rhIL-7. After 2 wk, γδ Vδ1 T cells were isolated and Vδ1 TCR transcripts from each individual were cloned and sequenced as described in Materials and Methods. These sequences are available from GenBank under accession numbers EF656621–EF656644.

FIGURE 4.

Junctional diversity of the Vδ1 T cell subpopulation. PBMC from a seropositive and a seronegative individual were stimulated with or without HHV-8 in the presence of rhIL-2 and rhIL-7. After 2 wk, γδ Vδ1 T cells were isolated and Vδ1 TCR transcripts from each individual were cloned and sequenced as described in Materials and Methods. These sequences are available from GenBank under accession numbers EF656621–EF656644.

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Identification of the Ags recognized by γδ Vδ1 T cells is critical for further understanding of their potential antiviral functions. This approach, however, requires a large number of effector cells and maintaining long-term T cell culture without losing their original functions. Therefore, we used HVS, an oncogenic tumor virus of New World monkeys, to immortalize different γδ Vδ1 T cell lines. Multiple lines of evidence have put forth the model that TCR γδ cell Ag reactivity is dependent on conformationally intact Ags, similar to Ig Ag recognition (13, 36, 37). As such, we further investigated whether different HHV-8 viral proteins known to be relevant serological Ags were able to induce stimulation of HVS-transformed γδ Vδ1 T cells. Immortalized Vδ1 T cells were cultured in the presence of the purified HHV-8-GST fusion proteins gB, ORF65, ORF73, and K8.1. Because γδ T cell activation has been shown to depend on costimulatory molecules and to require TCR cross-linking, PBMC were also added to the culture (38, 39). As shown in Fig. 5,D, viral protein stimulation of HVS-transformed γδ Vδ1 T cells from an HHV-8 seronegative individual induced a minimal amount of IFN-γ secretion. In contrast, the γδ Vδ1 line established from three different HHV-8 seropositive individuals produced significant amounts of IFN-γ following stimulation with the different viral proteins (Fig. 5, A–C).

FIGURE 5.

γδ T cells stimulation with HHV-8-purified viral proteins. Three HVS-immortalized γδ Vδ1 T cell lines derived from separate HHV-8+ patients (A–C) and an immortalized cell line derived from an HHV-8 seronegative person (D) were stimulated with different purified recombinant viral proteins (gB, K8.1, ORF 65, and ORF73) or control empty GST vector (vector). The specificity of a T cell clone established from the γδ Vδ1 line shown in A was also determined by stimulation in the presence of different purified viral proteins (E). HVS-immortalized γδ T cells (5 × 105 cells/ml) were cultured with irradiated (5,000 rad) heterologous PBMC (105 cells/ml) in the presence of different purified viral proteins (0.5 mg/ml) for 48 h. IFN-γ secretion was measured by ELISA in pooled supernatants from triplicate cultures. The coefficient of variation for cytokine titration was <10%. The data are representative of five independent experiments with similar results.

FIGURE 5.

γδ T cells stimulation with HHV-8-purified viral proteins. Three HVS-immortalized γδ Vδ1 T cell lines derived from separate HHV-8+ patients (A–C) and an immortalized cell line derived from an HHV-8 seronegative person (D) were stimulated with different purified recombinant viral proteins (gB, K8.1, ORF 65, and ORF73) or control empty GST vector (vector). The specificity of a T cell clone established from the γδ Vδ1 line shown in A was also determined by stimulation in the presence of different purified viral proteins (E). HVS-immortalized γδ T cells (5 × 105 cells/ml) were cultured with irradiated (5,000 rad) heterologous PBMC (105 cells/ml) in the presence of different purified viral proteins (0.5 mg/ml) for 48 h. IFN-γ secretion was measured by ELISA in pooled supernatants from triplicate cultures. The coefficient of variation for cytokine titration was <10%. The data are representative of five independent experiments with similar results.

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As shown in Fig. 5 there was a broad range of IFN-γ values observed, likely reflecting the heterogeneity of the different immortalized γδ Vδ1 T cell lines established from these HHV-8-infected individuals. In Fig. 3,A, different γδ Vδ1 T cell lines produced a broad range of IFN-γ and TNF-α secretions in response to the same stimulation with HHV-8-infected cell lines. Similarly, each immortalized γδ Vδ1 T cell line is heterogeneous. The highest value for IFN-γ secretion is observed for the γδ Vδ1 T cell line shown in Fig. 5,A. Analysis of the T cell repertoire diversity expressed by the PBMC used to establish this particular cell line is shown in Fig. 4. The Vδ1 transcripts were oligoclonal and a dominant clonotype was detected. The sequence of the dominant Vδ1 transcript was identical with the TCR sequence of a HHV-8-specific γδ T cell clone isolated from the same individual. The Vδ1 junctional sequence of the TCR expressed by this clone is shown in Fig. 4. As shown in Fig. 5 E, a significant amount of IFN-γ was detected following stimulation of this dominant T cell clone with the ORF65 purified fusion protein. Hence, the presence of a dominant clonotype is responsible for the high value for IFN-γ secretion observed for this particular immortalized Vδ1 cell line.

We next examined the ability of γδ T cells to inhibit HHV-8 propagation in vitro. Analysis of in vitro HHV-8 interaction with hosts cells and quantification of infection have been hampered by the absence of a lytic replication cycle and a reliable plaque assay. Therefore, we have developed a cell line made of JSC-1 cells that coexpress the HHV-8 recombinant virus rKSHV.219. We have previously shown that the fluorescent proteins expressed by rKSHV.219 can be used as indicators of viral entry and infection. As reported before, upon treatment with sodium butyrate the JSC/rKSHV.219 cell line was able to support viral replication with the release of infectious viral particles (23). The virus yield can easily be determined using a susceptible cell line by evaluating the number of GFP-expressing cells.

We initially evaluated whether the JSC/rKSHV.219 line was able to trigger γδ Vδ1 activation as well as the parental JSC-1 line. As shown in Fig. 6 A, the amounts of IFN-γ detected following stimulation by JSC-1 or JSC/rKSHV.219 were similar.

FIGURE 6.

γδ T cells antiviral activity. A, HVS-immortalized γδ T (1.5 × 106/ml) cells from a HHV-8-infected individual were cocultured in a 96-well plate with cells from JSC-1 or JSC/rKSHV.219 at various dilutions. After 48 h, pooled supernatants from triplicate cultures were recovered. The presence of IFN-γ was determined by ELISA. The coefficient of variation for cytokine titration was <10%. The data are representative of three independent experiments with similar results. B, Cells from the JSC/rKSHV.219 cell line were incubated with pooled culture supernatants from three different HVS-immortalized Vδ1 T cell lines previously stimulated for 48 h with JSC/rKSHV.219. Cells were cultured for 24 h in the presence of a blocking Ab specific for the IFN-γ receptor (10 μg/ml) or an isotype control before induction of lytic replication with sodium butyrate (1 μg/ml). Cell-free supernatants were recovered 48 h later. Infectious viral particles titration was assessed by counting GFP-positive 293 cells 48 h postinfection using an inverted fluorescent microscope. Data represent the mean value of three separate counts.

FIGURE 6.

γδ T cells antiviral activity. A, HVS-immortalized γδ T (1.5 × 106/ml) cells from a HHV-8-infected individual were cocultured in a 96-well plate with cells from JSC-1 or JSC/rKSHV.219 at various dilutions. After 48 h, pooled supernatants from triplicate cultures were recovered. The presence of IFN-γ was determined by ELISA. The coefficient of variation for cytokine titration was <10%. The data are representative of three independent experiments with similar results. B, Cells from the JSC/rKSHV.219 cell line were incubated with pooled culture supernatants from three different HVS-immortalized Vδ1 T cell lines previously stimulated for 48 h with JSC/rKSHV.219. Cells were cultured for 24 h in the presence of a blocking Ab specific for the IFN-γ receptor (10 μg/ml) or an isotype control before induction of lytic replication with sodium butyrate (1 μg/ml). Cell-free supernatants were recovered 48 h later. Infectious viral particles titration was assessed by counting GFP-positive 293 cells 48 h postinfection using an inverted fluorescent microscope. Data represent the mean value of three separate counts.

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Having established that both lines are as potent activator for γδ Vδ1 T cells, we then assessed whether immortalized Vδ1 T cells affected JSC/rKSHV.219 viral yield in vitro.

Previous studies have shown that IFN-γ treatment of HHV-8-infected cell lines reduced the amount of infectious virus released following the induction of lytic replication with 12-O-tetradecanoylphorbol-13-acetate (40). Therefore, we investigated whether IFN-γ secreted by different Vδ1 T cell lines in the presence of JSC/rKSHV.219 could inhibit HHV-8 propagation in vitro. To avoid Vδ1 T cell activation due to exposure to sodium butyrate, cells from the JSC/rKSHV.219 cell line were incubated with culture supernatants from different Vδ1 T cell lines previously stimulated with JSC/rKSHV.219. Cells were cultured in the presence of a blocking Ab specific for the IFN-γ receptor α-chain or isotype control before the induction of lytic replication with sodium butyrate. As shown in Fig. 6 B, viral particle production by JSC/rKSHV.219 in the presence of supernatants from stimulated γδ Vδ1 T cells was dramatically reduced. In contrast, the addition of a blocking Ab specific for the IFN-γ receptor restored the production of infectious viral particles.

These data demonstrate that IFN-γ production by γδ Vδ1 T cells established from HHV-8-infected patients has an antiviral effect by preventing infectious viral particle release.

Our analysis indicates that HHV-8 infection, while asymptomatic clinically in the HIV person, is associated with persistent phenotypic changes in different peripheral blood T cell subpopulations. In contrast with the relative percentage of αβ+CD3+ T cells that remained stable in both CD4+ and CD8+ lymphocyte populations, we observed an 8-fold expansion among γδ CD3+ T cells expressing the Vδ1 receptor chain. A specific expansion of γδ Vδ1 T cells was also observed in vitro following PBMC stimulation with HHV-8. The absence of consistent expansion of αβ+CD4+ or CD8+ T cell subpopulations is in agreement with a previous report showing low lymphoproliferative response to HHV-8 detectable in only 42% of a similar cohort of HIVHHV-8+ men who have sex with men (6).

We also demonstrated a 2.5- to 3-fold increase in the relative frequency of cells expressing an effector phenotype among the αβ+CD4+, CD8+, and γδ Vδ1 T cell subpopulations, suggesting immune activation. Significant increases in relative frequency of CD57+ T cells have been reported in situations of persistent viral infection and chronic inflammation (41, 42). Recently, CD57 expression on CD4 and CD8 lymphocytes was also associated with replicative senescence and terminal differentiation (31, 32). It is commonly accepted that this is the result of chronic activation due to persistent Ag exposure.

The observation of CD57 expression on a large fraction of αβ+CD4+, CD8+, and γδ Vδ1 T cells in asymptomatically infected patients with HHV-8 is of interest. Mucosal shedding of virus has been difficult to discern in many people and our data suggest that HHV-8 persists in a lytic state more frequently than currently appreciated. Moreover, the presence of an expanded pool of lymphocytes expressing CD57 in PBMC from HHV-8-infected individuals could lead to a lesser capacity to proliferate in vitro following viral stimulation. Therefore, the extent and potential of the cellular immune response to HHV-8 in previous studies may have been underestimated.

The most novel observation made during our study was the selective long-lasting expansion of γδ T cells specifically from the Vδ1 subtype in peripheral blood samples from HHV-8-infected individuals. To date, Vδ1 T cell expansion has been described following viral infection in two instances: transplant patients undergoing active CMV infection or HIV infected patients (17, 18). A similar Vδ1 T cell expansion has also been observed in patients with lyme arthritis in response to Borrelia burgdorferi stimulation (43). To our knowledge, HHV-8 is the only virus other than HIV for which a γδ Vδ1 T cell expansion is observed and the first to be described in immunocompetent individuals. We observed a long-term persistence of this expansion suggesting sustained γδ T cell activation.

One common peculiarity of the viral infections leading to Vδ1 T cell expansion is the involvement of mucosae (44). Our previous studies have shown that saliva is the major mucosal site of HHV-8 replication in humans, and our patients were selected as having salivary HHV-8 infection. Vδ1 cells are mainly located in intestinal epithelia where they represent 70–90% of the γδ T cells (45). Little is known about the replication of HHV-8 in the gastrointestinal tract and whether this is a source of Vδ1 (46). Our data also indicate that there appear to be an in vivo Ag-driven expansion of Vδ1 T cells during the course of HHV-8 infection. Consistent with this notion, we have demonstrated that Vδ1 T cell lines were specifically activated in the presence of HHV-8-infected cell lines.

Furthermore, our analysis of the γδ Vδ1 T cell repertoire from an infected individual provides the first evidence that the expanded peripheral γδ Vδ1 T cell population responsive to HHV-8 is reflective of a selective, clonally restricted response to viral Ags. Our observations support the model that the peripheral Vδ1 T cell repertoire is shaped by positive selection in response to an antigenic stimulation (47, 48).

γδ T cell specificity against HHV-8 was further demonstrated by their specific recognition of purified viral proteins. Only few examples of Ag-specific γδ T cells have been reported. The previously described Ag specificities identified included nonclassical class I MHC molecules, MHC-like molecules (CD1c, MICA, MICB), heat shock proteins, monoalkyl phosphate (49), and bacterial superantigens (38). A specific reactivity of γδ Vδ1 T cells to primary leukemia blasts has also been reported (50). Specific recognition of viral proteins by γδ T cells is infrequent; it is of interest that the other described virus was another herpesvirus (HSV-1 glycoprotein gI) (51).

Furthermore, we demonstrated that Vδ1 T cells are able to prevent in vitro the release of infectious particles from HHV-8-infected cells, suggesting a protective antiviral role. Previous studies have shown that IFN-γ treatment of HHV-8-infected cell lines reduced the amount of infectious virus that resulted when HHV-8 was induced into the lytic cascade using 12-O-tetradecanoylphorbol-13-acetate (40, 52, 53). This is in agreement with our data demonstrating a decrease in infectious viral particle release due to IFN-γ production by HHV-8 stimulated Vδ1 T cells.

In summary, our studies indicate a specific expansion of γδ Vδ1 T cells in the peripheral blood of asymptomatic HHV-8-infected individuals. Our results support the role of Vδ1 T cells specific for viral proteins in control of HHV-8 chronic infection. This situation represents a unique opportunity to better understand the γδ T cell physiology. Studies to determine whether alteration in γδ T cell formation is a factor in HHV-8-induced Kaposi’s sarcoma lesions in immunocompromised hosts are warranted.

We thank Jie Wang, Steve Kuntz, and Anna Wald (University of Washington, Seattle, WA) for specimen collection, Alan Fox for technical assistance, Bala Chandran (Rosalind Franklin University of Medicine and Science, Chicago, IL) for providing the different recombinant baculovirus, Ronald Desrosiers (New England Regional Primate Research Center, Southborough, MA) for kindly providing HVS, Michael Lagunoff (University of Washington, Seattle, WA) for providing the Bjab cell line, Veronika Groh (Fred Hutchinson Cancer Research Center, Seattle WA) for providing the C1R cell line, and Nancy LaCroix (Fred Hutchinson Cancer Research Center, Seattle, WA) for help in preparing this manuscript.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by National Institutes of Health Grants AI30731, R37 AI42528, RO1 DE016809, RO1 DE14149, and K23 AI054162-04. This research was also supported by the University of Washington Center for AIDS Research, a National Institutes of Health-funded program (P30 AI 27757).

3

Abbreviations used in this paper: HHV-8, human herpesvirus 8; HVS, Herpesvirus saimiri; KSHV, Kaposi’s sarcoma associated herpesvirus; ORF, open reading frame; PEL, primary effusion lymphoma; rh, recombinant human.

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