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Anti-viral T cells are thought to regulate whether hepatitis C virus (HCV) and HIV infections result in viral control, asymptomatic persistence, or severe disease, though the reasons for these different outcomes remain unclear. Recent genetic evidence, however, has indicated a correlation between certain natural killer (NK) cell receptors and progression of both HIV and HCV infection1–3, implying that NK cells are playing a role in these T cell-associated diseases. While direct NK cell-mediated lysis of virus-infected cells may contribute to anti-viral defense during some virus infections, especially murine cytomegalovirus (MCMV) infections in mice and perhaps HIV in humans4–5, NK cells have also been suspected as having immunoregulatory functions. For instance, NK cells may indirectly regulate T cell responses by lysing MCMV-infected antigen-presenting cells6–7. In contrast to MCMV, lymphocytic choromeningitis virus (LCMV) infection in mice seems resistant to any direct anti-viral effects of NK cells5,8. Here the roles of NK cells in regulating T cell-dependent viral persistence and immunopathology were examined in mice infected with LCMV, an established model for HIV and HCV infections in humans. We describe a three-way interaction, whereby activated NK cells cytolytically eliminate activated CD4 T cells that affect CD8 T-cell function and exhaustion. At high virus dose NK cells prevented fatal pathology while enabling T-cell exhaustion and viral persistence, but at a medium dose NK cells paradoxically facilitated lethal T cell-mediated pathology. Thus, NK cells can act as rheostats, regulating CD4 T cell-mediated support for the anti-viral CD8 T cells that control viral pathogenesis and persistence.
Intravenous (i.v.) inoculation of C57BL/6 mice with a low (5×104 PFU), medium (2×105 PFU), or high (2×106 PFU) dose of LCMV, strain Clone 13, resulted in different degrees of pathology, as indicated by weight loss (Fig. 1a) and by histological analysis of lung sections at day 15 p.i. (Fig. 1b). The high dose caused a precipitous drop in body weight during the first week of infection (Fig. 1a, right), but, thereafter, clonal exhaustion and deletion of LCMV-specific T cells resulted in a persistent infection9–10 associated with minimal lung pathology (Fig. 1b, Right) and 100% (77 of 77) survival (Fig. 1c, Top). Selective depletion of NK cells using 25 µg of anti-NK1.1 mAb (Supplemental Fig. 1) one day prior to high dose infection resulted in 58% (35 of 65) mortality between days 9 and 13 of infection (Fig. 1c, Top) associated with severe pulmonary edema (data not shown) and reduced viral titers by day 7 p.i. (Fig. 1d, Right). Under these high dose conditions, therefore, the presence of NK cells promoted persistence and prevented mortality.
In contrast to the beneficial role of NK cells during high dose infection, NK cell-depletion prevented the severe weight loss (Fig. 1a, Middle) and tissue pathology (Fig. 1b, Middle) associated with the medium dose of LCMV. Twenty-three percent (7 of 31) of control-treated mice succumbed to the medium dose during the second week of infection, and the lungs of surviving mice exhibited bronchus associated lymphoid tissue, pulmonary edema, and interstitial mononuclear infiltration. Lung pathology was absent in NK cell-depleted mice, which uniformly survived medium dose challenge (Fig. 1c, Bottom). Moreover, while high levels of replicating virus persisted in surviving control mice at day 15 p.i., NK cell-depletion resulted in complete viral clearance (Fig. 1d, Middle). In this case the presence of NK cells was detrimental for the host, as they promoted immune pathology and death.
Irrespective of the presence of NK cells, inoculation with a low dose of virus was uniformly non-lethal in 18 of 18 (100%) control and 18 of 18 (100%) of NK cell-depleted mice by >50 days p.i., with minimal weight loss (Fig. 1a, Left) and minimal lung pathology (Fig. 1b, Left). Virus was completely cleared in both groups of mice by day 15 of low dose infection (data not shown), but NK cell depletion resulted in more rapid elimination of LCMV in liver by day 7 p.i. (Fig. 1d, Left).
The weight loss, lung pathology, and mortality observed in medium dose-infected wild-type mice (Fig. 1a,b) did not occur after infection of αβ T cell-deficient (TCRβ−/−) mice, and NK cell depletion of TCRβ−/− mice did not alter weight loss or viral burden (Supplemental Fig. 2). Thus, NK cells regulate viral clearance and immunopathology during LCMV infection through a T cell-dependent mechanism.
As early as day 6 after medium dose infection, the proportion and number of IFN-γ+ LCMV-specific CD8 T cells was increased 2- to 6-fold in mice depleted of NK cells (Fig. 2a and Supplemental Fig. 3), and anti-viral T cells from these mice displayed an enhanced ability to co-produce TNF (Supplemental Fig. 3). The number of NP396–404 tetramer-binding CD8 T cells in the spleen on day 5 p.i. was increased 4- to 20-fold in NK cell-depleted mice relative to non-depleted control mice after infection with all doses of virus (Fig. 2b). The number of virus-specific IFN-γ+ CD4 T cells was also amplified 7-to 20-fold by NK cell depletion compared to control mice on different days after medium dose infection (Fig. 2c). Moreover, co-production of TNF and IL-2 by anti-viral CD4 T cells was augmented by NK cell depletion (Fig. 2d and Supplemental Fig. 3). The increased magnitude of the LCMV-specific T cell response in the absence of NK cells during medium dose infection correlated with rapid viral clearance (Fig. 2e). Depletion of NK cells using a carefully titrated dose of anti-asialo GM1 antibody, which eliminates NK cells but not CD8 T cells11, also enhanced anti-viral CD4 and CD8 T cell responses during medium dose infection (Supplemental Fig. 4).
The enhanced anti-viral T cell responses suggested that NK cell depletion may augment proliferation of LCMV-specific T cells. Transfer of CFSE-labeled Thy1.1+ T cells revealed a larger population of CFSElow donor CD4 (Fig. 2f) and CD8 (data not shown) T cells in multiple host tissues at day 6 p.i. of high dose infection in the absence of NK cells. There also was greater specific lysis of viral peptide-coated target cells as detected by a conventional in vivo cytotoxicity assay at day 4 of infection (Supplemental Fig. 5). Moreover, LCMV-specific Ly5.1+ TCR transgenic (P14) CD8 T cells (transfer 104) were recovered from tissues of NK cell-depleted recipient (Ly5.2+) mice at 2- to 9-fold greater numbers than control-treated mice 6 days after low dose infection (Supplemental Fig. 5). Together these results indicate that NK1.1+ cells repress the size of the anti-viral T cell response during LCMV infection.
The activities of CD4 T cells are important for maintaining CD8 T cell function during LCMV infection12–14. To assess whether CD4 T cells were involved in the NK cell suppression of LCMV-specific CD8 T cells, mice were treated with antibodies to concurrently deplete both NK and CD4 T cells. Whereas depletion of NK cells prior to medium dose LCMV infection resulted in a >200-fold reduction in splenic viral titers at day 7 p.i. relative to control and CD4-depleted (ΔCD4) mice (Fig. 3a), depletion of both NK and CD4 T cells (ΔNKΔCD4) had no effect on viral titers. The increased numbers and enhanced multiple cytokine production by anti-viral CD8 T cells caused by NK cell depletion were also prevented by co-depletion of CD4 T cells (Fig. 3b). In contrast, co-depletion of NK and CD8 T cells did not prevent an increase in IFN-γ+ GP61–80-specific CD4 T cells (Control: 3.8±0.5 % vs. ΔNK: 9.8±0.7 % vs ΔNK/ΔCD8: 10.7±1.6 %, n=3, p<0.05 vs control) at day 12 of medium dose infection.
Paradoxically, at the high virus dose, co-depletion of NK and CD4 T cells prevented the severe pulmonary edema (Fig. 3c) and heightened mortality (Fig. 3d) associated with depletion of NK cells alone. In this experiment, mice were harvested on day 12 p.i., when 3 surviving NK cell-depleted mice were moribund and required euthanasia, while all double-depleted mice displayed relatively normal vigor. The livers of NK/CD4 double-depleted mice contained 25-fold more PFU than livers from mice depleted of NK cells alone (NK: 5.7±0.2 PFU vs ΔNK/ΔCD4: 7.1±0.1 PFU, n=5, p<0.0001). Enhancement of LCMV-specific CD8 T cells in the absence of NK cells was also abrogated by concurrent depletion of CD4 T cells (Supplemental Fig. 6). Together these data indicate that CD4 T cells are needed for NK cell modulation of antiviral CD8 T cell responses associated with viral clearance, persistence, and immunopathology.
We utilized a modified in vivo cytotoxicity assay by injecting splenocytes from medium dose-infected NK cell-depleted mice (Ly5.1+, day 4 p.i.) into medium dose-infected NK cell-depleted (ΔNK) or isotype IgG2a-treated (Control) recipient mice (Ly5.2+, day 3 p.i.). After 5 hours, similar proportions of total donor T (Control: 0.16±0.03 % vs. ΔNK: 0.15±0.02 %, n=21, p=0.80) and B cells (Control: 1.8±0.2 % vs. ΔNK: 1.7±0.2 %, n=21, p=0.88) were recovered from infected recipients, regardless of NK cell depletion. Likewise, recovery of activated (CD44hi CD43(1B11)+) donor CD8 T cells was similar from spleens of Control and ΔNK mice, with minimal loss relative to uninfected control mice (Fig. 4a). In contrast, there was a substantial loss of activated donor CD4 T cells in infected relative to uninfected recipients, and this loss was prevented by depletion of NK cells (Fig. 4a). The magnitude of NK cell-dependent loss of activated donor CD4 T cells was similar in low-, medium-, and high dose-infected recipients (Fig. 4b). More activated CD4 T cells, both donor- and host-derived, in infected (Control) mice stained positively for the apoptosis indicator, AnnexinV, in comparison to naive donor CD4 T cells or to activated donor CD4 T cells in medium dose-infected ΔNK recipient mice (Fig. 4c). In contrast to activated donor CD4 T cells, the recoveries of naïve (CD44low) phenotype CD4 and CD8 donor T cells were not altered by NK cell-depletion (data not shown). These data indicate that NK cells in WT mice selectively and rapidly target activated CD4 T cells for elimination during LCMV infection.
We next examined the involvement of NK cell cytolytic mediators FasL, TNF, and perforin (Prf1) in this process. The loss of activated WT donor CD4 T cells in infected WT recipient mice (Fig. 4a,d) was seen when activated lpr (Fas mutant) mouse donor cells were transferred into WT recipient mice or when WT donor cells were transferred into TNF−/− recipient mice (Fig. 4d). In contrast, there was relatively little loss of activated WT donor CD4 T cells in Prf1−/− hosts (Fig. 4d), whose retention of activated donor CD4 cells was not significantly different (p > 0.1) from that in NK cell-depleted WT or Prf1−/− hosts. Thus, NK cell elimination of activated CD4 T cells is mediated through a perforin-dependent pathway that does not require Fas or TNF.
Previous work has implicated NKG2D in targeting of activated T cells by murine NK cells in vitro15–17, but we observed no differences in activated WT donor CD4 T cell survival in WT vs. NKG2D−/− recipients (Fig. 4d) or in WT mice treated with a blocking mAb to NKG2D18 (data not shown). Of note is that we did not observe the expression of ligands for activating NK cell receptors including NKG2D, NKp46, DNAM-1, and TRAIL on these early activated CD4 T cells, and NK cell-mediated elimination of activated donor CD4 T cells also occurred in antibody-deficient (µMT−/−) mice (data not shown), precluding a role for antibody-dependent mechanisms. Activated CD4 T cells did, however, express much higher levels of adhesion molecules than naïve cells, and these molecules have previously been shown to trigger NK cell cytotoxicity via LFA-119–20. Somewhat surprising was the observation that the activated CD4 T cells were far more susceptible than activated CD8 T cells to direct killing by the NK cells, even though both expressed high levels of adhesion molecules. We previously had shown that the presence of the negatively signaling receptor CD244 (2B4) on NK cells prevented NK cell-mediated lysis of activated CD8 T cells21. We found here that while expression of the CD244 ligand, CD48, was up-regulated on T cells after medium dose LCMV infection, expression levels of CD48 were much higher on activated CD8 than on activated CD4 cells (MFI of CD48: activated CD4, 3,423±147; activated CD8, 6,180±166; n=9, p<0.0001) (Supplemental Fig. 7).
In order to assess whether NK cell-mediated lysis of activated CD4 T cells is a general principle of virus infections, we examined the loss of LCMV-activated CD4 T cells following transfer into mice inoculated with an unrelated Arenavirus, Pichinde virus (PV), the Coronavirus mouse hepatitis virus (MHV), or the interferon inducer and NK cell activator poly I:C (pI:C). All three stimuli induced measureable loss of activated donor CD4 T cells that was dependent upon the presence of NK cells (Fig. 4e). In reciprocal experiments, CD4 T cells activated during infection with PV, MHV, vaccinia virus (VV), and MCMV were lost upon transfer into mice infected with medium dose LCMV when NK cells were present (Fig. 4f).
An analysis of the window of time at which NK cell regulation of T cells occurred in the LCMV medium dose model showed reduced frequencies of activated donor CD4 T cells relative to uninfected recipients by in vivo cytotoxicity assays after transfer into NK cell-sufficient mice one (43%), two (32%), three (26%), four (4%), and five (8%) days after infection (Supplemental Fig. 8). The frequencies of activated donor CD4 T cells were increased by NK cell depletion in recipient mice only at day 2 and day 3 p.i. These results suggest that NK cells target activated CD4 T cells mainly on the second and third day of infection, when the cytolytic activity of NK cells is at its peak22.
These results show that NK cells can play a crucial role in controlling virus-associated morbidity, mortality, and persistence in the absence of direct NK cell-mediated control of virus replication, and they do so by altering the numbers and polyfunctionality of virus-specific T cells. Their effect on activated CD4 T cells was presumably due to direct cytotoxicity, as demonstrated by rapid perforin-dependent elimination of activated CD4 T cells by NK cells in short term in vivo cytotoxicity assays and the observation of enhanced annexin reactivity of CD4 T cells in the presence of NK cells. The effect of activated NK cells on CD8 T cells and virus clearance appeared to be indirect, and depended on the presence of CD4 T cells, which are known to produce factors that preserve CD8 T-cell viability and functionality12–13,23–25. This suggests a three-way interaction, whereby NK cells suppress the CD4 T cell response, thereby preventing augmentation of the CD8 T cell response, which, in turn, directly regulates viral clearance and immunopathology in this system (Fig. 4f).
Our observation of direct NK cell-mediated lysis of T cells during virus infection is distinct from published accounts of NK cell regulation of anti-viral T cells during MCMV infection, in which NK cell-mediated lysis of virus-infected cells contributes to control of viral burden and persistence of MCMV-infected dendritic cells (DC) that in turn regulate activity of anti-viral T cells6–7,26. Moreover, we found no NK cell-dependent changes in the number and antigen-presenting function of splenic DCs during LCMV infection (Supplemental Fig. 9), consistent with our finding that NK cells directly regulated the T cells. A possible concern was that in vivo cellular depletion with antibodies against NK1.1 and CD4 have the potential to affect frequencies of NKT, γδ T, and regulatory T cells, but the frequencies of these lymphocytes in the spleen of LCMV-infected mice were not altered after 3 to 4 days of infection by the low concentration of anti-NK1.1 used in these studies (Supplemental Figs. 1 & 10). Moreover, depletion of NK cells in γδ T cell-deficient (TCRδ−/−) and NK T cell-deficient (CD1d−/−) mice during medium dose infection enhanced LCMV-specific T cell responses and reduced viral loads (Supplemental Fig. 9), similar to that in WT mice. Therefore, these lymphocyte lineages appear dispensable for NK cell immunoregulatory function during LCMV infection.
By adjusting the dose of the LCMV inocula, one can generate diverse patterns of CD8 T cell-regulated pathogenesis similar to the variety of pathogenic patterns a human HCV infection can take, including rapid viral clearance, severe T cell-dependent immunopathology, and long-term persistence. We show here that at a high dose of LCMV NK cells act beneficially by suppressing T cell responses, thereby preventing severe pathology and mortality while enabling the development of a persistent infection from which mice eventually recover and clear virus27. At the medium dose inoculum, NK cell suppression of T cells is detrimental to the host, as virus clearance is impaired due to the limited number and functionality of T cells. However, a medium dose of virus is not sufficient for complete clonal exhaustion of T cells, ultimately resulting in severe T cell-dependent immunopathology that can lead to death of the host.
These results suggest that NK cells can serve as rheostats or master regulators of anti-viral T cell responses. Consistent with the fact that many virus infections induce cytokines that potently activate NK cells28, we found that NK cell lysis of activated CD4 T cells was triggered by several viruses as well as after inoculation with pI:C, which induces interferon and activates NK cells. Although a previous study found that NK cell depletion did not alter the magnitude of antiviral T cell responses during infection with the Armstrong strain of LCMV29, we have observed enhanced anti-viral T cell responses and improved viral control at early time points after infection of NK cell-depleted mice with both LCMV Armstrong and Pichinde virus (SNW, unpublished observations). Thus, the timing and the type of evaluation may be important to detect detrimental effects of NK cells on T cells during more benign viral infections.
One day before infection, male C57BL/6 were selectively depleted of NK cells through a single i.p. injection of 25 µg anti-NK1.1 mAb (PK136) or a control rat IgG2a (both from Bio-X-Cell), as previously described21 (Supplementary Fig. 1). In some cases, mice were also depleted of CD4 T cells by i.p. injection of 100 µg anti-CD4 (GK1.5) at days −1 and +3 of infection. Mice were then infected i.v. with 5×104 (low dose), 2×105 (medium dose), or 2×106 (high dose) PFU of the clone 13 variant of LCMV. Virus was titrated by plaque assay on Vero cells. In some experiments, mice were inoculated i.p. with 1.5×107 PFU of PV, 8×105 PFU of MHV strain A59, 1×106 PFU of VV strain Western Reserve, 1×106 PFU of Smith strain MCMV, or 200 µg of pI:C (Invivogen).
The number of LCMV-specific T cells was measured by H-2Db-NP396–404 tetramer staining or by intracellular cytokine staining after 5 hour ex vivo stimulation with 1 µM viral peptide in the presence of brefeldin A. T cell cytolytic activity was measured in vivo as described previously21.
An unconventional in vivo cytotoxicity assay was previously established to determine NK cell killing of lymphocyte populations 21. Donor mice were depleted of NK cells and then infected i.v. or i.p. with different viruses. At day 4 p.i., single cell splenocyte suspensions were prepared from these mice, labeled with 2 µM CFSE, and transferred (2×107) into various strains of NK cell-depleted or control recipient mice which were either uninfected or had been infected with virus 1 to 5 days previously. Spleens of recipient mice were harvested 5 hours after transfer and assessed for survival of donor T cells.
C57BL/6, Thy1.1+, TCRδ−/−, TCRβ−/−, lpr, Prf1−/−, and µMT−/− mice were purchased from The Jackson Laboratories. Ly5.1+ mice were from Taconic Farms. NKG2D−/− and CD1d−/− mice on a C57BL/6 background were obtained from B. Polić30 and M. Exley31, respectively. Congenic (Ly5.1+) TCR transgenic P1432 mice and TNF−/− mice on a C57BL/6 background were bred at UMMS. Male mice at 6–16 weeks of age were routinely used in experiments. Mice were maintained under specific pathogen-free conditions, and experiments were performed in compliance with institutional guidelines as approved by the Institutional Animal Care and Use Committee of UMMS.
The clone 13 variant of LCMV was propagated in baby hamster kidney BHK21 cells9 and titrated by plaque assay on Vero cells. Mice were infected i.v. with 5×104 (low dose), 2×105 (medium dose), or 2×106 (high dose) PFU of LCMV. Selective depletion of NK cells was achieved through a single i.p. injection of 25 µg anti-NK1.1 mAb (PK136) or a control rat IgG2a produced by Bio-X-Cell, as previously described21 (Supplementary Fig. 1). Alternatively, mice received a carefully titrated dose of 10 µL of anti-asialo GM1 antibody (Wako Biochemicals) diluted in 200 µL PBS i.p. one day prior to virus infection. Anti-NKG2D mAb (CX5) was a gift of L. Lanier, and 200 µg was injected i.p. at the time of infection. Mice were depleted of T cells by i.p. injection of either 100 µg anti-CD4 (GK1.5) or 50 µg anti-CD8 (2.43) produced by Bio-X-Cell at day −1 and day +3 of infection. In some experiments, mice were inoculated i.p. with 1.5×107 PFU of PV, 8×105 PFU of MHV strain A59, 1×106 PFU of VV strain Western Reserve, 1×106 PFU of Smith strain MCMV, or 200 µg of polyI:C (Invivogen).
T-cell epitopes encoded by LCMV include NP396–404 (FQPQNGQFI), GP33–41 (KAVYNFATC), and GP61–80 (GLKGPDIYKGVYQFKSVEFD) 33–35. Peptides were purchased from 21st Century Biochemicals and purified by reverse phase-HPLC to 90% purity. H-2Db-NP396–404 tetramers were prepared as described36. CD1d-PBS57-allophycocyanin tetramers provided by NIAID Tetramer Facility were a generous gift of L. Berg.
Fluorescently labeled antibodies and reagents were purchased from BD Biosciences, eBioscience, BioLegend, and R&D Biosystems. Flow cytometric analyses of cells were performed on a LSR II cytometer (BD Biosciences) equipped with FACSDiva software and data were analyzed using FlowJo software (Tree Star).
Spleens from donor mice were mechanically disrupted, and erythrocytes were lysed using a 0.84% NH4Cl solution in order to generate single-cell leukocyte suspensions. Cells were labeled for 15 min at 37°C with the 2 µM fluorescent dye carboxyfluorescein diacetate, succinimidyl ester (CFSE) (CFDA-SE, Molecular probes), washed, and transferred i.v. (3×107 cell) to recipient mice.
T cell cytolytic activity was measured in vivo as described previously21. Briefly, single-cell suspensions were prepared from spleens of uninfected mice, and separate fractions of cells were then loaded with LCMV peptides (1 µM) for 45 minutes at 37°C before labeling with CFSE (2.5, 1, or 0.4 µM, Molecular Probes) for 15 minutes at 37°C. After washing, these populations were combined at equal ratios and transferred i.v. into either naïve or infected recipients. Survival of each transferred population in the spleens of recipient mice was assessed 16 hours after transfer. Specific lysis was calculated as follows: 100-([% LCMV target population in infected experimental/ % unlabeled population in infected experimental) ÷ (% LCMV target population in naïve control / % unlabeled population in naïve control)] × 100).
An unconventional in vivo cytotoxicity assay was previously established to determine NK cell killing of lymphocyte populations in vivo21. WT or lpr donor mice were depleted of NK cells and then infected with VV, MCMV, MHV, PV, or a medium dose of LCMV clone 13. At day 4 p.i., single cell splenocyte suspensions were prepared from these mice, labeled with CFSE, and then transferred (2×107) into experimental recipient mice on day 3 of medium dose LCMV infection, unless otherwise noted. Recipients included WT, Prf1−/−, TNF−/−, or NKG2D−/− mice that were administered anti-NK1.1 or isotype control antibodies one day prior to inoculation with PV, MHC, polyI:C, or various doses of LCMV clone 13. Some recipient mice were uninfected and served as controls. Spleens of recipient mice were harvested 5 hours after transfer and assessed for survival of donor T cells.
Stimulator cells were prepared by isolation of single-cell suspensions from the spleens of uninfected as well as isotype-treated or anti-NK1.1-treated mice infected 3 days previously with a medium dose of LCMV i.v. Following irradiation, stimulator cells (5×104) were plated at a 1:10 ratio with CFSE-labeled Ly5.1+ LCMV-specific P14 CD8 T cells (5×105) in T cell stimulation medium (RPMI supplemented with 100 U/ml penicillin G, 100 µg/ml streptomycin sulfate, 2 mM l-glutamine, 10 mM HEPES, 1 mM sodium pyruvate, 0.1 mM non-essential amino acids, 0.05 mM 2-mercaptoethanol, and 10% heat-inactivated (56°C, 30 min) FBS), which was refreshed every two days. P14 cells were enumerated and analyzed for dilution of CFSE as a measure of proliferation every 24 hours after initiation of co-culture.
Single-cell leukocyte suspensions from spleens, inguinal lymph nodes, lung and liver were prepared as described previously21 and were plated at 2×106 cells per well in 96 well plates. Cells were stimulated for 5 hours at 37°C with either 1 µM viral peptide or 2.5 µg/mL anti-CD3 mAb in the presence of brefeldin A and 0.2 U/mL rhIL-2. Stimulated cells were then pre-incubated with a 1:200 dilution of Fc Block (2.4G2) in FACS buffer (HBBS, 2% FCS, 0.1% NaN3) and stained for 20 min at 4°C with various combinations of fluorescently tagged mAbs. After washing, cells were permeabilized using BD Cytofix/Cytoperm solution and then stained in BD Permwash using mAbs specific for various cytokines. AnnexinV staining was performed in azide-free FACS buffer directly ex vivo according to manufacture’s instructions (BD Biosciences).
Results are routinely displayed as mean ± s.e.m, with statistical differences between experimental groups determined using a two-tailed unpaired Student’s t-test, where a p value of < 0.05 was deemed significant. Statistical differences in survival were determined by log rank (Mantel-Cox) analysis. Graphs were produced and statistical analyses were performed using GraphPad Prism.
We thank K. Hearn, C. Baer, J. Suschak and P. Afriyie for technical support; K. Daniels and M. Seedhom for insightful discussions; R. Taniguchi and V. Kumar for sharing unpublished observation; and H. Ducharme for mouse husbandry. We thank L. Berg for NKT tetramer, L. Lanier for anti-NKG2D blocking antibody (CX5), B. Polić for NKG2D−/− mice, and M. Exley for CD1d−/− mice. This work was supported by National Institutes of Health (NIH) training grant AI07349 (S.N.W.) and research grants AI-17672, AR-35506, CA34461 (R.M.W.), AI46578 (L.K.S.), a German Research Foundation fellowship CO310-2/1 (M.C.), and an institutional Diabetes Endocrinology Research Center (DERC) grant DK52530. The views expressed are those of the authors and do not necessarily express the views of the NIH.
Author contributions S.N.W. designed the study, performed experiments, analyzed data, and wrote the manuscript; R.M.W. designed the study, analyzed data, and wrote the manuscript; M.C. and L.K.S. were involved in study design, discussed results, and commented on the manuscript.
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The authors declare that no competing financial interests exist.