. 2012 Sep;86(17):9025–9034. doi: 10.1128/JVI.00942-12

FOXO3 Regulates the CD8 T Cell Response to a Chronic Viral Infection

Jeremy A Sullivan ¹, Eui Ho Kim ¹, Erin H Plisch ¹, M Suresh ^1,^✉

PMCID: PMC3416126 PMID: 22675000

Abstract

Chronic infections with viruses such as hepatitis B virus, hepatitis C virus, and HIV constitute a major global public health problem. Studies of chronic viral infections in humans and mice show that persistent antigenic stimulation induces dysregulation of T cell responses; virus-specific T cells either undergo clonal deletion or lose their ability to display the full spectrum of effector functions, a condition termed functional exhaustion. The ability to generate and retain sufficient numbers of functionally competent T cells, therefore, becomes vitally important in controlling chronic viral infections. Our understanding of the mechanisms governing T cell homeostasis during chronic viral infections, however, is poor. The phosphatidylinositol 3-kinase (PI3K)/Akt signaling pathway controls cell fate decisions in many cell types by modulating the activity of downstream effectors, including the FOXO family of transcription factors. We have observed dynamic, in vivo alterations in the phosphorylation levels of three key proteins (Akt, FOXO1/FOXO3 [FOXO1/3], and mammalian target of rapamycin [mTOR]) involved in this signaling cascade and have identified the transcription factor FOXO3 as a negative regulator of the magnitude and effector function of CD8 T cells during chronic lymphocytic choriomeningitis virus (LCMV) infection in mice. We report that ablation of FOXO3 in T cells reduced apoptosis, increased the abundance of polyfunctional virus-specific CD8 T cells, and improved viral control. Thus, FOXO3 is a promising candidate target for immunotherapies of chronic viral infection.

INTRODUCTION

More than 500 million people in the world are currently afflicted with chronic infections with viruses such as hepatitis B virus, hepatitis C virus, and HIV (24, 37, 41). A common denominator underlying viral persistence in these chronic viral infections (CVIs) is the dysregulation of virus-specific T cell responses; virus-specific T cells either undergo clonal deletion or lose their ability to express the full spectrum of effector functions, a condition termed functional exhaustion (8, 22, 29, 33, 41, 53, 56, 59). During CVIs, there is a continuum of T cell proliferation and apoptosis, and the balance between these cellular processes controls the abundance of virus-specific CD8 T cells (18, 48, 49, 53, 60). However, the homeostatic mechanisms that control the number of virus-specific T cells under conditions of protracted antigenic stimulation are poorly defined. This is an important issue, because the magnitude of the T cell response is a critical factor in determining viral control.

The FOXO family (FOXO1, FOXO3) of transcription factors plays a crucial role in regulating multiple facets of T cell homeostasis, including apoptosis, proliferation, trafficking, and differentiation (16, 20, 21, 34, 39, 47). In particular, FOXO3 is known to promote the apoptosis of T cells and to limit the clonal expansion of CD8 T cells during an acute viral infection (14, 47). Additionally, FOXO3 has been implicated in vitro as a target for modulating T cell dysregulation in human T cells during HIV infection (52). The principal objective of this study was to gain insight into the role of FOXO3 in orchestrating the dynamics of CD8 T cell responses during a chronic lymphocytic choriomeningitis virus (LCMV) (clone 13) infection in mice.

MATERIALS AND METHODS

Mice and viral infection.

The derivation of mice carrying floxed FOXO3 alleles has been described elsewhere (36, 50). Mice carrying floxed FOXO3 alleles were bred with CD4-Cre mice at the University of Wisconsin—Madison (UW-Madison) to generate T-cell-specific FOXO3-deficient (FOXO3L) mice. Wild-type (WT) littermates were used as controls with the FOXO3L mice. Mice used in experiments were between the ages of 6 and 8 weeks, and all experiments were performed in accordance with the protocols approved by the University of Wisconsin School of Veterinary Medicine Institutional Animal Care and Use Committee (IACUC). Mice were infected with 2 × 10⁶ PFU of the LCMV clone 13 strain by intravenous (i.v.) injection. Tissue viral titers were quantified by plaque assays with Vero cell monolayers (2).

General flow cytometry.

Single-cell suspensions of splenocytes were stained with antibodies for surface markers, including CD8, CD44, CD122, CD127, CD62L, LFA-1, and KLRG-1 (BD Biosciences, Franklin Lakes, NJ; eBioscience, San Diego, CA; or Southern Biotech, Birmingham, AL) in conjunction with major histocompatibility complex class I (MHC-I) tetramers (D^b) specific for the class I-restricted LCMV epitopes, NP396, GP33, and GP276 (30). Cells were fixed in 2% paraformaldehyde (PFA) and were acquired in a FACSCalibur or LSR II flow cytometer (BD Biosciences, Franklin Lakes NJ). To quantify intracellular cytokine production, splenocytes were incubated for 5 h at 37°C with LCMV epitope peptides in the presence of brefeldin A. After stimulation, cells were first incubated with antibodies for surface markers. Next, cells were permeabilized and stained for intracellular cytokines (gamma interferon [IFN-γ], interleukin 2 [IL-2], and tumor necrosis factor alpha [TNF-α]) using the Cytofix/Cytoperm kit (BD Biosciences, Franklin Lakes, NJ). The percentages of cytokine-producing cells were quantified by flow cytometry.

Phosphospecific flow cytometry.

Splenocytes were stained for cell surface markers as described above. After cell surface staining, cells were fixed and permeabilized using Phosflow lysis and Phosflow PermWash I reagents (BD Biosciences, Franklin Lakes, NJ) according to the manufacturer's recommendations. Next, cells were blocked for 30 min on ice in blocking buffer (10% normal goat serum in 2% bovine serum albumin [BSA]–phosphate-buffered saline [PBS]) and were subsequently stained with phosphospecific antibodies (against P-Akt [T308], P-FOXO1/3 [T24, T32], or P-mTOR [S2448]; Cell Signaling Technology, Danvers, MA). As negative controls for staining, antibodies were blocked by preincubation with their specific antigenic peptides for 1 h at room temperature before being added to the cells. Following incubation with the antibody or the peptide-blocked antibody, cells were washed twice and were incubated with a secondary antibody (Alexa 488-conjugated goat anti-rabbit antibody; Sigma-Aldrich, St. Louis, MO) for 40 min. Cells were washed and fixed with 2% PFA. The levels of phosphospecific staining were quantified by flow cytometry. Specific levels of staining (expressed as the corrected mean fluorescence intensity [MFI]) were calculated as the difference between the observed MFI for the phosphospecific protein and the observed MFI for the peptide-blocked control, divided by the observed MFI for the peptide-blocked control.

Ki67 and granzyme B staining.

Splenocytes were stained for surface markers and MHC-I tetramers as described above. After surface staining, cells were fixed and permeabilized using fluorescence-activated cell sorting (FACS) lysing solution and FACS permeabilization solution 2 reagents (BD Biosciences, Franklin Lakes, NJ) and were subsequently incubated with antibodies against Ki67 or granzyme B (BD Biosciences, Franklin Lakes, NJ) for 45 min at room temperature. Virus-specific CD8 T cells staining positive for Ki67 or granzyme B were visualized using a FACSCalibur flow cytometer. Data are expressed either as a percentage of antigen-specific CD8 T cells positive for the respective protein or as the MFI for the indicated protein.

Assessment of apoptosis ex vivo.

Splenocytes from WT and FOXO3L mice were isolated and stained with anti-CD8 and MHC-I tetramers on the indicated days postinfection (p.i.) as described above, except that no red blood cell lysis was performed. Annexin V staining (BD Biosciences, Franklin Lakes, NJ) was then carried out according to the manufacturer's protocol, except that all staining was performed on ice. The percentage of cells high in annexin V among antigen-specific CD8 T cells was determined by flow cytometry.

RESULTS

In vivo phosphorylation kinetics of Akt, FOXO1/3, and mTOR during chronic LCMV infection.

FOXO3, in its active hypophosphorylated state, localizes to the nucleus and drives the transcription of genes that oppose cell cycle entry (e.g., p27^Kip1) or promote cellular apoptosis (e.g., BIM) (15, 20, 23, 39, 46, 47). Since FOXO3 is part of a phosphatidylinositol 3-kinase (PI3K)-Akt signaling cascade, its hyperphosphorylation by kinases, including Akt, induces the relocalization of FOXO3 to the cytoplasm, thereby preventing the transcription of its target genes (7, 11, 27, 40). In T cells, signaling via the T cell receptor (TCR), CD28, and IL-2/7/15 receptors activates the PI3K/Akt pathway, leading to the phosphorylation of FOXOs, but the dynamics of Akt/FOXO phosphorylation in antigen-specific CD8 T cells in vivo during CVIs remains unknown (5, 9, 10, 13, 19, 42, 43, 54). For mice infected with LCMV clone 13, we used flow cytometry to define the phosphorylation kinetics of PI3K-associated signaling proteins, including Akt (T308), FOXO1/FOXO3 (FOXO1/3) (T24/T32), and mammalian target of rapamycin (mTOR) (S2448), in LCMV-specific CD8 T cells directly ex vivo (Fig. 1). The phosphorylation levels of Akt, FOXO1/3, and mTOR were readily detected in LCMV-specific CD8 T cells and displayed dynamic alterations during the course of a chronic LCMV infection; high levels of phosphorylation were detected at day 8 p.i., but levels declined progressively thereafter, a finding congruent with the circulating viral loads in LCMV clone 13-infected mice.

Fig 1 — *In vivo* phosphorylation kinetics of Akt, FOXO1/3, and mTOR during chronic LCMV infection. C57BL/6 mice were infected with the clone 13 strain of LCMV, and at the indicated days p.i., splenocytes were stained with anti-CD8, an MHC-I tetramer (D^b/GP33), and antibodies against either P-Akt (T308), P-FOXO1/3 (T24/T32), or P-mTOR (S2448) (Cell Signaling Technologies). To account for nonspecific antibody binding, parallel staining was performed using the antibody preincubated with its respective antigenic peptide. Representative histograms gated on GP33-specific CD8 T cells from days 8, 30, and 120 postinfection are shown for each phosphoprotein stain. Numbers on the right represent the difference between the observed MFI of antibody staining and the observed MFI of the specific peptide-blocked control sample, divided by MFI of the peptide-blocked control, at each day p.i. Data are representative of at least 3 individual experiments per time point.

FOXO3 deletion leads to an increase in the frequency of LCMV-specific CD8 T cells during the early stages of a chronic LCMV infection.

To explore whether FOXO3 regulates CD8 T cell responses by T-cell-intrinsic mechanisms during the early stages of chronic LCMV infection, we infected FOXO3L mice and WT littermates with LCMV clone 13, and LCMV-specific CD8 T cells were quantified at the peak of the expansion phase (day 8 p.i.) (4) using MHC-I tetramers. The low frequencies of NP396-specific CD8 T cells in WT mice were reminiscent of clonal deletion following infection with LCMV clone 13 (Fig. 2A). The frequencies and total numbers of NP396-specific CD8 T cells in FOXO3L mice were similar to those in WT mice, suggesting that FOXO3 deficiency did not mitigate deletion of these cells during a chronic LCMV infection (Fig. 2A and B). However, FOXO3 deficiency caused marked enhancements in the numbers of GP33- and GP276-specific CD8 T cells (Fig. 2A). Due to the low frequency and poor tetramer binding of NP396-specific CD8 T cells, studies from here on were focused on GP33- and GP276-specific CD8 T cells. Next, to determine whether the augmented accumulation of LCMV-specific CD8 T cells in FOXO3L mice resulted from altered proliferation and/or apoptosis, we measured Ki67 expression and annexin V binding directly ex vivo, as described previously (47). The increased expansion of LCMV-specific CD8 T cells cannot be explained by enhanced proliferation, since the percentages of Ki67-positive cells in WT and FOXO3L mice were similar (Fig. 2B). Instead, the enhanced number of LCMV-specific CD8 T cells in FOXO3L mice was linked to a significant reduction in apoptosis (Fig. 2C). Taken together, the data in Fig. 2 suggested that FOXO3 limited the accumulation of CD8 T cells during a CVI by promoting apoptosis. However, FOXO3 deficiency did not affect the expression of activation markers/cytokine receptors, including CD44, CD62L, LFA-1, CD122, and CD27, on LCMV-specific CD8 T cells (Fig. 3A). The population of effector CD8 T cells elicited by acute viral infections consists of at least two subsets based on the differential expression of CD127 and KLRG-1: the short-lived effector cells (SLECs) (KLRG-1^HI CD127^LO) and memory precursor effector cells (MPECs) (KLRG-1^LO CD127^HI) (19, 44). We observed no difference in the percentages of these two effector subsets among LCMV-specific CD8 T cells between WT and FOXO3L mice (Fig. 3B), suggesting that FOXO3 deficiency did not modulate the differentiation of effector subsets following a CVI. During CVIs, elevated levels of PD-1 and LAG-3 have been linked to impaired CD8 T cell effector function (functional exhaustion) (6, 8, 32, 55), but we found no alteration in the levels of either of these proteins on effector CD8 T cells in FOXO3L mice (Fig. 3C).

Fig 2 — FOXO3 deletion leads to an increase in the frequency of LCMV-specific CD8 T cells during the early stages of a chronic LCMV infection. (A) WT and FOXO3L mice were infected with LCMV clone 13, and at 8 days p.i., LCMV-specific CD8 T cells in spleens were quantified by staining with anti-CD8, anti-CD44, and either a D^b/NP396, D^b/GP33 (P < 0.01), or D^b/GP276 (P < 0.04) MHC-I tetramer. Representative FACS plots on the left are gated on total CD8 T cells. The percentages of tetramer-binding CD8 T cells in WT and FOXO3L mice are given above the plots. Flow cytometry data are representative of at least 4 independent experiments with 3 to 7 mice/group/experiment. (B) At day 8 p.i., splenocytes from WT and FOXO3L mice were stained with anti-CD8, MHC-I tetramers, and anti-Ki67. The percentage of LCMV-specific CD8 T cells positive for Ki67 staining was determined by flow cytometry. Data are representative of at least 4 independent experiments with 3 to 7 mice/group/experiment. (C) LCMV-specific CD8 T cells positive for annexin V were assessed at day 8 p.i. from splenocytes of WT and FOXO3L mice by staining for anti-CD8, MHC-1 tetramers, and annexin V directly *ex vivo*. Data are expressed as the percentages of annexin V-positive cells among GP33 (P < 0.02)- or GP276 (P < 0.05)-specific CD8 T cells. Data are representative of at least 2 independent experiments with 4 to 7 mice/group/experiment.

Fig 3 — Loss of FOXO3 does not affect the differentiation of effector CD8 T cells during the early period of chronic LCMV infection. (A) The expression of the indicated cell surface molecules was examined on LCMV-specific CD8 T cells from spleens at 8 days p.i. Representative histograms are gated on MHC-I tetramer-binding CD8 T cell populations from WT (black) or FOXO3L (gray) mice. The observed MFI or the percentage of cells positive (CD62L and CD27) for each indicated protein among tetramer-positive cells is given for WT and FOXO3L mice. (B) At day 8 p.i., splenocytes from WT and FOXO3L mice were stained with MHC-I tetramers, anti-CD8, anti-CD127, and anti-KLRG-1. The percentages of SLECs (KLRG-1^HI CD127^LO) and MPECs (KLRG-1^LO CD127^HI) among LCMV-specific CD8 T cells were determined by flow cytometry. (C) The expression levels of PD-1 and LAG-3 were analyzed on LCMV-specific CD8 T cells at 8 days p.i. Bars represent the MFI for PD-1 or LAG-3 staining on LCMV-specific CD8 T cells from WT or FOXO3L mice. Data are from at least 4 independent experiments with 3 to 7 mice/group/experiment.

Loss of FOXO3 does not affect effector function during the early stages of chronic LCMV infection.

Next, we investigated whether FOXO3 deficiency affected the effector function of CD8 T cells. For this purpose, we used intracellular cytokine staining (ICCS) to measure antigen-induced production of cytokines (IFN-γ, TNF-α, and IL-2) (Fig. 4). Notably, the frequencies and numbers of IFN-γ-producing LCMV-specific CD8 T cells were significantly greater in the spleens of FOXO3L mice than in those of WT mice. A small percentage of LCMV-specific IFN-γ-producing cells also produced TNF-α and IL-2 in both FOXO3L and WT mice (Fig. 4A). However, the numbers and frequencies of these polyfunctional triple-cytokine-producing (IFN-γ, TNF-α, and IL-2) CD8 T cells in FOXO3L mice were similar to those in WT mice. Likewise, quantification of granzyme B, as a surrogate marker of lytic effector function, in LCMV-specific CD8 T cells showed no differences between WT and FOXO3L mice (Fig. 4B). It is well established that maintenance of CD8 T cell responses and viral control following an LCMV clone 13 infection requires CD4 T cell help (28, 59). Therefore, we investigated whether FOXO3 deficiency affected CD4 T cell responses to the I-A^b-restricted LCMV CD4 epitope GP66 at day 8 p.i. Interestingly, the numbers of IFN-γ-producing GP66-specific CD4 T cells in the spleens of FOXO3L mice were not significantly different from those in WT mice (Fig. 4C). Despite a substantial enhancement in the accumulation of functional CD8 T cells in FOXO3L mice, we observed no differences in viral control in the serum, liver, or lung at day 8 p.i. (Fig. 4D).

Fig 4 — Deletion of FOXO3 does not affect effector function during the early stages of chronic LCMV infection. (A) On day 8 p.i., splenocytes from WT and FOXO3L mice were stimulated with LCMV epitope peptides for 5 h directly *ex vivo*. Following stimulation, cells were stained for cell surface CD8 and intracellular cytokines (IFN-γ, IL-2, and TNF-α; P < .02 for GP33; P < .03 for GP276).(Left) Representative dot plots gated on total splenocytes show the percentage of splenocytes (top number) or of CD8 T cells (bottom number) positive for IFN-γ at 8 days p.i. (Right) Bar graphs show the total number of IFN-γ-producing LCMV-specific CD8 T cells (P < .02 for GP33; P < .04 for GP276) or the percentages of IFN-γ-producing cells that also produced TNF-α and IL-2 (triple cytokine producers). Data are from at least 4 independent experiments with 3 to 7 mice/group/experiment. (B) Splenocytes were stained with MHC-I tetramers and anti-CD8, followed by intracellular staining for granzyme B. Representative FACS histograms for granzyme B MFIs are gated from GP33 (top)- or GP276 (bottom)-specific CD8 T cells. The observed MFI for granzyme B staining on epitope-specific CD8 T cells is given in each case. (C) Splenocytes from WT and FOXO3L mice were stimulated with the LCMV-specific CD4 T-cell-specific peptide GP66. Following stimulation, cells were stained for cell surface CD4 (anti-CD4) and intracellular IFN-γ (anti-IFN-γ). The bar graph indicates the total numbers of IFN-γ-positive CD4 T cells. Data are from at least 3 independent experiments with 3 to 6 mice/group/experiment. (D) Titers of LCMV clone 13 from the sera, livers, and lungs of WT and FOXO3L mice were assessed by a plaque assay utilizing a Vero cell monolayer.

FOXO3 limits virus-specific CD8 T cells by regulating apoptosis and not proliferation during the chronic phase of viral infection.

Next, we questioned whether FOXO3 regulated T cell responses during the chronic phase of LCMV infection (>day 8 p.i.). Strikingly, at day 30 p.i., the spleens of FOXO3L mice contained significantly higher numbers of LCMV-specific CD8 T cells than those of their WT counterparts (Fig. 5A). As on day 8 p.i. (Fig. 2), this increase in the magnitude of LCMV-specific CD8 T cells was the result of diminished apoptosis and not of increased proliferation (Fig. 5B). FOXO3 deficiency did not influence the expression of CD44, CD62L, CD27, CD122, or CD127 on the cell surface (Fig. 5C). Interestingly, however, the frequencies of the SLEC and MPEC subsets in FOXO3L mice were both increased over those of their WT counterparts (Fig. 5D), and the percentages of early effectors (EEs) (KLRG-1^LO CD127^LO) were reduced in FOXO3L mice. We theorize that the increased percentages of SLECs and MPECs among FOXO3L CD8 T cells might be due to reduced apoptosis of these effector subsets following differentiation from EEs. However, we cannot exclude the possibility that FOXO3 might also regulate the differentiation of effector CD8 T cells during a CVI.

Fig 5 — Increased numbers of LCMV-specific CD8 T cells during the chronic phase of LCMV infection in FOXO3L mice result from decreased apoptosis and not from enhanced proliferation. (A) On day 30 p.i., splenocytes from WT and FOXO3L mice were stained with anti-CD8, anti-CD44, and either a D^b/GP33 (P < 0.02) or a D^b/GP276 (P < 0.01) MHC-I tetramer. (Left) Representative FACS plots show tetramer-positive CD8 T cell populations gated on total CD8 T cells. The percentage of CD8 T cells positive for the indicated tetramer in WT or FOXO3L mice is shown above each plot. (Right) Quantified flow cytometry data are representative of at least 3 independent experiments with 3 to 6 mice/group/experiment. (B) The percentage of LCMV-specific CD8 T cells positive for Ki67 (top) or annexin V (bottom) (P, <0.03 for GP33; P, <0.04 for GP276) staining was determined by flow cytometry at 30 days p.i. Data are expressed as the percentage of Ki67- or annexin V-positive cells among GP33- or GP276-specific CD8 T cells. Data are representative of at least 2 independent experiments with 3 to 6 mice/group/experiment. (C) The expression of cell surface markers CD44, CD62L, CD27, CD122, and CD127 was examined on LCMV-specific CD8 T cells from spleens at 30 days p.i. Representative histograms are gated on double-positive (MHC-I tetramer and CD8) T cell populations from WT (black) or FOXO3L (gray) mice. Numbers above histograms represent the observed MFI or the percentage of cells positive (CD62L, CD27) for each indicated protein for WT or FOXO3L mice. (D) At day 30 p.i., splenocytes from WT and FOXO3L mice were stained with MHC-I tetramers, anti-CD8, anti-CD127, and anti-KLRG-1. The percentages of SLECs, MPECs, and early effectors (EE) (KLRG-1^LO CD127^LO) among LCMV-specific CD8 T cells were quantified by flow cytometry. Data are from at least 3 independent experiments with 3 to 6 mice/group/experiment (GP33, P < .05 for SLEC, P < .04 for MPEC, P < .04 for EE; GP276, P < .05 for SLEC, P < .04 for MPEC, P < .03 for EE).

FOXO3 deletion leads to enhanced CD8 T cell function and viral clearance during the chronic phase of LCMV infection.

Next, we enumerated the functional LCMV-specific CD8 T cells in the spleens of WT and FOXO3L mice. At 30 days p.i., we observed a significant augmentation in the number of IFN-γ-producing CD8 T cells in FOXO3L mice (Fig. 6A). As shown in Fig. 6A, only a small fraction of IFN-γ-producing CD8 T cells from WT mice were also able to produce TNF-α and IL-2; the inability to produce all three cytokines is a hallmark of functional exhaustion (45, 53). In contrast, there were substantial increases in the frequency and total number of triple-cytokine-producing cells in FOXO3L mice (Fig. 6A), suggesting that the FOXO3 deficiency might have muted the functional exhaustion of CD8 T cells. The improved functionality of CD8 T cells in FOXO3L mice was, however, not associated with alterations in the expression of the inhibitory receptors PD-1 and LAG-3 (Fig. 6B). Interestingly, levels of granzyme B in LCMV-specific CD8 T cells from FOXO3L mice were lower than those in their WT counterparts (Fig. 6C). In contrast to the enhanced CD8 T cell responses, the abundance of IFN-γ-producing GP66-specific CD4 T cells was minimally affected by FOXO3 deficiency (Fig. 6D). Notably, however, viral burdens in the circulation and livers of FOXO3L mice were significantly lower (∼10-fold) than those in WT mice (Fig. 6E). Since CD4 T cell responses were minimally affected by FOXO3 deficiency, improved viral control in FOXO3L mice can be attributed primarily to increased numbers of polyfunctional LCMV-specific CD8 T cell responses. Consistent with our data, maintenance of polycytokine-producing CD8 T cells has been linked with improved HIV and hepatitis C virus (HCV) control in humans (3, 12, 17, 26, 35, 51). Collectively, the data in Fig. 5 and 6 suggested that FOXO3 regulated CD8 T cell numbers, the composition of the effector subsets, the effector function of CD8 T cells, and viral control during the chronic phase of a viral infection.

Fig 6 — Enhanced CD8 T cell function and viral clearance in FOXO3L mice during the chronic phase of LCMV infection. (A) At 30 days p.i., splenocytes from WT and FOXO3L mice were stimulated with LCMV epitope peptides for 5 h directly *ex vivo*. Following stimulation, cells were stained for cell surface CD8 (with anti-CD8) and intracellular cytokines (with anti-IFN-γ, anti-IL-2, and anti-TNF-α). (Left) Total number of epitope-specific IFN-γ-producing CD8 T cells at day 30 p.i. (P, <0.01 for both GP33 and GP276). (Center) Percentage of IL-2- and TNF-α-positive cells among the IFN-γ-positive CD8 T cells (P, <0.02 for GP33 and <0.01 for GP276). (Right) Total number of triple-cytokine-producing CD8 T cells (P, <0.009 for GP33 and <0.01 for GP276). Data are from at least 3 independent experiments with 3 to 6 mice/group/experiment. (B) The expression levels of PD-1 (top) and LAG-3 (bottom) were analyzed on LCMV-specific CD8 T cells at 30 days p.i. Splenocytes from WT and FOXO3L mice were stained with MHC-I tetramers, anti-CD8, and either anti-PD-1 or anti-LAG-3. Values are MFIs for PD-1 or LAG-3 on LCMV-specific CD8 T cells from WT or FOXO3L mice. Data are from at least 3 independent experiments with 3 to 6 mice/group/experiment. (C) On day 30 p.i., splenocytes were stained with MHC-I tetramers and anti-CD8, followed by intracellular staining for granzyme B. Representative FACS histograms for granzyme B MFIs are gated from GP33 (P < 0.03)- or GP276 (P < 0.02)-specific CD8 T cells. The MFIs observed for the granzyme B staining on LCMV-specific CD8 T cells are given at the upper right. (D) At 30 days p.i., splenocytes from WT and FOXO3L mice were stimulated with the GP66 peptide. Following stimulation, cells were stained for cell surface CD4 (with anti-CD4) and intracellular IFN-γ (with anti-IFN-γ). Bars represent the total number of GP66-specific IFN-γ-producing CD4 T cells at day 30 p.i. Data are from at least 3 independent experiments with 3 to 6 mice/group/experiment. (E) LCMV clone 13 titers in the sera, livers, and lungs of WT and FOXO3L mice were assessed by a plaque assay utilizing a Vero cell monolayer. Titers in the serum (P < 0.02) and liver (P < 0.04) were significantly lower in FOXO3L mice.

FOXO3 deficiency leads to an increase in the level of LCMV-specific polyfunctional memory CD8 T cells after viral clearance.

Clearance of infectious LCMV clone 13 occurs in most tissues by day 60 to 90 p.i., and it was important to investigate whether FOXO3 regulated the establishment of “CD8 T cell memory” after a chronic viral infection (1, 2). At day 120 p.i., the numbers of LCMV-specific CD8 T cells were sustained at significantly higher levels in FOXO3L mice than in WT mice (Fig. 7A). Also, note that FOXO3 deficiency did not alter homeostatic proliferation (Fig. 7B) or the expression of adhesion molecules, cytokine receptors, and differentiation/activation markers on the surfaces of LCMV-specific memory CD8 T cells (Fig. 7C). The percentages of CD62L^HI (central memory) and CD27^HI cells among LCMV-specific CD8 T cells in FOXO3L mice were comparable to those in WT mice, which is suggestive of normal memory differentiation. Furthermore, the expression of inhibitory receptors PD-1 and LAG-3 on memory CD8 T cells was similar in FOXO3L and WT mice (Fig. 7D). The numbers of polyfunctional LCMV-specific memory CD8 T cells were prominently elevated in the spleens of FOXO3L mice relative to those in WT mice (Fig. 7E). In contrast, the numbers of GP66-specific CD4 T cells in the spleens of FOXO3L mice were similar to those in WT mice (Fig. 7F). Taken together, the data in Fig. 7 demonstrated that FOXO3L plays a T-cell-intrinsic role in restricting the number of high-quality memory CD8 T cells following a chronic LCMV infection.

Fig 7 — FOXO3 deletion leads to an increase in the level of LCMV-specific memory CD8 T cells and polyfunctional CD8 T cells with no decrease in quality after viral clearance. (A) On day 120 p.i., splenocytes from WT and FOXO3L mice were stained with anti-CD8, anti-CD44, and either D^b/GP33 (P < 0.01) or D^b/GP276 (P < 0.009) MHC-I tetramers. (Left) Representative FACS plots show tetramer-positive CD8 T cell populations gated on total CD8 T cells. The percentage of CD8 T cells positive for the indicated tetramer in WT and FOXO3L mice is given above each plot. (Right) Quantified flow cytometry data are representative of at least 3 independent experiments with 4 to 7 mice/group/experiment. (B) The percentage of LCMV-specific CD8 T cells positive for Ki67 staining was determined by flow cytometry at 120 days p.i. Data are expressed as the percentages of Ki67-positive cells among GP33- or GP276-specific CD8 T cells. Data are representative of at least 3 independent experiments with 4 to 7 mice/group/experiment. (C) The expression of cell surface markers CD44, CD62L, KLRG-1, CD127, CD122, and CD27 was examined on LCMV-specific CD8 T cells from spleens at 120 days p.i. Representative histograms are gated from double-positive (MHC-I tetramer and CD8) T cell populations from WT (black) or FOXO3L (gray) mice. The MFI or the percentage of cells positive (CD62L, CD27) for each indicated protein is given at the top of each panel. (D) The expression levels of PD-1 (top) and LAG-3 (bottom) were analyzed on LCMV-specific CD8 T cells at 120 days p.i. Splenocytes from WT and FOXO3L mice were isolated and stained with MHC-I tetramers, anti-CD8, and either anti-PD-1 or anti-LAG-3. Bars represent the MFIs for PD-1 or LAG-3 among LCMV-specific CD8 T cells from WT or FOXO3L mice. Data are from at least 3 independent experiments with 4 to 7 mice/group/experiment. (E) At 120 days p.i., splenocytes from WT and FOXO3L mice were stimulated with LCMV epitope peptides. Following stimulation, cells were stained for surface markers (with anti-CD8) and intracellular cytokines (with anti-IFN-γ, anti-IL-2, and anti-TNF-α). (Left) Bar graphs showing statistically significant increases in the total numbers of IFN-γ-positive CD8 T cells stimulated with GP33 (P < 0.02) or GP276 (P < 0.02) on day 120 p.i. (Right) Total numbers of triple-cytokine-producing CD8 T cells from WT and FOXO3L mice (P, <0.01 for GP33 and <0.02 for GP276). Data are from at least 3 independent experiments with 4 to 7 mice/group/experiment. (F) At 120 days p.i., splenocytes from WT and FOXO3L mice were stimulated with the GP66 peptide. Following stimulation, the percentages of GP66-specific IFN-γ-producing CD4 T cells were assessed by intracellular cytokine staining. Bars represent the total numbers of GP66-specific IFN-γ-producing CD4 T cells at day 120 p.i. Data are from at least 3 independent experiments with 4 to 7 mice/group/experiment.

DISCUSSION

The central finding in this study is that FOXO3 limited the accumulation of virus-specific CD8 T cells during a chronic LCMV infection by inducing apoptosis and not by increasing proliferation. By virtue of the increase in the number of functional virus-specific CD8 T cells, viral control was more effective in FOXO3L mice than in WT mice during the chronic phase of LCMV infection. This idea is consistent with the report that effective viral control depends on the ratio of the number of virus-infected cells to the number of antiviral CD8 T cells (25). In FOXO3L mice, not only were LCMV-specific CD8 T cells increased in number; they displayed polyfunctionality. Therefore, the increase in the abundance of nonexhausted polycytokine-producing CD8 T cells might have contributed to improved LCMV control in FOXO3L mice (3, 12, 17, 26, 35).

Other signaling intermediates, including the suppressor of cytokine signaling 3 (SOCS3), have been implicated in downregulating the function of CD8 T cells during a CVI (38). However, we found that the levels of SOCS3 mRNA in FOXO3L CD8 T cells were only modestly lower than those in WT cells (data not shown). In agreement with our findings with FOXO3L mice, IL-7 treatment also increased the abundance of polycytokine-producing CD8 T cells and accelerated viral control in LCMV clone 13-infected mice (31, 38). We have preliminary evidence that IL-7 treatment led to increased phosphorylation of FOXO1/3 in LCMV-specific CD8 T cells in vivo (not shown), and it is likely that at least some of the effects of IL-7 on CD8 T cells are mediated by FOXO3 inactivation.

How FOXO3 regulates CD8 T cell apoptosis in vivo is unclear. FOXO3 is known to induce the expression of BIM, a proapoptotic member of the Bcl-2 family, and lower levels of BIM in FOXO3-deficient CD8 T cells might have inhibited apoptosis (15, 23, 46, 57). We found that BIM mRNA levels in FOXO3L CD8 T cells were ∼2-fold lower than those in WT CD8 T cells (data not shown). Additionally, FOXO3 could promote the apoptosis of CD8 T cells by inducing the proapoptotic molecule PUMA (58). It is less likely that FOXO3 induces apoptosis by Fas ligand-dependent mechanisms, because accumulation of virus-specific CD8 T cells is minimally affected by Fas deficiency during a chronic LCMV infection (60). Nevertheless, further studies to determine how FOXO3 regulates T cell apoptosis during a CVI are warranted.

The results of this study have provided important insights into the mechanisms of T cell homeostasis by identifying the transcription factor FOXO3 as a potential negative regulator of virus-specific CD8 T cell responses during a CVI. This finding has implications for the exploration of new therapeutic avenues to enhance CD8 T cell responses by targeting FOXO3 in patients with CVIs.

ACKNOWLEDGMENTS

We thank Ron Depinho for the generous gift of the FOXO3L mouse.

This work was supported by PHS grant AI048785 from the NIH to M. Suresh. Jeremy A. Sullivan was supported by a postdoctoral fellowship from the American Heart Association (10POST4580038).

Footnotes

Published ahead of print 6 June 2012

REFERENCES

1. Ahmed R, et al. 1991. Molecular basis of organ-specific selection of viral variants during chronic infection. J. Virol. 65:4242–4247 [DOI] [PMC free article] [PubMed] [Google Scholar]
2. Ahmed R, Salmi A, Butler LD, Chiller JM, Oldstone MB. 1984. Selection of genetic variants of lymphocytic choriomeningitis virus in spleens of persistently infected mice. Role in suppression of cytotoxic T lymphocyte response and viral persistence. J. Exp. Med. 160:521–540 [DOI] [PMC free article] [PubMed] [Google Scholar]
3. Almeida JR, et al. 2007. Superior control of HIV-1 replication by CD8⁺ T cells is reflected by their avidity, polyfunctionality, and clonal turnover. J. Exp. Med. 204:2473–2485 [DOI] [PMC free article] [PubMed] [Google Scholar]
4. Badovinac VP, Porter BB, Harty JT. 2002. Programmed contraction of CD8⁺ T cells after infection. Nat. Immunol. 3:619–626 [DOI] [PubMed] [Google Scholar]
5. Barata JT, et al. 2004. Activation of PI3K is indispensable for interleukin 7-mediated viability, proliferation, glucose use, and growth of T cell acute lymphoblastic leukemia cells. J. Exp. Med. 200:659–669 [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Barber DL, et al. 2006. Restoring function in exhausted CD8 T cells during chronic viral infection. Nature 439:682–687 [DOI] [PubMed] [Google Scholar]
7. Birkenkamp KU, Coffer PJ. 2003. FOXO transcription factors as regulators of immune homeostasis: molecules to die for? J. Immunol. 171:1623–1629 [DOI] [PubMed] [Google Scholar]
8. Blackburn SD, et al. 2009. Coregulation of CD8⁺ T cell exhaustion by multiple inhibitory receptors during chronic viral infection. Nat. Immunol. 10:29–37 [DOI] [PMC free article] [PubMed] [Google Scholar]
9. Bradley LM, Haynes L, Swain SL. 2005. IL-7: maintaining T-cell memory and achieving homeostasis. Trends Immunol. 26:172–176 [DOI] [PubMed] [Google Scholar]
10. Brennan P, et al. 1997. Phosphatidylinositol 3-kinase couples the interleukin-2 receptor to the cell cycle regulator E2F. Immunity 7:679–689 [DOI] [PubMed] [Google Scholar]
11. Calnan DR, Brunet A. 2008. The FoxO code. Oncogene 27:2276–2288 [DOI] [PubMed] [Google Scholar]
12. Ciuffreda D, et al. 2008. Polyfunctional HCV-specific T-cell responses are associated with effective control of HCV replication. Eur. J. Immunol. 38:2665–2677 [DOI] [PubMed] [Google Scholar]
13. Colombetti S, Basso V, Mueller DL, Mondino A. 2006. Prolonged TCR/CD28 engagement drives IL-2-independent T cell clonal expansion through signaling mediated by the mammalian target of rapamycin. J. Immunol. 176:2730–2738 [DOI] [PubMed] [Google Scholar]
14. Dejean AS, et al. 2009. Transcription factor Foxo3 controls the magnitude of T cell immune responses by modulating the function of dendritic cells. Nat. Immunol. 10:504–513 [DOI] [PMC free article] [PubMed] [Google Scholar]
15. Dijkers PF, Medema RH, Lammers JW, Koenderman L, Coffer PJ. 2000. Expression of the pro-apoptotic Bcl-2 family member Bim is regulated by the forkhead transcription factor FKHR-L1. Curr. Biol. 10:1201–1204 [DOI] [PubMed] [Google Scholar]
16. Fabre S, et al. 2008. FOXO1 regulates l-Selectin and a network of human T cell homing molecules downstream of phosphatidylinositol 3-kinase. J. Immunol. 181:2980–2989 [DOI] [PubMed] [Google Scholar]
17. Ferrando-Martinez S, et al. 2012. Differential Gag-specific polyfunctional T cell maturation patterns in HIV-1 elite controllers. J. Virol. 86:3667–3674 [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Grayson JM, Weant AE, Holbrook BC, Hildeman D. 2006. Role of Bim in regulating CD8⁺ T-cell responses during chronic viral infection. J. Virol. 80:8627–8638 [DOI] [PMC free article] [PubMed] [Google Scholar]
19. Hand TW, Morre M, Kaech SM. 2007. Expression of IL-7 receptor alpha is necessary but not sufficient for the formation of memory CD8 T cells during viral infection. Proc. Natl. Acad. Sci. U. S. A. 104:11730–11735 [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Hedrick SM. 2009. The cunning little vixen: Foxo and the cycle of life and death. Nat. Immunol. 10:1057–1063 [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Kerdiles YM, et al. 2009. Foxo1 links homing and survival of naive T cells by regulating l-selectin, CCR7 and interleukin 7 receptor. Nat. Immunol. 10:176–184 [DOI] [PMC free article] [PubMed] [Google Scholar]
22. Klenerman P, Hill A. 2005. T cells and viral persistence: lessons from diverse infections. Nat. Immunol. 6:873–879 [DOI] [PubMed] [Google Scholar]
23. Kurtulus S, et al. 2011. Bcl-2 allows effector and memory CD8⁺ T cells to tolerate higher expression of Bim. J. Immunol. 186:5729–5737 [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Letvin NL, Walker BD. 2003. Immunopathogenesis and immunotherapy in AIDS virus infections. Nat. Med. 9:861–866 [DOI] [PubMed] [Google Scholar]
25. Li Q, et al. 2009. Visualizing antigen-specific and infected cells in situ predicts outcomes in early viral infection. Science 323:1726–1729 [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Macal M, et al. 2008. Effective CD4⁺ T-cell restoration in gut-associated lymphoid tissue of HIV-infected patients is associated with enhanced Th17 cells and polyfunctional HIV-specific T-cell responses. Mucosal Immunol. 1:475–488 [DOI] [PubMed] [Google Scholar]
27. Martinez-Gac L, et al. 2004. Phosphoinositide 3-kinase and Forkhead, a switch for cell division. Biochem. Soc. Trans. 32:360–361 [DOI] [PubMed] [Google Scholar]
28. Matloubian M, Concepcion RJ, Ahmed R. 1994. CD4⁺ T cells are required to sustain CD8⁺ cytotoxic T-cell responses during chronic viral infection. J. Virol. 68:8056–8063 [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Moskophidis D, Lechner F, Pircher H, Zinkernagel RM. 1993. Virus persistence in acutely infected immunocompetent mice by exhaustion of antiviral cytotoxic effector T cells. Nature 362:758–761 [DOI] [PubMed] [Google Scholar]
30. Murali-Krishna K, et al. 1998. Counting antigen-specific CD8 T cells: a reevaluation of bystander activation during viral infection. Immunity 8:177–187 [DOI] [PubMed] [Google Scholar]
31. Nanjappa SG, Kim EH, Suresh M. 2011. Immunotherapeutic effects of IL-7 during a chronic viral infection in mice. Blood 117:5123–5132 [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Odorizzi PM, Wherry EJ. 2012. Inhibitory receptors on lymphocytes: insights from infections. J. Immunol. 188:2957–2965 [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Ou R, Zhang M, Huang L, Moskophidis D. 2008. Control of virus-specific CD8⁺ T-cell exhaustion and immune-mediated pathology by E3 ubiquitin ligase Cbl-b during chronic viral infection. J. Virol. 82:3353–3368 [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Ouyang W, Beckett O, Flavell RA, Li MO. 2009. An essential role of the Forkhead-box transcription factor Foxo1 in control of T cell homeostasis and tolerance. Immunity 30:358–371 [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Owen RE, et al. 2010. HIV⁺ elite controllers have low HIV-specific T-cell activation yet maintain strong, polyfunctional T-cell responses. AIDS 24:1095–1105 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Paik JH, et al. 2007. FoxOs are lineage-restricted redundant tumor suppressors and regulate endothelial cell homeostasis. Cell 128:309–323 [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Pantaleo G, Koup RA. 2004. Correlates of immune protection in HIV-1 infection: what we know, what we don't know, what we should know. Nat. Med. 10:806–810 [DOI] [PubMed] [Google Scholar]
38. Pellegrini M, et al. 2011. IL-7 engages multiple mechanisms to overcome chronic viral infection and limit organ pathology. Cell 144:601–613 [DOI] [PubMed] [Google Scholar]
39. Peng SL. 2008. Foxo in the immune system. Oncogene 27:2337–2344 [DOI] [PubMed] [Google Scholar]
40. Peng SL. 2007. Immune regulation by Foxo transcription factors. Autoimmunity 40:462–469 [DOI] [PubMed] [Google Scholar]
41. Rehermann B, Nascimbeni M. 2005. Immunology of hepatitis B virus and hepatitis C virus infection. Nat. Rev. Immunol. 5:215–229 [DOI] [PubMed] [Google Scholar]
42. Riou C, et al. 2007. Convergence of TCR and cytokine signaling leads to FOXO3a phosphorylation and drives the survival of CD4⁺ central memory T cells. J. Exp. Med. 204:79–91 [DOI] [PMC free article] [PubMed] [Google Scholar]
43. Rubinstein MP, et al. 2008. IL-7 and IL-15 differentially regulate CD8⁺ T-cell subsets during contraction of the immune response. Blood 112:3704–3712 [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Sarkar S, et al. 2008. Functional and genomic profiling of effector CD8 T cell subsets with distinct memory fates. J. Exp. Med. 205:625–640 [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Shin H, Wherry EJ. 2007. CD8 T cell dysfunction during chronic viral infection. Current Opin. Immunol. 19:408–415 [DOI] [PubMed] [Google Scholar]
46. Stahl M, et al. 2002. The forkhead transcription factor FoxO regulates transcription of p27Kip1 and Bim in response to IL-2. J. Immunol. 168:5024–5031 [DOI] [PubMed] [Google Scholar]
47. Sullivan JA, Kim EH, Plisch EH, Peng SL, Suresh M. 2012. FOXO3 regulates CD8 T cell memory by T cell-intrinsic mechanisms. PLoS Pathog. 8:e1002533 doi:10.1371/journal.ppat.1002533 [DOI] [PMC free article] [PubMed] [Google Scholar]
48. Suresh M, Gao X, Fischer C, Miller NE, Tewari K. 2004. Dissection of antiviral and immune regulatory functions of tumor necrosis factor receptors in a chronic lymphocytic choriomeningitis virus infection. J. Virol. 78:3906–3918 [DOI] [PMC free article] [PubMed] [Google Scholar]
49. Tinoco R, Alcalde V, Yang Y, Sauer K, Zuniga EI. 2009. Cell-intrinsic transforming growth factor-beta signaling mediates virus-specific CD8⁺ T cell deletion and viral persistence in vivo. Immunity 31:145–157 [DOI] [PMC free article] [PubMed] [Google Scholar]
50. Tothova Z, et al. 2007. FoxOs are critical mediators of hematopoietic stem cell resistance to physiologic oxidative stress. Cell 128:325–339 [DOI] [PubMed] [Google Scholar]
51. Trautmann L, et al. 2006. Upregulation of PD-1 expression on HIV-specific CD8⁺ T cells leads to reversible immune dysfunction. Nat. Med. 12:1198–1202 [DOI] [PubMed] [Google Scholar]
52. van Grevenynghe J, et al. 2008. Transcription factor FOXO3a controls the persistence of memory CD4⁺ T cells during HIV infection. Nat. Med. 14:266–274 [DOI] [PubMed] [Google Scholar]
53. Virgin HW, Wherry EJ, Ahmed R. 2009. Redefining chronic viral infection. Cell 138:30–50 [DOI] [PubMed] [Google Scholar]
54. Wan Q, et al. 2011. Cytokine signals through PI-3 kinase pathway modulate Th17 cytokine production by CCR6⁺ human memory T cells. J. Exp. Med. 208:1875–1887 [DOI] [PMC free article] [PubMed] [Google Scholar]
55. Wherry EJ. 2011. T cell exhaustion. Nat. Immunol. 12:492–499 [DOI] [PubMed] [Google Scholar]
56. Wherry EJ, Blattman JN, Murali-Krishna K, van der Most R, Ahmed R. 2003. Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J. Virol. 77:4911–4927 [DOI] [PMC free article] [PubMed] [Google Scholar]
57. Wojciechowski S, et al. 2007. Bim/Bcl-2 balance is critical for maintaining naive and memory T cell homeostasis. J. Exp. Med. 204:1665–1675 [DOI] [PMC free article] [PubMed] [Google Scholar]
58. You H, et al. 2006. FOXO3a-dependent regulation of Puma in response to cytokine/growth factor withdrawal. J. Exp. Med. 203:1657–1663 [DOI] [PMC free article] [PubMed] [Google Scholar]
59. Zajac AJ, et al. 1998. Viral immune evasion due to persistence of activated T cells without effector function. J. Exp. Med. 188:2205–2213 [DOI] [PMC free article] [PubMed] [Google Scholar]
60. Zhou S, Ou R, Huang L, Moskophidis D. 2002. Critical role for perforin-, Fas/FasL-, and TNFR1-mediated cytotoxic pathways in down-regulation of antigen-specific T cells during persistent viral infection. J. Virol. 76:829–840 [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

FOXO3 Regulates the CD8 T Cell Response to a Chronic Viral Infection

Jeremy A Sullivan

Eui Ho Kim

Erin H Plisch

M Suresh

Abstract

INTRODUCTION