γδT Cells Are Required for CD8+ T Cell Response to Vaccinia Viral Infection

Vaccinia virus (VV) is the most studied member of the poxvirus family, is responsible for the successful elimination of smallpox worldwide, and has been developed as a vaccine vehicle for infectious diseases and cancer immunotherapy. We have previously shown that the unique potency of VV in the activation of CD8+ T cell response is dependent on efficient activation of the innate immune system through Toll-like receptor (TLR)-dependent and -independent pathways. However, it remains incompletely defined what regulate CD8+ T cell response to VV infection. In this study, we showed that γδT cells play an important role in promoting CD8+ T cell response to VV infection. We found that γδT cells can directly present viral antigens in the context of MHC-I for CD8+ T cell activation to VV in vivo, and we further demonstrated that cell-intrinsic MyD88 signaling in γδT cells is required for activation of γδT cells and CD8+ T cells. These results illustrate a critical role for γδT cells in the regulation of adaptive T cell response to viral infection and may shed light on the design of more effective vaccine strategies based on manipulation of γδT cells.


HIGHLIGHTS
Targeting the immune systems has powerful potentials to treat many disorders, such as cancers and viral infections. By understanding how the immune system responds to model infections, we can better determine strategies to manipulate our immune systems. Vaccinia virus is responsible for the worldwide elimination of smallpox and produces one of the longest immune responses known in humans. We know from previous findings that NK cells are required for initial immune response and CD8 + T cells are required for the elimination of the virus. How CD8 + T cells are activated in response to vaccinia virus is not fully understood. This manuscript found that gdT cells activate CD8 + T cells in response to vaccinia virus infection through MyD88 pathway.

INTRODUCTION
Vaccinia virus (VV), an enveloped double-stranded DNA virus, is a member of the Orthopoxvirus genus of the Poxviridae family. It has approximately a 200kb genome that encodes all the proteins required for cytoplasmic viral replication in host cells (1). It is responsible for the worldwide elimination of smallpox, and as a result has been developed as recombinant vaccine vehicle for infectious diseases and cancer immunotherapy (2). It is unique among viral agents to be able to elicit both potent and long-lasting immunity (3). Though its natural route of infection is via the skin, many studies have noted that intraperitoneal, intravenous, and intramuscular modes of VV inoculation provides similar clinical efficacy in both mice and humans (4)(5)(6).
We have previously shown that the unique potency of intraperitoneal VV inoculation in the activation of CD8 + T cell responses is dependent on efficient activation of the innate immune system through Toll-like receptor (TLR)-dependent and -independent pathways (7,8). Specifically, we have demonstrated that intrinsic TLR2-MyD88 (myeloid differentiation factor 88) signaling in CD8 + T cells is critical for clonal expansion and longlived memory formation (9). In addition, TLR-independent production of type I interferons (IFNs) is also important for efficient CD8 + T cell responses (7,10). However, despite these advances, the mechanisms by conventional antigen-presenting cells are unable to fully explain the unique potency of VV in the activation of CD8 + T cell responses.
gdT cells are a unique population of lymphocytes that exert a strong influence on the immune system (11). Previous studies have also shown that there are several subpopulations of gdT cells with distinct functions. However, a definitive system to categorize the different subpopulations has remained elusive. Initial proposals to define gdT cells based on their different TCR expression in both mice and humans have had to be revisited (12)(13)(14)(15)(16)(17). As a result, we assessed the effects of gdT cells as a population in this study.
In this study, we found that gdT cells play a critical role in promoting CD8 + T cell response to VV infection in vivo. We showed that activation of gdT cells by VV presented viral antigens in the context of MHC class I for CD8 + T cell activation. We further demonstrated that cell-intrinsic MyD88 signaling in gdT cells is required for gdT cell activation and CD8 + T cell responses. These results demonstrated a critical role for gdT cells in the regulation of CD8 + T cell response to viral infection and may shed light on the design of more effective vaccine strategies based on manipulation of gdT cells.

Animals
Eight-to ten-week-old C57BL/6, dTCR -/-, OT-1, and b2m -/mice were purchased from The Jackson Laboratory. MyD88 -/on C57BL/6 background were kindly provided by Shizuo Akira (Osaka University, Osaka, Japan). All experiments involving the use of mice were done in accordance with protocols approved by the Animal Care and Use Committee at Duke University and the Ohio State University.

Vaccinia Virus
Western Reserve (WR) strain of VV was purchased from American Type Culture Collection (Manassas, VA). Recombinant VV-OVA was provided by Jonathan Yewdell at NIH. The viruses were grown in TK-143B cells and purified by centrifugation through a 35% sucrose cushion as previously described (36). The titer was determined by plaque assay on TK-143B cells and subsequently stored at -80˚C until use. For in vivo studies, 5x10 6 pfu of live VV in 0.1mL Tris-Cl was injected into mice intraperitoneally, unless otherwise specified.

DC Culture
Femurs and tibiae of mice were harvested and bone marrow cells were flushed with DC medium (RPMI-1640 with 5% fetal bovine serum [FBS], 2mM L-glutamine, 10mM HEPES, 50µM bmercaptoethanol, 100 IU/mL penicillin, and 100 IU/mL streptomycin), as previously described (36). After lysis of red blood cells with ACK lysis buffer (Gibco Life Technologies, Waltham, MA), the bone marrow cells were cultured in 6-well plates at density of 3x10 6 cells/mL in 3mL DC medium in the presence of mouse granulocyte macrophage-colony stimulating factor (GM-CSF; 1000 U/mL; R&D Systems, Minneapolis, MN) and interleukin 4 (IL-4; 500 U/mL; R&D Systems). GM-CSF and IL-4 were replenished on day 2 and 4. On day 5, DCs were harvested, and CD11c + DCs were transferred onto a new 24-well plate at a density of 0.85 x 10 6 cells/mL in 2mL DC media.

Isolation of gdT Cells
Splenocytes were harvested from C57BL/6 mice 2 days after peritoneal inoculation with VV. gdT cells were isolated from harvested splenocytes with pan-T cell microbeads, followed by anti-gdTCR microbeads (Miltenyi Biotec, Auburn, CA). The isolated gdT cells were assessed via flow cytometry for confirmation.

Isolation of Immune Cells From Peritoneal Cavity
On the day of harvest, mice were euthanized in accordance with protocols approved by the Animal Care and Use Committee at Duke University and the Ohio State University. A midline incision was made along the Linea alba, without disrupting the peritoneal membrane. After exposing the outer peritoneum, a 18G needle and syringe with 4mL of 1x phosphate buffered saline (PBS) was injected into the abdomen, followed by gentle agitation. The intraperitoneal fluid was then isolated for further analysis. CD8 + T Cell Proliferation Assay CD8 + T cells were isolated from splenocytes of OT-I mice on C57BL/6 background using anti-CD8a microbeads (Miltenyi Biotec), and then fluorescently labeled with carboxyfluorescein succinimidyl ester (CFSE). Labeled CD8 + T cells and OVA-I peptide were then cocultured with matured DCs or VV-activated gdT cells at 1:1 ratio in 96 well plates. The cells were incubated at 37˚C for 72 hours, and then assessed via flow cytometry.

Adoptive Transfer of gdT Cells
Naïve gdT cells were isolated from pooled spleens and lymph nodes of wild-type or MyD88 -/mice on C57BL/6 background, with pan-T cell microbeads, followed by anti-gdTCR microbeads (Miltenyi Biotec). The isolated cells were confirmed via flow cytometry and suspended in 1xPBS. The cells were then injected intravenously via the tail vein into dTCR -/or MyD88 -/mice on C57BL/6 background at 1x10 6 cells/mouse, unless otherwise specified.

Antibodies and Flow Cytometry Analysis
The list of used antibodies is provided in Table 1. Cells were suspended in 1xPBS buffer with 2% heat-inactivated FBS and 0.1% sodium azide. After staining, cells were washed twice, and analyzed with FACSCanto flow cytometer (BD Biosciences) using FlowJo software (BD Biosciences).

Intracellular Cytokine Staining
Splenocytes were re-stimulated specifically for CD8 + T cells with 2 µg/mL B8R peptide (TSYKFESV, MBL International) with 5 µg/mL Brefeldin A (Invitrogen) for 5 hours at 2 µg/mL at 37°C. Splenocytes or mesenteric lymph node cells were stimulated specifically for gdT cells with 50 ng/mL Ionomycin, 100 ng/mL PMA, and 5 µg/mL Brefeldin A for 3 hours at 37°C. After staining with cell surface markers, the cells were fixed and permeabilized with Cytoperm/ Cytofix solution (BD Biosciences) for 20 minutes and incubated with anti-IFN-g antibodies for 30 minutes. The cells were washed twice with Permeabilization buffer (BD Biosciences) and analyzed with a FACSCanto flow cytometer using FlowJo software (BD Biosciences).

MHC/Peptide Tetramer
The VV-specific epitope B8R 2 0 -2 7 , TSYKFESV, is a synthetic peptide based on modified vaccinia virus Ankara (MVA) sequence (37). Peptide MHC I tetramers consisting of B8R 2 0 -2 7 /K b conjugated to phycoerythrin (PE) Mallophycocyanin were obtained from the NIH Tetramer Core Facility (Emory University, Atlanta, GA, USA). Cells were stained with the tetramer for 30 minutes at room temperature in the dark together with surface staining and subsequently analyzed by flow cytometry.

Plaque Assay
Viral load in the peritoneum is measured by plaque-forming assay as described (38). Mice were euthanized 3 days after infection, and the peritoneum is washed with PBS, and stored at -80˚C. Peritoneum washings were homogenized with bead homogenizer (MP Biomedical, Irvine, CA), and serial dilutions were performed to determine virus titers by plaque assay on confluent TK-143B cells.

VV Quantitative Real-Time PCR
3 days post-VV inoculation, total DNA was isolated from peritoneal fluid as previously described (39). Real-time quantitative PCR was used to analyzed VV E3L gene in duplicates using SYBR Green Real-Time PCR Master Mix (Bio-Rad). PCR conditions were 50˚C for 2 minutes, 95˚C for 10 minutes, 40 cycles of 95˚C for 15 seconds, and 60˚C for 50 seconds, followed by a melt curve capture on CFX96 Touch Real-Time PCR Detection System (Bio-Rad, Hercules, CA). Primer sequences are provided in Table 2. Relative gene expression levels for each respective gene were calculated using threshold cycle (2 -DDCT ) and normalized to GAPDH.

Statistical Analysis
Results are expressed as mean ± SEM. Differences between groups were examined for statistical significance using ANOVA with post-hoc t-test, Mann-Whitney test, or unpaired student t-test. P-values less than 0.05 are significant.

gdT cells Are Required for CD8 + T Cell Response to VV
To address whether gdT cells play a role in regulating CD8 + T cell responses, we first examined the activation status of gdT cells in response to VV infection in vivo. C57BL/6 mice were injected with VV intraperitoneally, and at different time points after infection, gdT cells were examined for IFN-g production. We found that in both spleen ( Figure 1A) and peritoneal cavity ( Figure 1B), IFN-g + gdT cell count reached its peak around day 4 following VV infection, with subsequent decline in the days following. This contrasts with VV-specific CD8 + T cell response in that B8R + IFN-g + CD8 + T cell count reached its peak around day 7 ( Figure 1). These results indicated that the activation of gdT cells peaked prior to that of CD8 + T cells. We found that naïve gdT cells secreted IFN-g after stimulation, therefore we subsequently determined IFN-g gating within each experiment against fluorescence minus one (FMO) controls ( Figures 1C, D), and between inoculated versus naïve mice (data not shown).
We next determined if gdT cells play a role in CD8 + T cell response to VV. We inoculated wild-type (WT) and dTCR -/-C57BL/6 mice intraperitoneally with VV and assessed for VVspecific B8R + CD8 + T cell activation 7 days post-inoculation. B8R is a VV epitope that is recognized by VV-specific CD8 + T cells; B8R + CD8 + T cells are specifically activated by VV. At 3 days posttransfer, we could only minimally detect the adoptively transferred gdT cells on flow cytometry, compared to dTCR -/mice without adoptive transfer (Figures 2A, B; Supplementary Figure 1). However, we found that there was a significant decrease in peritoneal VV titer in dTCR -/mice adoptively transferred with WT gdT cells, compared to VV-inoculated dTCR -/mice alone (Supplementary Figure 2; P < 0.01). Similarly, we found that there is a significant decrease in VV-specific B8R + (Figures 2C, D) and functional IFN-g + (Figures 2E, F) CD8 + T cells in dTCR -/mice that lack gdT cells at 7 days post-transfer, compared to that of WT mice (P < 0.005). We subsequently found that this defect can be rescued with adoptively transferred WT gdT cells. However, VV inoculation of dTCR -/mice with adoptive transfer of WT gdT cells had significantly greater VV-specific B8R + and IFN-g + CD8 + T cells, compared to that of dTCR -/mice with VV inoculation alone (P < 0.005). VV-specific B8R + and IFN-g + CD8 + T cell response in dTCR -/with adoptive transfer of WT gdT cells following VV inoculation also approximated the same response as WT mice with VV inoculation alone (P not significant). This suggested that gdT cells play a critical role in CD8 + T cell activation following VV infection.

VV Activates gdT Cells to Present MHC-I Peptide and Upregulate CD80 and CD86
To determine how gdT cells promote the activation CD8 + T cells to VV infection, we explored whether gdT cells contributed to signals that are required to activate CD8 + T cells: 1) direct presentation of VV-specific peptide on MHC-I, 2) costimulation with CD80 and CD86 ligands, and 3) cytokines release (40)(41)(42). To assess peptide presentation on MHC-I, we inoculated WT mice with VV or VV encoded with OVA (VV-OVA). We then assessed gdT cells for expression of H2K b specific for SIINFEKL peptide on MHC-I. We found that there is an increase in H2K b SIINFEKL + gdT cells in mice inoculated with VV-OVA, compared to that of naïve or mice inoculated with VV ( Figure 3A; P < 0.05).
We also found that following VV infection, there is an increase in CD80 and CD86 expression on the surface of gdT cells by flow cytometry. CD86 is expressed first as the initial co-stimulatory ligand, and CD80 is expressed after antigen-presenting-cell activation (43). We found that 4 days post-inoculation, there is a corresponding increase in CD80 ( Figure 3B; P < 0.05), and a significant increase in CD86 ( Figure 3C; P < 0.01). These results suggests that gdT cells could provide the necessary signals for CD8 + T cell activation after VV infection.
Signal 3 of effector CD8 + T cell activation is mainly associated with type I interferon, IL-1, and IL-12 (40,44,45). 4 days post-VV inoculation, we found that there is a significant increase in expression of IL-1 and IFN-a in gdT cells compared to that of naïve gdT cells ( Figure 3D; P < 0.001). gdT cells also secrete a basal level of IL-12 that does not change following VV inoculation but is significantly decreased following depletion of MyD88 (Supplementary Figure 3; P < 0.001). This suggests that gdT cells can provide the necessary signals for CD8 + T cell activation after VV infection.
gdT Cells Also Directly Activate CD8 + T Cells via MHC-I We next explored whether gdT cells directly activates CD8 + T cell following VV infection in vivo. To determine if VV can activate gdT cells to act as antigen presenting cells, we isolated gdT cells from WT C57BL/6 mice that had been inoculated intraperitoneally with VV 48 hours prior. CD8 + T cells were obtained from OT-I mice on C57BL/6 background and pulsed with CFSE. The CFSE-labeled CD8 + T cells were then cocultured with the VV-activated gdT cells and OVA-I peptide. 72 hours after co-incubation, CD8 + T cell proliferation was assayed by CFSE dilution. As a control, the CD8 + T cells were also incubated with matured DCs and OVA-I peptide. We found that CD8 + T cells proliferated at a similar magnitude when cocultured with VV-activated gdT cells as with matured DCs ( Figure 4A). The same proliferation is not seen if CD8 + T cells are incubated with gdT cells or OT-I peptide alone. This suggests that VV can activate gdT cells to become antigen presenting cells. We then determined if gdT cells functionally acts as professional APCs via MHC-I by assessing if CD8 + T cell activation could be rescued via adoptive transfer of b2m -/-gdT cells. b2m is a necessary component of MHC-I. We found that adoptive transfer of WT gdT cells into dTCR -/followed by VV infection significantly increased percentages and cell count of B8R + and IFN-g + CD8 + T cells compared to dTCR -/with VV infection alone (Figures 4B-E; P < 0.001). However, b2m -/-gdT cells adoptive transfer into dTCR -/followed by VV infection resulted in similar percentages and cell count of B8R + and IFN-g + CD8 + T cells as dTCR -/with VV infection alone (Figures 4B-E; P < 0.001). We found that adoptive transfer of b2m -/-gdT cells was also unable to rescue the change in absolute CD8 + T cell count, compared to adoptive transfer of WT gdT into dTCR -/mice (Supplementary Figure 4; P < 0.05). This suggests that gdT cells also activate CD8 + T cells via presentation of epitope on MHC-I for CD8 + T cell recognition. defective signaling pathways. This suggests that MyD88 signaling is required for VV activation of gdT cells. We next addressed if intrinsic MyD88 activation in gdT cells is sufficient for VV activation, opposed to signaling from other cells, we adoptively transferred WT gdT cells into MyD88 -/mice and assessed for gdT cell activation. We found that adoptive transfer of WT gdT cells into MyD88 -/mice is sufficient to rescue gdT cell expansion and IFN-g + secretion ( Figure 6; P < 0.001), suggesting intrinsic MyD88 signaling is required for activation of gdT cells by VV.
To investigated if MyD88 signaling in gdT cells is needed for subsequent CD8 + T cell activation, we adoptively transferred WT or MyD88 -/-gdT cells into dTCR -/mice and found that there was similarly gdT cell reconstitution ( Figure 7A; nonsignificant), but there was a significant decrease in VV-specific B8R + and IFN-g + CD8 + T cell expansion in dTCR -/adoptively transferred with MyD88 -/-gdT cells when compared to that of WT gdT cells (Figures 7B-E; P < 0.005).

DISCUSSION
Here we showed that VV can activate gdTcells via the MyD88 signaling pathway. We further showed that VV-activated gdT cells can present antigens to activate and induce VV-specific CD8 + T cell response.
Our results further demonstrate MyD88 has a critical role in VV activation of gdT cells to promote specific CD8 + T cell response.
gdT cells represents approximately 0.7% of the peripheral blood T cells and play an important role in the integration of the innate and adaptive immune system (18). Previous studies have shown that although activation of NK cells is critical for the initial control of VV infection (7,8), efficient activation of CD8 + T cell response is required for the eradication of VV infection (9). What promotes the activation of CD8 + T cell response to VV infection remains incompletely defined. Understanding how gdT cells activate CD8 + T cells will better elucidate the mechanisms that govern CD8 + T cell activation and how to better employ them for future strategies in vaccination or immunotherapy.
gdT cells play an active role in the control of parasitic, bacterial, and viral infections, such as malaria, Listeria monocytogenes, Salmonella, EBV, and HSV (22,(24)(25)(26)(27)(46)(47)(48). Unlike other innate immune cells, gdT cells require activation by various antigens prior to exhibiting cytotoxic characteristics (49). Currently, most strategies that target gdT cells employ phosphoantigens to activate gdT cells. However, recent evidence suggests that phosphoantigen activation is nonspecific, and induce both inflammatory and anti-inflammatory functions in the targeted cells (50)(51)(52)(53). The goal is therefore to investigate a method that would preferentially activate one subpopulation of gdT cells. Previous studies have found that different infections result in 2 main differential responses in IFN-g and IL-17 expression (54,55). However, categorization of gdT cell subpopulations remains controversial (13,16). Given the long-lasting immunity that VV produce in clinical evaluations, understanding how VV activates gdT cells to produce IFNg could provide insights into strategies to shifting gdT cells towards cytotoxic immunity overall.
In this study, we demonstrate that gdT cells is required for the full activation of CD8 + T cells after VV infection. dTCR -/mice have deficient VV-specific CD8 + T cell proliferation and functional response. We find that deficiency of gdT cells results in over 3-fold decrease in CD8 + T cell response. Given previous studies that demonstrate gdT cells' influence on the immune system, it is possible that gdT cells may be directly responsible for CD8 + T cell activation following VV infection (19,20,56). To investigate this possibility, we examined whether VV upregulates the 3 conventional signals of CD8 + T cell activation in gdT cells. We found that following infection with VV-OVA, SIINFEKL peptide is present on MHC-I on the surface of gdT cells. We also found that there is a significant decrease in CD8 + T cell activation in dTCR -/mice adoptively transferred with deficient MHC-I gdT cells, compared to that transferred with wild-type  gdT cells. Similarly, there is a significant increase in expression of CD86, IFN-a, and IL-1 in gdT cells following VV infection for signals 2 and 3 that are required for CD8 + T cell activation. This suggests that gdT cells can provide all 3 signals necessary for CD8 + T cell activation, and that gdT cells directly influence CD8 + T cell activation following VV infection.
To determine how VV activates gdT cells for antigen presentation, we assessed gdT cell activity in mice with deficiencies in IFN-ab, IFNg, TNF-a, and MyD88 signaling. We found that only mice with global deficient MyD88 signaling presented with impaired gdT cell response to VV infection. To determine if MyD88 expression in gdT cells (cellintrinsic) or in other cells (cell-extrinsic) is responsible for gdT cell activation, we investigated whether the gdT cell deficiency in MyD88 -/mice are due to signaling from other cells that require MyD88 signaling or MyD88 signaling in gdT cells alone. We found that MyD88 -/mice that were adoptively transferred with WT gdT cells exhibited normalized gdT cells response to VV. This indicates that VV could activate gdT cells via MyD88-associated PRRs. Additionally, when MyD88 -/-gdT cells are adoptively transferred to dTCR -/mice, there is a deficient CD8 + T cell response. This means that MyD88 activation on gdT cells is required for CD8 + T cell response. In conclusion, our study reveals that VV activates gdT cells via MyD88 signaling pathway, which leads to direct antigen presentation to CD8 + T cells. In vivo, this bridge between the innate and adaptive immune pathways plays a critical role in the activation of CD8 + T cell response to VV. Furthermore, we demonstrate that cell-intrinsic MyD88 signaling in gdT cells is required for activation of CD8 + T cells. These results demonstrate a critical role for gdT cells in the regulation of adaptive T cell response to viral infection and may shed light on the design of more effective vaccine strategies based on manipulation of gdT cells.

DATA AVAILABILITY STATEMENT
The raw data supporting the conclusions of this article will be made available by the authors, without undue reservation.

ETHICS STATEMENT
The animal study was reviewed and approved by The Animal Care and Use Committees at Duke University and The Ohio State University.

AUTHOR CONTRIBUTIONS
YY conceived the study and supervised the work. RD performed the experiments. YY, RD, and XH designed experiments, analyzed and interpreted data. RD and YY wrote the manuscript, and all authors reviewed and revised the manuscript, and approved the submitted version.

FUNDING
This work was supported by National Institutes of Health grants CA136934, CA186973 and CA193167 (to YY).