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This information is current as of August 12, 2017. Specific Recognition of Mycobacterial Protein and Peptide Antigens by γδ T Cell Subsets following Infection with Virulent Mycobacterium bovis Jodi L. McGill, Randy E. Sacco, Cynthia L. Baldwin, Janice C. Telfer, Mitchell V. Palmer and W. Ray Waters Supplementary Material References Subscription Permissions Email Alerts http://www.jimmunol.org/content/suppl/2014/02/14/jimmunol.130256 7.DCSupplemental This article cites 79 articles, 31 of which you can access for free at: http://www.jimmunol.org/content/192/6/2756.full#ref-list-1 Information about subscribing to The Journal of Immunology is online at: http://jimmunol.org/subscription Submit copyright permission requests at: http://www.aai.org/About/Publications/JI/copyright.html Receive free email-alerts when new articles cite this article. Sign up at: http://jimmunol.org/alerts The Journal of Immunology is published twice each month by The American Association of Immunologists, Inc., 1451 Rockville Pike, Suite 650, Rockville, MD 20852 All rights reserved. Print ISSN: 0022-1767 Online ISSN: 1550-6606. Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 J Immunol 2014; 192:2756-2769; Prepublished online 14 February 2014; doi: 10.4049/jimmunol.1302567 http://www.jimmunol.org/content/192/6/2756 The Journal of Immunology Specific Recognition of Mycobacterial Protein and Peptide Antigens by gd T Cell Subsets following Infection with Virulent Mycobacterium bovis Jodi L. McGill,* Randy E. Sacco,* Cynthia L. Baldwin,† Janice C. Telfer,† Mitchell V. Palmer,‡ and W. Ray Waters‡ M ycobacterium bovis, a member of the Mycobacterium tuberculosis complex, causes tuberculosis (TB) in cattle and zoonotic infections in people, resulting in considerable economic hardship to the livestock industry (1) and a significant public health threat worldwide (2). Bovine TB parallels human TB in several aspects of disease pathogenesis and the development of innate and adaptive immune responses (3, 4). In particular, the M. bovis–specific gd T cell response is uniquely similar to that described in human TB patients (3, 4). Thus, the study of virulent M. bovis infection in cattle both increases our understanding of the bovine immune response to TB and represents an excellent model for understanding M. tuberculosis infection in humans. *Ruminant Diseases and Immunology Research Unit, National Animal Disease Center, Agricultural Research Service, U.S. Department of Agriculture, Ames, IA 50010; † Department of Veterinary and Animal Sciences, University of Massachusetts, Amherst, MA 01003; and ‡Infectious Bacterial Diseases Research Unit, National Animal Disease Center, Agricultural Research Service, U.S. Department of Agriculture, Ames, IA 50010 Received for publication September 24, 2013. Accepted for publication January 12, 2014. This work was supported by Agriculture and Food Research Initiative Competitive Grant 2011-67015-30736 from the U.S. Department of Agriculture National Institute of Food and Agriculture. Address correspondence and reprint requests to Dr. Jodi L. McGill, Ruminant Diseases and Immunology Research Unit, National Animal Disease Center, Agricultural Research Service, U.S. Department of Agriculture, 1920 Dayton Avenue, Ames, IA 50010-0070. E-mail address: [email protected] The online version of this article contains supplemental material. Abbreviations used in this article: BCG, bacille Calmette–Guérin; cRPMI, complete RPMI; E:C, ESAT6:CFP10; LAM, lipoarabinomannan; LN, lymph node; mAGP, mycolyl-arabinogalactan peptidoglycan; p.i., postinfection; PK-WCS, proteinase K–digested whole-cell sonicate; PPD-B, purified protein derivative from Mycobacterium bovis; PRR, pattern recognition receptor; TB, tuberculosis; WC1, Workshop Cluster 1; WCS, whole-cell sonicate. www.jimmunol.org/cgi/doi/10.4049/jimmunol.1302567 gd T cells are particularly recognized for their ability to respond to Mycobacterium; both human and murine gd T cells proliferate and secrete cytokines in recall response to protein and nonprotein phosphoantigens of M. tuberculosis (5, 6) and expand significantly in patients with active TB (7–9). Mice deficient in gd T cells exhibit significantly larger and less-organized granulomas following infection with Mycobacterium (10, 11), suggesting a role for granuloma formation. In humans and rodent species, gd T cells represent ∼5% of circulating lymphocytes (12), making it difficult to experimentally dissect their role in the immune response to TB. In contrast, gd T cells constitute a significant population in ruminants, representing as much as 30–60% of PBLs in young cattle (13, 14). This high frequency indicates a critical role for gd T cells in the immune system of the ruminant and makes it an excellent model for investigating their role in the response to TB. In cattle, gd T cells are among the first cells to accumulate at the site of delayed-type hypersensitivity reactions following injection with purified protein derivative from M. bovis (PPD-B) (15) and at the site of lesions in the lungs and lymph nodes (LNs) of virulent M. bovis–infected animals (16). In vitro, bovine gd T cells proliferate and produce IFN-g in recall responses to complex Ags, such as PPD-B (17, 18), and to specific Ags, such as the protein complex ESAT6:CFP10 (E:C) (17, 19), and the nonprotein Ag mycolylarabinogalactan peptidoglycan (mAGP), a component of the mycobacterial cell wall (20). From these studies, it is clear that gd T cells respond to mycobacterial infection; however, much remains to be understood about the specific Ags and Ag-presenting molecules mediating this recognition. Bovine gd T cells are divided into subpopulations based upon their expression of the scavenger receptor cysteine-rich superfamily member Workshop Cluster 1 (WC1), which is also known as CD163L1, or by expression of markers, such as CD8 and CD2 (14). Functionally, recent reports (21) suggest that WC1 molecules act as Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 Promoting effective immunity to Mycobacterium bovis infection is a challenge that is of interest to the fields of human and animal medicine alike. We report that gd T cells from virulent M. bovis–infected cattle respond specifically and directly to complex, protein, and nonprotein mycobacterial Ags. Importantly, to our knowledge, we demonstrate for the first time that bovine gd T cells specifically recognize peptide Ags derived from the mycobacterial protein complex ESAT6:CFP10 and that this recognition requires direct contact with APCs and signaling through the T cell Ag receptor but is independent of MHC class I or II. Furthermore, we show that M. bovis infection in cattle induces robust IL-17A protein responses. Interestingly, in contrast to results from mice, bovine CD4 T cells, and not gd T cells, are the predominant source of this critical proinflammatory mediator. Bovine gd T cells are divided into subsets based upon their expression of Workshop Cluster 1 (WC1), and we demonstrate that the M. bovis–specific gd T cell response is composed of a heterogeneous mix of WC1-expressing populations, with the serologically defined WC1.1+ and WC1.2+ subsets responding in vitro to mycobacterial Ags and accumulating in the lesions of M. bovis–infected animals. The results described in this article enhance our understanding of gd T cell biology and, because virulent M. bovis infection of cattle represents an excellent model of tuberculosis in humans, contribute to our overall understanding of the role of gd T cells in the mycobacterial-specific immune response. The Journal of Immunology, 2014, 192: 2756–2769. The Journal of Immunology Materials and Methods 10-7428 (n = 8 in experiment 1, n = 8 in experiment 2) or were mock infected (n = 7 in experiment 1). Aerosol inoculations were performed as previously described (40). Briefly, animals were fitted with an aerosol mask (Trudell Medical International, London, ON, Canada) that was modified with a rubber gasket to ensure a secure fit on the muzzle, and 2 ml bacterial inoculum in PBS was delivered via nebulization. Preparation of PBMCs PBMCs were isolated by density centrifugation from buffy coat fractions of peripheral blood collected from the jugular vein into 23 acid citrate dextrose. Contaminating RBCs were removed using hypotonic lysis. Cells were washed and resuspended in complete RPMI (cRPMI) composed of RPMI 1640 (Life Technologies, Carlsbad, CA) supplemented with 2 mM L-glutamine, 25 mM HEPES buffer, 1% antibiotic-antimycotic solution, 50 mg/ml gentamicin sulfate, 1% nonessential amino acids, 2% essential amino acids, 1% sodium pyruvate, 50 mM 2-ME, and 10% (v/v) FBS. For PBMC-proliferation assays, cells were labeled with 2.5 mM CFSE and then cultured for 6 d at 37˚C with 2 3 105 cells/well in 96-well plates. Results were corrected for background proliferation by subtracting the frequency of cells that divided in mock-stimulated cultures. Abs and reagents The following primary mAbs were used in these studies: mouse antibovine gd TCR (TCR1-N24, d-chain specific; clone GB21A, isotype IgG2b), WC1-N3 (WC1.2, clone CACTB32A, isotype IgG1), WC1-N4 (WC1.1, clone BAG25A, isotype IgM), CD14 (clone CAM36A, isotype IgG1), MHC class I (clone H58A, isotype IgG2a), MHC class II (BoLA-DRa, clone TH14B; BoLA-DQa, clone TH22A, both isotype IgG2a), WC1 (clone ILA29, isotype IgG1), and CD4 (clone IL-A11A, isotype IgG2a) (all from Washington State University mAb Center, Pullman, WA); IFN-g (clone CC302; AbD Serotec, Raleigh, NC); and anti-human CD68 (clone EBM11; from Dako USA, Carpinteria, CA). Polyclonal rabbit anti-bovine IL-17A was obtained from Kingfisher Biotech (St. Paul, MN). The following secondary Abs were used in this study: goat anti-mouse IgG1-AF488, IgG1allophycocyanin, IgM-AF594, IgM-allophycocyanin, IgG2b-PE-Cy7, IgG2bAF350, and IgG2b-Cy5 (all from Southern Biotech, Birmingham, AL). Several Ags derived from Mycobacterium were used in these studies. PPD-B (5 mg/ml) was obtained from Prionics. Recombinant E:C (2 mg/ml) was a kind gift from Dr. F. Chris Minion (Iowa State University, Ames, IA) and was prepared as previously described (41). Recombinant MPB59 (also known as Ag85B) was a kind gift from Dr. Jim McNair (Agri-Food and Biosciences Institute, Stormont, Belfast, Northern Ireland) and was used at 5 mg/ml (42). Purified mAGP from M. tuberculosis strain H37Rv, NR-14851 and purified lipoarabinomannan (LAM) from M. tuberculosis strain H37Rv, NR-14848 were obtained through the National Institutes of Health Biodefense and Emerging Infections Research Resources Repository. Both mAGP and LAM were used at a concentration of 10 mg/ml. Peptides of 14-mer length originating from E:C (Prionics) were used at a concentration of 1 mg/ml (43). Irrelevant control peptides were synthesized by New England Peptide and used at a concentration of 1 mg/ml. Whole-cell sonicates (WCSs) and proteinase K–digested WCSs (PKWCSs) from M. bovis 95-1315 were used at 10 mg/ml (44). WCSs from L. borgpetersenii serovar Hardjo, Brucella ovis, and Treponema phagedenis were used at 10 mg/ml as negative-control Ags (45, 46). The specific protein and nonprotein Ags used in this study were .98% pure and were tested for endotoxin levels using the ToxinSensor Chromogenic LAL Endotoxin Assay Kit (GenScript). Levels of endotoxin were ,3 endotoxin U/mg, a concentration similar to that found in mycobacterial Ags used in other studies (47). Animals and M. bovis challenge Flow cytometry A total of 31 male Holstein steers ∼6 mo of age were used in two independent experiments (n = 23 animals in experiment 1, n = 8 animals in experiment 2). Animals were obtained from a TB-free herd and selected based upon negative reactivity to PPD-B using the BOVIGAM IFN-g release assay (Prionics, Schlieren, Switzerland). Animals were housed in a biosafety level-3 facility at the National Animal Disease Center. All procedures were conducted according to federal and institutional guidelines and were approved by the National Animal Disease Center Animal Care and Use Committee. M. bovis strain 95-1315 was isolated from a white-tailed deer in Michigan in 1995 (38). M. bovis strain 10-7428 was isolated from a dairy cow in Colorado in 2010. Bacterial inoculums were propagated and prepared as previously described (39). Animals were infected by aerosol inoculation with 104 CFU M. bovis 95-1315 (n = 8, experiment 1) or M. bovis For surface staining, cells were resuspended at 107 cells/ml in FACS buffer and incubated for 20 min on ice with 10 mg/ml primary Abs. Cells were washed once and resuspended at 107 cells/ml with 5 mg/ml secondary Abs: goat anti-mouse IgG2b-Cy5, goat anti-mouse IgG1-PE, and goat antimouse IgM-FITC (all from Southern Biotech). PBMCs were incubated for 20 min on ice, washed, and fixed in BD FACS lysis buffer. For intracellular cytokine staining, cells were incubated with Ag for 2–3 h, and brefeldin A (5 mg/ml) was added to the cultures for the remaining 18-h incubation. Cells were surface stained as described and then permeabilized and stained for intracellular IFN-g (clone CC302; 10 mg/ml) using the BD Cytofix/Cytoperm Fixation and Permeabilization Solution kit, per the manufacturer’s instructions. All flow cytometry data were collected on a BD LSR II flow cytometer and analyzed using FlowJo software (TreeStar, San Carlos, CA). Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 pattern recognition receptors (PRRs) on gd T cells, similar to TLR. WC1+ gd T cells are CD22CD82 and are the predominant subset in circulation, whereas WC12 gd T cells are CD2+CD8+ and are prevalent in tissues such as the spleen, intestinal mucosa, and mesenteric LNs (22–24). There are 13 WC1 genes (25, 26), and differential expression of these gene products divides WC1+ gd T cells into the serologically defined subpopulations: WC1.1+, WC1.2+, and WC1.3+ (27, 28). WC1.1+ gd T cells are proinflammatory and produce IFN-g in response to Leptospira borgpetersenii (21, 29, 30), whereas the WC1.2+ gd T cell subset produces little IFN-g in response to mitogen stimulation but substantial amounts of IL-10 and can suppress CD4 T cell proliferation (31, 32). Interestingly, however, WC1.2+ gd T cells produce IFN-g in specific response to the bovine rickettsial pathogen Anaplasma marginale (33), suggesting that this subset has some functional plasticity that may be dependent upon PRR stimulation with the TCR. With regard to TB, a recent report by Price et al. (34) demonstrated that WC1.1+ gd T cells are more highly recruited to the lungs and pulmonary LNs of animals inoculated intranasally with M. bovis bacille Calmette– Guérin (BCG), although both WC12 and WC1.2+ subsets were also present. Thus, several questions remain, including the ability of individual WC1-expressing gd T cell subsets to respond specifically and directly to M. bovis Ags and their functions during infection, the types of Ags that may be recognized, and the role of WC1 itself in this recognition. In this study, we demonstrate that both WC1.1+ and WC1.2+ bovine gd T cell subsets from virulent M. bovis–infected animals proliferate and produce IFN-g in specific and direct response to complex, as well as defined, nonprotein and protein mycobacterial Ags but that WC1.2+ gd T cells predominate in the lung lesions of animals infected with virulent strains of M. bovis. Importantly, to our knowledge, we demonstrate for the first time that bovine gd T cells respond directly to peptide Ags derived from the E:C protein complex of M. bovis and that this recognition requires direct contact with APCs and recognition via the gd TCR but occurs independently of Ag-presentation via MHC class I or II. We further show that significant numbers of bovine gd T cells produce and secrete IL-17 in response to stimulation with Mycobacterium; however, in contrast to reports from mice, the dominant producers of IL-17A protein are CD4 T cells during virulent M. bovis infection in cattle. Our result is particularly interesting, given that the primary source of IL-17 in human TB patients remains controversial (35–37). Together, our results suggest that multiple functional subsets of gd T cells respond to mycobacterial Ags in both the peripheral blood and tissues of M. bovis–infected animals, and they indicate a critical role for gd T cell subsets in the immune response to TB. 2757 2758 BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION gd T cell purification and culture Real-time PCR Total RNA was extracted using the RNeasy Mini RNA Isolation Kit (QIAGEN), according to the manufacturer’s instructions. Contaminating genomic DNA was removed using the RNase-Free DNase digestion set (QIAGEN), per the manufacturer’s instructions. Total eluted RNA was reverse transcribed into cDNA using Superscript III Reverse Transcriptase and Random Primers (both from Invitrogen, Life Technologies), per the manufacturer’s instructions. Real-time PCR was performed using Power SYBR Green PCR Master Mix (Applied Biosystems). Forward and reverse primers for WC1 genes and amplification conditions were described previously (26). Primers and amplification conditions for bovine cytokines were described previously (52). Reactions were performed on a 7300 Real-Time PCR System (Applied Biosystems, Life Technologies). Relative gene expression was expressed as 22DCt (53), with RPS9 as the reference housekeeping gene (52). IL-17A ELISA and ELISPOT Cell culture supernatants were collected after 72 h of stimulation with PPD-B or E:C. IL-17A protein concentrations in the culture supernatants were determined using a commercial ELISA kit (Kingfisher Biotech), per the manufacturer’s instructions. The protocol for the IL-17A ELISPOT assay was adapted from previously described methods (54). Briefly, 2 3 105 PBMCs, purified gd T cells, or purified CD4 T cells (plus APCs) were added to polyvinylidene difluoride 96-well assay plates (Millipore, Watford, U.K.), coated with anti-bovine IL17A polyclonal Ab (5 mg/ml; Kingfisher Biotech), and incubated in the presence or absence of PPD-B or E:C. Plates were incubated for 18 h, washed, and incubated for 2 h with biotinylated anti-bovine IL-17A detec- tion Ab (5 mg/ml; Kingfisher Biotech). Plates were washed and developed using VECTASTAIN ABC-AP Standard Kit and Vector Blue Alkaline Phosphatase Substrate Kit (both from Vector Laboratories, Burlingame, CA), per the manufacturer’s instructions. Plates were read and analyzed using a standard ELISPOT Reader (Cellular Technology). IFN-g ELISA Cell culture supernatants were collected on day 6 of culture and stored at 280˚C until thawing for analysis. Concentrations of IFN-g were assessed by commercial ELISA kit (BOVIGAM; Prionics), per the manufacturer’s instructions. Concentrations of IFN-g in test samples were determined by comparing absorbances of test samples with absorbances of standards within a linear curve fit. Tissue sections At necropsy, tissues were fixed by immersion in 10% neutral-buffered formalin or snap-frozen in liquid nitrogen–cooled isopentane. For microscopic examination, formalin-fixed tissues were processed by routine paraffin-embedment techniques, cut into 5-mm sections, and stained with H&E. Lesion scoring and analysis were performed as previously described (55). For immunofluorescence, samples of lung were cut into 6-mm sections and fixed in 50% acetone/50% methanol. Blocking and staining were carried out at room temperature in a humidified chamber using 0.05 M Tris buffer. Samples were blocked for 1 h using 10% goat serum, stained with primary Ab for 1–2 h, washed, and stained with secondary Ab for 1 h. Primary Abs were used at a concentration of 0.5 mg/ml. Secondary antimouse IgG2b-AF350 was used at 1 mg/ml; all other secondary Abs were used at 0.1 mg/ml. Slides were mounted using Prolong Gold Anti-Fade Reagent (Life Technologies). To quantify the number of gd T cell subsets infiltrating pulmonary M. bovis lesions, the number of each subset present in a single highmagnification field was counted based upon expression of WC1.1 and the gd TCR (WC1.1+), WC1.2 and the gd TCR (WC1.2+), or the gd TCR but not WC1.1 or WC1.2 (WC12). WC1.1+WC1.2+ double-positive gd T cells that stained positive for the gd TCR, WC1.1, and WC1.2 and were yellow on the merged images were excluded from the analysis. Four to six high-magnification lesion fields were counted for each animal, and a total of 11 infected animals was examined. Statistics Data were analyzed using a paired one-way ANOVA with Bonferroni posttest analysis or one-tailed Student t test, when appropriate. Results are expressed as mean 6 SEM. Statistical analysis was performed using Prism v5.0 software (GraphPad). For proliferation and intracellular cytokinestaining data, background (mock-stimulated) responses were subtracted from the response to Ag and presented as change over mock. Results Bovine gd T cells from virulent M. bovis–infected animals respond to protein and nonprotein mycobacterial Ags gd T cells from infected individuals were described to proliferate and produce IFN-g in response to mycobacteria (17, 18, 20); however, the specific Ags promoting gd T cell responsiveness during virulent M. bovis infection remain poorly defined. Thus, we first set out to characterize the capacity of gd T cells from virulent M. bovis–infected animals to respond to complex, protein, or nonprotein Ags derived from Mycobacterium spp. Animals were infected via aerosol inoculation with the virulent strains of M. bovis 10-7428 or 95-1315 or were mock inoculated. Following infection, PBMCs were collected, labeled with CFSE, and cultured in the presence or absence of mycobacterial Ags of interest. On day 6, cultures were analyzed by flow cytometry for gd TCR-expressing cells that had proliferated in response to Ag, as measured by CFSE dilution. Results were gated on total live cells, total lymphocytes, and total cells expressing the gd TCR. Fig. 1A displays representative FACS plots from cultures of PBMCs from an uninfected and M. bovis–infected animal on week 3 postinfection (p.i.). Summarized in Fig. 1B, gd T cells from M. bovis–infected animals exhibited robust proliferation in response to stimulation with the Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 For MACS isolation, PBMCs were resuspended at 107 cells/ml in MACS buffer (0.5% BSA, 2 mM EDTA in PBS) and labeled with 10 mg/ml antibovine gd TCR (GB21A) for 25 min at 4˚C. Cells were washed and resuspended in anti-mouse IgG2a+b MicroBeads (Miltenyi Biotec, Auburn, CA), and cell isolation was performed per the manufacturer’s instructions. When appropriate, purified gd T cells were resuspended at 2 3 107 cells/ml and labeled with 2.5 mM CFSE. Isolated gd T cells were cultured at 2 3 106 cells/ml, and autologous monocytes were added to cultures at a ratio of 5:1. Purified gd T cells and monocytes were cultured with 10 U/ml recombinant human IL-2 (Sigma, Poole, U.K.). For TCRblocking experiments, purified gd T cells and monocytes were cultured for 1 h with 20 mg/ml anti-bovine gd TCR (GB21A) or mouse IgG2b isotype control (BD Biosciences, San Jose, CA) prior to the addition of Ag (48). For MHC class I– and MHC class II–blocking experiments, monocytes were cultured for 1 h in 20 mg/ml each anti-bovine MHC class I (H58A) (49), anti-bovine MHC class II (TH14B and TH22A) (50), or mouse IgG2a isotype control (BD Biosciences) prior to the addition of purified gd T cells. For Transwell experiments, 2 3 105 purified gd T cells were cultured in the bottom chamber of a 0.4-mM pore-size Transwell plate (Corning Life Sciences), with autologous monocytes cultured at a 5:1 ratio in the top chamber. Monocytes were isolated to 75–80% purity by MACS or by plastic adherence, as previously described (51). Briefly, PBMCs were suspended in cRPMI and allowed to adhere to plastic petri dishes for 2 h at 37˚C. Nonadherent cells were removed by washing with warm cRPMI. Adherent cells were collected by washing with ice-cold PBS and gentle scraping. In some instances, monocytes were isolated by MACS. PBMCs were resuspended at 107 cells/ml in MACS buffer (0.5% BSA, 2 mM EDTA in PBS) and labeled with 10 mg/ml anti-bovine CD14 (CAM36A, IgG1) for 25 min at 4˚C. Cells were washed and resuspended in anti-mouse IgG1 MicroBeads (Miltenyi Biotec), and cell isolation was performed per the manufacturer’s instructions. For FACS purification of the WC1.1+ and WC1.2+ gd T cell subsets, PBMCs were labeled with CFSE and cultured for 6 d with the indicated Ag. On day 6, cells were resuspended at 107 cells/ml in cRPMI and incubated for 20 min on ice with 10 mg/ml anti-bovine gd TCR and antiWC1.1 or anti-WC1.2. Cells were washed once and resuspended at 107 cells/ml in cRPMI with 5 mg/ml of the appropriate secondary Abs. PBMCs were incubated for 20 min on ice, washed once, and resuspended at ∼108 cells/ml in cRPMI for FACS sorting. gd T cell subsets were sort purified based on surface expression of the gd TCR and WC1.1 or WC1.2 and then based on dilution of CFSE in response to PPD-B or E:C. Subsets were sorted to .90% purity using a BD FACSAria Cell Sorting System (BD Biosciences). Samples were sorted directly in Buffer RLT (QIAGEN, Valencia, CA) in preparation for RNA isolation and stored at 280˚C. The Journal of Immunology 2759 complex Ags PPD-B and M. bovis 95-1315 WCSs, as well as the nonprotein Ags mAGP and LAM (Fig. 1B), both major components of the mycobacterial cell wall, and in response to PK-WCSs. These responses were expected, given the propensity of gd T cells to recognize unprocessed and nonprotein Ags. Furthermore, we observed robust proliferation in response to the protein Ags E:C and Ag85B (Fig. 1B), which were previously shown by us (19, 41, 57) and other investigators (43, 56) to be potent stimulators of M. bovis– specific CD4 T cell responses. gd T cells also divided in response to the protein Ag MPB83, to levels similar to that of Ag85B (data not shown). The responses by gd T cells were dependent upon M. bovis infection, because we did not observe significant proliferation by gd T cells from uninfected control animals, except to WCSs and PKWCSs (Fig. 1A, 1B). Moreover, we did not observe significant proliferation by gd T cells from the virulent M. bovis–infected animals in response to stimulation with negative-control Ags from other bacterial species, including L. borgpetersenii, B. ovis, or T. phagedenis (Supplemental Fig. 1A). gd T cells are potent producers of IFN-g and may be critical in promoting Th1-type immunity during M. bovis infection (58). Thus, we next chose to determine whether gd T cells from M. bovis–infected animals had the capacity to produce IFN-g in response to selected mycobacterial Ags. To this end, we performed intracellular cytokine staining for IFN-g production by gd T cells following overnight stimulation with or without the indicated mycobacterial Ags. As seen in Fig. 1C and 1D, gd T cells from M. bovis–infected animals, but not uninfected animals, produced IFN-g in response to PPD-B and the protein Ag E:C, as well as in response to mAGP, LAM, and Ag85B (Fig. 1D). Fig. 1C displays representative FACS plots from cultures of PBMCs from an uninfected and M. bovis–infected animal on week 6 p.i. M. bovis– specific gd T cell proliferation and IFN-g production were measurable as early as 2 wk p.i. and continued for the duration of the experiment. We did not observe any significant changes in the magnitude or phenotype of the gd T cell response over time. We also did not detect any significant differences in the response of gd T cells between animals infected with the M. bovis strain 95-1315 or the M. bovis strain 10-7428 or any significant differences in clinical signs, lesion scores, or histopathologic changes (M.V. Palmer, unpublished observations). Bovine gd T cells from virulent M. bovis–infected animals respond specifically and directly to mycobacterial Ags Given the complicated cytokine milieu that can exist in mixed PBMC cultures, we next wanted to determine whether gd T cells can be activated directly by mycobacterial Ags or instead respond as bystander cells. We purified gd T cells out of PBMCs from uninfected or virulent M. bovis–infected animals using MACS. Purified gd T cells were labeled with CFSE and cultured with or without IL-2 and the indicated Ags. gd T cells require contact with an APC to respond to Ag; however, because gd T cell Ag recognition is not MHC restricted, both autologous and allogeneic APCs are known to mediate gd T cell activation (48, 59, 60). We chose to include autologous peripheral blood monocytes in the cultures at a 1:5 ratio to purified gd T cells. As seen in Fig. 2A, gd T cells divided in response to M. bovis Ags, even in the absence of Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 1. gd T cells from M. bovis–infected animals divide and produce IFN-g in specific response to complex, nonprotein, and protein mycobacterial Ags. (A and B) PBMCs from uninfected or virulent M. bovis 10-7428–infected animals were labeled with CFSE, and 2.5 3 106 cells/ml were cultured for 6 d in the presence or absence of 5 mg/ml PPD-B, 2 mg/ml E:C, 10 mg/ml mAGP, 10 mg/ml LAM, 5 mg/ml recombinant Ag85B, 10 mg/ml WCSs from M. bovis 95-1315, or 10 mg/ml PK-WCSs. Cells were labeled with anti-bovine gd TCR and analyzed by flow cytometry for CFSE dilution. (A) Representative CFSE profiles from an uninfected and infected animal, gated on total live cells and total cells expressing the gd TCR. (B) The percentage of gd T cells that proliferated in response to mycobacterial Ags, as measured by CFSE dilution. (C and D) PBMCs from uninfected, virulent M. bovis 95-1315– infected, or virulent M. bovis 10-7428–infected animals were cultured at 2.5 3 106 cells/ml overnight in the presence of brefeldin A and mycobacterial Ags, as indicated above. Cells were then stained for anti-bovine gd TCR and intracellular IFN-g and analyzed by flow cytometry. (C) Representative flow plots from an uninfected control animal and an infected animal, gated on total live cells. (D) The percentage of IFNg+ cells of total gd T cells. (B and D) The background (mock-stimulated) proliferation or IFN-g production was subtracted, and results represent change over mock. n = 5–8 animals/group. Data are mean 6 SEM. Results are representative of two or three independent experiments. †p # 0.1, *p # 0.05, **p # 0.01, versus uninfected animals. 2760 BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION additional lymphocyte populations. Importantly, this proliferative response was significantly greater for gd T cells from M. bovis– infected animals than from uninfected animals following stimulation with complex (PPD-B), nonprotein (mAGP and LAM), and specific protein (E:C) Ags. To demonstrate further that this proliferation was specific for M. bovis Ags and in response to infection with M. bovis, we also stimulated purified gd T cells from infected animals with Ags derived from other bacterial species, including L. borgpetersenii, B. ovis, or T. phagedenis (Supplemental Fig. 1B). Importantly, we did not observe significant proliferation by gd T cells from M. bovis–infected animals in response to any of these irrelevant bacterial Ags. Purified gd T cells from M. bovis– infected, but not uninfected, animals also exhibited the capacity to produce IFN-g in direct response to both nonprotein and protein mycobacterial Ags, as measured by ELISA using cell culture supernatants (Fig. 2B) and by intracellular cytokine staining following overnight culture (Fig. 2C). Together, our results suggest that gd T cells from M. bovis– infected animals proliferate and produce cytokine specifically and directly in response to stimulation with defined protein and nonprotein mycobacterial products. Importantly, addition of exogenous IL-2 was required for significant gd T cell proliferation (data not shown). These results are in agreement with previous reports (61, 62) and suggest that gd T cells likely require additional cytokine signals, such as IL-2 from ab T cells, to mount a full effector response. CD4 T cells are the predominant producers of IL-17 in cattle during M. bovis infection IFN-g is a key cytokine in the immune response against M. bovis and is known to be produced by both CD4 and gd T cells following infection (3). However, accumulating evidence also suggests the importance of additional inflammatory cytokines, particularly IL-17, as M. tuberculosis and BCG induce significant IL-17 responses in both mice and humans (36, 63–65). In mice, although there is a minor contribution by IL-17+ CD4 T cells, the Mycobacteriumspecific IL-17 response appears to be dominated by gd T cells (64, 65). In humans with active TB, the result is less clear, with some reports citing dominant IL-17 production by gd T cells (37) and others citing CD4 T cells (35, 36). In the bovine, there is evidence for increased IL-17 mRNA (66, 67), but it has not been confirmed that IL-17 protein secretion is a component of the M. bovis–specific response. Furthermore, although it has been assumed that, like mice, gd T cells are the dominant producers of IL-17 during bovine TB, this has not been proven. Thus, we next chose to confirm that IL-17 protein is produced by M. bovis–specific T lymphocytes and to elucidate the individual contribution of gd T cells and CD4 T cells to this response in cattle. PBMCs from uninfected and M. bovis–infected animals were cultured for 72 h in the presence or absence of PPD-B or E:C, and supernatants were analyzed by ELISA for IL-17A protein. As seen in Fig. 3, PBMCs from M. bovis–infected animals secreted significant levels of IL-17A in response to both PPD-B and E:C (Fig. 3A), whereas those from uninfected animals did not. We confirmed these results by ELISPOT assay, and measured significant numbers of Ag-specific IL-17A–secreting cells in the blood of infected animals (Fig. 3B). We next chose to examine the individual contribution of bovine CD4 T cells and gd T cells to the M. bovis–specific IL-17 response. CD4 and gd T cells from M. bovis–infected cattle both produced IL-17A protein; however, we observed significantly more in the supernatants from purified CD4 T cells compared with purified gd T cells (Fig. 3C). We obtained similar results by ELISPOT assay, with ∼2-fold more IL-17+ CD4 T cells than IL17+ gd T cells following stimulation with either PPD-B or E:C (Fig. 3D). Similar results also were observed by intracellular cytokine staining (data not shown). It is important to note that, although the population of CD4 T cells producing IL-17A is significantly greater than the population of gd T cells, the numbers of IL-17–producing cells from both cell types remain quite large, indicating the presence of a particularly robust IL-17 response in these animals. We did not observe any pronounced difference in spot size between the CD4 and gd T cell ELISPOTs (data not Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 2. gd T cells from M. bovis–infected animals respond directly to complex, nonprotein, and protein mycobacterial Ags. (A and B) Total gd T cells and autologous monocytes were enriched from uninfected or virulent M. bovis 10-7428–infected animals, as described in Materials and Methods. Purified cells were labeled with CFSE, and 2.5 3 106 T cells/ml were cultured for 6 d at a 5:1 ratio with autologous APCs and IL-2 in the presence or absence of mycobacterial Ags, as in Fig. 1. (A) On day 6 of culture, cells were labeled with anti-bovine gd TCR and analyzed for CFSE dilution by flow cytometry. Results shown in (A) were gated on total live cells and total gd T cells and represent the percentage of total gd T cells that proliferated in response to mycobacterial Ags, as measured by CFSE dilution. (B) Supernatants from (A) were analyzed by ELISA for IFN-g. (C) MACS-purified gd T cells from uninfected, virulent M. bovis 95-1315–infected, or virulent M. bovis 10-7428–infected animals were cultured at 2.5 3 106 T cells/ml overnight in the presence of brefeldin A and the indicated Ags. Cells were stained for anti-bovine gd TCR and intracellular IFN-g and analyzed by flow cytometry. Results are gated on total live cells and total gd T cells and represent the percentage of gd T cells that are positive for IFN-g. (A and C) Background (mockstimulation) proliferation or IFN-g production was subtracted; results represent change over mock. n = 5–8 animals/group. Data are mean 6 SEM. Results are representative of two or three independent experiments. †p # 0.1, *p # 0.05, **p # 0.01, versus uninfected control animals. The Journal of Immunology shown) or in mean fluorescence intensity by flow cytometry (data not shown), indicating that the two cell types likely produced similar amounts of IL-17 on a per-cell basis. Importantly, our results demonstrate a previously unrecognized similarity in the IL-17 response of humans and cattle against Mycobacterium and reinforce the importance of using cattle as a model for human TB. Bovine gd T cell subsets respond to mycobacterial peptide Ags via a mechanism that requires direct cell–cell contact and the gd TCR Our observation that gd T cells respond specifically to M. bovis protein Ags, particularly E:C, was unexpected, given that gd T cells are primarily thought to recognize unprocessed and nonprotein Ags. In a study by Li and Wu (68), ESAT6 responsiveness by human gd T cells occurred independently of Ag processing and presentation, suggesting that recognition may be based upon conserved protein three-dimensional conformation as opposed to unique protein sequence. From this hypothesis, gd T cells would be expected to respond to whole recombinant E:C via recognition of its three-dimensional structure but not respond to peptide Ags derived from it. Thus, we purified gd T cells from the blood of virulent M. bovis–infected or uninfected animals as in Figs. 2 and 3, labeled them with CFSE, and placed them in culture with IL-2 and autologous APCs in the presence or absence of whole recombinant E:C, a peptide mixture derived from E:C, or a mixture of irrelevant control peptides. On day 6 of culture, we measured gd T cell proliferation by flow cytometry (Fig. 4). Similar to our results in Fig. 2, we observed specific and robust proliferation in response to whole E:C by cells from M. bovis–infected, but not uninfected, animals (Fig. 4A, 4B). Surprisingly, however, we also observed significant gd T cell division in response to peptides derived from E:C. This proliferation was specific to M. bovis infection, because gd T cells from uninfected animals did not divide in response to the E:C peptide mixture (Fig. 4A, 4B), nor did gd T cells proliferate in response to a mixture of irrelevant control peptides (Fig. 4B). We also measured IFN-g secretion by ELISA on cell culture supernatants and detected significant cytokine production by gd T cells in response to both E:C protein and peptides (Fig. 4B, right panel). To our knowledge, this is the first demonstration of bovine gd T cells proliferating and producing cytokine in response to peptide Ags specific to M. bovis. Despite the growing evidence for gd T cell recognition of peptide/ protein Ag, the mechanisms of this recognition continue to remain unclear (69). In our experience, bovine gd T cells must interact with monocytes or other APCs to respond to mycobacterial Ags. However, it is unclear whether this interaction requires direct cell–cell contact (i.e., Ag presentation) or relies on signals from soluble cytokine mediators (e.g., IL-2). Thus, we next chose to determine whether gd T cells require direct cell–cell interactions with APCs to recognize peptide/protein Ags derived from Mycobacterium. gd T cells were purified out of PBMCs from virulent M. bovis–infected animals as in Figs. 2 and 3, labeled with CFSE, and cultured with IL-2 in the presence or absence of E:C protein or peptides at a 5:1 ratio, with autologous purified monocytes as APCs. APCs were placed with gd T cells in the same culture (normal) or on opposite sides of a 0.4-mM pore-size Transwell. The Transwell allows secreted cytokines and soluble mediators to pass freely, but it prevents direct cell–cell interactions (i.e., presentation of mycobacterial proteins or peptides). On day 6 of culture, we analyzed gd T cell proliferation by flow cytometry. As shown in Fig. 4C, when placed together in cultures with autologous APCs (normal), we observed significant proliferation by gd T cells in response to E:C protein and peptides but not irrelevant control peptides (Fig. 4C). However, when gd T cells were separated from APCs by Transwell, we observed no significant proliferation in response to either protein or peptide mixture (Fig. 4C). We also observed that purified gd T cells failed to proliferate in response to the nonprotein Ag mAGP when separated from monocytes by Transwell (data not shown). Together, our results suggest that bovine gd T cells require direct cell–cell contact with autologous APCs to recognize both protein and nonprotein Ags from Mycobacterium. gd T cells respond to stimulation via their TCR, but they also can recognize microbial products through PRRs, such as WC1 and TLR (14, 48, 70, 71). Thus, we next chose to determine whether gd T cell recognition of mycobacterial protein and peptide Ags requires signaling through the TCR. gd T cells and autologous monocytes were purified from virulent M. bovis–infected animals as above, labeled with CFSE, and cultured with IL-2 in the presence or absence of E:C protein or peptides and 20 mg/ml anti-bovine gd TCR or isotype Ab. We then analyzed T cell proliferation on day 6 of culture. As seen in Fig. 4D, gd T cells cultured in the presence of isotype Ab divided robustly in response to both protein and peptides, whereas those cultured in the presence of the TCR-blocking Ab were significantly inhibited, indicating that gd T cell recognition of mycobacterial Ags requires signaling through the TCR. As further confirmation, purified gd T cells also failed to proliferate in response to the nonprotein Ag mAGP in the presence of the TCRblocking Ab (data not shown). Our results are in agreement with several previous reports (14, 21, 48, 52, 70) suggesting that gd T cell functions, such as proliferation and IFN-g production, re- Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 3. CD4 T cells are the predominant producers of IL-17A during virulent M. bovis infection in cattle. (A) PBMCs from uninfected or virulent M. bovis 95-1315–infected animals were cultured for 72 h at 2.5 3 106 cells/ml, with or without 5 mg/ml PPD-B or 2 mg/ml recombinant E:C. Culture supernatants were assessed for IL-17A, as measured by ELISA. (B) PBMCs from M. bovis 10-7428–infected animals were stimulated with 5 mg/ml PPD-B or 2 mg/ml recombinant E:C in ELISPOT plates (2 3 105 PBMC/well) for 18 h prior to spot development and counting. Plates were developed, as described in Materials and Methods. Results are expressed as spot-forming units (sfu)/2 3 105 cells. (C) Total gd T cells, total CD4 T cells, and APCs were enriched from M. bovis 10-7428–infected animals. gd T cells and autologous monocytes or CD4 T cells and autologous monocytes (5:1 ratio) were cultured for 72 h at 2.5 3 105 cells/well with 5 mg/ml PPD-B or 2 mg/ml recombinant E:C. Culture supernatants were assessed for IL-17A protein by ELISA. (D) Total gd T cells, total CD4 T cells, and autologous APCs were enriched as in (C) and stimulated for 18 h with PPD-B or recombinant E:C in ELISPOT plates (2 3 105 T cells/ well). Plates were developed as in Materials and Methods. Results are expressed as sfu/2 3 105 T cells. n = 5–8 animals/group. Data are mean 6 SEM. Results are representative of two independent experiments. *p # 0.05, **p # 0.01, versus uninfected animals or as indicated. 2761 2762 BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION quire stimulation through both the TCR and additional PRRs, such as WC1 or TLR. Bovine gd T cell subsets respond to mycobacterial peptide Ags via a mechanism that is independent of Ag presentation via MHC class I or MHC class II gd T cells, unlike ab T cells, are not MHC restricted (59). However, given our results suggesting a requirement for TCR signaling in the activation of M. bovis–specific gd T cells, we next chose to confirm that recognition of M. bovis–derived proteins and peptides did not require Ag presentation via MHC class I or II. We again isolated gd T cells and monocytes from M. bovis–infected animals and cultured these cells with IL-2 in the presence or absence of E:C protein or peptides and 20 mg/ml anti-bovine MHC class I (Fig. 5A), anti-bovine MHC class II (Fig. 5B), or the appropriate isotype Ab. We then analyzed T cell proliferation (Fig. 5) and IFN-g secretion (data not shown) on day 6 of culture. As seen in Fig. 5A and 5B, gd T cells from virulent M. bovis–infected animals proliferated in response to E:C protein and peptides, and this division was not inhibited by the addition of the anti-bovine MHC class I– or anti-bovine MHC class II–blocking Abs. Importantly, in parallel cultures, we observed a significant reduction in CD8 T cell proliferation in response to PPD-B when cultured with the MHC class I–blocking Ab (Supplemental Fig. 1C) and a significant reduction in CD4 T cell proliferation in response to E:C protein and peptide in the presence of the anti-bovine MHC class II–blocking Ab mixture (Supplemental Fig. 1D), confirming that our blockade was effective. Together, our results suggest that bovine gd T cells respond specifically to both whole recombinant E:C protein and the peptides derived from it and that this recognition requires direct cell–cell contact with an APC and signaling through the gd TCR but is independent of Ag presentation via MHC class I or II. Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 4. gd T cells from M. bovis–infected animals respond specifically to peptides from M. bovis in a manner dependent upon direct T cell–APC contact and the gd TCR. Total gd T cells and monocytes were enriched from uninfected or virulent M. bovis 10-7428–infected animals as in Fig. 2. Purified cells were labeled with CFSE, and 2.5 3 106 T cells/ml were cultured for 6 d at a 5:1 ratio with autologous APCs and IL-2 in the presence or absence of E:C, a peptide mixture from E:C, or an irrelevant control peptide mixture. (A and B) On day 6 of culture, cells were labeled with anti-bovine gd TCR and analyzed for CFSE dilution by flow cytometry. Representative division profiles are shown in (A) and were gated on total live cells and total gd T cells. (B) Percentage of gd T cells that divided in response to mycobacterial Ags, as measured by CFSE dilution (left panel). Culture supernatants were analyzed by ELISA for IFN-g (right panel). n = 5–8 animals/group. Data are mean 6 SEM. Results are representative of two independent experiments. †p # 0.01, *p # 0.05, **p # 0.01, versus uninfected animals. (C) gd T cells and autologous monocytes from M. bovis 10-7428–infected animals were prepared and cultured as in (B) (normal), or gd T cells and monocytes were placed on opposite sides of a 0.4-mM Transwell and incubated for 6 d. Cells were analyzed by flow cytometry for CFSE dilution. n = 4 animals/group. Data are mean 6 SEM. Results are representative of two independent experiments. †p # 0.1, *p # 0.05, versus normal cultures. (D) gd T cells and autologous monocytes from M. bovis 10-7428–infected animals were prepared as in (B). Cells were cultured for 6 d at 2 3 106 cells/ml with IL-2 in the presence of 20 mg/ml isotype Ab or anti-bovine gd TCR (GB21A)-blocking Ab, as outlined in Materials and Methods. gd T cells were analyzed by flow cytometry for CFSE dilution. n = 7 animals/group. Data are mean 6 SEM. Results are representative of two independent experiments. †p # 0.1, *p # 0.05, **p # 0.01, versus isotype cultures. The Journal of Immunology 2763 FIGURE 5. gd T cells from M. bovis–infected animals respond specifically to proteins and peptides from M. bovis in a manner that is independent of MHC class I or MHC class II. gd T cells and autologous monocytes from M. bovis 10-7428–infected animals were prepared as in Fig. 4. Cells were cultured for 6 d at 2 3 106 cells/ml with IL-2 in the presence of 20 mg/ml isotype Ab or 20 mg/ml anti-bovine MHC class I–blocking Ab (A) or anti-bovine MHC class II–blocking Ab (B), as outlined in Materials and Methods. gd T cells were analyzed by flow cytometry for CFSE dilution. n = 7 animals/group. Data are mean 6 SEM. Results are representative of two independent experiments. †p # 0.1, *p # 0.05, versus isotype cultures. Both WC1.1+ and WC1.2+ gd T cells respond to M. bovis WC1-expressing subsets contribute to the M. bovis–specific gd T cell response in cattle There are 13 WC1 genes whose distribution defines unique subsets of bovine gd T cells (25, 26). Although several subpopulations Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 WC1 has been hypothesized to act as a PRR on bovine gd T cells, binding pathogen components and lending specificity or amplification to TCR-mediated Ag recognition (21). In support of this, the serologically defined WC1.1+ gd T cell subpopulation responds specifically to Leptospira, and WC1 plays a direct role in this recall response (29, 30), whereas the serologically defined WC1.2+ gd T cell subpopulation responds specifically to infection with A. marginale (48). Serologically defined WC1.1+ gd T cells selectively accumulate in the lungs and draining LNs of animals vaccinated with M. bovis BCG (34); however, it has not been clearly proven that WC1.1+ gd T cells are the dominant subset mediating the M. bovis–specific gd T cell response. To this end, we again purified gd T cells from the blood of uninfected and M. bovis–infected cattle, labeled them with CFSE, and cultured them in the presence of autologous APCs, IL-2, and selected mycobacterial Ags. On day 6, we examined the proliferation of WC1.1+, WC1.2+, and WC12 gd T cells by flow cytometry (representative gating, Supplemental Fig. 2). As shown in Fig. 6, both WC1.1+ (Fig. 6A, 6C) and WC1.2+ (Fig. 6B, 6D) gd T cell subsets proliferated in response to stimulation with complex (PPD-B), protein (E:C), and nonprotein (mAGP and LAM) Ags. WC12 gd T cells have not been previously shown to respond to mitogen or experimental infection. In agreement, WC12 gd T cells did not proliferate in response to either protein or nonprotein mycobacterial Ags (Supplemental Fig. 2C, 2D). We observed significant IFN-g production, as measured by intracellular cytokine staining, from both WC1.1+ and WC1.2+ gd T cell subsets in response to mycobacterial Ags (Supplemental Fig. 3). It is important to note that, in some animals, the overall frequency of WC1.1+ cells in circulation is greater—sometimes up to 2-fold more—than the overall frequency of the WC1.2+ gd T cell subset. Thus, although the absolute number of WC1.1+ gd T cells responding in these cultures is likely greater than the number of WC1.2+ cells, the frequency of responding cells is remarkably similar between the two subsets. In agreement with our results by intracellular cytokine staining (Supplemental Fig. 3), purified WC1.1+ and WC1.2+ gd T cell subsets significantly upregulated expression of IFN-g mRNA in response to stimulation with mycobacterial Ags (Supplemental Fig. 4A). Additionally, we observed increased message for IL-17A (Supplemental Fig. 4B) and, although not significant, a trend for increased expression of IL-10 mRNA (Supplemental Fig. 4C) by both subsets. exist, serologically, gd T cells can only be divided into three major subsets: WC1.1+, WC1.2+, and WC1.3+. With respect to nomenclature, the WC1 genes are referred to as WC1-1, WC1-2, and so forth. There is no correlation between the naming of the WC1 genes and the serological-based nomenclature of the WC1 groups (i.e., WC1.2+ are not named because they express WC1-2). Recently, Wang et al. (21) showed by mRNA silencing that the gd T cell response to bacterial Ags was dependent upon WC1 expression, and they hypothesized that WC1 may act as a PRR, directly binding Ag and lending increased specificity to gd T cell Ag recognition. It is of great interest to identify unique targets for the 13 WC1 genes and to elucidate how signals through individual WC1s and the gd TCR are integrated to regulate Ag-specific responses. Thus, given our results demonstrating that both WC1.1+ and WC1.2+ gd T cells respond to Mycobacterium, we next chose to determine which WC1 genes were expressed by the M. bovis–responsive gd T cell subsets. PBMCs from four M. bovis–infected animals were labeled with CFSE and cultured in the presence or absence of PPD-B or E:C. On day 6, gd T cells were FACS purified based upon their serologic expression of WC1.1 or WC1.2, as well as their ability to respond to M. bovis, as measured by CFSE dilution. FACS sorting was performed using a gating strategy similar to that outlined in Supplemental Fig. 2A and 2B. The purified cells were then analyzed by quantitative real-time PCR for expression of the 13 individual WC1 genes, as described (C. Chuang, H. Hsu, J.C. Telfer, C.L. Baldwin, submitted for publication). Results were normalized to expression of the housekeeping gene RPS9 and expressed as 2-DCt. Serologically defined WC1.1+ T cells that responded to PPD-B (Fig. 7A) or E:C (Fig. 7B) expressed mRNA for the genes WC1-1, WC1-2, WC1-3, WC1-8, WC1-10, and WC1-11. The presence of these multiple WC1 genes suggests that the M. bovis–specific gd T cell response may not be restricted to a single population of WC1-expressing gd T cells but rather is composed of a heterogeneous mix of subsets. As additional support, analysis of the serologically defined WC1.2+ population indicated that gd T cells responding to PPD-B expressed mRNA for WC1-1, WC1-4, WC17, and WC1-9 (Fig. 7C), again indicating a heterogeneous response, because it was shown that WC1-4, WC1-7, and WC1-9 are expressed by the serologically designated WC1.2+ cells, whereas WC1-1 is expressed by a population defined by mAb reactivity as WC1.1+/WC1.2+ (C. Chuang et al, submitted for publication). Together, our results demonstrate that the Ag-specific gd T cell response in virulent M. bovis–infected cattle is made up of a diverse population of WC1-expressing gd T cell subsets. 2764 BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION Bovine gd T cells accumulate in pulmonary granulomas during M. bovis infection Thus far, we demonstrated robust Mycobacterium-specific responses by gd T cells isolated from the peripheral blood of virulent M. bovis–infected cattle. However, the role of gd T cells in tissues and lesions during in vivo M. bovis infection remains unclear. Therefore, we next chose to examine the localization and distribution FIGURE 7. Both WC1.1+ and WC1.2+ gd T cells respond to M. bovis Ags. PBMCs from four M. bovis 10-7428–infected animals were labeled with CFSE, and 2.5 3 106 cells/ml were cultured for 6 d in the presence of PPD-B (A, C) or recombinant E:C (B, D). Cells were FACS purified based upon CFSE dilution (i.e., responsiveness to mycobacterial Ags), expression of the gd TCR, and serologic detection of WC1.1 (A, B) or WC1.2 (C, D). Purified gd T cell subsets were analyzed by quantitative real-time PCR for expression of the individual WC1 genes (WC1-1 through WC1-13). Results are normalized to expression of the housekeeping gene RPS9 and are expressed as 22DCt. n = 4 individual animals. of gd T cells in the lesion sites of our M. bovis–infected cattle. Animals were euthanized ∼3.5 mo p.i., and the lungs and pulmonary LNs were collected for frozen and paraffin-embedded preservation. Gross and microscopic lesions were observed in the mediastinal and tracheobronchial LNs and lungs of all cattle examined (M.V. Palmer, unpublished observations). By these parameters, no significant differences were observed between cattle infected with M. bovis Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 6. Both WC1.1+ and WC1.2+ gd T cells respond to M. bovis Ags. Total gd T cells and monocytes were enriched from uninfected or virulent M. bovis 10-7428–infected animals, as in Fig. 2. T cells were labeled with CFSE, and 2 3 106 cells/ml were cultured with autologous APCs and IL-2 and stimulated with PPD-B, E:C, mAGP, or LAM for 6 d. On day 6, cells were labeled with anti-bovine gd TCR and anti-bovine WC1.1 (clone BAG25A) or antibovine WC1.2 (clone CACTB32A). CFSE dilution was analyzed by flow cytometry. Representative flow plots from an infected animal are shown for WC1.1+ gd T cells (A) and WC1.2+ gd T cells (B). Plots are gated on total live cells and total gd T cells (upper panels) and WC1.1 or WC1.2, respectively (lower panels). Percentage of WC1.1+ (C) or WC1.2+ (D) gd T cells that have diluted CFSE in response to stimulation with the indicated mycobacterial Ags. Background (mock) proliferation in (C) and (D) was subtracted from each stimulation condition, and results represent change over mock. n = 5–8 animals/ group. Data are mean 6 SEM. Results are representative of two or three independent experiments. †p # 0.1, *p # 0.05, compared with uninfected animals. The Journal of Immunology FIGURE 8. gd T cells accumulate in M. bovis lesions in the lungs of infected cattle. The lungs of uninfected or virulent M. bovis 95-1315– or M. bovis 10-7428–infected animals were analyzed 3.5 mo after aerosol challenge for M. bovis lesion formation. Paraffin-embedded lung sections from uninfected (A) and infected (B) animals were stained with H&E (original magnification 320). Frozen lung sections from control uninfected (C) and infected (D) animals were stained for immunofluorescence with anti-bovine CD68 (green) and anti-bovine gd TCR (red), as described in Materials and Methods (original magnification 320). No significant differences were observed between animals infected with M. bovis 95-1315 or 10-7428. Results are representative of four uninfected animals and nine infected animals. were significantly increased over WC1.1+ and WC12 subsets (Fig. 9E). The frequency of circulating WC1.1+ gd T cells declines with age, with the most significant changes in their numbers occurring prior to 6 mo of age (30). The animals used in these experiments were infected at 6 mo old and necropsied at 10 months (study 1) or 13 months (study 2) of age, and thus displayed relatively little change in the frequency of circulating WC1.1+ and WC1.2+ subsets (data not shown). Thus, our observation in M. bovis lesions is made even more interesting when we consider that the WC1.1+ population is larger than the WC1.2+ population in the majority of these animals, suggesting that the increased presence of WC1.2+ cells in the lesions is quite pronounced. Together, our results suggest that both WC1.1+ and WC1.2+ gd T cells respond to mycobacterial Ags in vitro; however, WC1.2+ gd T cells may in fact be the dominant subset responding to M. bovis at the site of infection. Discussion We report in this article that gd T cells from virulent M. bovis– infected animals proliferate and produce IFN-g in specific and direct response to the complex mycobacterial Ag PPD-B, the nonprotein mycobacterial Ags LAM and mAGP, and the protein Ag E:C. We further demonstrate for the first time, to our knowledge, that bovine gd T cells recognize peptide Ags derived from the mycobacterial protein E:C via a mechanism that requires direct cell–cell contact with an APC and signaling through the gd TCR, but it is independent of MHC class I– or class II–mediated Ag presentation. Interestingly, the bovine gd T cell response is composed of a heterogeneous mix of WC1-expressing subsets, because we observe accumulation of both WC1.1+ and WC1.2+ gd T cell subsets in vivo at the site of pulmonary M. bovis lesions, as well as proliferation and cytokine production by both WC1.1+ and WC1.2+ gd T cell subsets in vitro in response to stimulation with mycobacterial Ags. Finally, to our knowledge, we demonstrate for the first time that virulent M. bovis infection induces a significant IL-17A protein response in cattle and that CD4 T cells are the major source of IL-17 protein following stimulation with mycobacterial Ags. Our results are a key step in understanding the role of WC1-expressing gd T cell subsets in the immune system of the bovine, but also more broadly, because cattle make an excellent model of mycobacterial infections in humans and Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 strain 95-1315 and those infected with strain 10-7428. Uninfected lungs displayed few macrophages and no pathology (Fig. 8A), whereas lungs from infected animals exhibited granulomas composed of macrophages and multinucleated giant cells (Fig. 8B). Immunofluorescence staining for CD68+ macrophages (green) and gd TCR+ lymphocytes (red) showed only sporadic cells present in the lungs of uninfected controls (Fig. 8C). Numerous CD68+ macrophages and multinucleated giant cells (green) were observed infiltrating the center and periphery of granulomas in infected animals (Fig. 8D). Interestingly, frequent gd TCR+ lymphocytes (red) also were observed accumulating in the lymphoid mantle around the periphery of granulomatous lesions. Based on previous reports (55, 66), we know that these peripheral areas of gd T cell infiltration are sites of lymphoid cell accumulation and, thus, are also sites of CD4 T cell infiltration and, to a lesser extent, CD8 T cell infiltration. Immunofluorescence staining for CD4 T cells and gd T cells confirmed that the two subsets colocalized to similar areas in the periphery of lung granulomas (data not shown). We also observed gd T cell accumulation in the lymphoid mantle of tracheobronchial LN lesions (data not shown); however, we had difficulty discerning lymphoid follicles from granulomas by immunofluorescence microscopy; thus, we chose to examine only lung sections, because lesions could be visualized clearly in these tissues. Having demonstrated that gd T cells accumulate in the lung lesions of M. bovis–infected animals, we next chose to determine which serologically defined WC1 subsets were predominant. To this end, we stained lung granuloma sections from M. bovis– infected animals with Abs to the gd TCR (Fig. 9A, blue), WC1.2 (Fig. 9B, green), and WC1.1 (Fig. 9C, red). A merged image of blue, green, and red is shown in Fig. 9D. We quantified the number of WC1.1+, WC1.2+, and WC12 gd T cells infiltrating the lesion sites by counting high-magnification fields, as outlined in Materials and Methods. In cattle, there is a minor population of WC1.1+WC1.2+ double-positive gd T cells. In the M. bovis lesions, we observed a very small number of cells that were positive for both WC1.1 and WC1.2 and that showed up as yellow on our merged images. The number of cells was too few to quantify, and these cells were excluded from our analysis. As seen in Fig. 9, although all three subsets of gd T cells accumulated in pulmonary M. bovis lesions, the numbers of WC1.2+ gd T cells 2765 2766 BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION other species, our results enhance our understanding of the role of gd T cells in the immune response to mycobacterial infections in general. Despite emerging evidence that differential expression of WC1 molecules correlates with immunologic function, few studies have examined the distribution of WC1-expressing subsets responding to M. bovis infection. A recent report by Price et al. (34) demonstrated that aerosol inoculation of calves with M. bovis BCG induced a selective accumulation of WC1.1+ gd T cells in the lungs and LNs of the head and neck, suggesting that specifically this subset responded to M. bovis infection. In our experiments, we observed M. bovis–specific proliferation and IFN-g production by the WC1.1+ gd T cell subset; however, we also noted robust cell division and cytokine production by the WC1.2+ gd T cell subset in response to stimulation with mycobacterial products. In the lungs of our virulent M. bovis–infected animals, we observed increased frequencies of the WC1.1+, WC1.2+, and WC12 gd T cell subsets; however, in contrast to the results of Price et al. (34), we observed the most significant accumulation with the WC1.2+ subset. There are several potential factors that may contribute to these disparate observations. Price et al. (34) examined M. bovis–specific immunity that develops following vaccination; thus, their animals were inoculated with the attenuated BCG strain of M. bovis, whereas we used two strains of virulent M. bovis. The kinetics of infection may also be playing a key role in the differences observed by us compared with the study by Price et al. (34). During our experimental infection, we analyzed tissues at ∼3.5 mo p.i.; at this time point, the animals have progressed well into the chronic stage of disease, with robust M. bovis–specific adaptive immune responses and the development of late-stage granulomatous lesions in the lungs and draining LNs. In contrast, Price et al. (34) examined their animals 1 wk after experimental challenge with BCG. At this time, calves have not yet developed a robust M. bovis–specific adaptive immune response; further, with BCG inoculation, these animals will not develop pulmonary lesions. It is interesting to hypothesize that, given the proinflammatory role of WC1.1+ cells and their propensity for IFN-g production, this subset may act as the first line of defense against M. bovis challenge, being recruited in the first 1–2 wk p.i., whereas additional WC1-expressing subsets (i.e., members of the serologically defined WC1.2+ population) may then be recruited into the later M. bovis–specific response. However, in the peripheral blood of our animals, we did comparisons of the bovine gd T cell response early postinfection (i.e., 2 wk p.i.) and much later in infection (i.e., week 12 p.i. in experiment 1 and week 16 p.i. in experiment 2) and did not observe any notable differences in the overall magnitude of the response or the responding proportions of WC1.1+ and WC1.2+ gd T cell subsets (data not shown). The expression of IL-17, an inflammatory cytokine involved in granulopoiesis and neutrophil recruitment, is significantly induced in response to infection with Mycobacterium (36, 37, 65, 72). In cattle, there were reports (66, 73) of IL-17 mRNA expression following infection with M. bovis; however, to our knowledge, ours is the first confirmation that lymphocytes from M. bovis– infected animals produce IL-17A protein (Fig. 3). In our animals, the numbers of both CD4 and gd T cells producing IL-17 were quite large, indicating that the IL-17 response is a major component of the Mycobacterium-specific response in cattle. Studies from mice suggest that gd T cells are the predominant producers Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 FIGURE 9. WC1.2+ gd T cells are the predominant subset in the lung lesions of M. bovis–infected cattle. Frozen sections of the pulmonary lesions from virulent M. bovis 95-1315–infected or M. bovis 10-7428– infected animals were analyzed by immunofluorescence for WC1.1+, WC1.2+, or WC12 gd T cell subsets. Sections were stained for the gd TCR (A, blue), WC1.2 (B, green), and WC1.1 (C, red), and the images were merged (D). Shown is an animal infected with M. bovis 95-1315 (original magnification 340). No significant differences were observed between animals infected with M. bovis 95-1315 and M. bovis 10-7428. (E) The number of each gd T cell subset/high-magnification field was counted, and four to six fields were counted per animal to determine the predominant gd T cell subsets infiltrating M. bovis lesion sites. n = 11 infected animals. Data are mean 6 SEM. **p # 0.01. The Journal of Immunology of M. bovis–infected animals, as well as responses by both subsets in vitro. A recent report by Wang et al. (21) demonstrates that expression of gene products encoding for WC1.1, specifically genes WC1-1, WC1-3, and WC1-2, is necessary for the Ag-specific activation of gd T cells by Leptospira, suggesting that WC1 is acting as a PRR similar to TLRs. From these results, one can infer that different WC1 molecules are binding specific microbial products; thus, differential expression of WC1 can contribute to determining the Ag-specific activation of the cell. Given our results demonstrating the expression of several WC1 gene products by M. bovis–responsive gd T cell subsets, it is possible that WC1 is recognizing several conserved motifs in mycobacterial Ags, although this seems unlikely, given the diverse array of complex, protein, and nonprotein Ags contributing to both WC1.1+ and WC1.2+ gd T cell subset activation. It is also possible that WC1 is not contributing specifically (i.e., in the fashion of a PRR) to the M. bovis–specific response. Ultimately, the question of how signaling through the gd TCR and the various forms of WC1 contribute to Ag-specific responses will be essential to our understanding of overall gd T cell biology in cattle. In conclusion, we provide evidence in this article for the critical contribution of gd T cell subsets both in vitro and in vivo to the immune response to virulent M. bovis infection. In the future, it is imperative that we further our understanding of the functions of this poorly understood immune cell population to properly harness the gd T cell response in the development of novel intervention strategies for both human and bovine TB. Acknowledgments We thank Bruce Pesch, Kristin Bass, Molly Stafne, Jessica Pollock, Emma Frimml-Morgan, Dr. Mayara Maggioli, Tracy Porter, and Theresa Waters for excellent technical assistance. We are grateful to Dr. Jennifer Wilson-Welder and Dr. Steven Olsen for the kind gift of reagents. We also thank the animal care staff for attentive care of the animals. Disclosures The authors have no financial conflicts of interest. References 1. Waters, W. R., M. V. Palmer, B. M. Buddle, and H. M. Vordermeier. 2012. Bovine tuberculosis vaccine research: historical perspectives and recent advances. Vaccine 30: 2611–2622. 2. Michel, A. L., B. M€uller, and P. D. van Helden. 2010. Mycobacterium bovis at the animal-human interface: a problem, or not? Vet. Microbiol. 140: 371–381. 3. Waters, W. R., M. V. Palmer, T. C. Thacker, W. C. Davis, S. Sreevatsan, P. Coussens, K. G. Meade, J. C. Hope, and D. M. Estes. 2011. Tuberculosis immunity: opportunities from studies with cattle. Clin. Dev. Immunol. 2011: 768542. 4. Van Rhijn, I., J. Godfroid, A. Michel, and V. Rutten. 2008. Bovine tuberculosis as a model for human tuberculosis: advantages over small animal models. Microbes Infect. 10: 711–715. 5. Haregewoin, A., G. Soman, R. C. Hom, and R. W. Finberg. 1989. Human gamma delta+ T cells respond to mycobacterial heat-shock protein. Nature 340: 309– 312. 6. Born, W., L. Hall, A. Dallas, J. Boymel, T. Shinnick, D. Young, P. Brennan, and R. O’Brien. 1990. Recognition of a peptide antigen by heat shock–reactive gamma delta T lymphocytes. Science 249: 67–69. 7. Meraviglia, S., S. El Daker, F. Dieli, F. Martini, and A. Martino. 2011. gd T cells cross-link innate and adaptive immunity in Mycobacterium tuberculosis infection. Clin. Dev. Immunol. 2011: 587315. 8. Balbi, B., M. T. Valle, S. Oddera, D. Giunti, F. Manca, G. A. Rossi, and L. Allegra. 1993. T-lymphocytes with gamma delta+ V delta 2+ antigen receptors are present in increased proportions in a fraction of patients with tuberculosis or with sarcoidosis. Am. Rev. Respir. Dis. 148: 1685–1690. 9. Ito, M., N. Kojiro, T. Ikeda, T. Ito, J. Funada, and T. Kokubu. 1992. Increased proportions of peripheral blood gamma delta T cells in patients with pulmonary tuberculosis. Chest 102: 195–197. 10. D’Souza, C. D., A. M. Cooper, A. A. Frank, R. J. Mazzaccaro, B. R. Bloom, and I. M. Orme. 1997. An anti-inflammatory role for gamma delta T lymphocytes in acquired immunity to Mycobacterium tuberculosis. J. Immunol. 158: 1217–1221. 11. Okamoto Yoshida, Y., M. Umemura, A. Yahagi, R. L. O’Brien, K. Ikuta, K. Kishihara, H. Hara, S. Nakae, Y. Iwakura, and G. Matsuzaki. 2010. Essential Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 of the inflammatory cytokine IL-17 during the response to Mycobacterium (37, 64, 65, 72). In a mouse model of M. tuberculosis infection, Lockhart et al. (64) provided convincing evidence that gd T cells are the first source of IL-17 and are present in the lungs and spleen prior to ab T cell priming or measurable IFN-g production. Although IL-17+ CD4 T cells were present in the tissues at later time points p.i., the investigators went on to demonstrate that gd T cells continued to be the predominant source of IL-17 throughout the course of infection. In contrast, we demonstrate in this study that, in cattle infected with M. bovis, CD4 T cells are in fact the predominant source of IL-17A following stimulation with mycobacterial Ags; almost 2-fold more IL-17+ CD4 T cells than IL-17+ gd T cells are observed at both early (weeks 4 and 5) and late (week 16) time points after infection (data not shown). M. bovis–specific immune responses are undetectable prior to week 2–3 p.i. Our results are particularly interesting and may suggest an inherent difference in the Mycobacterium-specific immune response of cattle compared with rodents. However, when considering these possibilities, it is important to note that we measured IL-17–producing cells that were circulating in the blood of M. bovis–infected animals, whereas Lockhart et al. (64) examined T cells isolated from the lungs and spleen. Interestingly, reports by Jurado et al. (36) and Basile et al. (35), which examined IL-17 production by human PBLs isolated from active TB patients, concur with our results in cattle, with CD4 T cells being the dominant source of IL-17 cytokine following restimulation in vitro. However, the source of IL-17 in human TB patients remains debatable, as yet another report (37) showed predominant IL-17 production by gd T cells. Our observation that bovine gd T cells respond to peptide Ags derived from E:C was particularly unexpected. Classically, it is described that gd T cells respond to unprocessed and nonproteinaceous Ags via a mechanism that is TCR dependent but not MHC restricted (14, 74, 75). However, this dogma has changed because gd T cells have been described to respond to an incredibly diverse array of Ags, including prenyl pyrophosphates, cell surface molecules, and soluble proteins (reviewed in Ref. 76). Our results, and those of other investigators, previously suggested that bovine gd T cells have the capacity to recognize protein Ags, specifically the mycobacterial protein E:C (17, 19); however, we assumed that the nature of this recognition depended upon protein conformation and three-dimensional structure, rather than recognition of a protein epitope, as was suggested for human gd T cell recognition of ESAT6 (68, 77). Thus, we were surprised at the ability of an E:C peptide mixture to directly activate M. bovis– specific bovine gd T cells (Fig. 4). However, this result is not without precedence, because bovine gd T cells are known to respond to peptides derived from the major surface protein 2 of A. marginale (48), and a growing number of peptide Ags also have been described for murine and human gd T cells (69). Of particular interest, it was demonstrated that gd T cells isolated from humans vaccinated with BCG recognize BP3, a peptide derived from the oxidative stress response regulatory protein of M. bovis BCG (78) and, more recently, that gd T cells isolated from pulmonary TB patients expand and secrete cytokines in response to two unique M. tuberculosis peptides, one derived from 1-deoxyD-xylulose 5-phosphate synthase 2 and one derived from the extracellular region (extracellular peptide) of a mycobacterial transmembrane protein Rv2272 (79). Many questions remain concerning the role of WC1 molecules in gd T cell signaling and activation, as well as the significance of differential expression of these molecules by bovine gd T cell subsets. We report in this article the accumulation of WC1.1+ and WC1.2+ gd T cells within the pulmonary granulomatous lesions 2767 2768 12. 13. 14. 15. 16. 17. 18. 19. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. role of IL-17A in the formation of a mycobacterial infection-induced granuloma in the lung. J. Immunol. 184: 4414–4422. Kabelitz, D. 2011. gd T-cells: cross-talk between innate and adaptive immunity. Cell. Mol. Life Sci. 68: 2331–2333. Hein, W. R., and C. R. Mackay. 1991. Prominence of gamma delta T cells in the ruminant immune system. Immunol. Today 12: 30–34. Jutila, M. A., J. Holderness, J. C. Graff, and J. F. Hedges. 2008. Antigenindependent priming: a transitional response of bovine gammadelta T-cells to infection. Anim. Health Res. Rev. 9: 47–57. Doherty, M. L., M. L. Monaghan, H. F. Bassett, P. J. Quinn, and W. C. Davis. 1996. Effect of dietary restriction on cell-mediated immune responses in cattle infected with Mycobacterium bovis. Vet. Immunol. Immunopathol. 49: 307–320. Cassidy, J. P., D. G. Bryson, J. M. Pollock, R. T. Evans, F. Forster, and S. D. Neill. 1998. Early lesion formation in cattle experimentally infected with Mycobacterium bovis. J. Comp. Pathol. 119: 27–44. Rhodes, S. G., R. G. Hewinson, and H. M. Vordermeier. 2001. Antigen recognition and immunomodulation by gamma delta T cells in bovine tuberculosis. J. Immunol. 166: 5604–5610. Smyth, A. J., M. D. Welsh, R. M. Girvin, and J. M. Pollock. 2001. In vitro responsiveness of gammadelta T cells from Mycobacterium bovis-infected cattle to mycobacterial antigens: predominant involvement of WC1(+) cells. Infect. Immun. 69: 89–96. Maue, A. C., W. R. Waters, M. V. Palmer, B. J. Nonnecke, F. C. Minion, W. C. Brown, J. Norimine, M. R. Foote, C. F. Scherer, and D. M. Estes. 2007. An ESAT-6:CFP10 DNA vaccine administered in conjunction with Mycobacterium bovis BCG confers protection to cattle challenged with virulent M. bovis. Vaccine 25: 4735–4746. Vesosky, B., O. C. Turner, J. Turner, and I. M. Orme. 2004. Gamma interferon production by bovine gamma delta T cells following stimulation with mycobacterial mycolylarabinogalactan peptidoglycan. Infect. Immun. 72: 4612–4618. Wang, F., C. T. Herzig, C. Chen, H. Hsu, C. L. Baldwin, and J. C. Telfer. 2011. Scavenger receptor WC1 contributes to the gd T cell response to Leptospira. Mol. Immunol. 48: 801–809. Machugh, N. D., J. K. Mburu, M. J. Carol, C. R. Wyatt, J. A. Orden, and W. C. Davis. 1997. Identification of two distinct subsets of bovine gamma delta T cells with unique cell surface phenotype and tissue distribution. Immunology 92: 340–345. Wilson, E., M. K. Aydintug, and M. A. Jutila. 1999. A circulating bovine gamma delta T cell subset, which is found in large numbers in the spleen, accumulates inefficiently in an artificial site of inflammation: correlation with lack of expression of E-selectin ligands and L-selectin. J. Immunol. 162: 4914–4919. Wijngaard, P. L., N. D. MacHugh, M. J. Metzelaar, S. Romberg, A. Bensaid, L. Pepin, W. C. Davis, and H. C. Clevers. 1994. Members of the novel WC1 gene family are differentially expressed on subsets of bovine CD42CD82 gamma delta T lymphocytes. J. Immunol. 152: 3476–3482. Herzig, C. T., and C. L. Baldwin. 2009. Genomic organization and classification of the bovine WC1 genes and expression by peripheral blood gamma delta T cells. BMC Genomics 10: 191. Chen, C., C. T. Herzig, L. J. Alexander, J. W. Keele, T. G. McDaneld, J. C. Telfer, and C. L. Baldwin. 2012. Gene number determination and genetic polymorphism of the gamma delta T cell co-receptor WC1 genes. BMC Genet. 13: 86. Chen, C., C. T. Herzig, J. C. Telfer, and C. L. Baldwin. 2009. Antigenic basis of diversity in the gammadelta T cell co-receptor WC1 family. Mol. Immunol. 46: 2565–2575. Rogers, A. N., D. G. Vanburen, B. Zou, K. K. Lahmers, C. T. Herzig, W. C. Brown, J. C. Telfer, and C. L. Baldwin. 2006. Characterization of WC1 coreceptors on functionally distinct subpopulations of ruminant gamma delta T cells. Cell. Immunol. 239: 151–161. Rogers, A. N., D. G. VanBuren, E. Hedblom, M. E. Tilahun, J. C. Telfer, and C. L. Baldwin. 2005. Function of ruminant gammadelta T cells is defined by WC1.1 or WC1.2 isoform expression. Vet. Immunol. Immunopathol. 108: 211–217. Rogers, A. N., D. G. Vanburen, E. E. Hedblom, M. E. Tilahun, J. C. Telfer, and C. L. Baldwin. 2005. Gammadelta T cell function varies with the expressed WC1 coreceptor. J. Immunol. 174: 3386–3393. McGill, J. L., B. J. Nonnecke, J. D. Lippolis, T. A. Reinhardt, and R. E. Sacco. 2013. Differential chemokine and cytokine production by neonatal bovine gd Tcell subsets in response to viral toll-like receptor agonists and in vivo respiratory syncytial virus infection. Immunology. 139: 227–244. Hoek, A., V. P. Rutten, J. Kool, G. J. Arkesteijn, R. J. Bouwstra, I. Van Rhijn, and A. P. Koets. 2009. Subpopulations of bovine WC1(+) gammadelta T cells rather than CD4(+)CD25(high) Foxp3(+) T cells act as immune regulatory cells ex vivo. Vet. Res. 40: 6. Lahmers, K. K., J. F. Hedges, M. A. Jutila, M. Deng, M. S. Abrahamsen, and W. C. Brown. 2006. Comparative gene expression by WC1+ gammadelta and CD4+ alphabeta T lymphocytes, which respond to Anaplasma marginale, demonstrates higher expression of chemokines and other myeloid cell-associated genes by WC1+ gammadelta T cells. J. Leukoc. Biol. 80: 939–952. Price, S., M. Davies, B. Villarreal-Ramos, and J. Hope. 2010. Differential distribution of WC1(+) gammadelta TCR(+) T lymphocyte subsets within lymphoid tissues of the head and respiratory tract and effects of intranasal M. bovis BCG vaccination. Vet. Immunol. Immunopathol. 136: 133–137. Basile, J. I., L. J. Geffner, M. M. Romero, L. Balboa, C. Sabio Y Garcı́a, V. Ritacco, A. Garcı́a, M. Cuffré, E. Abbate, B. López, et al. 2011. Outbreaks of mycobacterium tuberculosis MDR strains induce high IL-17 T-cell response in patients with MDR tuberculosis that is closely associated with high antigen load. J. Infect. Dis. 204: 1054–1064. 36. Jurado, J. O., V. Pasquinelli, I. B. Alvarez, D. Peña, A. I. Rovetta, N. L. Tateosian, H. E. Romeo, R. M. Musella, D. Palmero, H. E. Chuluyán, and V. E. Garcı́a. 2012. IL-17 and IFN-g expression in lymphocytes from patients with active tuberculosis correlates with the severity of the disease. J. Leukoc. Biol. 91: 991–1002. 37. Peng, M. Y., Z. H. Wang, C. Y. Yao, L. N. Jiang, Q. L. Jin, J. Wang, and B. Q. Li. 2008. Interleukin 17-producing gamma delta T cells increased in patients with active pulmonary tuberculosis. Cell. Mol. Immunol. 5: 203–208. 38. Schmitt, S. M., S. D. Fitzgerald, T. M. Cooley, C. S. Bruning-Fann, L. Sullivan, D. Berry, T. Carlson, R. B. Minnis, J. B. Payeur, and J. Sikarskie. 1997. Bovine tuberculosis in free-ranging white-tailed deer from Michigan. J. Wildl. Dis. 33: 749–758. 39. Palmer, M. V., W. R. Waters, and D. L. Whipple. 2003. Aerosol exposure of white-tailed deer (Odocoileus virginianus) to Mycobacterium bovis. J. Wildl. Dis. 39: 817–823. 40. Palmer, M. V., W. R. Waters, and D. L. Whipple. 2002. Aerosol delivery of virulent Mycobacterium bovis to cattle. Tuberculosis (Edinb.) 82: 275–282. 41. Waters, W. R., B. J. Nonnecke, M. V. Palmer, S. Robbe-Austermann, J. P. Bannantine, J. R. Stabel, D. L. Whipple, J. B. Payeur, D. M. Estes, J. E. Pitzer, and F. C. Minion. 2004. Use of recombinant ESAT-6:CFP-10 fusion protein for differentiation of infections of cattle by Mycobacterium bovis and by M. avium subsp. avium and M. avium subsp. paratuberculosis. Clin. Diagn. Lab. Immunol. 11: 729–735. 42. Roche, P. W., P. W. Peake, H. Billman-Jacobe, T. Doran, and W. J. Britton. 1994. T-cell determinants and antibody binding sites on the major mycobacterial secretory protein MPB59 of Mycobacterium bovis. Infect. Immun. 62: 5319–5326. 43. Vordermeier, H. M., A. Whelan, P. J. Cockle, L. Farrant, N. Palmer, and R. G. Hewinson. 2001. Use of synthetic peptides derived from the antigens ESAT-6 and CFP-10 for differential diagnosis of bovine tuberculosis in cattle. Clin. Diagn. Lab. Immunol. 8: 571–578. 44. Waters, W. R., M. V. Palmer, D. L. Whipple, M. P. Carlson, and B. J. Nonnecke. 2003. Diagnostic implications of antigen-induced gamma interferon, nitric oxide, and tumor necrosis factor alpha production by peripheral blood mononuclear cells from Mycobacterium bovis-infected cattle. Clin. Diagn. Lab. Immunol. 10: 960–966. 45. Zuerner, R. L., D. P. Alt, M. V. Palmer, T. C. Thacker, and S. C. Olsen. 2011. A Leptospira borgpetersenii serovar Hardjo vaccine induces a Th1 response, activates NK cells, and reduces renal colonization. Clin. Vaccine Immunol. 18: 684–691. 46. Zuerner, R. L., M. Heidari, M. K. Elliott, D. P. Alt, and J. D. Neill. 2007. Papillomatous digital dermatitis spirochetes suppress the bovine macrophage innate immune response. Vet. Microbiol. 125: 256–264. 47. Mustafa, A. S., Y. A. Skeiky, R. Al-Attiyah, M. R. Alderson, R. G. Hewinson, and H. M. Vordermeier. 2006. Immunogenicity of Mycobacterium tuberculosis antigens in Mycobacterium bovis BCG-vaccinated and M. bovis-infected cattle. Infect. Immun. 74: 4566–4572. 48. Lahmers, K. K., J. Norimine, M. S. Abrahamsen, G. H. Palmer, and W. C. Brown. 2005. The CD4+ T cell immunodominant Anaplasma marginale major surface protein 2 stimulates gammadelta T cell clones that express unique T cell receptors. J. Leukoc. Biol. 77: 199–208. 49. Davis, T. L., and J. L. Pate. 2007. Bovine luteal cells stimulate proliferation of major histocompatibility nonrestricted gamma delta T cells. Biol. Reprod. 77: 914–922. 50. Brown, W. C., T. C. McGuire, W. Mwangi, K. A. Kegerreis, H. Macmillan, H. A. Lewin, and G. H. Palmer. 2002. Major histocompatibility complex class II DR-restricted memory CD4(+) T lymphocytes recognize conserved immunodominant epitopes of Anaplasma marginale major surface protein 1a. Infect. Immun. 70: 5521–5532. 51. Stabel, J. R., M. E. Kehrli, Jr., T. A. Reinhardt, and B. J. Nonnecke. 1997. Functional assessment of bovine monocytes isolated from peripheral blood. Vet. Immunol. Immunopathol. 58: 147–153. 52. McGill, J. L., B. J. Nonnecke, J. D. Lippolis, T. A. Reinhardt, and R. E. Sacco. 2013. Differential chemokine and cytokine production by neonatal bovine gd Tcell subsets in response to viral toll-like receptor agonists and in vivo respiratory syncytial virus infection. Immunology 139: 227–244. 53. Livak, K. J., and T. D. Schmittgen. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(2Delta Delta C(T)) Method. Methods 25: 402–408. 54. Waters, W. R., M. V. Palmer, B. J. Nonnecke, T. C. Thacker, C. F. Scherer, D. M. Estes, R. G. Hewinson, H. M. Vordermeier, S. W. Barnes, G. C. Federe, et al. 2009. Efficacy and immunogenicity of Mycobacterium bovis DeltaRD1 against aerosol M. bovis infection in neonatal calves. Vaccine 27: 1201–1209. 55. Palmer, M. V., W. R. Waters, and T. C. Thacker. 2007. Lesion development and immunohistochemical changes in granulomas from cattle experimentally infected with Mycobacterium bovis. Vet. Pathol. 44: 863–874. 56. Vordermeier, H. M., M. A. Chambers, P. J. Cockle, A. O. Whelan, J. Simmons, and R. G. Hewinson. 2002. Correlation of ESAT-6-specific gamma interferon production with pathology in cattle following Mycobacterium bovis BCG vaccination against experimental bovine tuberculosis. Infect. Immun. 70: 3026– 3032. 57. Waters, W. R., T. C. Thacker, B. J. Nonnecke, M. V. Palmer, I. Schiller, B. Oesch, H. M. Vordermeier, E. Silva, and D. M. Estes. 2012. Evaluation of gamma interferon (IFN-g)-induced protein 10 responses for detection of cattle infected with Mycobacterium bovis: comparisons to IFN-g responses. Clin. Vaccine Immunol. 19: 346–351. 58. Kennedy, H. E., M. D. Welsh, D. G. Bryson, J. P. Cassidy, F. I. Forster, C. J. Howard, R. A. Collins, and J. M. Pollock. 2002. Modulation of immune Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 20. BOVINE gd T CELL SUBSETS RESPOND TO M. BOVIS INFECTION The Journal of Immunology 59. 60. 61. 62. 63. 64. 65. 67. 68. Li, L., and C. Y. Wu. 2008. CD4+ CD25+ Treg cells inhibit human memory gammadelta T cells to produce IFN-gamma in response to M tuberculosis antigen ESAT-6. Blood 111: 5629–5636. 69. Born, W. K., L. Zhang, M. Nakayama, N. Jin, J. L. Chain, Y. Huang, M. K. Aydintug, and R. L. O’Brien. 2011. Peptide antigens for gamma/delta T cells. Cell. Mol. Life Sci. 68: 2335–2343. 70. Hedges, J. F., K. J. Lubick, and M. A. Jutila. 2005. Gamma delta T cells respond directly to pathogen-associated molecular patterns. J. Immunol. 174: 6045–6053. 71. Kerns, H. M., M. A. Jutila, and J. F. Hedges. 2009. The distinct response of gammadelta T cells to the Nod2 agonist muramyl dipeptide. Cell. Immunol. 257: 38–43. 72. Khader, S. A., J. E. Pearl, K. Sakamoto, L. Gilmartin, G. K. Bell, D. M. JelleyGibbs, N. Ghilardi, F. deSauvage, and A. M. Cooper. 2005. IL-23 compensates for the absence of IL-12p70 and is essential for the IL-17 response during tuberculosis but is dispensable for protection and antigen-specific IFN-gamma responses if IL-12p70 is available. J. Immunol. 175: 788–795. 73. Blanco, F. C., M. V. Bianco, V. Meikle, S. Garbaccio, L. Vagnoni, M. Forrellad, L. I. Klepp, A. A. Cataldi, and F. Bigi. 2011. Increased IL-17 expression is associated with pathology in a bovine model of tuberculosis. Tuberculosis (Edinb.) 91: 57–63. 74. Hayday, A. C. 2000. [gamma][delta] cells: a right time and a right place for a conserved third way of protection. Annu. Rev. Immunol. 18: 975–1026. 75. Chen, Z. W. 2011. Immune biology of Ag-specific gd T cells in infections. Cell. Mol. Life Sci. 68: 2409–2417. 76. Born, W. K., M. Kemal Aydintug, and R. L. O’Brien. 2013. Diversity of gd Tcell antigens. Cell. Mol. Immunol. 10: 13–20. 77. Casetti, R., A. Martino, A. Sacchi, C. Agrati, D. Goletti, and F. Martini. 2008. Do human gammadelta T cells respond to M tuberculosis protein antigens? Blood 112: 4776–4777, author reply 4777. 78. Xi, X., X. Zhang, B. Wang, J. Wang, H. Huang, L. Cui, X. Han, L. Li, W. He, and Z. Zhao. 2011. A novel strategy to screen Bacillus Calmette-Guérin protein antigen recognized by gd TCR. PLoS ONE 6: e18809. 79. Xi, X., X. Han, L. Li, and Z. Zhao. 2013. Identification of a new tuberculosis antigen recognized by gd T cell receptor. Clin. Vaccine Immunol. 20: 530–539. Downloaded from http://www.jimmunol.org/ by guest on August 12, 2017 66. responses to Mycobacterium bovis in cattle depleted of WC1(+) gamma delta T cells. Infect. Immun. 70: 1488–1500. Bonneville, M., R. L. O’Brien, and W. K. Born. 2010. Gammadelta T cell effector functions: a blend of innate programming and acquired plasticity. Nat. Rev. Immunol. 10: 467–478. Daubenberger, C. A., E. L. Taracha, L. Gaidulis, W. C. Davis, and D. J. McKeever. 1999. Bovine gammadelta T-cell responses to the intracellular protozoan parasite Theileria parva. Infect. Immun. 67: 2241–2249. Collins, R. A., P. Sopp, K. I. Gelder, W. I. Morrison, and C. J. Howard. 1996. Bovine gamma/delta TcR+ T lymphocytes are stimulated to proliferate by autologous Theileria annulata-infected cells in the presence of interleukin-2. Scand. J. Immunol. 44: 444–452. Fikri, Y., O. Denis, P. Pastoret, and J. Nyabenda. 2001. Purified bovine WC1+ gamma delta T lymphocytes are activated by staphylococcal enterotoxins and toxic shock syndrome toxin-1 superantigens: proliferation response, TCR V gamma profile and cytokines expression. Immunol. Lett. 77: 87–95. Khader, S. A., and A. M. Cooper. 2008. IL-23 and IL-17 in tuberculosis. Cytokine 41: 79–83. Lockhart, E., A. M. Green, and J. L. Flynn. 2006. IL-17 production is dominated by gammadelta T cells rather than CD4 T cells during Mycobacterium tuberculosis infection. J. Immunol. 177: 4662–4669. Umemura, M., A. Yahagi, S. Hamada, M. D. Begum, H. Watanabe, K. Kawakami, T. Suda, K. Sudo, S. Nakae, Y. Iwakura, and G. Matsuzaki. 2007. IL-17-mediated regulation of innate and acquired immune response against pulmonary Mycobacterium bovis bacille Calmette-Guerin infection. J. Immunol. 178: 3786–3796. Aranday-Cortes, E., N. C. Bull, B. Villarreal-Ramos, J. Gough, D. Hicks, A. Ortiz-Peláez, H. M. Vordermeier, and F. J. Salguero. 2013. Upregulation of IL-17A, CXCL9 and CXCL10 in early-stage granulomas induced by Mycobacterium bovis in cattle. Transbound. Emerg. Dis. 60: 525–537. Vordermeier, H. M., B. Villarreal-Ramos, P. J. Cockle, M. McAulay, S. G. Rhodes, T. Thacker, S. C. Gilbert, H. McShane, A. V. Hill, Z. Xing, and R. G. Hewinson. 2009. Viral booster vaccines improve Mycobacterium bovis BCG-induced protection against bovine tuberculosis. Infect. Immun. 77: 3364–3373. 2769