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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
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http://www.jimmunol.org/content/suppl/2014/02/14/jimmunol.130256
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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
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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).
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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
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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
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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
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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-
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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.
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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
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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
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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
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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
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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.
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