Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2019 Nov 25.
Published in final edited form as: Immunohorizons. 2019 Jun 12;3(6):208–218. doi: 10.4049/immunohorizons.1900032

Characterization of γδ T cell effector/memory subsets based on CD27 and CD45R expression in response to Mycobacterium bovis infection

Mariana Guerra-Maupome *, Mitchell V Palmer †, W Ray Waters †, Jodi L McGill *
PMCID: PMC6875775  NIHMSID: NIHMS1059623  PMID: 31356167

Abstract

Tuberculosis (TB) remains a leading cause of death from infectious diseases worldwide. Mycobacterium bovis is the causative agent of bovine tuberculosis and zoonotic TB infection. γδ T cells are known to participate in the immune control of mycobacterial infections. Data in human and non-human primates suggest that mycobacterial infection regulates memory/effector phenotype and adaptive immune functions of γδ T cells. To date, the impact of M. bovis infection on bovine γδ T cells and their effector and memory differentiation remains unknown. Here, we show for the first time that circulating γδ T cells from M. bovis infected cattle can be differentiated based on the expression of CD27, which is indicative of their capacity to respond to virulent M. bovis infection: CD27+ γδ T cells proliferated in response to M. bovis antigen and thus, may comprise the adaptive γδ T cell compartment in cattle. We further show that bovine M. bovis-specific γδ T cells express surface markers characteristic of TCM cells (CD45R-CD27+CD62Lhi) and that M. bovis-specific CD4 and γδ T cells both upregulate the expression of the tissue-homing receptors CXCR3 and CCR5 during infection. Our studies contribute significantly to our understanding of γδ T cell differentiation during TB infection and provide important insights into the link between phenotypic and functional subsets in the bovine. Accurate characterization of γδ T cell effector and memory-like responses induced during mycobacterial infection will contribute to improved strategies for harnessing the γδ T cell response in protection against TB for humans and animals.

INTRODUCTION

Tuberculosis (TB) is a leading cause of deaths related to an infectious disease worldwide (1). Mycobacterium bovis is a member of the M. tb complex and is the causative agent of bovine tuberculosis (bTB) and zoonotic TB infection. Despite continuous efforts to control the disease, bTB is a significant cause of economic loss to the livestock industry, and a major public health risk to populations in developing countries (2–4). Furthermore, bTB parallels human TB in several aspects of disease pathogenesis and the development of innate and adaptive immune responses (2, 5, 6). Thus, experimental studies of bTB are an excellent model to understand the immune response to M. tb infection in humans.

Cell-mediated Th-1 immune responses are essential for controlling TB (7, 8). Importantly, however, γδ T cells been shown to contribute to the immune response against TB. M. bovis infection elicits a marked in vitro expansion and robust production of IFNγ by γδ T cells suggesting that bovine γδ T cells can mount memory-like responses upon restimulation (9–14). Expression of cell surface memory markers CD45RA and CD27 is linked to functional segregation of γδ T cell memory subsets in non-human primates and humans (15, 16). Naïve CD45RA+CD27+ γδ T cells represent ~10–20% of the circulating γδ T cell population in healthy human adults. CD45RA−CD27+ central memory (TCM) γδ T cells are plentiful in peripheral blood and exhibit robust proliferative capacity, but limited effector functions. TCM cells may express CXCR3 and CCR5 but retain expression of CD62L (15). CD45RA−CD27− effector memory (TEM) and CD45RA+CD27− effector memory RA (TEMRA) γδ T cells are generally recognized to be fully differentiated subsets. They are infrequent in the blood, but abundant in tissues and sites of inflammation. Both TEM and TEMRA cells have low proliferative capacity, but robust effector functions. Consistent with their differential homing capacity, the TEM pool expresses high levels of CXCR3 and CCR5 and loses expression of the lymph node homing receptors CD62L and CCR7.

In non-human primates, TEM Vγ9Vδ2 T cell expansion after a secondary M. tb challenge correlates with protection against fatal TB (17–19). Importantly, however, serious TB disease results in a progressive loss of TEM and TEMRA γδ T cell subsets from the peripheral blood of humans with active TB (16, 20, 21). The implications of these changes on control of the disease is not well defined, and there are limitations for assessing the biological significance of γδ T cells in the response to TB in humans. Cattle have abundant γδ T cells and, as a physiologic model of TB infection, can be use to investigate γδ T cell biology in the context of Mycobacterium infection (2, 5, 22). Currently, little is known about bovine γδ T cell differentiation during TB infection. Characterization of γδ T cell effector and memory-like responses induced by mycobacterial infection will contribute to our understanding of their role in protection against TB for both humans and animals. To this end, cattle were infected via aerosol with virulent M. bovis and the effector and memory phenotypes of M. bovis-specific γδ T cells were examined by flow cytometry. We hypothesized that infection with virulent M. bovis would result in the development of circulating memory-like γδ T cell populations. M. bovis-specific γδ T cell effector and memory subsets were identified by their expression of CD27, CD45R and CD62L, and their expression of the activation and homing molecules CXCR3 and CCR5. Our results show, that CD27 and CD45R can differentiate functional (i.e. proliferating), M. bovis-specific γδ T cell subsets in bovine peripheral blood and suggest that bovine γδ T cells differentiate into effector and memory T cell subsets that are comparable to those populations which have been defined in humans with active TB infection.

MATERIALS AND METHODS

ANIMAL USE ETHICS

All animal studies were conducted according to federal and institutional guidelines and approved by the National Animal Disease Center Animal Care and Use committee and performed under appropriate project licenses. A total of 16 Holstein steers (~3 months of age) were used in the following experiments. Animals were housed in temperature-and humidity-controlled biosafety label-3 (BSL-3) containment rooms based upon treatment group, at the National Animal Disease Center in Ames, Iowa. Animals were acquired from a M. bovis-free herd in Sioux Center, IA.

Mycobacterium bovis

M. bovis strain 10–7428 isolated from a dairy farm in Colorado, was used for challenge inoculum. Low passage (≤ 3) cultures were prepared using standard techniques in Middlebrook 7H9 liquid media (Becton Dickinson, Franklin Lakes, NJ) supplemented with 10% oleic acid-albumin-dextrose 67 complex (OADC) plus 0.05% Tween 80 (Sigma, St. Louis, Missouri).

AEROSOL CHALLENGE PROCEDURES

Treatment groups consisted of non-infected steers (n =6) and animals receiving 104 colony-forming units (CFU) of M. bovis 10–7428 (n = 10). M. bovis challenge inoculum was delivered to restrained calves by aerosol as described by Palmer et al. (23). Briefly, inoculum was nebulized into a mask (Trudell Medical International, London, ON, Canada) covering the nostrils and mouth, allowing regular breathing and delivery of the bacterial inoculum to the upper and lower respiratory tract via the nostrils. The process continued until the inoculum, a 1 ml PBS wash of the inoculum tube, and an additional 2 ml PBS were delivered-a process taking ~10 min. Strict biosafety protocols were followed to protect personnel from exposure to M. bovis throughout the study, including BSL-3 containment upon initiation of M. bovis challenge in animal rooms and standard laboratory practices for handling M. bovis cultures and samples from M. bovis-infected animals.

PBMC ISOLATION

Peripheral blood was drawn from the jugular vein into 2 × acid-citrate-dextrose solution. For peripheral blood mononuclear cell (PBMC) isolation, blood was diluted 1:1 in phosphate-buffered saline (PBS) and cells were isolated from buffy coat fractions, and PBMCs were isolated by density centrifugation on Histopaque (Sigma). PBMCs were collected and washed twice in PBS to remove platelets. Residual red blood cells (RBC) were removed by adding RBC lysis buffer. Finally, PBMCs were resuspended in complete Roswell Park Memorial Institute (RPMI) 1640 medium (GIBCO, Grand Island) supplemented with 2 mM L-glutamine, 25 mM HEPES buffer, 1% antibiotic-antimycotic (Sigma, St. Louis, MO), 1% non-essential amino acids (Sigma), 2% essential amino acids (Sigma), 1% sodium pyruvate (Sigma), 50 μM 2-mercaptoethanol (Sigma), and 10% (v/v) heat-inactivated fetal bovine sera (FBS).

PROLIFERATION ASSAY

To asses proliferation, PBMC were labeled using CellTrace Violet (Invitrogen, Carlsbad, CA) prior to cell culture following manufacturer’s instructions. Briefly, freshy isolated cells were resuspended at 1×107 cells/mL in PBS containing 10 μM/ml of the CellTrace dye. After gently mixing, PBMCs were incubated for 20 min at 37°C in a water-bath. Labeling was quenched by using an equal volume of FBS, and cells were washed three times with RPMI medium. Subsequently, cells (5×105 /well) were plated in round-bottom 96-well plate in duplicates, cultured for 6-days at 37°C, in the presence of 5% CO2, in the presence of M. bovis PPD (PPD-b, 200 IU/mL, Prionics Ag, Schlieren, Switzerland), and recombinant ESAT-6/CFP-10 fusion protein (2 μg/ml, LIONEX Diagnostics and Therapeutics GmbH). Pokeweed mitogen (PWM, 1 μg/ml, Sigma) or complete RPMI medium were used as positive and negative control, respectively. After six days, cells were surface stained (see Flow cytometry section) and analyzed for proliferation and surface marker expression by flow cytometry.

FLOW CYTOMETRY

Following the appropriate culture duration, cells were stained with primary and secondary monoclonal antibodies (mAbs) listed on Table 1. All incubation steps for staining were performed in FACS buffer (PBS with 10% FBS and 0.02% NA-azide) and incubated for 25 min at 4°C. Cells were washed and fixed with BD FACS lysis buffer (BD Biosciences, Mountain View, CA) for 10 min at room temperature, washed and resuspended in FACS buffer until analysis. Samples were acquired using a BD LSR Fortessa flow cytometer (BD Biosciences). Data were analyzed using Flowjo software (Tree Star Inc., San Carlos, CA). Electronic gates were set using Fluorescence minus one (FMO) controls.

Table 1.

Primary and secondary antibodies and staining reagents.

Reagent or
antibody
clone		 Specificity, Source Secondary antibodies, Source
ILA11 Bovine CD4, Washington State University Allophycocyanin
GB21A Bovine TCR1 delta chain, Washington State University APC-Cy7, SouthernBiotech
ILA116 Bovine CD45RO, Washington State University Alexa-fluor 488, Life Technologies
BAQ92A Bovine CD62L, Washington State University Allophycocyanin, Life technologies
GC6A Bovine CD45R, Washington State University PercpCy5.5, Life technologies
M-T271 Human CD27, Biolegend Allophycocyanin or Pe-Cy7, Life technologies
G025H7 Human CXCR3 FITC, Biolegend Not applicable
HM-CCR5 Human CCR5 PerCP/Cy5.5, Biolegend Not applicable
Live Dead Aqua Dead cells, Invitrogen Not applicable
CellTrace Violet Not applicable, Life technologies Not applicable
Open in a new tab

STATISTICAL ANALYSIS

Results are expressed as mean ± standard errors of the mean. Statistical significance was deterTmined by one-way Analysis of Variance (ANOVA) followed by Bonferroni test, or Student’s t test using Prism software (GraphPad, La Jolla, CA).

RESULTS

Virulent M. bovis infection induces antigen-specific γδ T cell responses

We and others have previously demonstrated that γδ T cells from virulent M. bovis-infected cattle respond to both the complex mycobacterial antigen, PPD-b, and to the specific protein antigen, ESAT-6/CFP-10 (10, 11, 24). To corroborate our prior studies, animals were infected via aerosol inoculation with virulent M. bovis strain 10–7428 (n=10) or non-infected group (n=6). Following infection, PBMC were labeled with CellTrace dye and cultured for 6 days in the presence or absence of PPD-b or recombinant ESAT-6/CFP-10. On day 6, cells were analyzed by flow cytometry for the frequency of γδ and CD4 T cells that divided in response to the mycobacterial antigens (Supplemental Fig. 1). Representative proliferative responses of γδ and CD4 T cells following mycobacterial stimulation in vitro are depicted (Fig 1A and B). Robust proliferative responses to PPD-b and ESAT-6/CFP-10 were observed by γδ T cells at 4 weeks post infection and persisted until at least 12-weeks post infection (Fig. 1C). Consistent with prior reports, the mean CD4 T cell proliferative response to PPD-b and ESAT-6/CFP-10 peaked by 8 weeks post infection and persisted for at least 12 weeks post infection (Fig. 1D) (25). Recall responses from both γδ and CD4 T cell populations were M. bovis-specific, as neither CD4 nor γδ T cells from non-infected cattle responded to the specific or complex mycobacterial antigens.

Fig. 1. Proliferative responses from M. bovis-infected and non-infected cattle in response to in vitro restimulation with mycobacterial antigens.

Fig. 1.

Open in a new tab

PBMCs from uninfected (n= 6) or virulent M. bovis–infected animals (n= 10) were labeled with CellTrace, and 5 × 106 cells/ml were cultured for 6 days in the presence or absence of 200 UI/ml PPD-b or 2 μg/ml ESAT-6/CFP-10. Cells were labeled with anti-bovine γδ TCR or CD4 and analyzed by flow cytometry for CellTrace dilution (Supplemental Fig.1). Representative histograms of proliferative responses from a M. bovis-infected and control animal, gated on total live cells, lymphocytes (SSC-A vs FSC-A), and cells expressing the γδ TCR (A) or CD4 (B) Percentage of γδ T cells (C) and CD4 T cells (D) from M. bovis infected and uninfected animals that proliferated in response to mycobacterial antigens, as measured by CellTrace dilution, at 4-, 8-or 12-weeks post aerosol challenge. Analysis was performed with Flowjo software. Background proliferation was subtracted, and results represent change over mock. Data are mean ± SEM and are representative of one independent experiment.

Virulent M. bovis infection results in phenotypic changes on M. bovis-specific γδ and CD4 T cells

Similar to humans, bovine naïve and memory CD4 T cells can be distinguished based on the expression of CD45 isoforms. Naïve T cells express the high CD45RA isoform and upon antigenic recognition, T cells switch to the expression of the low CD45RO isoform (26, 27). In the bovine CD45RA and CD45RB isoforms are not yet defined; thus, CD45R is used as a marker for naïve lymphocytes in cattle (26). To evaluate if CD45 isoform expression correlated with the bovine γδ T cell effector/memory population that responded to mycobacteria, we analyzed the CD45 phenotype of expanding M. bovis-specific γδ and CD4 T cells by flow cytometry following a 6-day in vitro restimulation with PPD-b (Fig. 2A). In accordance with previous data (25, 28), the mycobacterial-driven proliferative responses to PPD-b were within the CD4+ CD45RO+ T cell subset, and the antigen-responsive cells downregulated CD45R expression compared to non-responding cells (Fig. 2B). In contrast to CD4 T cells, γδ T cells from M. bovis-infected cattle showed no significant changes in cell surface expression of CD45RO, between antigen-specific (proliferating) and non-responding γδ T cell populations. However, antigen-responsive γδ T cells from infected cattle downregulated CD45R expression in response to PPD-b stimulation (Fig. 2C). Changes in expression of either CD45R or CD45RO were not detected in γδ T cells from noninfected animals after 6 days of culture. From these results it can be concluded that CD45Rneg γδ and CD4 T cell subset participates in the in vitro recall responses against M. bovis.

Fig. 2. Evaluation of CD45RO and CD45R expression on M. bovis-specific γδ and CD4 T cells.

Fig. 2.

Open in a new tab

Approximately 8 weeks after aerosol challenge with M. bovis, PBMCs from M. bovis–infected animals (n= 10) were labeled with CellTrace, and 5×106 cells/ml were cultured for 6 days in the presence or absence of PPD-b. Cells were labeled with anti-bovine CD4, or γδ TCR; and CD45RO or CD45R expression by flow cytometry within proliferative subsets as determined by CellTrace dilution. (A) Gating hierarchy (gating sequence as depicted by the arrows): lymphocytes (gate 1), live cells (gate 2), γδ or CD4 T cells (gate 3), proliferating (1) and non-proliferating cells (2) and CD45RO or CD45R expression (gate 5). (B and C) The expression of CD45RO or CD45R was determined for individual T cell subsets CD4 T cells (B) or γδ T cells (C) or based on CellTrace staining intensity (i.e., bright or dim). Data are presented as means (± SEM) and are representative of one independent experiment. p values for differences from responding and non-responding fractions for the respective T cell subset (i.e., comparisons between black and grey bars for each graph) were as follows: *, p < 0.05; **, p < 0.01, p ***<0.001, p****<0.0001 as determined by Student’s t test.

Peripheral γδ T cells from M. bovis infected cattle possess an activated/memory phenotype based on CD27 expression

CD27, a costimulatory molecule, has been commonly used to identify stages of T cell differentiation (29). In humans, two subsets of γδ T cells develop in response to antigen stimulation and are identified based upon their expression of CD27. These subsets exhibit unique functions during mycobacterial infection that parallel the functions attributed to differentiated subsets of CD8 and CD4 T cells (15, 29). Moreover, flow-cytometric analysis of CD27 expression on circulating MTB-specific T cells can help to discriminate disease progression (16, 30).

CD27 expression has been reported in bovine PBMCs, however, its expression on bovine γδ and CD4 T cells has not been determined (31). Therefore, we performed ex vivo staining on PBMC from healthy cattle and observed that ~70% of CD4 T cells, and ~50% of γδ T cells express CD27 as measured by flow cytometry. Following mitogen stimulation, both CD4 and γδ T cells CD27+ displayer higher clonogenic potential (Fig. 3). Thus, CD27 expression was abundantly expressed on bovine peripheral T cells, and can be used to identify CD4 and γδ T cell subsets based on their capacity to proliferate (16, 32).

Fig. 3. CD27 expression on bovine γδ and CD4 T cells ex vivo and in response to mitogen.

Fig. 3.

Open in a new tab

Using flow cytometry, bovine PBMCs from healthy cows were analyzed for CD4 or γδ TCR expression. Gating hierarchy (gating sequence as depicted by the arrows): live cells (gate 1), lymphocytes (gate 2), γδ or CD4 T cells (gate 3). Each lymphocyte population was further analyzed for CD27 expression following mock or PWM stimulation for 6 days (dot-plots). Data are representative of n=12 animals from 6 independent experiments.

CD27 expression on bovine γδ T cells identifies effector/memory subsets following virulent M. bovis infection

To determine whether mycobacteria-specific T cells could be defined by the expression of CD27, we analyzed the expression on M. bovis-specific γδ and CD4 T cells following in vitro restimulation with PPD-b and ESAT-6/CFP-10 (Fig. 4A and B). After 6 days in culture with PPD-b or ESAT-6/CFP-10, the majority of antigen-specific γδ T cells from M. bovis infected animals proliferated and upregulated expression of CD27 compared to non-responding γδ T cells in the same culture (Fig. 4A and Supplemental Fig. 2). Similarly, CD27 expression was significantly greater on the antigen-specific CD4 T cell population from M. bovis-infected animals compared to non-specific CD4 T cells in the same culture (Fig. 4B). Both γδ and CD4 T cells from infected animals exhibited significant increases in CD27 expression compared cells from noninfected control calves. Thus, from our results, increased expression of CD27 correlates with the capacity for bovine γδ T cells to proliferate in response to mycobacterial antigens, suggesting that bovine γδ T cell effector/memory phenotypes are consistent with those reported for human γδ T cells (16).

Fig. 4. CD27+ expression correlates with robust antigen-specific responses to M. bovis.

Fig. 4.

Open in a new tab

Approximately 8 weeks after virulent M. bovis infection, PBMCs (5×106 cells/ml) were labeled with CellTrace violet and restimulated in vitro for 6 days in the presence or absence of PPD-b, ESAT-6/CFP-10 or left unstimulated. Cells were then labeled with anti-bovine CD4 or γδ TCR, and CD27. CellTrace dilution (proliferative responses) and CD27 cell surface expression was determined for (A) γδ T cells and (B) CD4 T cells (Supplemental Fig. 2). Data are presented as means ± SEM and are representative of one independent experiment.; n = 10. p values indicate differences between CellTrace dim and positive fractions for the respective T cell subset (i.e., comparisons between black and grey bars for each graph) as follows: *, p < 0.05; **, p < 0.01, p ***<0.001, p****<0.0001 as determined by Student’s t test.

M. bovis-specific γδ T cells exhibit a TCM phenotype based on CD27 and CD45R expression

Expression of CD45RA and CD27 defines four subsets of memory γδ T cells in humans (15). Here, we sought to determine if the combination of both markers could be used to identify subsets of antigen-specific γδ T cells in cattle (Supplemental Fig. 3). PBMCs from M. bovis-infected animals were stimulated in vitro with PPD-b for 6 days. As shown in Fig. 5A, M. bovis-specific γδ T cells can be divided in four subsets: naïve (CD27+CD45R+), TCM (CD27+CD45R−), TEM (CD27−CD45R−), TEMRA (CD27−CD45R+) following in vitro stimulation with PPD-b. CD27+CD45R− TCM γδ T cells exhibited a change in distribution and in absolute numbers relative to the other subsets following in vitro restimulation with mycobacterial antigens, followed by the CD27−CD45R− TEM γδ T cell subset (Fig. 5B and C). Previous reports have shown that antigen-specific bovine CD4 T cells can also be divided by their expression of CD27 and CD45R (25). To confirm these previous results and to validate our results of our γδ T cell analysis, we examined parallel cultures to determine the distribution of effector/memory CD4 subsets responding to M. bovis infection. As seen in Fig. 5D and E, after 6 days, the CD4 TCM and TEM subset exhibited the highest change in distribution and in absolute numbers relative to the other subsets following in vitro restimulation with mycobacterial antigens compared to non-responding CD4 T cells. Our results suggest that the TCM subset comprises the majority of the in vitro proliferative response to mycobacterium antigens for γδ T cells.

Fig. 5. Phenotype of TCM, TEM and effector γδ and CD4 T cells proliferating in response to M. bovis.

Fig. 5.

Open in a new tab

PBMCs were isolated from calves ~ 8 weeks after challenge with virulent M. bovis. Cells were stained with CellTrace dye and incubated with M. bovis PPD-b for 6 days. A flow cytometric-based proliferation assay was used to study CD45R and CD27 expression on M. bovis-specific CD4 and γδ T cells. (A) Stimulated PBMCs were analyzed for proliferative responses. Live CD4 and γδ T cells were gated based on response to M. bovis (i.e., proliferating cells) and analyzed for CD27 versus CD45R expression within CellTrace bright (i.e., non-proliferative fraction; black bars) or CellTrace dim (i.e., proliferative fraction; grey bars) fractions (Supplemental Fig 3). Relative distribution and absolute count measurements of γδ T cells (B and C) and CD4 T cells (D and E) to proliferative response to PPD-b within CD45R/CD27 defined cell populations (CD45R+CD27+; CD45R-CD27+; CD45R-CD27-; CD45R+CD27-). Data are presented as mean (± SEM) and are representative of one independent experiment, with n = 10. *, p < 0.05; **, p < 0.01, p ***<0.001, p****<0.0001 as determined by One-way ANOVA.

Increased expression of CD27 on bovine peripheral γδ T cells correlates with memory-type responses

TCM cells are highly proliferative, abundant in lymph nodes, and retain their expression of the homing receptors CCR7 and CD62L (33). The results described in Fig. 5 suggest that peripheral M. bovis-specific γδ T cells share functional similarities (i.e. high proliferative capacity) with central memory T cells (16). To address this hypothesis, we evaluated the expression of CD62L on peripheral M. bovis-specific γδ T cells. As seen in Fig. 6A, following a 6-day stimulation with PPD-b, CD27+ M. bovis-responsive γδ T cells co-expressed CD62L compared to γδ T cells that did not proliferate to M. bovis antigens. In contrast, M. bovis-specific CD27+ CD4 T cells were low for CD62L expression compared to non-M. bovis-specific CD4 cells. These data are consistent with previous descriptions identifying the majority of M. bovis-specific CD4 T cells in the peripheral blood as TEM cells (Fig. 6A, D and E) (25). Therefore, similar to human γδ T cells, M. bovis-responsive bovine γδ T cells display phenotypic and functional properties of TCM memory cells (Fig. 6B and C)(25, 33).

Fig. 6. Memory marker expression of CD27 and CD62L T cell subsets of M. bovis-infected cattle.

Fig. 6.

Open in a new tab

PBMCs were isolated from calves ~ 8 weeks after challenge with virulent M. bovis. Cells were stained with CellTrace dye and incubated with M. bovis PPD-b for 6 days. Flow cytometry was used to study both CD27 and CD62L expression on M. bovis-specific T cells. Live lymphocytes, CD4 or γδ T cells were gated based on responsiveness to M. bovis (i.e., proliferating cells) and analyzed for CD27 versus CD62L expression. (A) Representative dot plots from an M. bovis infected animal, gated on dividing and nondividing (non-specific) CD4 or γδ T cells. Data are presented as CD27 versus CD62L expression. Relative distribution and absolute count measurements of CD27 and CD62L expressing γδ T cell subsets (B and C) and CD4 T cell subsets (D and E) by dividing (dark bars) and nondividing (grey) cells are shown for all infected animals. Data are presented as mean (± SEM) and are representative of one independent experiment with n = 10. *, p < 0.05; **, p < 0.01, p ***<0.001, p****<0.0001 as determined by One-way ANOVA.

Bovine M. bovis-specific γδ T cells upregulate expression of the tissue-associated chemokine receptors CXCR3 and CCR5

Chemokine receptor expression is modulated during differentiation of both CD4 and γδ T cells (15, 33, 34). We hypothesized that antigen responsive γδ T cells would modify the expression of certain homing receptors that would allow cells to migrate to tissues and sites of inflammation. PBMCs from infected cattle were stimulated with PPD-b, as described above. As seen in Fig. 7A and 7B, M. bovis-responsive bovine γδ T cells express higher surface levels of both CCR5 (RANTES/macrophage inflammatory protein-1α/−1β receptor) and CXCR3 compared to non-responding γδ T cells. As seen in Fig. 7C, antigen-specific CD4 T cells also express higher levels of CCR5 and CXCR3 compared to nonresponsive CD4 T cells. The increased expression of CCR5 and CXCR3 on M. bovis-responsive CD4 and γδ T cells is expected to enable these populations to migrate into the inflamed tissue.

Fig. 7. Surface expression of chemokine receptors on M. bovis-specific peripheral γδ and CD4 T cells.

Fig. 7.

Open in a new tab

PBMCs were isolated from calves ~ 8 weeks after challenge with virulent M. bovis. Cells were stained with CellTrace dye and incubated with PPD-b for 6 days. Flow cytometry was used to study CXCR3 and CCR5 expression on M. bovis-specific CD4 and γδ T cells. Cells were surface stained and then analyzed by flow cytometry for CXCR3 and CCR5 expression. (A) Representative histograms of CXCR3 and CCR5 expression on antigen-responsive (blue solid line) and non-responsive CD4 and γδ T cells (red solid line). Cumulative results from live γδ (B) and CD4 T cells (C), gated based on response to M. bovis (i.e., proliferating cells) and analyzed for CXCR3 or CRR5 expression. The FMO control is shown in grey shaded histograms. Data are presented as mean ± SEM MFI expression of cell surface markers. Results are representative of one independent experiment with n=10. *, p < 0.05; **, p < 0.01, p ***<0.001 indicates a significant difference (p< 0.05) from antigen-responsive cells compared to non-responsive cells as determined by students t-test.

DISCUSSION

Following M. bovis infection, a marked in vitro expansion suggests that bovine γδ T cells can mount memory-like responses upon restimulation with mycobacterial antigen (Fig. 1) and [reviewed (35)]. However, to date, a combination of surface markers that effectively identify effector/memory-like subsets of bovine antigen-specific γδ T cells have not been reported. Here, we show that compared to CD4 T cells, proliferating M. bovis-specific γδ T cells do not significantly alter CD45RO expression after restimulation with mycobacterial antigens in vitro. Our results agree with reports showing that γδ T cells from human TB patients do not modulate CD45RO expression compared to γδ T cells from non-infected subjects (16, 17). Thus, γδ T cells may acquire a ‘pre-activated’ state early in their development and thus, CD45RO is not useful to identify antigen-experienced γδ T cells. In contrast to CD45RO expression, we found that proliferating γδ T cells from M. bovis infected cattle, downregulated the expression of CD45R following antigen restimulation in vitro compared to non-responsive γδ T cells (Fig. 2) (28, 36). These observations indicate that mycobacteria-specific γδ T cells that can be identified using CD45R.

In humans, two subsets of γδ T cells develop in response to antigen stimulation and are identified based upon their expression of CD27 (22). These subsets exhibit unique functions during mycobacterial infection that parallel the functions attributed to differentiated subsets of CD8 and CD4 T cells (15, 29). Moreover, flow-cytometric analysis of CD27 expression on circulating MTB-specific T cells can help to discriminate disease progression (30). Thus, assessing the changes in γδ T cell effector/memory phenotype based on CD27 might be helpful to monitor changes related to TB disease progress and vaccine-induce protection as previously reported (16, 18). Although the expression of CD27 in bovine cells has been reported before, its expression on bovine γδ and CD4 T cells has not been determined (31). We observed that in healthy adult cattle ~70% of CD4 T cells, and ~50% of γδ T cells express CD27. Importantly, we observed that a subpopulation of γδ +CD27+ effector/memory-like T cells from M. bovis infected cattle develop in response to Ag stimulation and exhibit high proliferative capacity after restimulation with either PPD-b or the M. bovis protein antigen, ESAT-6/CFP-10. Our observations are consistent with findings in humans, where patients with acute pulmonary TB, and BCG vaccinated individuals, demonstrate increases in circulating CD27+ Vγ9Vδ2+ T cells, and these cells exhibit enhanced proliferative activity upon stimulation with mycobacterial antigens (16, 37). Thus, our results suggest that CD27 expression may be potentially used to monitor γδ T cell responses to M. bovis infection in cattle.

Coexpression of cell surface markers CD45RA and CD27 has been used to functional segregate subsets of human and non-human primate γδ T cells (15, 16). In non-human primates, TEM (CD45RA-CD27+) Vγ9Vδ2 T cells expansion after a secondary M. tb challenge correlates with protection against fatal TB (17–19). Interestingly, there is a significant reduction in effector γδ cells (CD27− TEM and TEMRA cells) in patients with active infection, but the population rebounds after successful clinical treatment (38). Our data show that M. bovis-specific γδ T cells can be divided into four distinct subsets: naïve (CD27+CD45R+), TCM (CD27+CD45R−), TEM (CD27−CD45R−), TEMRA (CD27−CD45R+). Consistent with reports from humans, our data show that virulent M. bovis infection is associated with an expansion in primarily γδ TCM subset in the blood (Fig. 5) (39). Although effector functions were not assessed in this study, it has been shown that γδ T effectors develop functional defects and progressively lose the ability to produce IFNγ compared with γδ T cells from healthy tuberculin positive adults (40). Whether the loss of γδ T cell effectors impacts the ability to acquire memory T cell properties is not known. Together, our observations, coupled with reports from human TB patients, demonstrate a negative correlation between γδ T effector (CD27− CD45R−) phenotype and disease. Further studies should address the functional implications of these phenotypic changes following M. bovis experimental challenge and vaccine efficacy studies in the bovine.

Lastly, we show that M. bovis-specific bovine γδ T cells exhibited higher CCR5 and CXCR3 compared to non-responding γδ T cells. Similarly, a higher expression of CXCR3 and CCR5 is preferentially found on circulating γδ T cells from TB patients (38, 41). The expression of CCR5 and CXCR3 on M. bovis-specific γδ and CD4 T cells suggests that these cell populations are memory/effector T cells (15, 38, 41). Similar to our results, Blumerman et al. showed that bovine γδ T cells participating in recall responses against Leptospira antigens exhibited increased transcription of CCR5 and CXCR3 (42). γδ T cells are amongst the first cells to localize to the granuloma following M. bovis infection (43). The increased expression of CCR5 and CXCR3 on M. bovis-responsive γδ T cells presumably enables this population to migrate into the inflamed peripheral tissue. We did not investigate the functionality of CCR5 and CXCR3 on this population, however studies in human γδ T cells have confirmed their capacity to migrate towards the inflammatory chemokines RANTES and IP-10 (38). Importantly, the chemokine receptor profile of peripheral γδ T cells may differ from that of γδ T cells at the site of infection; thus, further studies will address the functionality of CXCR3 and CCR5 in bovine γδ T cells, and the expression of tissue homing receptors such as CXCR3 and CCR5 on cells within granuloma lesions of M. bovis-infected cattle.

In conclusion, our results show that, like humans, γδ T cells from M. bovis-infected cattle can be divided into phenotypic and functional subsets based upon their surface expression of CD27, CD45R, and the homing molecules, CXCR3 and CCR5. Further elucidation of the pathway of γδ T cell differentiation and the functional implications will contribute significantly to our ability to engage the γδ T cell response in effective immune responses to mycobacterial diseases of both humans and animals.

Supplementary Material

Supplemental Data

ACKNOWLEDGMENTS

We thank Maegan Rabideau for her excellent technical assistance.

Footnotes

CONFLICT OF INTEREST

The authors declare no conflicts of interest

REFERENCES

  • 1.World Health Organization. 2018. Global Tuberculosis Report 2018. 231. [Google Scholar]
  • 2.Waters WR, Palmer MV, Buddle BM, and Vordermeier HM. 2012. Bovine tuberculosis vaccine research: historical perspectives and recent advances. Vaccine. 30: 2611–2622. [DOI] [PubMed] [Google Scholar]
  • 3.Muller B, Durr S, Alonso S, Hattendorf J, Laisse CJ, Parsons SD, van Helden PD, and Zinsstag J. 2013. Zoonotic Mycobacterium bovis-induced tuberculosis in humans. Emerg Infect Dis. 19: 899–908. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Schiller I, Oesch B, Vordermeier HM, Palmer MV, Harris BN, Orloski KA, Buddle BM, Thacker TC, Lyashchenko KP, and Waters WR. 2010. Bovine tuberculosis: a review of current and emerging diagnostic techniques in view of their relevance for disease control and eradication. Transbound Emerg Dis. 57: 205–220. [DOI] [PubMed] [Google Scholar]
  • 5.Endsley JJ, Waters WR, Palmer MV, Nonnecke BJ, Thacker TC, Jacobs WR Jr., Larsen MH, Hogg A, Shell E, McAlauy M, Scherer CF, Coffey T, Howard CJ, Villareal-Ramos B, and Estes DM. 2009. The calf model of immunity for development of a vaccine against tuberculosis. Vet Immunol Immunopathol. 128: 199–204. [DOI] [PubMed] [Google Scholar]
  • 6.Waters WR, Palmer MV, Thacker TC, Davis WC, Sreevatsan S, Coussens P, Meade KG, Hope JC, and Estes DM. 2011. Tuberculosis immunity: opportunities from studies with cattle. Clin Dev Immunol. 2011: 768542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Diedrich CR, and Flynn JL. 2011. HIV-1/mycobacterium tuberculosis coinfection immunology: how does HIV-1 exacerbate tuberculosis? Infect Immun 79: 1407–1417. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Cooper AM, Solache A, and Khader SA. 2007. Interleukin-12 and tuberculosis: an old story revisited. Curr Opin Immunol. 19: 441–447. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Rhodes SG, Hewinson RG, and Vordermeier HM. 2001. Antigen recognition and immunomodulation by gamma delta T cells in bovine tuberculosis. J Immunol. 166: 5604–5610. [DOI] [PubMed] [Google Scholar]
  • 10.Smyth AJ, Welsh MD, Girvin RM, and Pollock JM. 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. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.McGill JL, Sacco RE, Baldwin CL, Telfer JC, Palmer MV, and Waters WR. 2014. Specific recognition of mycobacterial protein and peptide antigens by gammadelta T cell subsets following infection with virulent Mycobacterium bovis. J Immunol. 192: 2756–2769. [DOI] [PubMed] [Google Scholar]
  • 12.Price S, Davies M, Villarreal-Ramos B, and Hope J. 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. [DOI] [PubMed] [Google Scholar]
  • 13.Vesosky B, Turner OC, Turner J, and Orme IM. 2004. Gamma interferon production by bovine gamma delta T cells following stimulation with mycobacterial mycolylarabinogalactan peptidoglycan. Infect Immun. 72: 4612–4618. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Kabelitz D, Bender A, Prospero T, Wesselborg S, Janssen O, and Pechhold K. 1991. The primary response of human gamma/delta + T cells to Mycobacterium tuberculosis is restricted to V gamma 9-bearing cells. J Exp Med. 173: 1331–1338. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Dieli F, Poccia F, Lipp M, Sireci G, Caccamo N, Di Sano C, and Salerno A. 2003. Differentiation of effector/memory Vdelta2 T cells and migratory routes in lymph nodes or inflammatory sites. J Exp Med. 198: 391–397. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Gioia C, Agrati C, Casetti R, Cairo C, Borsellino G, Battistini L, Mancino G, Goletti D, Colizzi V, Pucillo LP, and Poccia F. 2002. Lack of CD27-CD45RA-V gamma 9V delta 2+ T cell effectors in immunocompromised hosts and during active pulmonary tuberculosis. J Immunol. 168: 1484–1489. [DOI] [PubMed] [Google Scholar]
  • 17.Shen Y, Zhou D, Qiu L, Lai X, Simon M, Shen L, Kou Z, Wang Q, Jiang L, Estep J, Hunt R, Clagett M, Sehgal PK, Li Y, Zeng X, Morita CT, Brenner MB, Letvin NL, and Chen ZW. 2002. Adaptive immune response of Vgamma2Vdelta2+ T cells during mycobacterial infections. Science. 295: 2255–2258. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Shen L, Frencher J, Huang D, Wang W, Yang E, Chen CY, Zhang Z, Wang R, Qaqish A, Larsen MH, Shen H, Porcelli SA, Jacobs WR Jr., and Chen ZW. 2019. Immunization of Vgamma2Vdelta2 T cells programs sustained effector memory responses that control tuberculosis in nonhuman primates. Proc Natl Acad Sci U S A. 116: 6371–6378. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Chen CY, Yao S, Huang D, Wei H, Sicard H, Zeng G, Jomaa H, Larsen MH, Jacobs WR Jr., Wang R, Letvin N, Shen Y, Qiu L, Shen L, and Chen ZW. 2013. Phosphoantigen/IL2 expansion and differentiation of Vgamma2Vdelta2 T cells increase resistance to tuberculosis in nonhuman primates. PLoS Pathog. 9: e1003501. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Meraviglia S, El Daker S, Dieli F, Martini F, and Martino A. 2011. gammadelta T cells cross-link innate and adaptive immunity in Mycobacterium tuberculosis infection. Clin Dev Immunol. 2011: 587315. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Dieli F, Ivanyi J, Marsh P, Williams A, Naylor I, Sireci G, Caccamo N, Di Sano C, and Salerno A. 2003. Characterization of lung gamma delta T cells following intranasal infection with Mycobacterium bovis bacillus Calmette-Guerin. J Immunol. 170: 463–469. [DOI] [PubMed] [Google Scholar]
  • 22.Buddle BM, Pollock JM, Skinner MA, and Wedlock DN. 2003. Development of vaccines to control bovine tuberculosis in cattle and relationship to vaccine development for other intracellular pathogens. Int J Parasitol. 33: 555–566. [DOI] [PubMed] [Google Scholar]
  • 23.Palmer MV, Waters WR, and Whipple DL. 2002. Aerosol delivery of virulent Mycobacterium bovis to cattle. Tuberculosis (Edinb) 82: 275–282. [DOI] [PubMed] [Google Scholar]
  • 24.Waters WR, Rahner TE, Palmer MV, Cheng D, Nonnecke BJ, and Whipple DL. 2003. Expression of L-Selectin (CD62L), CD44, and CD25 on activated bovine T cells. Infect Immun. 71: 317–326. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Maggioli MF, Palmer MV, Thacker TC, Vordermeier HM, and Waters WR. 2015. Characterization of effector and memory T cell subsets in the immune response to bovine tuberculosis in cattle. PLoS One. 10: e0122571. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Michie CA, McLean A, Alcock C, and Beverley PC. 1992. Lifespan of human lymphocyte subsets defined by CD45 isoforms. Nature. 360: 264–265. [DOI] [PubMed] [Google Scholar]
  • 27.Bembridge GP, MacHugh ND, McKeever D, Awino E, Sopp P, Collins RA, Gelder KI, and Howard CJ. 1995. CD45RO expression on bovine T cells: relation to biological function. Immunology. 86: 537–544. [PMC free article] [PubMed] [Google Scholar]
  • 28.Maue AC, Waters WR, Davis WC, Palmer MV, Minion FC, and Estes DM. 2005. Analysis of immune responses directed toward a recombinant early secretory antigenic target six-kilodalton protein-culture filtrate protein 10 fusion protein in Mycobacterium bovis-infected cattle. Infect Immun. 73: 6659–6667. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Hintzen RQ, de Jong R, Lens SM, Brouwer M, Baars P, and van Lier RA. 1993. Regulation of CD27 expression on subsets of mature T-lymphocytes. J Immunol. 151: 2426–2435. [PubMed] [Google Scholar]
  • 30.Streitz M, Tesfa L, Yildirim V, Yahyazadeh A, Ulrichs T, Lenkei R, Quassem A, Liebetrau G, Nomura L, Maecker H, Volk HD, and Kern F. 2007. Loss of receptor on tuberculin-reactive T-cells marks active pulmonary tuberculosis. PLoS One. 2: e735. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Sopp P, Kwong LS, and Howard CJ. 2001. Cross-reactivity with bovine cells of monoclonal antibodies submitted to the 6th International Workshop on Human Leukocyte Differentiation Antigens. Vet Immunol Immunopathol. 78: 197–206. [DOI] [PubMed] [Google Scholar]
  • 32.DeBarros A, Chaves-Ferreira M, d’Orey F, Ribot JC, and Silva-Santos B. 2011. CD70-CD27 interactions provide survival and proliferative signals that regulate T cell receptor-driven activation of human gammadelta peripheral blood lymphocytes. Eur J Immunol. 41: 195–201. [DOI] [PubMed] [Google Scholar]
  • 33.Sallusto F, Lenig D, Forster R, Lipp M, and Lanzavecchia A. 1999. Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature. 401: 708–712. [DOI] [PubMed] [Google Scholar]
  • 34.Caccamo N, Meraviglia S, Ferlazzo V, Angelini D, Borsellino G, Poccia F, Battistini L, Dieli F, and Salerno A. 2005. Differential requirements for antigen or homeostatic cytokines for proliferation and differentiation of human Vgamma9Vdelta2 naive, memory and effector T cell subsets. Eur J Immunol. 35: 1764–1772. [DOI] [PubMed] [Google Scholar]
  • 35.McGill JL, Sacco RE, Baldwin CL, Telfer JC, Palmer MV, and Waters WR. 2014. The role of gamma delta T cells in immunity to Mycobacterium bovis infection in cattle. Vet Immunol Immunopathol. 159: 133–143. [DOI] [PubMed] [Google Scholar]
  • 36.Howard CJ, Sopp P, Parsons KR, McKeever DJ, Taracha EL, Jones BV, MacHugh ND, and Morrison WI. 1991. Distinction of naive and memory BoCD4 lymphocytes in calves with a monoclonal antibody, CC76, to a restricted determinant of the bovine leukocyte-common antigen, CD45. Eur J Immunol. 21: 2219–2226. [DOI] [PubMed] [Google Scholar]
  • 37.Hoft DF, Brown RM, and Roodman ST. 1998. Bacille Calmette-Guerin vaccination enhances human gamma delta T cell responsiveness to mycobacteria suggestive of a memory-like phenotype. J Immunol. 161: 1045–1054. [PubMed] [Google Scholar]
  • 38.Glatzel A, Wesch D, Schiemann F, Brandt E, Janssen O, and Kabelitz D. 2002. Patterns of chemokine receptor expression on peripheral blood gamma delta T lymphocytes: strong expression of CCR5 is a selective feature of V delta 2/V gamma 9 gamma delta T cells. J Immunol. 168: 4920–4929. [DOI] [PubMed] [Google Scholar]
  • 39.Martino A, Casetti R, Sacchi A, and Poccia F. 2007. Central memory Vgamma9Vdelta2 T lymphocytes primed and expanded by bacillus Calmette-Guerin-infected dendritic cells kill mycobacterial-infected monocytes. J Immunol. 179: 3057–3064. [DOI] [PubMed] [Google Scholar]
  • 40.Swaminathan S, Gong J, Zhang M, Samten B, Hanna LE, Narayanan PR, and Barnes PF. 1999. Cytokine production in children with tuberculous infection and disease. Clin Infect Dis. 28: 1290–1293. [DOI] [PubMed] [Google Scholar]
  • 41.Eberl M, Engel R, Beck E, and Jomaa H. 2002. Differentiation of human gamma-delta T cells towards distinct memory phenotypes. Cell Immunol. 218: 1–6. [DOI] [PubMed] [Google Scholar]
  • 42.Blumerman SL, Wang F, Herzig CT, and Baldwin CL. 2007. Molecular cloning of bovine chemokine receptors and expression by WC1+ gammadelta T cells. Dev Comp Immunol. 31: 87–102. [DOI] [PubMed] [Google Scholar]
  • 43.Pollock JM, Pollock DA, Campbell DG, Girvin RM, Crockard AD, Neill SD, and Mackie DP. 1996. Dynamic changes in circulating and antigen-responsive T-cell subpopulations post-Mycobacterium bovis infection in cattle. Immunology. 87: 236–241. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental Data
NIHMS1059623-supplement-Supplemental_Data.pdf (340.4KB, pdf)

RESOURCES