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Department of Animal Sciences, The Ohio State University/Ohio Agricultural Research and Development Center, Wooster, Ohio 44691
ABSTRACT
Luteal cells are potent activators of T cell proliferation in vitro. The purpose of this study was to determine which subset of T cells is stimulated by luteal cells and whether luteal cell-induced T cell activation elicits a proinflammatory or anti-inflammatory T cell response. The first objective was to determine if luteal cell-stimulated T cell proliferation was mediated by class I or II major histocompatibility complex (MHC) molecules. T cell proliferation was inhibited by anti-MHC class I but not anti-MHC class II antibodies. The second objective was to determine which T cell subtype proliferates when cultured with luteal cells. The proportions of CD4+ and CD8+ cells were unchanged, but the number of gamma delta T cells was increased by coculture with luteal cells. Immunohistochemistry confirmed the presence of gamma delta T cells in midcycle and regressing corpus luteum. The final objective was to characterize T cell cytokine production stimulated by luteal cells. The concentrations of interferon-gamma (IFNG) and interleukin 10 (IL10) were increased in luteal cell-T cell cocultures, whereas IL4 was undetectable, and IL12 was barely detectable in culture medium. It was concluded that coculture of luteal cells and T cells resulted in activation of a somewhat unique T cell subset, gamma delta T cells, as well as production of both pro- and anti-inflammatory cytokines. To our knowledge, this is the first report of gamma delta T cell activation by luteal parenchymal cells of any species, raising the possibility that tissue-resident gamma delta T cells are involved in regulating the balance between tissue homeostasis and luteolysis.
corpus luteum, immunology, ovary
The bovine corpus luteum (CL) contains both lymphocytes and macrophages that are resident within the tissue. Interactions between these immune cells and the parenchymal cells (steroidogenic and endothelial) that constitute the bulk of luteal tissue may be important for both luteal regression and normal function of the CL. Luteal cells are capable of stimulating T cell proliferation in vitro, with cells obtained from regressing CL eliciting a greater T cell response than cells obtained from midcycle CL [1]. The first signal for T cell proliferation is the interaction of the major histocompatibility complex (MHC) molecules on the antigen-presenting cell with the T cell receptor [2]. Cells within the bovine CL express class I and II molecules [3], and the percentage of cells that express MHC class II molecules increases at the late stage of the luteal phase or after the administration of a luteolytic dose of prostaglandin (PG)F2
[4]. The proinflammatory cytokine, interferon-gamma (IFNG), stimulates an increase of class I and II MHC molecules on cultured bovine luteal cells [3], inhibits LH-stimulated progesterone production [5], and decreases luteal cell viability [6–8].
The CD4+ T cells form a complex with antigen-presenting cells via the MHC class II molecule [9], while the CD8+ T cells recognize antigens presented via MHC class I molecules [10, 11]. Both CD4+ and CD8+ T cells are present in the bovine CL during the luteal phase and luteolysis [12–14]. Addition of a class II MHC antibody cocktail attenuated luteal cell-stimulated T cell proliferation when the luteal cells were derived from regressing CL [1], suggesting that regressing luteal cells elicit a class II MHC-dependent T cell response in vitro by interacting with CD4+ T cells. To our knowledge, the possibility that CD8+ T cells are stimulated by luteal cells via class I MHC has not been addressed, nor have any studies examined a potential interaction between luteal cells and non-MHC-restricted T cells, the 
T cells. In fact, the localization of T cells in luteal tissue has not been previously reported. Unlike CD4+ and CD8+ cells, T cells that express the T cell receptor recognize cell surface molecules independently of antigen processing and MHC-mediated presentation [15–17]. The CD4+ T helper 1 (TH1) [18], CD8+ cytolytic [19], and T [20, 21] cells produce IFNG, but CD4+ TH2 cells as well as T cells produce anti-inflammatory cytokines such as interleukin (IL)10 and IL4. The overall objective of the present study was to determine which subset of T cells was stimulated by bovine luteal cells in vitro, if the response was MHC class I or II restricted, and which cytokines were produced during luteal cell-induced T cell activation.
Culture medium RPMI 1640, Ham F-12, L-glutamine, gentamicin, streptomycin, penicillin, and heat-inactivated fetal calf serum were purchased from Invitrogen Life Technologies (Carlsbad, CA). Collagenase was obtained from Worthington Biochemical Corp. (Lakewood, NJ). Ficoll-Hypaque Plus was purchased from Amersham Biosciences (Uppsala, Sweden). Antibodies specific for bovine cell surface molecules were purchased from VMRD Inc. (Pullman, WA) or ABD-Serotec (Raleigh, NC). Bovine immunoglobulin G (IgG) was obtained from Jackson ImmunoResearch Laboratories (West Grove, PA). Secondary antibodies conjugated with fluorescein isothiocyanate (FITC) or Cy5 were purchased from Caltag Laboratories (Burlingame, CA), and the phycoerythrin (PE)-conjugated antibody was purchased from Southern Biotechnology Associates, Inc. (Birmingham, AL). For immunofluorescence, FITC-conjugated secondary antibody was purchased from ABD-Serotec, and Alexa Fluor 546 was purchased from Invitrogen. The antibodies used for ELISA were obtained from Serotec. The IL4 ELISA kit and recombinant human IL12 were acquired from eBioscience (San Diego, CA). The ELISA specific for bovine IFNG was purchased from Biosource International (Camarillo, CA). The avidin-biotin-peroxidase kit, avidin-biotin blocking kit, and diaminobenzidine tetrahydrochloride (DAB) kit, as well as the mouse IgG used for immunohistochemistry, were purchased from Vector Laboratories, Inc. (Burlingame, CA). The 96-well culture plates were obtained from Corning (Corning, NJ). The [3H]thymidine was obtained from ICN Pharmaceuticals, Inc. (Costa Mesa, CA). All other reagents were purchased from Sigma-Aldrich Co. (St. Louis, MO).
Collection and Dissociation of Luteal Tissue
The procedure to collect CL was approved by the Institutional Laboratory Animal Care and Use Committee of The Ohio State University in compliance with National Institutes of Health guidelines. Cows that exhibited normal estrous cycles were used in the present study. Corpora lutea (n = 5) were collected during the midluteal phase (Days 10–12) of the estrous cycle (Day 0 = estrus), 8 h after a 25-mg i.m. injection of PGF2 (Lutalyse; Upjohn Co., Kalamzaoo, MI). The time of collection after PGF2 administration was chosen because it has previously been determined in this laboratory to be a time at which luteal cells very effectively stimulate T cell proliferation [1]. A blood sample was taken prior to and 8 h after PGF2 administration to determine if plasma progesterone concentrations declined in response to PGF2. Progesterone concentrations were measured by ELISA as previously described [1].
Luteal tissue was dissociated as previously described [22]. Briefly, luteal tissue was finely minced and placed in Ham F-12 medium containing gentamicin at 20 ng/ml, 0.5% BSA, and collagenase (2000 U/g of tissue). After a 1-h incubation at 37°C, medium containing the dispersed luteal cells was decanted and placed on ice. The remaining tissue was dissociated for an additional hour with new medium containing collagenase. After the second dissociation, luteal cells were washed three times in BSA-free medium and resuspended in RPMI 1640 medium containing 10% fetal calf serum. To prevent proliferation of luteal cells in culture, luteal cells (7 x 106) were pretreated with mitomycin C (50 µg/ml) for 20 min at 37°C and washed three times with RPMI 1640. Luteal cell number was determined by trypan blue dye exclusion. For all experiments, cells were cultured in RPMI 1640 containing 10% heat-inactivated fetal calf serum, 2 mM L-glutamine, 100 IU of penicillin, and streptomycin at 100 µg/ml.
After CL removal, blood was collected from the jugular vein into a sterile bottle containing acid citrate dextrose-A and placed on ice for transport to the laboratory. Blood was aliquoted into sterile glass tubes and centrifuged at 1000 x g for 10 min at 4°C. Peripheral blood mononuclear cells (PBMCs) were isolated from the lymphocyte-rich layer by centrifugation through Ficoll-Hypaque Plus as previously described [1].
T cells were purified from PBMCs by the MACS Cell Separation System (Miltenyi Biotec, Inc., Auburn, CA) as previously described [23]. Briefly, class II MHC-positive cells were labeled with a cocktail of mouse anti-bovine MHC class II monoclonal antibodies (TH14B, TH81A, and H42A; VMRD) and then incubated with paramagnetic MACS (magnetic cell sorting) rat anti-mouse IgG microbeads. The anti-MHC class II-labeled cells were retained within the MACS mass spectrometry column, over which the cells were passed, and the CD3+ lymphocytes were depleted and captured in the eluate. The purity of the cell population obtained by cell separation was 
96% for CD3+ cells, as determined by flow cytometry.
Experiment 1: Effect of blocking MHC class I and II on T cell proliferation. Mitomycin C-treated luteal cells (3.2 x 104) and autologous T cells (1 x 105) were cocultured for 72 h in the presence of 0, 1, 5, 10, or 20 µg/ml of anti-MHC class I (H58A), anti-MHC class II (TH14B, TH81A, and H42A), or a combination of these antibodies. To stabilize the interaction of the MHC molecules with the T cell receptor complex, staphylococcal enterotoxin B (SEB; 1 µg/ml) was included in all cultures, with the exception of the control luteal cell-T cell cultures. Other controls for this experiment included luteal cells or T cells in the presence of SEB and a nonspecific anti-bovine IgG antibody to ensure the specificity of any effect of anti-MHC class I or II antibodies. All treatments were performed in duplicate, and the experiment was replicated four times with luteal cells and autologous T cells from four separate animals. At 66 h of culture, 0.5 µCi [3H]thymidine was added to the culture wells. At the termination of the culture, the plate was placed at –80°C until the cells were harvested with a cell harvester (Skatron Instruments, Sterling, VA). Cells were washed twice prior to harvesting to remove unincorporated [3H]thymidine. Incorporation of [3H]thymidine, an indicator of cell proliferation, was measured by liquid scintillation counting.
Experiment 2: Effect of anti-CD4, anti-CD8, and anti- on T cell proliferation.
Mitomycin C-treated luteal cells (3.2 x 104) and T cells (1.0 x 105) were cultured in the presence of SEB (1 µg/ml) for 72 h in 96-well culture plates as described above. The cells were treated with antibodies specific for the CD4 coreceptor, the CD8 -chain, or the T cell receptor chain at 0, 1, 5, 10, or 20 µg/ml final concentration. The cells were pulsed with 0.5 µCi [3H]thymidine during the last 6 h of culture. Incorporation of [3H]thymidine was measured as previously described. Controls for this experiment included luteal cell-T cell cultures in the absence of SEB, luteal cells, and T cells cultured separately, both in the presence of SEB, and the addition of anti-bovine IgG. All treatments were performed in duplicate, and the experiment was replicated four times with luteal and autologous T cells from four separate animals.
Experiment 3: Determination of T cell types and cytokine production. Cultures containing T cells, T cells with SEB (1 µg/ml), or mitomycin C-treated luteal cells (3.2 x 104) and T cells (1.0 x 105) with SEB were incubated for 72 h in 96-well plates. After the 72-h incubation, medium containing the nonadherent T cells was removed from the adherent luteal cells. The medium was centrifuged for 10 min at 400 x g at 4°C, decanted, and stored at –80°C until assayed for cytokine content. The T cells were resuspended in PBS (4°C) and prepared for flow cytometry. All treatments were performed in duplicate, and the experiment was replicated five times with luteal and autologous T cells from four or five separate animals.
T cells were prepared for three-color flow cytometric analysis by indirect immunofluorescence. All incubations with antibodies and washes were performed at 4°C. To determine the purity of the cell population analyzed by flow cytometry, the cells were labeled with the antibody to CD3, which is present on CD4+, CD8+, and T cells. Cells were incubated with mouse anti-bovine CD3, mouse anti-bovine CD4, and mouse anti-bovine chain or with mouse anti-bovine CD3, mouse anti-bovine CD8, and mouse anti-bovine chain (2.5 µg/ml each) for 30 min, washed with PBS (4°C), and incubated for 30 min with the appropriate secondary antibodies. Goat anti-mouse IgG1 conjugated to PE (62.5 ng/ml) was used to detect CD3+ T cells; goat anti-mouse IgG2a conjugated to FITC (1 µg/ml) was used to detect CD4+ or CD8+ cells; and goat anti-mouse IgG2b conjugated to Cy5 (1 µg/ml) was used to detect T cells. The concentration for each secondary antibody that gave the least background was determined prior to analysis. Following incubation with the secondary antibody, cells were washed twice, fixed in 0.5% paraformaldehyde, and analyzed within 3 days. Flow cytometric data were collected with a FACSCalibur and analyzed by the CellQuest program. Prior to each analysis, unlabeled T cells were used to determine autofluorescence, and each secondary antibody was analyzed separately to compensate for spectral overlap. Two separate cell populations were detected by forward and side scatter properties that were considered smaller resting T cells or larger activated T cells. A total of 10 000 cells were analyzed for the region of activated T cells determined by forward and side scatter properties.
Interferon-Gamma ELISA. Medium was collected from cultures containing T cells plus luteal cells with SEB at 48 h of culture to assess if cytokine production was different from 48 to 72 h. The concentration of IFNG in culture supernatants was measured with a commercially available ELISA specific for bovine IFNG according to the procedure supplied by the manufacturer (Biosource). Briefly, 100 µl of sample was added to the anti-bovine IFNG-coated well and incubated for 1 h at room temperature. The wells were washed three times, and 100 µl of anti-bovine IFNG horseradish peroxidase conjugate solution was added. After the incubation with conjugate, the wells were washed as before, and IFNG was detected with 3,3',5,5'-tetramethylbenzidine chromagen solution. Values of duplicate samples are given as the mean optical density (OD) value (450 nm). IFNG was measured in luteal cell-T cell cocultures from five separate animals.
Interleukin 4 ELISA. A recombinant human IL4 ELISA assay (eBioscience) was used, because neither a kit nor antibodies specific for bovine IL4 were available. Plates were coated with anti-human IL4 antibody overnight at 4°C. Standards and culture supernatants (100 µl) were added to the appropriate wells and incubated for 1 h at room temperature. The standard curve was generated from serial dilutions of rhuIL4. After washing, 100 µl of biotin-conjugated anti-human IL4 was added to the wells and detected with avidin-horseradish peroxidase. Concentrations of IL4 are reported as picograms per milliliter. Because it has not been reported if the rhuIL4 kit used in the present study would cross-react with bovine IL4, medium collected from Concanavalin A (ConA)-stimulated bovine PBMCs was used as a control. The PBMCs were isolated as previously described. The PBMCs (1.0 x 106) were treated with ConA (1 mg/ml) for 48 h, at which time the medium was collected. Serial dilutions of medium collected from ConA-stimulated PBMCs and unstimulated PBMCs were included as positive controls to assess cross-reactivity.
Interleukin 10 ELISA. The concentration of IL10 in culture supernatants was measured with a pair of monoclonal antibodies that recognize bovine IL10 [24]. Plates were coated with 6 µg/ml of mouse anti-bovine IL10 capture antibody (CC318) in coating buffer overnight at room temperature. Plates were washed four times with PBS containing 0.01% Tween 20 (v:v) after each incubation. All samples and buffers were added in 100-µl aliquots. Nonspecific binding sites were saturated with PBS containing sodium casein (1 mg/ml) for 1 h. Culture supernatants were added to the appropriate wells in duplicate. Because rboIL10 was commercially unavailable and rhuIL10 did not cross-react with the antibodies specific for bovine IL10, serial dilutions of medium from ConA-stimulated PBMCs were used to generate a standard curve. Biotin-conjugated mouse anti-bovine IL10 (CC320; 2.0 µg/ml) was added and incubated at room temperature for 1 h. The streptavidin horseradish peroxidase enzyme (1:500) was used for the detection of IL10. Values are reported as the mean OD (450 nm). IL10 was measured in luteal cell-T cell cocultures from five separate animals.
Interleukin 12 ELISA. Concentrations of IL12 in culture supernatants were measured by an ELISA procedure as described above, except that individual samples were analyzed, because there was insufficient remaining culture medium to measure concentrations of IL12 in duplicate. The standard curve (nanograms per milliliter) was generated from serial dilutions with rhuIL12. The mouse anti-bovine IL12 capture antibody (CC301; 8.0 µg/ml) and mouse anti-bovine IL12 biotin-conjugated detection antibody (CC326; 8.0 µg/ml) have been reported to react with bovine IL12 in an ELISA [25] and cross-react with the rhuIL12. The streptavidin horseradish peroxidase enzyme (1:500) was used for the detection of IL12. The IL12 antibodies recognize IL12B and the IL12 heterodimer (IL12A+IL12B), and so the protein recognized in these culture samples is herein referred to as IL12. The IL12 assay was performed on media collected from luteal cell-T cell cocultures with autologous cells from three separate animals.
Immunohistochemical Localization of T Cells in CL
An immunoperoxidase system was used to detect T cells in luteal tissue during the midluteal phase and 8 h after PGF2 administration. Immediately after removal, luteal tissue was sliced into small pieces and frozen over chilled isopentane in OCT. Frozen sections (7 µm) were fixed in ice-cold 95% ethanol. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide in methanol for 10 min on ice. All remaining incubations were performed at room temperature. Nonspecific binding sites were saturated with a 10% dilution of normal goat serum in PBS for 1 h. Endogenous biotin was saturated with avidin solution for 15 min and then incubated with the biotin solution for 15 min, prior to addition of the primary antibody. A monoclonal antibody specific for the µ chain of the T cell receptor (GB21A; 1:500) was used to examine the presence of T cells in luteal tissue. This antibody is specific for two populations of bovine T cells having the phenotype WC1+CD2–CD4–CD8– and WC1–CD2+CD8+/– [26, 27]. Substitution of the primary antibody with mouse IgG (1:500) was used as a negative control. After incubation with the primary antibody (1 h), sections were treated with biotinylated anti-mouse IgG (1:200) for 30 min and then incubated for 30 min with avidin and biotinylated horseradish peroxidase H solution. Localization of peroxidase was determined with DAB as the peroxidase substrate. Sections were counterstained with hematoxylin and viewed by light microscopy.
Plasma progesterone concentrations before and after PGF2 administration were compared by a paired t-test. Cell proliferation and flow cytometry data were analyzed by a two-way analysis of variance followed by the Tukey honestly significant difference test. Cell proliferation data were normalized with the square-root transformation function. Mean OD values for IFNG and IL10 were analyzed by a two-way analysis of variance, and all pairwise comparisons were determined by the Tukey honestly significant difference test. Mean OD values for IFNG were presented as log units. All statistical analyses were performed by SigmaStat software (Jandel Corporation, San Rafael, CA) or SAS (Statistical Analysis Systems Institute, Cary, NC).
Although the standard luteolytic dose was used, the PGF2 injection effectively decreased plasma progesterone concentrations in only two of the five cows used in this study. The experiment had been designed to use regressing luteal cells, because T cell proliferation in coculture is greater following PGF2 compared with no PGF2 treatment in vivo [1]. Separating the present data on the basis of the in vivo response to PGF2 did not show obvious differences; hence, data from all cows were combined, regardless of progesterone status. Consistent with previous data from this laboratory [1], minimal proliferation occurred when T cells were cultured alone, but T cell proliferation was significantly stimulated by coculture with luteal cells (20 843 ± 6003 vs. 149 310 ± 56 710 disintegrations per minute, T cells alone vs. T cells + luteal cells, respectively). To determine if T cell proliferation was induced via MHC class I or II, increasing concentrations of mouse anti-bovine MHC class I and mouse anti-bovine MHC class II antibodies were added to the cocultures. Proliferation of T cells was attenuated by the addition of anti-MHC class I antibody at 5 or 20 µg/ml but was not affected by any concentration of anti-MHC class II antibody (Fig. 1). The combination of anti-class I and anti-class II antibodies at concentrations of 10 or 20 µg/ml decreased T cell proliferation, but T cell proliferation was never completely inhibited by antibodies to MHC. Addition of nonspecific IgG at concentrations the same as for the MHC antibodies had no effect on T cell proliferation (data not shown).
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One approach to determine which T cell population was stimulated to proliferate when cultured with luteal cells was to use antibodies to compete with the CD8 or CD4 T cell coreceptors, which interact with the MHC class I and II molecules, respectively. Although the lower concentrations of mouse anti-bovine CD4 and CD8 did not inhibit T cell proliferation, each of these antibodies at a concentration of 20 µg/ml effectively decreased luteal cell-induced T cell proliferation (Fig. 2). Addition of mouse anti-bovine chain antibody had no effect on luteal cell-induced T cell proliferation (Fig. 2).
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The proliferative response of T cell subsets after 72 h of coculture with luteal cells was assessed by flow cytometry. Representative scatter diagrams for one CL are shown in Figure 3. CD4+ and CD8+ cells are those in the right two quadrants in columns 1 and 2, respectively. The T cells are those in the upper two quadrants in column 2. For quantitative analyses, the number of cells that were positive for either CD4, CD8, or per 10 000 cells analyzed was compared among cultures of T cells alone (TC), T cells + luteal cells (TC+LC), and similar cultures with the addition of SEB (TC+SEB; TC+LC+SEB). Coculture with luteal cells or the addition of SEB had no effect on the number of CD4+ or CD8+ T cells per 10 000 total cells (Fig. 4, panels A and B, respectively). The proportion of CD4+ cells was less in cultures that contained TC+LC+SEB than in cultures that contained T cells alone (P < 0.05; Fig. 4, panel A) but was not different from its respective controls (TC+LC, TC+SEB). In contrast, the proportion of T cells was greater when T cells were cultured with luteal cells than with T cells alone (P < 0.05; Fig. 4, panel C), and this effect of luteal cells on T cell proliferation occurred in both the presence and absence of SEB. The majority of T cells in the cultures were CD8–, and it was these cells that proliferated in response to luteal cells. The very small number of CD8+ T cells in the cultures did not proliferate in response to luteal cells, although there was a stimulatory effect of SEB on these cells (Fig. 4, panel D). Indirect immunofluorescence imaging was used to visualize CD4+ and T cells after coculture with luteal cells (Fig. 5). When T cells were cultured alone, cells and CD4+ were similar in size (Fig. 5A). However, after coculture with luteal cells, the T cells were larger in size than the CD4+ cells (Fig. 5B), indicating that these cells had been stimulated by the luteal cells. An increase in size of the proliferating cells, as assessed by forward scatter properties of the cells in flow cytometry, was significant (P < 0.05) in the LC+TC+SEB compared with the TC+SEB cultures in the absence of luteal cells. Side scatter, a reflection of granularity of the T cells analyzed, was also increased (P < 0.05) after culture with luteal cells (Fig. 6). Because there were no previous reports of T cells in the CL of any species, immunohistochemistry was used to determine if T cells were present in the bovine CL. The T cells were identified in the bovine CL collected on Day 10 of the estrous cycle and 8 h post-PGF2 injection (Fig. 7).
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Cytokine production by activated T cells may be used to determine whether the nature of the T cell response is proinflammatory or anti-inflammatory. All T cell cultures produced detectable concentrations of the proinflammatory cytokine, IFNG, after 72 h of culture (Fig. 8). Further, IFNG production was stimulated by coculture with luteal cells (TC vs. TC+LC; TC+SEB vs. TC+LC+SEB) or by SEB (TC vs. TC+SEB; TC+LC vs. TC+LC+SEB). Analysis of culture medium collected at 48 h from separate culture wells with TC+LC+SEB indicated there was no increase in IFNG concentration in the medium from 48 to 72 h of culture (2.386 ± 0.158 vs. 2.646 ± 0.158 OD value). The other proinflammatory cytokine measured, IL12, was detectable in some, but not all, cocultures. Although the numeric means appear to indicate increased IL12 when T cells alone were cultured with luteal cells, the high degree of variability in IL12 production resulted in a lack of significant differences (Fig. 8).
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IL4 was not detected in any of the T cell cultures, despite clear stimulation of IL4 production in ConA-treated PBMCs (data not shown). IL10 was detected in all of the T cell cultures but was elevated in the TC+LC+SEB cultures compared with all others (Fig. 8). As with IFNG, there was no difference in the amount of IL10 between 48 and 72 h of culture in medium collected from T cells cultured with luteal cells in the presence of SEB (0.30 ± 0.04 vs. 0.356 ± 0.04 OD value).
The results of the present study support previous reports from our laboratory that luteal cells can elicit a T cell response, manifested as a proliferation of T cells in an in vitro assay [1, 23]. Superantigens, including SEB, are produced by Staphylococcus aureus and are characterized by their ability to stabilize the interaction of MHC class II and the Vβ segment of the β T cell receptor [28]. Superantigens bypass antigen specificity by binding outside the MHC peptide groove, stimulating large numbers of T cells [29]. Luteal cells stimulate T cell proliferation in either the absence or presence of SEB; however, because luteal cell-induced T cell proliferation was greater in the presence of SEB [1], it was hypothesized that T cell activation was primarily a result of interaction with MHC molecules expressed on the surface of the luteal cells.
To determine if T cell proliferation induced by luteal cells was MHC class I or II mediated, antibodies specific for these molecules were added to the cultures to block the interaction between luteal cells and T cells. The addition of anti-MHC class I attenuated but did not completely inhibit luteal cell-induced T cell proliferation, implying that a small percentage of the cells were class I MHC restricted, presumably CD8+ cells. The addition of anti-MHC class II antibodies did not inhibit T cell proliferation in the present study. In previous studies from this laboratory, expression of class II MHC molecules was increased in regressing compared with midcycle CL [4], and in vivo exposure to PGF2 enhanced the ability of luteal cells to stimulate T cell proliferation [23]. Further, antibodies to MHC class II attenuated T cell proliferation when luteal cells were obtained from regressing, but not midcycle, CL [1]. Because PGF2 injection did not result in a decrease in plasma progesterone concentrations in all animals in the present study, it is likely that the luteal cells obtained from these animals were more representative of midcycle, fully functional luteal cells, and the lack of response to class II antibody was therefore consistent with previous results. The lack of inhibition of T cell proliferation by the MHC class II antibody in midcycle cells raises the interesting possibility that cells from the midcycle CL activate a different type of T cell (presumably cells other than CD4+ cells) than do cells from the regressing CL.
In the second experiment, antibodies specific for the T cell coreceptors, CD4, CD8, or the chain of the T cell receptor, were added to the cocultures in an effort to prevent interaction of the coreceptor with the MHC molecule on the antigen-presenting cell. Again, only the highest concentration of antibody suppressed CD4+ or CD8+ cells, and cell proliferation was not inhibited by any concentration of the chain antibody. However, the greatest inhibition was observed in the presence of the CD8+ antibody, consistent with the results of experiment 1 with anti-MHC class I.
Data obtained with antibodies to inhibit the interaction of cell surface molecules must be interpreted with some caution, because the antibodies themselves can sometimes elicit signaling through the cell surface receptor and hence a response. Also, the antibodies may not prevent interaction of the coreceptor with the stimulating cell, causing a false-negative response, as perhaps observed with the anti- chain antibody. Because the results obtained with the antibody blocking approach were not entirely convincing, the proportion of T cell types that existed in the cultures following stimulation by luteal cells was next assessed by flow cytometry. No difference was observed in the proportion of CD4+ T cells after exposure to luteal cells, consistent with the results of experiments 1 and 2 with MHC class II or CD4+ blocking antibodies. The TC+LC+SEB group had a lesser proportion of CD4+ cells than did the T cell-alone group, but this is not really a relevant comparison to make. Similarly, the proportion of CD8+ T cells was not altered by the presence of luteal cells.
Whereas there was no effect of luteal cells on the proportion of CD4+ or CD8+ cells, the stimulatory effect of luteal cells on T cell proliferation was clearly manifested as an increase in the proportion of T cells, regardless of the presence or absence of SEB. The stimulation of T cells was also apparent under visual observation, with the increased size of the cells, representative of BLAST cells. Flow cytometric analysis detected not only an increase in cell size, but also an increased granularity after coculture with luteal cells, perhaps due to the stimulation of cytokine production and increased secretory granules in the cells. The T cells are a unique population of T cells that do not require antigen processing or presentation via MHC molecules to be activated. It has been proposed that these cells recognize stress-induced self-antigens defined as MHC class I-related molecules, MICA/MICB [15–17], and nonclassical MHC class Ib [30], as well as nonpeptidic molecules such as phosphoantigens and lipids [31]. T cells have received far less attention than β cells, primarily because they compose a minor percentage of circulating T cells in human adults. However, cows have proportionately more T cells than do humans or mice [32, 33], and T cells are more commonly found within tissues of most species, where it is believed they contribute as much to immunoregulation and tissue repair as to immunoprotection against pathogens. A distinct subset of cells composes about one-half of all T cells within reproductive epithelia [34, 35]. These cells are found within the reproductive tract and decidua and are thought to be important for nonrejection of the fetal allograft. In fact, cells within the reproductive tract are increased nearly 100-fold in pregnant animals compared with nonpregnant animals [36], and greater numbers of cells are found in the peripheral blood of healthy pregnant women compared with nonpregnant women or recurrent aborters [37]. After observation of cell stimulation by luteal cells, it was of interest to determine if cells were present in the CL. Although they were sparsely scattered throughout the tissue amid the parenchymal cells, it was evident that T cells were present within the bovine CL. Interactions between luteal parenchymal cells and T cells might determine if lymphocytes within the tissue exist in a proinflammatory or regulatory state.
The response of T cells to luteal cells observed in this study was unexpected but raises intriguing questions about the potential role of these cells in the regulation of normal luteal function, particularly because a number of distinct subpopulations of cells with diverse functions exist. A unique cell surface receptor referred to as workshop cluster (WC) 1 has been described in ruminant T cells. T cells that express WC1 but that are negative for CD2, CD4, and CD8 were predominantly in the peripheral blood, whereas WC1–/CD2+/CD8+/– cells tended to be localized in tissues [38]. WC1+/CD8– cells express genes consistent with inflammatory functions, whereas genes associated with regulatory functions are more frequently expressed in WC1–/CD8+ cells [39–41]. The latter cell type was found with greater frequency within tissues than in blood. Luteal cells stimulated the proliferation of CD8–, but not CD8+, T cells in the present study, but it should be noted that relatively few CD8+ T cells were placed into the cocultures; thus, these data should be interpreted with caution. Constitutive expression of CD25, a marker for T-regulatory cells, has also been described on cells [42]. As T-regulatory cells, these cells produce the immunosuppressive molecules IL-10 and transforming growth factor β, and their depletion results in increased tumoricidal activity of cytolytic T cells and natural killer cells [43]. To add to the complexity of T cell functional subsets, there are three well-known variants of the WC1 molecule (WC1.1, WC1.2, and WC1.4), and Rogers et al. [44] found 20 biochemically distinguishable WC1 spots by two-dimensional Western blotting as well as 13 different cDNA intracytoplasmic tail sequences. This demonstrates the remarkable diversity of T cell subtypes and functions. Current efforts in this laboratory are to determine which class of T cells exist in the bovine CL and are stimulated by luteal cells in vitro.
The antibody blocking experiments indicated that some CD8+ cells and perhaps even CD4+ cells were activated by luteal cells, whereas the flow cytometry data demonstrated that luteal cells stimulated only the proliferation of T cells. Functional interactions exist between cells and β (CD4+ and CD8+) cells. Chiodini and Davis [45] have reported the nonresponsiveness of CD4+ T cells that was mediated by the presence of T cells. In the presence of bacterial antigens, cells suppress CD4+ T cells [46–48]. It is also possible that interaction with luteal cells caused CD4+ and/or CD8+ cells to produce cytokines that stimulated cell proliferation, without proliferating themselves. This would explain the decrease in T cell proliferation in the presence of antibodies to CD4, CD8, or MHC molecules, causing an indirect inhibitory effect on cell proliferation.
Stimulation of T cells by luteal cells resulted in a clear increase in IFNG as well as a significant increase in IL10. Interferon-gamma and IL10 have opposing functions; IFNG is a proinflammatory cytokine, whereas IL10 is anti-inflammatory and immunosuppressive. Activated CD4+ TH1 cells [18], CD8+ T cells [19], and T cells [20, 21] produce IFNG. Fikri et al. [49] reported that activated T cells express IFNG but not IL10; however, some subsets of cells produce IL10 [50]. Distinct clonal populations of CD4+ T cells have been thought to produce either IFNG or IL10 [51, 52], but some CD4+ T cells can coexpress these cytokines [52]. Because luteal cells stimulated the production of both IFNG and IL10, it is not yet possible to characterize the T cell response to luteal cells as inflammatory or regulatory. Because IL4 was not detectable in the cultures and the production of IL12 was highly variable, these cytokines were not useful indicators of the functional status of the T cells. IL12 is typically produced by monocytes, macrophages, and dendritic cells, but there have also been reports of IL12 production by nonimmune cells [53, 54], as well as a role for intrinsic renal cell-derived IL12 in renal inflammation [55]. The source of IL12 in the T cell-luteal cell cocultures may have been from a few contaminating macrophages or dendritic cells, although the purity of the cell preparations used might argue otherwise, raising the intriguing possibility that IL12 is produced by parenchymal cells within the CL. Studies with purified populations of T cell subsets, expression of additional markers that distinguish cell function, intracellular staining of specific cells for cytokines, and temporal changes in cytokine secretion will provide more insight into the nature of the luteal cell-T cell interaction.
In conclusion, it was demonstrated that the coculture of luteal cells and T cells results in the activation of a somewhat unique T cell subset, the T cells. To our knowledge, this is the first report of the existence of T cells within luteal tissue and their activation by luteal parenchymal cells of any species. Although it was surprising to find that CD4+ and CD8+ cells did not proliferate in response to luteal cells, it might be inferred from the experiments with antibodies that those cells were involved in a paracrine manner with the proliferation response of the T cells. Because both a proinflammatory (IFNG) and an anti-inflammatory (IL10) cytokine were increased when T cells were cultured with luteal cells, it remains unknown whether luteal cells elicit primarily a TH1- or a TH2-type response. Future studies are required to determine if T cells have a role in the maintenance of tissue homeostasis within the CL or participate in an inflammatory response during luteal regression.
ACKNOWLEDGMENTS
The authors thank Ms. Jodi Winkler for her assistance in collecting corpora lutea and Mr. Justin Fear for his assistance with fluorescence imaging.
FOOTNOTES
1Supported by NIH (National Institutes of Health) research grant HD37550 and National Research Initiative Competitive Grant no. 2004-35203-14789 from the USDA Cooperative State Research, Education, and Extension Service, Animal Reproduction Program to J.L.P. Salaries and research support also provided by state and federal funds appropriated. 
Correspondence: 2FAX: 330 263 3949; e-mail: pate.1{at}osu.edu
3Current address: Animal Reproduction and Biotechnology Laboratory, Department of Biomedical Sciences, Colorado State University, Fort Collins, CO 80523.
Received: 28 December 2006.
First decision: 5 February 2007.
Accepted: 17 August 2007.
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