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The Journal of Immunology, 2007, 178, 5524 -5532
Copyright © 2007 by The American Association of Immunologists, Inc.
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CD56 Marks an Effector T Cell Subset in the Human Intestine1

Offer Cohavy and Stephan R. Targan2

Cedars-Sinai Inflammatory Bowel Disease Center, Los Angeles, CA 90048


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
T cells are key mediators of intestinal immunity, and specific T cell subsets can have differing immunoregulatory roles in animal models of mucosal inflammation. In this study, we describe human CD56+ T cells as a morphologically distinct population expressing a mature, nonproliferative phenotype that is frequent in the gut. Enhanced potential for IFN-{gamma} and TNF synthesis suggested a proinflammatory function, and we directly demonstrate effector function mediated by direct T-T interaction with responder cells in vitro. CD56+ T cells from peripheral blood responded to the gut-related CD2 signal, and were necessary for effective CD2-mediated proliferation of peripheral blood CD56– T cells. Our findings associate CD56+ T cells with the intestinal immune compartment and suggest a putative effector function in human mucosal immunity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Analysis of human genetics and mouse models of mucosal inflammation indicated perturbations at the regulation of innate, as well as adaptive, immune responses (1) and point to a complex multigene pathology (2), although specific mechanisms of pathogenesis are yet to be discovered. The intestinal immune compartment is tightly regulated and its antigenic repertoire is independently shaped to accommodate the heavy antigenic load characteristic of the gut environment (1, 3). In inflammatory bowel disease (IBD),3 tolerance to intestinal Ags is perturbed (4, 5), and strong evidence implicates a skewed T cell-mediated Th1 response in Crohn’s disease (CD) (6), as well as mouse models of IBD (7). Contrary to the periphery, intestinal T cells largely express CD45RO, reflective of Ag-driven differentiation (8). Yet, TCR-mediated response to antigenic stimulation is attenuated, pointing to gut-specific mechanisms of T cell signaling and activation (9). Following success in other autoimmune disease, soluble TNF and TNF-expressing T cells were targeted by anti-TNF therapy with benefit to a subset of CD (10), and ulcerative colitis (UC) patients (11). However, the partial success of blocking TNF emphasized the complexity of mucosal immune regulation and demonstrated the limits of targeting one T cell subset in the mucosal compartment.

The role of specific effector cell subsets in mucosal immune regulation has been extensively investigated in the mouse. Analysis of adaptive cell transfers into SCID mice revealed a CD4+/CD45RBhigh T cell subset with a pathogenic proinflammatory effector function, and a second CD4+/CD25+ regulatory T cell subset (12, 13). Cong et al. (14) further established a requirement for effector T cell "priming" by intestinal commensal bacteria, and recent studies reported key roles for B and NKT cells in the maintenance of intestinal immune homeostasis (15) and immune suppression (13). In humans, several functionally distinct T cell populations have been reported in the gut mucosa, such as T cells expressing NK markers (16). For instance, intestinal T cells expressing CD161 were recently reported to express proinflammatory cytokines (17), whereas a second population of mucosal T cells expresses CD56 (18), a marker which may identify proinflammatory T cells in the liver (19, 20).

CD56, also known as neural cell adhesion molecule (21), D2-CAM (22), Leu19 (23), or NKH-1 (24) is the first characterized member of the Ig superfamily of cell adhesion molecules, which now also includes cadherins, selectins, and integrins (25). Initially, CD56 was identified in the nervous system, where it mediates cell-cell interactions of neural cells during embryogenesis (21, 22), and recently proposed as a receptor for glial cell line-derived neurotrophic factor (26). In addition, CD56 expression was reported in a variety of normal and abnormal tissues including skin, muscle, small cell carcinoma, neuroblastoma, neurons, astrocytes, Schwann cells, NK cells, and a subset of T cells or T cell lymphomas. CD56 is a calcium-independent adhesion molecule (27), primarily mediating homophilic binding to CD56 on adjacent cells (21). Although mediating NK-target cell contact is a likely function in some contexts (28), CD56 does not appear essential for cell-mediated cytotoxicity (29), and its role in immune regulation remains an enigma.

Within the hemopoietic lineage, CD56 serves as an important marker defining functionally distinct subsets of NK and T lymphocytes. In the NK compartment, bright CD56 staining (CD56bright) defines elevated potential for cytokine production (30), whereas weaker fluorescence intensity (CD56dim) defines enhanced cytotoxicity associated with a mature differentiation state (31). On T cells, CD56 is expressed on the NKT subset, initially proposed to carry a cytotoxic function (32), and more recently a regulatory function (15). NKT cells can rapidly produce large amounts of cytokines (33), which can mediate effector as well as suppressor functions (34). However, these studies primarily focused on the mouse as a model, whereas NKT cells are scarce in the human and may differ functionally from those in mouse (35). In humans, CD56 is expressed on a second, more abundant, T cell subset, which is often confused with classical NKT (36). Although sharing some characteristics with NK or T cells, our analysis as well as others clearly define CD56+ T cells as distinct in function and morphology (37). Most importantly, these are not mono- or oligoclonal and are not CD1 restricted (19), thus suggesting conservative mechanisms of Ag repertoire shaping and a role in adaptive immunity.

In this study, we examined the distribution and function characteristics of CD56+ T cells in association with the human intestinal compartment. Mucosal CD56+ T cells express a mature phenotype and can produce high amounts of IFN-{gamma} and TNF. Peripheral CD56+ T cells express gut-homing integrins and respond to engagement of the gut-associated CD2 signaling pathway. Finally, we directly demonstrate an effector potential for CD56+ T cells in vitro. Our findings associate the CD56+ population of T cells with the intestinal immune compartment and suggest a functional role for these cells in mucosal immunity.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Cohort and specimen procurement

Blood leukocytes were obtained by venipuncture from healthy adult volunteers. Intestinal specimens were obtained from patients undergoing intestinal resection for clinical reasons at the Cedars-Sinai IBD Center. Patient diagnosis was defined as CD, UC, or non-IBD using standard clinical, radiographic, and endoscopic criteria (38), and gross tissue involvement was validated microscopically. Patients treated with cyclosporin A were omitted from the study, and patients with indeterminate colitis were omitted from analysis of disease associations. Procedures for subject recruitment, informed consent, and specimen procurement were in accordance with protocols approved by the Institutional Review Board for Human Subject Protection of the Cedars-Sinai Medical Center.

Lymphocyte isolation and T cell subset purification

PBMC were isolated from uncoagulated blood by standard Ficoll-Hypaque density gradient centrifugation. Mononuclear cells from the intestinal lamina propria (LPMC) were isolated as described previously (8). Briefly, the epithelial layer (including intraepithelial lymphocytes) was removed by washing in EDTA, followed by enzymatic disruption of the lamina propria (LP) matrix, mincing and density gradient purification of LPMC. Mononuclear cells were isolated from mesenteric lymph nodes (MLNs) by mechanical sheering of the node into culture medium. LPMC, lymph node (LN) cells, and PBMC preparations were cultured for 16–20 h in plastic plates, and nonadherent cells were removed for further purification or experimentation. CD3+ T cell subsets (CD56+/– or CD161+/–) and NK cells were purified or depleted from PBMC by flow cytometry (FACStar (BD Biosciences) or MoFlow (DakoCytomation)) gating on viable CD3+, lymphocyte-size cell subsets. Purity of enriched populations was consistently >99% for the gated markers with <0.2% of depleted cell subsets remaining when reanalyzed by flow cytometry (FACScan (BD Biosciences) or Cyan (DakoCytomation)).

Cell culture

Lymphocytes were cultured at 0.25–1 x 106 cells/ml in RPMI 1640 containing 2 mM L-glutamine and 25 mM HEPES buffer (Mediatech), supplemented with 10% heat-inactivated FBS (Atlanta Biologicals), 50 µg/ml gentamicin (Omega Scientific), and LPMCs with additional 0.25 µg/ml amphotericin B (Gemini Bio-Products). Where indicated, lymphocytes were stimulated by 40 ng/ml PMA and 1 µg/ml ionomycin (Sigma-Aldrich); or by Ab cross-linking of cell surface CD2 used at 0.4 µg/ml. For coculture experiments, cells were washed thrice in culture medium to deplete the stimulant before coculture with responder cells and a total of 106 cells/ml medium were cocultured at 1:1 ratio with CFDA-SE (CFSE)-labeled responder cells (Molecular Probes). To achieve physical separation, effector or responder cells were placed in Transwells (Transwell; Corning) providing 0.25-µm pore-sized membrane for small molecule diffusion between cell culture compartments. Absence of cell transfer across the Transwell membrane was validated by flow cytometry with consistently <3% non-CFSE-stained effector cells in the responder cell compartment.

Ab reagents

The anti-CD2 mAb pair, clones GD10 and CB6, was a gift from Dr. C. D. Benjamin (Biogen, Cambridge, MA). The ascites was purified over a protein G column and quantified by ELISA. Additional chromophore-conjugated Abs specific for human CD3, CD4, and CD8, IL-10, IL-4, IL-5, TNF, and IFN-{gamma} were obtained from Caltag Laboratories, and anti-human CD56 was from Beckman Coulter.

Cell staining and flow cytometry

For intracellular cytokine analysis, cells were incubated in the presence of Golgi inhibitor, brefeldin A (Calbiochem), for the last 5–6 h of stimulation. Cells were then washed and stained for surface markers as described above, followed by light fixation and permeabilization in the presence of anti-cytokine, or isotype control Abs using the Fix and Perm, intracellular staining kit (Caltag Laboratories). Cells were then washed, fixed in 1% paraformaldehyde, and stabilized at 4°C for 16–20 h before flow cytometric analysis. Nonspecific staining by control isotypes or staining of unstimulated cells was subtracted from the percentage of staining for each cell subset to determine specific mean fluorescence.

CFSE was diluted to 5 µM in PBS with 0.1% BSA and used to stain cells at 25 x 106/ml for 8 min at 37°C. Staining reaction was stopped by dilution into cold medium, and cells were washed thrice to eliminate excess CFSE stain. Flow cytometric analysis included at least 2 x 104 events on a FACScan (BD Biosciences) or Cyan (DakoCytomation) and analyzed with the respective CellQuest or Summit software. Percentages of cytokine-producing or proliferating cells represent the fraction of total CD56+ or CD56– T cells that stain intracellularly for cytokine, or localize to the CFSE g1–6 gate. Proliferation indexes were calculated for CD3+ cells in the responder population gate using the ModFit Cell Cycle Analysis software (Verity Software House).

Statistical analysis

CD56+ T cell frequencies were calculated as a fraction of total T cells for each donor. Differences in CD56+ T cell frequency were evaluated for multiple patient groups, anatomic locations, or clinical diagnosis using the Kruskal-Wallis test. Considering nonnormal distribution shape, Wilcoxon two-sample test was applied as nonparametric statistical analysis to further elucidate differences between pairs of groups, and p values (rounded up) that remain significant following Bonferroni adjustment are reported in Results. The Student t test was applied when comparing frequencies of cytokine-producing cells.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
CD56dim marks T cells that are increased in frequency in the human intestinal mucosa

CD56+ T cells may represent a functionally distinct cell subset with a significant immunoregulatory role. Hence, we characterized the CD56+ population of T cells in the intestinal immune compartment. CD56 T cells were detected by surface immunostaining and flow cytometric analysis of lymphocytes isolated from the human intestinal LP (LP lymphocytes), MLNs, or peripheral blood (PB). Scatter plot analysis of gated live lymphocytes resolved CD56+/CD3+ T cells from CD56–/CD3+ T cells or CD3–/CD56+ NK cells (Fig. 1A), with true NKT cells (CD3+/CD56+/CD161+) representing <20% of the CD56+ T cell subset (16). CD56 staining intensity in CD3+ T cell population was relatively low and almost exclusively corresponded to the CD56dim profile, characteristic of a functionally distinct subset of NK cells (31), whereas CD3– NK cells included both, the CD56dim and CD56bright profiles (Fig. 1A).


Figure 1
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  		FIGURE 1. CD56dim expression marks a subset of T cells in the human intestinal LP. Lymphocytes were isolated from the intestinal LP, MLNs, or PB and surface stained with fluor-conjugated anti-CD56 and anti-CD3 Abs, followed by flow cytometric analysis. A, Gated viable, lymphocyte-size, scatter plots are shown for representative samples, indicating CD56+ staining profiles for T (CD3+) and NK (CD3–) cells from small bowel (uninvolved UC), colon (non-IBD), MLNs (small bowel CD), and PBL (healthy donor). Data are representative of at least five experiments with similar results. B, The frequency of intestinal CD56+ T cells was computed in a cohort of IBD patients as a fraction of total T cells from the indicated lymphoid tissue location. Pileup data points represent unique individuals with value bar indicating group mean frequency. C, Pileup diagram represents frequencies of colonic CD56+ T cells analyzed for colonic IBD specimens defined as inflamed or uninvolved based on gross and microscopic mucosal pathology vs cells from non-IBD colons. Indicated p values are for the Wilcoxon exact test for frequencies compared with PBL (B), or inflamed vs uninvolved (C).

 
Statistical analysis of intestinal CD56+ T cell frequencies as a fraction of total CD3+ T cells in a cohort of IBD patients indicated elevated frequency of CD56+ T cells in small bowel (mean frequency = 18.9%; p < 0.001) and colonic (mean frequency = 11.0%; p < 0.0002) specimens but lower frequency in MLNs (mean frequency = 1.3%; p < 0.0007) when compared with PBL (mean frequency = 5.8%) (Fig. 1B). Significant variation in CD56+ T cell frequency was detected in the colon and small bowel (ranging 2–36 and 5–33%, respectively), in agreement with a study by Pittet et al. (39), although lower frequency variations were reported in a second smaller cohort (16). Further analysis of colonic specimens from inflamed and uninvolved intestinal sites indicated reduced frequencies of CD56+ T cells in inflamed sites (p < 0.0001 vs non-IBD and p < 0.0028 vs uninvolved), whereas frequencies in uninvolved sites did not significantly differ from non-IBD mucosa (Fig. 1C). However, CD56+ T cell frequency in colonic specimens did not significantly differ between UC and CD patients in this size cohort (p = NS for uninvolved 9 UC and 4 CD, or for inflamed 17 UC and 4 CD specimens). Reduced CD56+ T cell-relative frequency in the inflamed mucosa correlates with the lower relative frequency in LN (procured from inflamed sites).

CD56+ T cell morphology and surface marker characteristics

Substantial presence of CD56+ T cells in the gut mucosa suggests CD56+ as a marker for gut-associated T cells circulating in the periphery. We therefore examined peripheral CD56+ T cells for expression of the {alpha}4beta7 integrin as a marker for gut homing. Indeed, peripheral CD56+ T cells costained for the integrin beta7 subunit more frequently than the CD56– subset (67 vs 42%), thus associating CD56 expression with the integrin {alpha}4beta7 gut-homing pathway (Fig. 2A). However, nonexclusive beta7 expression on CD56+ T cells implies diversity within the peripheral CD56+ T cell population and may suggest additional tissue specificities for these cells (20, 40). In addition, we assessed CD45RO expression as a marker of Ag encounter and T cell maturation, which is expressed by the majority of gut-derived T cells (8). CD45RO was more frequently, although not exclusively, expressed on CD56+ T cells from the PB (Fig. 2A), or from the gut (Fig. 2B), further associating the peripheral CD56+ T cell subset with the intestinal compartment and suggesting more frequent exposure to Ag-mediated stimulation.


Figure 2
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  		FIGURE 2. CD56+ T cell morphology and surface marker expression characteristics. A, PBL from healthy donor blood were surface stained with anti-CD56, anti-CD3, and anti-beta7 integrin or anti-CD45RO Abs and analyzed by flow cytometry. Scatter plots are shown for live CD56+ and CD56– gates. B, Lymphocytes were isolated from the intestinal LP and surface stained for CD56, CD3, and CD45RO or CD8 followed by flow cytometry. C, Forward scatter/side scatter flow cytometric analysis is shown for intestinal CD56+ T cells vs CD56– T cells or NK cells. Data are representative of at least five experiments with similar results.

 
Because CD56+ T cells appear functionally distinct from T, NK, and NKT cell populations, we tested for unique phenotypic characteristics in addition to CD56 expression on intestinal T cells. The frequency of gut CD56+/CD8+ T cells approximated that of CD56– T cells (Fig. 2B), similar to blood (data not shown), or profiles recently reported for CD56+ T cells in the liver (20). Nonexclusive CD8 expression thus differentiates intestinal CD56+ T cells from traditional NKT cells, which uniformly express the CD8 accessory molecule (32). Additional analysis of density vs size (side scatter vs forward scatter by flow cytometry) indicated a distinctive bimodal profile in gut CD56+ T cells, representing small and less granular T lymphocyte morphology, as well as a broader size/density profile characteristic of NK cells (Fig. 2C). Although granular phenotype may suggest cytolytic potential, significant CD56+ T cell cytotoxicity was not detected in our experiments (data not shown) or other studies (37, 41).

Mucosal CD56+ T cells are nonproliferating and express a Th1-like cytokine profile

Mucosal T cells are characterized by a mature stage of differentiation associated with a reduced capacity to proliferate (8). We therefore assessed mucosal CD56+ T cell proliferation potential in a 3-day culture following PMA and ionomycin (P/I) stimulation. Proliferation was measured as a function of CFSE stain dilution, and revealed compromised proliferation potential in the intestinal CD56+ T cell population when compared with the CD56– subset (Fig. 3A). No excessive CD56+ cell death was detected during the 3-day incubation (<20%; data not shown). The compromised proliferation potential of CD56+ T cells is consistent with reduced relative frequency in the inflamed mucosa or LNs, which are sites of extensive T cell proliferation (Fig. 1).


Figure 3
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  		FIGURE 3. Mucosal CD56+ T cells have a limited proliferation potential and express a Th1-like cytokine profile. LPMC were isolated from the intestinal mucosa using a standard enzymatic method. A, Isolated LPMC were CFSE stained and cultured in medium for 3 days following P/I activation. Cells were surface stained for CD3 and CD56, and gated live CD3+ lymphocyte-size histograms are presented for activated (shaded) vs nonactivated (unshaded) cells. Inset values indicate percentage of stimulated CD56+ or CD56– T cells at the g1–6 CFSE gate. B, Cells were rested in culture for 24 h and then stimulated for an additional 6 h with P/I in the presence of Golgi inhibitor, brefeldin A. Activated cells were surface stained with anti-CD3 and anti-CD56 Abs followed by intracellular staining for the indicated cytokines and analyzed by flow cytometry. Histograms show intracellular staining profiles for activated (shaded) CD56+ or CD56– T cells in comparison with the respective nonactivated cells (unshaded). Inset values indicate percentage of stimulated CD56+ or CD56– T cells expressing intracellular cytokine. Data for A and B are representative of five experiments with similar results.

 
Compromised proliferation and the mature state of differentiation prompted us to evaluate the effector capacity of CD56+ T cells. Specialized T cell effector functions can be determined based on specific cytokine expression profiles; hence, we assessed cytokine expression potential for intestinal CD56+ T cells in vitro. Isolated human LP lymphocyte preparations were activated for 6 h with P/I in the presence of Golgi maturation inhibitor, brefeldin A, and cytokine production was detected by intracellular immunostaining. T cells expressing inflammatory cytokines, IFN-{gamma} and TNF, were consistently more frequent in the CD56+ population than in the CD56– subset (~2-fold), in CD (data not shown) as well as UC or non-IBD guts (Fig. 3B). Fluorescence intensity was comparable in the cytokine-producing cells in the CD56+ and CD56– T population, implying similar cytokine levels per responsive cell. Analysis of IL-13 expression indicated low frequency of CD56+ producing T cells from UC guts (UC shown for IL-13 and IL-4), and undetectable IL-13 expression in CD or non-IBD specimen, not supporting a significant role for CD56+ T cells in a Th2-type response. In addition, marginal expression of IL-10, IL-4, and IL-5 clearly distinguished CD56+ T cells from classical NKT cells (32, 33) and did not support suppression or a regulatory function associated with expression of these cytokines. Finally, cytokine expression in gut CD56+ T cells agree with cytokine production profiles reported for CD56+ T cells in the liver, thus suggesting that CD56 identifies a functionally distinct T cell subset (20). Taken together, these data suggest mucosal CD56+ T cells as a poorly proliferating, cytokine-producing T cell subset, in accord with a mature CD45RO phenotype (Figs. 2 and 3) (8).

CD56+ T cells respond to CD2 signaling in the periphery

Preferential expression of integrin {alpha}4beta7 and maturation marker, CD45RO, phenotypically associated peripheral CD56+ T cells with the gut compartment. At the functional level, we tested for cytokine production in response to CD2 engagement in vitro, because responsiveness to CD2 signaling is associated with mucosal T cells (8, 42). Intracellular cytokine staining analysis following CD2 cross-linking for 6 h in the presence of brefeldin A indicated robust IFN-{gamma} and TNF production by PB CD56+ T cells. Cytokine-producing cells were significantly more frequent in the CD56+ T cell subset (35–51% IFN-{gamma}+, 45–63% TNF+) when compared with CD56– T cells (1.5–3% IFN-{gamma}+, 4–7% TNF+) (Fig. 4). The magnitude of CD2-mediated cytokine production by CD56+ T cells, measured as cytokine-producing cell frequency or staining intensity was comparable to the universal P/I stimulus, thus suggesting very effective CD2 signaling in responsive CD56+ T cells (Figs. 4 and 3A, respectively). Minimal CD2 responsiveness in CD56– T cells was not a consequence of mature T cell depletion, because CD56+ T cells constitute a small fraction of total mature (CD45RO) T cells in blood (1.7% of total T cells in Fig. 5). CD2 is also expressed on CD3–/CD56+ NK cells, but although one study reported potentiation of NK cytotoxicity by CD2 (43), only marginal cytokine production was detected in NK cells within the time frame of our experiments.


Figure 4
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  		FIGURE 4. Peripheral CD56+ T cells respond to CD2-mediated signaling. Isolated PBL were stimulated for 6 h by CD2 cross-linking in the presence of Golgi inhibitor, brefeldin A, and surface stained with anti-CD3 and anti-CD56 Abs followed by intracellular immunostaining for the indicated cytokines. Flow cytometry histograms show intracellular staining profiles for activated (shaded) CD56+ or CD56– T cells and NK cells in comparison with the respective nonactivated cells (unshaded). Inset values indicate percentage of CD2-stimulated CD56+ or CD56– T cells expressing intracellular cytokine. Spontaneous cytokine production by unstimulated cells was <1% (IFN-{gamma}) or <2% (TNF). Data are representative of at least five experiments with similar results.

 

Figure 5
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  		FIGURE 5. CD56+ T cells facilitate CD2-mediated proliferation of peripheral T cells. A, Indicated cell populations were purified by flow sorting (>99% by flow reanalysis) from isolated PBL. Pure cell preparations were CFSE stained and cultured for 5 days following activation by P/I or CD2 cross-linking as indicated. Cell proliferation is presented as a function of CFSE dilution in histograms for activated (shaded) vs nonactivated (unshaded) T cells. Inset values indicate percentage of stimulated T cells at the g1–6 CFSE gate. B, Isolated PBL were CFSE stained and cultured for 5 days following activation with CD2 and then surface stained for CD3 and CD56. CFSE dilution histograms of CD56+ and CD56– gated T cells are shown for activated (shaded) vs nonactivated (unshaded) cells. Data are representative of three experiments with similar results.

 
Remarkably, despite the CD56+ T cell subset representing only a small percentage of the total PB T cell population (1–10%), CD56+ T cells can comprise a significant fraction of IFN-{gamma} (65%) or TNF (46%) producing T cells following CD2-mediated activation. Robust CD56+ T cell responsiveness to CD2 could imply that CD56– T cell responses are secondary and mediated by a cue from a CD2-responsive cell. To ascertain whether such interaction occurs, we depleted PBL preparations of candidate cell subsets before CD2 cross-linking and evaluated proliferative responses as a function of CFSE dilution. Depletion of either CD56+ or CD161+ T cells partially inhibited CD2-mediated proliferation, while concurrent depletion of both these subsets effectively abrogated the response (Fig. 5A). In addition, partial inhibition of P/I-mediated proliferation resulted from concurrent depletion of both CD56+ and CD161+ T cells, suggesting that both subsets are capable of providing proliferation-enhancing signals following global activation. By contrast, depletion of non-T cells did not have a significant effect on proliferation, ruling out a response mediated by NK cells that can also respond to CD2 signaling (43). Both CD56+ and CD161+ T cells stimulated by CD2 proliferated less than the respective CD56– and CD161– cells, indicating that compromised proliferation does not reflect depletion of a highly proliferative cell subset (Fig. 5B). These data demonstrate that T cells expressing CD56 or CD161 are necessary for optimal PB T cell proliferation in response to CD2 stimulation, and thus suggest that T cell responses to CD2 stimulation are mediated by a small subset of responsive cells.

CD56+ T cells induce responder T cell activation

To directly assess CD56+ T cell effector function, we used a cell culture system that permits analysis of minimally perturbed effector-responder T cell interactions (Fig. 6). Adherent cell-depleted PBL were used as responder cells and labeled with CFSE stain, to permit identification following coculture with unstained effector cell populations. Effector cell subsets were purified from the autologous PBL preparation by flow sorting (>99% pure by flow cytometry), activated with P/I for 6 h, and washed. Effector and responder cells were cocultured at varied effector:responder (E:R) ratios for responder cell activation, followed by flow cytometric analysis of CD3-stained T cells in the responder population. Responder T cell responses were evaluated by measurement of IFN-{gamma} synthesis detected at 24 h by intracellular staining following 6-h culture in the presence of brefeldin A, and proliferation was assessed as a function of CFSE stain dilution over 5 days in coculture.


Figure 6
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  		FIGURE 6. Experimental setup for the analysis of effector-responder interaction. PBL were depleted of adherent cells and used as responder cells when labeled with CFSE stain. Effector cell subsets were excluded from the CFSE staining step and purified by flow sorting (>99% pure by flow cytometric reanalysis). Effector cells were then activated with P/I for 6 h, washed, and cocultured with responder cells at varying E:R ratios. Responder T cell responses were evaluated by measurement of IFN-{gamma} synthesis detected at 24 h by intracellular staining following 6-h culture in the presence of brefeldin A. In addition, responder T cells cell proliferation was assessed as a function of CFSE stain dilution over 5 days in coculture.

 
Responder T cell proliferation was enhanced when purified CD3+/CD56+ T cells were used as the effector population, in comparison with either PBL or purified CD3+/CD56– T cells (Fig. 7A). Specifically, CD56+ T cells were more effective when cultured at a lower E:R ratio, with significant responder proliferation detected at 0.25 E:R. Higher frequency of cells proliferating through g2 and proliferation index indicated more efficacious CD56+ T cell induction of responder cell proliferation. Calculating the fraction of responder cells initially induced to proliferate did not suggest a significantly higher fraction responding to effector CD56+ T cells. Particularly at 1:1 E:R ratio, a similar fraction of initial responder cell population was induced to proliferate by PBL, CD56–, or CD56+ T cells, whereas the proliferation index was significantly higher for CD56+ T cells (Fig. 7A).


Figure 7
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  		FIGURE 7. CD56+ T cells provide enhanced proliferative and cytokine-inducing signals via T-T interactions. Adherent cell-depleted PBL, flow-sorted CD3+/CD56–, or CD3+/CD56+ T cells to be tested as effectors cells were activated with P/I for 6 h and washed. Activated effector cells were then cocultured with adherent cell-depleted, CFSE-stained PBL as responder cells. A, Histograms represent effector-induced CFSE dilution profiles for a CD3+ gate at day 5. Inset values indicate percentage of stimulated responder T cells at the g1–6 CFSE gate (top) or g2–6 gate (bottom). Graphs indicate the proliferation index calculated for CD3+ cell in the responder population (left) or calculated the initial responder cell fraction induced to proliferate (right), at varying E:R ratios. B, CD56+ T cells induced enhanced IFN-{gamma} synthesis in responder CD3+ T cells (bars represent mean IFN-{gamma}+ cell percentage of total CD3+ responder cell gate at 24 h for three experiments). C, CFSE dilution histograms at day 5 for CD56+ T cell induced responder T cell proliferation in the absence (top) or presence (bottom) of a Transwell semipermeable membrane separating effector from responder cell populations. Data for A and C are representative of a minimum of three experiments.

 
In addition, effector-cell-mediated activation at 1:1 E:R ratio consistently induced IFN-{gamma} production by a small fraction of responder T cells (2.85 ± 0.73% by PBL). Pure CD56+ T cells induced a higher frequency of IFN-{gamma}-producing cells in the responder T cell population (6.75 ± 2.53%; p < 0.029) when compared with CD56– T cells (2.84 ± 2.33%) (Fig. 7B). No significant CD56+ T cell-mediated cytotoxicity was detected in these experiments (<15% apoptosis of responder cells incubated with either CD56+ or CD56– effector T cells) (41). Although depletion experiments suggested a similar effector function for the CD161+ T cell subset, we did not detect consistent effector function for this subset in our coculture experimental setting (data not shown).

Efficient cytokine production by CD56+ T cells suggested cytokine-mediated activation of responder T cells (Fig. 3). We tested this possibility by introducing a "Transwell" membrane into the coculture experiment, physically separating the effector from the responder cells, while permitting diffusion of soluble factors under 0.25 µm in diameter. Interestingly, responder T cell proliferation was completely abrogated as a result of cell contact inhibition, with similar inhibition for effector cells activated either by P/I or CD2 (Fig. 7C). Taken together, these experiments demonstrate the potency of CD56+ T cells as an effector population and reveal a requirement for cell-cell contact in responder T cell activation.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
CD56+ T cells represent a unique population with a differentiated effector function. Single-positive CD4+ or CD8+ profile and CD45RO expression suggest a mature T cell phenotype both in the gut and in the periphery (Fig. 2). Efficient cytokine production but compromised proliferation potential following P/I or CD2 stimulation further implies a fully differentiated developmental state (Fig. 3). Moreover, CD56 expression profile on T cells conforms to the CD56dim profile (Fig. 1), which in NK cells defines a functionally unique subset (31), initially proposed more mature (44). Preferential IFN-{gamma} and TNF expression profiles, but marginal expression of regulatory cytokines such as IL-13, IL-4, IL-5, and IL-10 (Fig. 3) (36), or significant CTL activity (41), specifically support a proinflammatory effector function, and clearly distinguish CD56+ T cells from the NKT subset (33). Notably, we directly demonstrate an enhanced effector potential for CD56+ T cells by using a unique in vitro culture system that permits analysis of effector-responder interactions. CD56+ T cells effectively induced responder T cell proliferation and IFN-{gamma} production (Fig. 7) while not decreasing responder cell survival (data not shown) (37, 41). Interestingly, responder T cell activation was not driven by enhanced proinflammatory cytokine production by CD56+ T cells (Fig. 3) but was contact dependent (Fig. 7C), suggesting additional and more complex cell-cell interaction mechanisms. For instance, surface expression of gut-associated proinflammatory molecules such as CD40L (45), LIGHT (46), or TL1A (47) could potentially mediate T cell activation.

Enhanced CD56+ effector T cell potential implies an important immunoregulatory function, but is such function tissue specific? Morphology and cytokine expression profiles for CD56+ from the gut or the periphery were similar and in agreement with a recent study of liver CD56+ T cells (20), suggesting effector potential to be independent of the tissue environment. However, immunoregulatory activity could be mediated by signaling mechanisms, which are tissue specific (9), or simply by increased presence of proinflammatory CD56+ T cells. In support of the latter, our data as well as others indicate localization to the intestinal mucosa (Fig. 1) (16), or the liver (19, 20), where CD56+ T cells can constitute a significant fraction of the resident T cell population. Although we do not directly elucidate cell trafficking and compartmentalization, intestinal CD56+ T cells may circulate through the periphery (48), because expression of CD45RO and a compromised proliferation potential are both characteristics of mucosal T cells (8). Similarly, preferential expression of gut-homing integrin, {alpha}4beta7 (Fig. 2), suggests a mechanism for CD56+ T cell transport to the intestinal mucosa (25). Although, CD56+ T cells share phenotypic characteristics with the CD45RO and beta7+ subsets, our data suggest that CD56 marks an independent cell that is not just a subset of the CD45RO or beta7+ populations. CD2 responsiveness could not be only reflective of property associated with the CD45RO and beta7 markers, because CD56– T cells do not respond to CD2-mediated stimulation despite frequent CD45RO and beta7 expression in this subset. Moreover, cytokine expression cannot be exclusively linked to CD45RO or beta7 expression in the CD56+ T cell population because the frequency of cytokine-producing CD56+ T cells stoichiometrically exceeds the CD45RO or beta7+ subset frequency.

Mucosal T cells are highly responsive to CD2-mediated signaling, which offers an effective alternative to strictly regulated Ag-dependent activation pathways in the gut compartment (8). In general, peripheral T cells are only marginally responsive to CD2-mediated signaling; however, peripheral CD56+ T cells respond to CD2 cross-linking and, in fact, can account for a significant fraction of IFN-{gamma}-producing PB T cells following CD2 activation (Fig. 4). We further demonstrate that although CD56+ T cells are marginally proliferative, they are necessary for CD2-mediated proliferative responses of peripheral T cells (Fig. 5). The fact that CD2 responsiveness is unique to gut-associated T cells is intriguing, because most T cells express CD2 and an apparently intact CD2 signal transduction machinery (49). Our finding of a peripheral CD56+ T cell population responsive to CD2 signaling further associates this T cell subset with the intestinal immune compartment and may facilitate further analysis of CD2 signaling mechanisms.

Collectively, our data indicate peripheral CD56+ T cells to reflect the activity of intestinal T cells, which could be immunopathogenic. Further analysis of this peripheral CD56+ T cell population is thus likely to provide important insight into mechanisms of mucosal immune regulation or associated pathology. For instance, elevated frequency of peripheral CD56+ T cells was reported in other inflammatory conditions such as uveitis, which is often associated with IBD (40). These observations link peripheral CD56+ T cells to mucosal immune dysregulation and suggest pathological significance for mechanisms modulating CD56+ T cell frequency (50). Although perturbations in CD56+ T cell frequency may not be unique to IBD, specific modulation of CD56+ T cell trafficking or effector function may represent targets for gut-specific therapeutic intervention (51).

Although the molecular immune function of CD56 is unclear, its expression on a specific T cell subset associated with the mucosal immune compartment is noteworthy. Extensive exposure to Ag at the intestinal mucosa is consistent with the mature (CD45RO) and nonproliferative phenotypes, which are primary hallmarks of intestinal T cells (8, 9), shared by CD56+ T cells (Figs. 2 and 3). Yet, Ag specificity repertoires unique to the gut compartment have been described, which suggested mucosal T cell differentiation and selection independent of the thymus (4). A recent study reported "leakage" of T cell progenitors from the thymus to the gut as a novel mechanism for thymus-independent T cell selection (52). More specifically, Gunther et al. (53) reported CD56 expression on CD7+/CD3– T cell progenitors in the gut that are capable of differentiating into mature T cells, and Musha et al. (54) proposed extrathymic development of CD56+ T cells in studies of human neonate cord blood. NK markers such as CD56 or CD161 are expressed early in lymphoid cell development before the T-NK split, but lost during T cell selection and maturation in the thymus. Hence, the occurrence of fully differentiated T cells expressing CD56 in the adult gut is intriguing and consistent with the concept of thymus-independent differentiation (55). Although we do not directly demonstrate a role in pathological inflammation, CD56+ T cell robust effector potential combines with a likely thymus-independent immune repertoire to suggest a significant role in Ag-specific responses in the intestines. Enhanced proinflammatory cytokine production by CD56+ T cells and induction of additional T cell subsets may contribute to elevated cytokine concentrations in the inflamed gut (6), whereas circulating CD56+ T cells may instigate extraintestinal manifestations of IBD through the periphery (40).

In conclusion, CD56+ T cells are a significant fraction of intestinal T cells that may well play a significant immunoregulatory role mediated by a robust effector potential and enhanced expression of proinflammatory cytokines. Peripheral CD56+ T cells share phenotypic characteristics with mucosal T cells and respond to the gut-associated CD2 signaling pathway. Thus, CD56 expression may discern circulating mucosal T cells. Further analysis of the CD56+ T cell role in mucosal immunity will better our understanding of pathological intestinal inflammation and could identify new targets for gut-specific intervention.


   Acknowledgments
 
We are grateful to our surgical colleague Phillip Fleshner for facilitating intestinal specimen procurement, Grace Kim and Svetlana Jovicic for technical assistance, Patricia Lin for help with flow cytometry, John L. Prehn for helpful discussions, and Carl F. Ware for critical reading of this manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported in part by National Institutes of Health Grants F32DK10139 (to O.C.) and R01DK57328/R01DK43211 (to S.R.T.). Back

2 Address correspondence and reprint requests to Dr. Stephan R. Targan, Cedars-Sinai Inflammatory Bowel Disease Center, 8700 Beverly Boulevard, Suite D4063, Los Angeles, CA 90048. E-mail: targans{at}cshs.org Back

3 Abbreviations used in this paper: IBD, inflammatory bowel disease; CD, Crohn’s disease; UC, ulcerative colitis; LPMC, mononuclear cells from the intestinal lamina propria; LP, lamina propria; MLN, mesenteric lymph node; LN, lymph node; PB, peripheral blood; P/I, PMA and ionomycin; E:R, effector:responder. Back

Received for publication February 21, 2006. Accepted for publication February 12, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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