Skip to main content
Access keys NCBI Homepage MyNCBI Homepage Main Content Main Navigation
Clin Exp Immunol. 1998 May; 112(2): 226–236.
doi: 10.1046/j.1365-2249.1998.00566.x
PMCID: PMC1904972
PMID: 9649184

Interactions between peripheral blood CD8 T lymphocytes and intestinal epithelial cells (iEC)

F A Arosa, C Irwin,* L Mayer,† M De Sousa, and D N Posnett*
Author information Article notes Copyright and License information PMC Disclaimer

Abstract

Intestinal intraepithelial lymphocytes (iIEL) are primarily CD8 cells and most of them have a CD28− phenotype, the phenotype of effector cytotoxic T cells. We asked whether the predominance of CD8+ CD28− T cells in the gut may result from peripheral blood T cells preferentially migrating to the iIEL compartment and adhering to iEC. Compared with CD4 cells, adhesion of resting CD8+ T cells to iEC cell lines was significantly higher. Adhesion could be blocked with a MoAb to gp180, a molecule expressed on iEC which is known to interact with CD8/lck. No significant difference in the level of adhesion was observed between CD8+ CD28+ and CD8+ CD28− T cells. Thus CD8 cells may preferentially migrate to the iIEL compartment, but loss of CD28 expression could occur in situ after migration. Consistent with this hypothesis, the CD8+ CD28− cells became enriched after co-culturing T cells with iEC cell lines and primary iEC. Induction of the CD8+ CD28− phenotype in cord blood and adult T cells was observed in co-cultures with iEC and also with mitogens and superantigens. In the latter case, CD28 down-modulation was seen specifically in the Vβ subset targeted by the superantigen, indicating that loss of CD28 expression is a direct result of T cell receptor (TCR)-mediated stimulation. The combined results suggest that CD8+ CD28− T cells are antigen experienced T cells, and that they may have a survival advantage in the presence of gut epithelial cells in vitro. This may contribute to the predominance of CD8+ CD28− T cells in the iIEL compartment.

Keywords: intestinal intraepithelial lymphocytes, CD8 T cells, CD28 expression

INTRODUCTION

CD28 is an essential costimulatory molecule of T cells known to interact with the ligands CD80 (B7-1) or CD86 (B7-2) on antigen-presenting cells (APC). Stimulation of a T cell via the T cell antigen receptor (TCR) and simultaneously via CD28 results in T cell responses such as proliferation, IL-2 secretion and activation of cytolytic effector function. By contrast, TCR stimulation without CD28 stimulation, for instance by APC that lack CD80/86, can lead to anergy [1,2]. Not all T cells express CD28. In the peripheral blood lymphocytes (PBL) of adult humans there is a major population of CD28− CD8+ cells which is absent at birth and ranges from 6% to 59% of total CD8+ CD3+ T cells in normal adults [3]. Many CD4− CD8− TCRαβ+ or TCRγδ+ T cells also lack CD28 expression [4–6], but CD4+ CD28− cells are usually rare.

CD8+ CD28− cells are thought to include cytolytic effector cells, as they are highly enriched for cytolytic activity in redirected cytotoxic T lymphocyte (CTL) assays [3]. These cells contain a high content of granzyme B and perforin. They express high levels of FasL and produce interferon-gamma (IFN-γ) and tumour necrosis factor-alpha (TNF-α) consistent with an effector cell phenotype [7]. This same subset may also be responsible for antigen non-specific suppresser functions in co-culture assays [8–13]. By contrast, CD8+ CD28+ cells are thought to include naive and memory CD8 subsets. CD8+ CD28− T cells characteristically give weak proliferative responses [3]. They are usually CD57+, CD11ahi, CD11b+, CD49d (VLA-4)+, and CD27−, representative of the phenotype of effector CD8 T cells [3,7]. CD8+ CD28− T cells frequently contain clonal expansions [14,15], indicating that they are antigen-driven. High percentages of peripheral blood CD8+ CD28− T cells are a common feature in certain chronic inflammatory diseases, during viral and parasitic infections and in hereditary haemochromatosis [16–20]. However, these expansions can also be found in apparently healthy individuals [14,15].

Many of the features of CD8+ CD28− peripheral blood T cells are shared with human CD8 intestinal intra-epithelial lymphocytes (iIEL). In the iIEL compartment most cells express CD8 but lack CD28, particularly in the small intestine, both in humans and in mice [21–23]. In mice a major subset of iIEL tend to use the CD8αα homodimer rather than the CD8αβ heterodimer, and may represent T cells that have matured in an extrathymic environment [24,25].

The origin of CD8+ CD28− T cell expansions in the PBL is unclear. It has been suggested that this subset may perhaps represent circulating CD8+ T cells derived from the iIEL compartment, but they may also represent effector CTL reactive with ubiquitous viral antigens like herpesvirus antigens [14,26]. Evidence for a role of human iEC in activating CD8+ T cells has been reported, and this may in part be due to stimulation via non-classical MHC class I molecules like CD1b, CD1c and CD1d interacting with CD8 [4,27,28]. In CD28 knockout mice humoral responses were lacking, but the CTL response to lymphocytic choriomeningitis virus and allograft rejection were preserved [29,30], providing indirect evidence for CD28-independent costimulatory pathways in some cases. CD28/B7 was not a costimulatory requirement in the case of two peripheral blood-derived CD4− CD8− T cell clones restricted by CD1 [4], but in this case the costimulation was provided by blood monocytes, not iEC. CDw101 is a candidate molecule expressed by iIEL that may transmit a costimulatory signal in CD28− T cells [31].

Regarding the lack of CD28 expression by CD8 iIEL, there are several possible hypotheses. CD8+ CD28− iIEL may represent a separate lineage of extrathymic cells that matures in the gut and does not require CD28 ligation as a costimulus. Alternatively, they may be recruited to the iIEL compartment where they might adhere particularly well to iEC. Finally, they may enter the gut as CD28+ CD8+ cells, become activated in response to antigens presented by APC and mature into effector CTL with a CD28− phenotype. The data presented here favour the latter possibility.

MATERIALS AND METHODS

Cell lines

The human colonic epithelial cell lines HT-29 and T-84 (American Type Culture Collection (ATCC), Rockville, MD) were used. They were grown in Dulbecco's modified Eagles' medium (DMEM)/F12 media supplemented with 10% fetal calf serum (FCS), 1% glutamine, 1% pen/strep and 1% fungizone (culture media 1 (CM1)). Cells were placed in 75-cm2 Falcon flasks and media changed every 2–3 days. When cells reached confluence, 10 ml of a 1% solution of trypsin/EDTA were added for 20–30 min. After harvesting, cells were washed twice with PBS and split into two new flasks. In some experiments human epithelial cell lines from the brain (HTB-16), duodenum (HTB-40), kidney (HTB-34), liver (CCL-13), lung (HTB-53) and pancreas (HTB-134) were used, all obtained from ATCC. They were grown as indicated above. Human skin fibroblast cell lines from a patient with Gaucher's disease and one relative were kindly donated by Dr Clara Sa Miranda (Instituto de Genetica Medica, Porto, Portugal). They were maintained in culture media 2 (CM2, see below).

Peripheral blood T lymphocytes

PBL were obtained from buffy coats or from heparinized blood from healthy volunteers after centrifugation over Ficoll–Hypaque. PBL were used unfractionated or were further depleted of non-T cells (monocytes and B lymphocytes) after rosetting with neuraminidase-treated sheep erythrocytes yielding T lymphocytes that were > 95% CD3+. T lymphocytes were placed in RPMI media supplemented with 10% FCS, 1% glutamine, 1% pen/strep and 1% fungizone (CM2).

Isolation of T cell subsets

CD4+ and CD8+ T cells were obtained by negative selection by conventional indirect panning techniques using rabbit anti-mouse immunoglobulin-coated Petri dishes and culture supernatants of mouse anti-human CD4 or CD8 antibodies. The CD4+ and CD8+ selected populations usually contained < 10% contaminating CD8+ and CD4+ cells, respectively, as determined by flow cytometry. CD4+ CD28+, CD8+ CD28+ and CD8+ CD28− T cell subsets were negatively selected on a cell sorter (Becton Dickinson) after enrichment of CD4+ and CD8+ T cell subsets by panning. The mouse anti-human MoAbs CD8-PE, CD4-PE and CD28-PE in combination with a cocktail of CD11b, CD16 and CD57 followed by goat anti-mouse-FITC as second step reagent were used. For example, to obtain CD8+ CD28+ cells by negative selection we sorted for CD4−, CD11b−, CD16− and CD57− cells. To obtain CD8+ CD28−, cells were sorted for CD4−, CD28− and CD16− cells. The negatively sorted populations were > 85% pure and were maintained in CM2.

Adhesion assays

HT-29 and T-84 cells were seeded at 1 × 105 cells/well in 96-well flat-bottomed microtitre plates and grown until confluence in CM1. Before each assay, confluent monolayers were washed twice with PBS to remove dead or floating cells. One to two million cells/ml were labelled for 1 h at 37°C with 150 μCi of 51Cr in RPMI media supplemented with 1% FCS, 1% glutamine, 1% pen/strep and 1% fungizone (CM3). Cells were extensively washed and resuspended in CM3. Labelled cells (0.5–2 × 105) were added on to confluent monolayers or empty wells (negative control) in a final volume of 200 μl CM3 in 96-well plates. Assays were done in triplicate. After 2 h at 37°C, non-adherent lymphocytes were harvested by gentle pipetting followed by two additional gentle washes with CM3. Remaining cells (epithelial cells plus adherent lymphocytes) were harvested after a 30-min treatment with 200 μl of a 1% solution of trypsin/EDTA, followed by two more washes with CM3. The same procedure was done with the empty wells. Radioactivity on the adherent and non-adherent fractions was quantified in a γ-counter. The percentage of cell adhesion was calculated according to the following formula: % adhesion = ((a/b) −(c/d)) × 100, where a = ct/min in adherent fraction (wells with monolayers), b = ct/min in adherent fraction + ct/min in non-adherent fraction (wells with monolayers), c = ct/min in adherent fraction (empty wells), and d = ct/min in adherent fraction + ct/min in non-adherent fraction (empty wells). This method allows subtraction of background adherence that was variable (2–10%) from experiment to experiment.

Co-culture assays

Resting T cells or PBL at 1 × 106 cells/ml were cultured either alone, with equal numbers of the colonic epithelial cell lines HT-29 and T-84 (1:1 ratio) in CM1 or with 2–3 day culture supernatants (50% final dilution) of HT-29 and T-84 cells and placed at 37°C for 1–7 days. In two experiments human epithelial cells from six different origins were also used (see above). After the culture period T lymphocytes were harvested by vigorous pipetting and washed three times with PBS. Microscopic visualization of the co-cultures confirmed that few lymphocytes remained in the wells. T cells or PBL were then stained for CD4, CD8, CD3 and CD28, using optimal dilutions of the following MoAbs: CD4-TriColour (Caltag), CD3-PE and CD8-FITC (Dakopatts) and CD8-FITC/CD28-PE simultest (from Becton Dickinson). Propidium iodide (PI) was used as a marker of dead cells. Cells were analysed by flow cytometry on a FACScan (Becton Dickinson) and 10 000 cells (viable and non-viable) were usually acquired and analysed using the Lysys II software. PI-negative cells were gated and the respective percentage as well as cell numbers of CD3+, CD4+, CD8+ and subsets determined. Co-cultures of primary ex vivo iEC with allogeneic PBL-derived T cells were performed as described elsewhere [32]. Primary iEC were isolated from the tumour-free margins of small intestinal resections after dispase digestion and centrifugation over Hypaque.

Stimulation of cord blood

PBL isolated from normal cord blood were cultured at 5 × 105 cells/well in 2-ml wells with the superantigen toxic shock syndrome toxin-1 (TSST-1; 1 ng/ml), or phytohaemagglutinin (PHA) 2 μg/ml. Cultures were fed with IL-2 (50 U NIH-BRMP/ml) containing CM2 medium × 2/week starting on day 2. Three-colour staining was performed on days 9–11 with TCR-FITC (Pharmingen, San Diego, CA) or Vβ2-FITC (Immunotech/Coulter), CD28-PE (Pharmingen) and CD8-biotin (Olympus). Predetermined optimal dilutions of the antibodies were used. First the cells were stained with the FITC-conjugated antibodies for 30 min at room temperature, then the CD8-biotin was added for 30 min and then CD28-PE was added with streptavidin Tri-Colour (Caltag) for 30 min. Each step was followed by three washes with Hanks', 1% FCS, 0.02% sodium azide, the staining buffer. Samples were read on a Coulter EPICS-XL gated on live cells by forward and side scatter characteristics.

The CD8+ CD28 − and CD8+ CD28+ subsets after co-culture or stimulation

Normal adult and cord blood PBL were placed in culture in CM2 at 1 × 106 cells/ml each under the following conditions; no treatment, IL-2 (50 U NIH-BRMP/ml), phorbol myristate acetate (PMA) 10 ng/ml and ionomyocin 1 μm, PHA 10 μg/ml, TSST-1 1 ng/ml, co-culture on a layer of an equal number of confluent HT-29 cells, and co-culture with an equal number of HT-29 cells that were physically separated in a transwell. The cultures were not supplemented with any media between days 0 and 5.

Statistical analysis

The significance of differences between group means was assessed by Student's t-test.

RESULTS

Adhesion of purified T cell populations to monolayers of iEC

We tested the level of adhesion of purified resting CD4+ and CD8+ T cells as well as unseparated T lymphocytes from several unrelated healthy individuals to monolayers of two different human intestinal epithelial cell lines, HT-29 and T-84. To avoid any possible in vitro effects on the T cells that might affect binding to the monolayers (such as partial stimulation occurring during positive selection), only negatively selected populations were used. All adhesion assays were performed with resting PBL or purified subsets obtained within 24 h of blood collection. The resting status of the cells was confirmed by flow cytometry based on forward and side scatter characteristics. As shown in Fig. 1a, the level of adhesion of purified CD8+ T cells to HT-29 monolayers was three-fold higher than the level of adhesion of purified CD4+ T cells (P < 0.04). This difference was five-fold when T-84 monolayers were used (P < 0.001). Unseparated T cell preparations showed a level of adhesion intermediate between CD8+ and CD4+ T cells. On the other hand, adhesion of negatively sorted CD8+ CD28+ and CD8+ CD28− T cells did not differ from unseparated CD8+ T cell preparations, and both were statistically significantly higher than the level of adhesion of negatively sorted CD4+ CD28+ T cells (Fig. 1b).

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f1.jpg

Adhesion of T lymphocyte subsets to monolayers of colonic epithelial cell lines. 51Cr-labelled T cells or T cell subsets (0.5–2 × 105) were added in triplicate to wells containing monolayers of the epithelial cell lines HT-29 and T-84. After 2 h at 37°C, the percentage of adherent cells was determined. (a) Level of adhesion (mean ± s.d., n = 5 experiments) of unseparated T cells, CD4+ and CD8+ T cells to either HT-29 (left) or T-84 (right). (b) Level of adhesion (mean ± s.d., n = 2) of negatively sorted CD8+ CD28+, CD8+ CD28− and CD4+ CD28+ T cells to either HT-29 (left) or T-84 (right). (c) Level of adhesion (mean ± s.d., n = 2) of CD4+ T cells or CD8+ T cells to monolayers of T-84 cells in the presence (anti-gp180 B9 or anti-CD2 in saturating concentrations) or absence (None) of the indicated antibodies. ND, Not done.

To show that binding of CD8 cells to the iEC cell lines was specific and inhibitable, we tested the ability of anti-gp180 antibody to inhibit binding. Gp180 is a molecule expressed on iEC, and antibodies to gp180 inhibit the activation of CD8-associated p56lck in iEC/T cell co-cultures [33]. MoAb to gp180, but not a MoAb to CD2, blocked adhesion of CD8 cells to T-84 (Fig. 1a). Similar results were observed with HT-29 cells (data not shown).

T cell phenotypic changes after co-culture of PBL with iEC

In order to characterize phenotypic changes in T lymphocytes that have been in contact with dividing iEC, we performed co-cultures for different periods of time. In preliminary experiments using co-cultures of PBL and iEC cell lines, an increase in cells with different light scatter characteristics, as defined by side and forward scatter, was observed after the third day of co-culture. In subsequent 5–6 day co-culture experiments a similar and more prominent population was observed (R2 in Fig. 2). Using PI, a cell dye that enters cells with damaged plasma membranes, this population stained positive and therefore represented dying cells. PI+ cells were positive for CD3 expression, confirming that these were T cells rather than contaminating epithelial cells. Similar results were obtained when PBL were co-cultured with skin fibroblasts or with human epithelial cell lines from duodenum, liver, brain, kidney, pancreas and lung (data not shown).

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f2.jpg

T cell death induced in co-cultures with iEC, fibroblast cell lines, or iEC supernatant. (a) Peripheral blood lymphocytes (PBL) at 106 cells/ml were cultured either alone or with equal cell numbers of HT-29 or fibroblast cells (FBL) in culture media 2 (CM2). Cells were stained after 5 days with 20 μg/ml propidium iodide (PI) and 10 000 cells were acquired on a FACScan and analysed with Lysys II software. Cells were gated according to forward (FSC) and side scatter (SSC) and the percentage of PI+ viable cells in gates R1 and R2. (b) PBL at 106/ml were cultured either alone or with supernatant of 3 day HT-29 cultures. Supernatants were centrifuged at 3000 rev/min for 10 min before use. PBL were harvested at the indicated time points and the percentage of PI− cells determined. Total cell numbers (PI− and PI+) remained approximately the same for both cultures during the observation period. One of two similar experiments is shown.

The phenotype of viable T cells (PI−) resulting from co-cultures with either epithelial cell lines or skin fibroblasts was characterized by a decrease in the percentage of CD8+ T cells and an increase in the percentage of CD4+ and CD3+ T cells, in comparison with cultures of PBL alone (Fig. 3,Table 1). Although total viable cell numbers in all three populations were decreased, the reduction in total CD8+ T cell numbers was most striking (Fig. 3). A preferential decrease in CD8+ T cells after prolonged co-cultures with epithelial cells was observed in most of the individuals studied (Table 1). To determine whether the decrease in CD8+ T cell numbers affected CD8+ CD28+ or CD8+ CD28− T cells, we performed double labelling after the co-culture. Both the percentage and total cell numbers of CD8+ CD28+ T cells among viable lymphocytes were reduced after co-culture (Fig. 4). By contrast, CD8+ CD28− T cells were hardly affected and the mean percentage of CD8+ CD28−/total CD8+ cells rose from 26.2 ± 8.4% to 45.7 ± 14.7% in the presence of co-cultured HT-29 cells (calculated from the data in Fig. 4a, subjects 1–3). This pattern was observed in most of the individuals studied, indicating that CD8+ CD28− T cells are more resistant to the T cell death induced in these cultures. Similar results were obtained in co-cultures with several different epithelial cell lines and with skin fibroblasts.

Table 1

Changes in T cell subsets after co-cultures with iEC

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-t1.jpg
An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f3.jpg

Changes in T cell subsets induced by co-culture with iEC. Peripheral blood lymphocytes (PBL) were cultured for 5 days as in the legend to Fig. 2 and then stained with propidium iodide (PI) and antibodies to CD3, CD4 and CD8. The percentages (and total cell numbers) are indicated for the gated PI− viable cells.

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f4.jpg

Changes in CD8 T cell subsets induced by co-cultures with iEC. Peripheral blood lymphocytes (PBL) from three subjects were cultured either alone or in the presence of HT-29 cells (a). Cultures were always paired for comparisons between control PBL cultured for 5 days versus experimental conditions. (b) PBL were cultured alone, with HT-29 pretreated with mitomycin C or with 3 day supernatant of HT-29 cells. After 5 days PBL were stained for propidium iodide (PI), CD8-FITC and CD28-PE. The data were obtained on PI− viable cells. The decrease in CD8+ CD28+ cells was significant: PBL alone PBL + HT-29 CD8+ CD28+ Mean ± s.d. (n = 5)17.8 ± 5.0 7.9 ± 4.1 Paired t-test (P = 0.009) CD8+ CD28−Mean ± s.d. (n = 5)11.7 ± 5.611.4 ± 8.5 Paired t-test (P = 0.94)

T cell death induced by iEC

To test whether cell–cell contact was required or whether cell death was mediated by secreted factor(s), we performed experiments in which PBL were co-cultured with HT-29 supernatants. Epithelial cell supernatants induced time-dependent T cell death that was maximal by day 7 (Fig. 2b). Similar to the co-cultures with intact iEC, CD8+ CD28− T cells co-cultured with supernatant were resistant to cell death (Fig. 4b). In contrast, co-cultures of mitomycin-treated iEC and PBL induced no significant T cell death in comparison with cultures of PBL alone (Fig. 4b and data not shown). In summary, T cell death is mediated at least in part by factor(s) contained in the supernatant of epithelial cell lines.

In preliminary experiments aimed at characterization of the putative factor(s) involved in this process, we observed that anti-transforming growth factor-beta (TGF-β) antibodies, even at high titres, did not abolish T cell death. Whether the putative supernatant factor(s) are at all related to the immunosuppressive factor described by Ebert and others [34,35] is not clear. In addition to iEC-derived immunosuppressive factors, CD8+ CD28− cells themselves can exert suppression in various assays [8,9].

Ex vivo obtained iEC also induce increased percentages of CD8+ CD28− T cells in co-cultured PBL

In two experiments PBL were co-cultured with freshly isolated ex vivo irradiated iEC. Under these culture conditions there is a proliferative response of the PBL [32], unlike the co-cultures with epithelial cell lines. After 5 days of culture nearly all CD8 cells had switched from a CD28+ to a CD28− phenotype (Table 2). A large percentage also expressed CD25, the IL-2 receptor α-chain. Thus the phenotypic switch of CD8 cells from CD28+ to CD28− also occurs in co-cultures with natural iEC obtained directly ex vivo.

Table 2

Co-culture with iEC results in enrichment of CD8+ CD28− T cells

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-t2.jpg

CD8+ CD28− phenotype specifically induced by TCR-mediated stimulation

We asked whether T cell activation in the co-cultures with allogeneic iEC might be the cause of the switch from a CD28+ to a CD28− phenotype in CD8+ cells. Cord blood cells are devoid of CD28− T cells [3] and are therefore useful to show induction of this phenotype. Cord blood PBL were stimulated with the Vβ2-specific superantigen, TSST-1, or with PHA. By day 9 of culture the CD8 T cells had down-regulated CD28 expression (Fig. 5). These cells resembled the ex vivo CD8+ CD28− cells of a normal adult which were stained simultaneously as a control. In other experiments with TSST-1, the down-regulation of CD28 expression occurred primarily in the Vβ2 CD8+ subset and not in bystander CD8 cells expressing other Vβ. Thus, emergence of CD28− cells occurs in those cells targeted by the superantigen and is therefore directly related to TCR-mediated activation.

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f5.jpg

De novo generation of CD8+ CD28− T cells from cord blood. Nine day cultures of two cord blood PBL with toxic shock syndrome toxin-1 (TSST-1) or phytohaemagglutinin (PHA) are compared with the corresponding ex vivo PBL and with adult ex vivo PBL. By day 9 the TSST-1-targeted Vβ2 subset had increased from 6% to > 50% of CD3+ T cells. The bottom row of quadrant histograms depicts TCRαβ or Vβ2 versus CD8-PE staining with the positive gated population indicated. The top row of quadrant histograms depicts CD28-PE versus CD8-biotin staining in the gated populations with the cell percentages indicated in each quadrant. The one-dimensional histogram for the adult donor is gated on TCR+ CD8+ cells and shows the clearly distinct CD28− and CD28+ subsets.

In Fig. 6 cord blood and adult PBL were stimulated with the superantigen TSST-1, the mitogens PHA or PMA, or by the cultured colonic cell line HT-29 in direct contact or in a transwell. Compared with PBL stained ex vivo or PBL maintained in culture without IL-2 for 5 days, all other conditions for the adult PBL showed evidence of down-modulated CD28 expression in the CD8 subset. The down-modulation of CD28 was most dramatic in cells exposed to HT-29 separated by a transwell. The cord blood showed variable degrees of CD28 modulation. Primary cord blood PBL ex vivo when stained within 6 h of isolation contained very few CD28− cells in either CD8+ or CD8− subsets. By contrast, adult PBL, stained in tandem as a positive control, contained CD28− CD8+ TCRαβ+ T cells (Fig. 6) as previously described [3,14]. Down-regulation of CD28 expression on TCRαβ+ T cells occurred not only in co-cultures with iEC, but also with various other means of T cell activation and with simple culture in FCS-containing media.

An external file that holds a picture, illustration, etc. Object name is cei0112-0226-f6.jpg

CD28 expression on CD8 T cells under different conditions of stimulation. Peripheral blood lymphocytes (PBL) were cultured under different conditions of stimulation to observe the effects on CD28 expression of CD8+ T cells. The cells were stained with TCRαβ-FITC, CD28-PE, CD8-biotin and streptavidin-TriColour, to allow quantification of the percentage of CD8+ CD28+ and CD8+ CD28− cells per total TCRαβ+ cells. A characteristic stain of cord blood and a 34-year-old adult are shown in (a) with the difference in TCRαβ+ CD28− subsets highlighted. The results on day 5 of stimulation and on day 0 are shown for both adult and cord blood in (b), demonstrating an increase in TCRαβ+ CD8+ CD28− cells in culture. Two independent determinations were made with the ex vivo cells.

DISCUSSION

Both in humans and mice the iIEL compartment is populated by a majority of T cells with the characteristic CD8+ CD28− phenotype [21–23]. In mice these cells often express the CD8αα homodimer, thought to indicate their extrathymic origin, but in adult humans most iIEL express the CD8αβ heterodimer [22]. They express the integrin αEβ7 which allows adhesion to epithelial cells by binding to E-cadherin [36,37]. A small percentage of T cells in the peripheral blood also express αEβ7. Are these cells, or the CD8+ CD28− T cells in the blood, destined for the iIEL compartment? Herein we found no difference between CD28+ and CD28− CD8+ PBL-derived T cells in their ability to adhere to iEC. Thus, preferential adhesion is not a likely reason for the dominant CD28− phenotype in the iIEL compartment. However, PBL-derived CD8 T cells adhered three to five times better that resting CD4+ T cells to monolayers of iEC. This adhesion could be blocked with an antibody to gp180, a surface molecule of iEC which may bind to CD8 and activate p56lck [33]. The fact that αEβ7 is expressed by > 95% of iIEL but < 5% of resting PBL [38] appears to rule out the possible role of this integrin in the increased adhesion of the CD8 subsets examined here. The possibility that αEβ7 might have been induced on some cells during the adhesion assay seems unlikely, as maximal expression occurs 5–7 days after antigenic or mitogenic stimulation [39,40]. In the present experiments the purified T cell subsets were tested within 24 h after blood collection and the adhesion assay lasted 2 h. Experiments with PHA-activated CD4 and CD8 cells were also performed (data not shown), but the increased levels of adhesion observed did not differ for any of the subsets tested. Thus resting CD8 cells appear to express adhesion molecules that allow increased adhesion to iEC cell lines compared with resting CD4 cells. A full characterization of these molecules will require further studies.

Our current studies were performed with PBL T cells, not with iIEL T cells. They imply that there is a peripheral blood CD8+ T cell subset which can interact with gut-derived epithelial cells. Binding of resting as well as activated PBL T cells to a variety of different epithelial cell monolayers has previously been described [22,36,37,41,42], but the T cell subsets involved were not characterized. The higher binding level of resting CD8+ T cells to intestinal epithelial cell monolayers observed here reflects the natural in vivo tropism of CD8+ T cells for the iIEL compartment, but there was no difference between CD8+ CD28+ and CD8+ CD28− T cells in this assay, indicating perhaps that both subsets express the required set of adhesion molecules and that loss of CD28 expression may be a consequence of CD8 cell activation in the iIEL compartment rather than a marker of the type of CD8 cells that migrates to the iIEL compartment.

There is evidence that CD8+ CD28− iIEL T cells are antigen-driven cells that have been activated in vivo. This is suggested by their increased size compared with CD28+ CD8+ cells [21], as well as their expression of activation markers [23,41,43–45]. Their spontaneous cytotoxicity [46], transient proliferative hyporesponsiveness to various TCR stimuli [43,47], and the dramatic oligoclonality of CD8+ CD28− T cells that is observed extending throughout the gut all add weight to this argument [27,48–50]. Although the nature of the antigen(s) remains uncharacterized, it was recently reported that signalling through the TCR/CD8-p56lck complex is triggered by non-classical MHC class I molecules, including CD1d, expressed by epithelial cells [28,33]. In a series of recent studies with allogeneic human iEC obtained ex vivo, preferential activation and subsequent proliferation on PBL T cells with the CD8 phenotype was seen [28,51]. These data are now extended by showing that the T cells that accumulate are CD8+ CD28−. In this system T cells proliferate vigorously, which differs from the cultures with epithelial cell lines, where T cell proliferation was not observed. A lack of proliferation was also observd in other studies using iEC cell lines [52,53]. Nevertheless, both systems involve co-cultures of T cells with iEC and in both systems there is accumulation of CD8+ CD28− cells.

Perhaps CD8+ CD28− cells outnumber the CD8+ CD28+ subset among iIEL because they express an alternative costimulatory cell surface molecule for interaction with epithelial cells. A good candidate was CDw101, but this molecule is expressed on practically all iIEL regardless of CD28 expression [31], and CDw101 is primarily expressed on CD28+, not CD28−, T cells in PBL [54]. It is also conceivable that alternative costimulatory molecules are not required. Intestinal epithelial cells from normal or inflamed human intestinal tissue do not express CD80 or CD86 [55], the ligands for CD28. Thus, there would be no need for CD28 expression among iIEL, but perhaps there is a need for CD28 at an earlier stage of maturation, i.e. in the lamina propria where CD80/86+ monocytes and B cells are present. In this case CD8+ CD28+ precursors would give rise to CD8+ CD28− cells after appropriate stimulation. Evidence that lamina propria T cells give rise to iIEL after TCR stimulation in situ has been presented [56].

In this context, we show for the first time that CD8+ CD28− PBL T cells, which are completely absent in cord blood, derive from CD8+ CD28+ precursors after TCR-mediated stimulation. De novo generation of this subset was seen 9 days post-TCR-mediated activation using either superantigens or mitogens. Down-regulation of CD28 expression by mitogenic or superantigen-mediated stimulation was even more efficient than in co-cultures of cord blood cells with iEC.

The small intestine is the site of epithelial transport which may be regulated by inflammatory cells [52,57], and in this context it may be of interest that PBL CD8+ CD28− T cells are increased in haemochromatosis, a disease characterized by lack of regulation of iron transport [58]. In addition, TCR/CD8 signalling is regulated by non-classical MHC class I molecules in the iIEL compartment [33]. A novel non-classical MHC class I molecule has been found to be mutated in familial haemochromatosis, at a position that disrupts interaction with β2-microglobulin, thus abrogating surface expression [59,60]. Might these findings be relevant to the expansions of CD8+ CD28− cells observed in HLA-A3+ haemochromatosis patients?

Here we show that co-cultures of PBL T cells with iEC in two different systems as well as mitogenic stimulation and stimulation with superantigens resulted in gradual enrichment of the CD8+ CD28− subset. Possibly, CD8+ CD28− T cells are less susceptible to activation-induced apoptosis in these cultures. We propose that CD8+ CD8− T cells, in the blood or in the iIEL compartment, arise as a consequence of chronic antigenic stimulation. The nature of this antigenic signal still needs to be characterized.

Acknowledgments

Part of this work was performed at the Laboratory of Human Molecular Immunology, Cornell University Medical College (New York, USA) thanks to a Visiting Fellowship from Junta Nacional de Investigacion Cientifica e Tecnologica (JNICT) granted to F.A.A. We thank Dr Tim McCaffrey for providing anti-TGF-β antibodies. F.A.A. is a PhD Fellow from JNITC (PRAXIS XXI Program, BD 3209/94).

References

1. Linsley PS. Distinct roles for CD28 and cytotoxic T lymphocyte-associated molecule-4 receptors during T cell activation? J Exp Med. 1995;182:289–92. [PMC free article] [PubMed] [Google Scholar]
2. Bluestone JA. New perspectives of CD28- B7-mediated T cell costimulation. Immunity. 1995;2:555–9. [PubMed] [Google Scholar]
3. Azuma M, Phillips JH, Lanier LL. CD28− T lymphocytes. Antigenic and functional properties. J Immunol. 1993;150:1147–59. [PubMed] [Google Scholar]
4. Behar SM, Porcelli SA, Beckman EM, Brenner MB. A pathway of costimulation that prevents anergy in CD28− T cells: B7-independent costimulation of CD1-restricted T cells. J Exp Med. 1995;182:2007–18. [PMC free article] [PubMed] [Google Scholar]
5. Groh V, Porcelli S, Fabbi M, et al. Human lymphocytes bearing T cell receptor γ/δ are phenotypically diverse and evenly distributed throughout the lymphoid system. J Exp Med. 1989;169:1277. [PMC free article] [PubMed] [Google Scholar]
6. Testi R, Lanier LL. Functional expression of CD28 on T cell antigen receptor gamma/delta-bearing T lymphocytes. Eur J Immunol. 1989;19:185–8. [PubMed] [Google Scholar]
7. Hamann D, Baars PA, Rep MHG, Hooibrink B, Van Lier RAW. Phenotypical and functional separation of memory and effector human CD8+ T cells. J Exp Med. 1997;186:1407–18. [PMC free article] [PubMed] [Google Scholar]
8. Lum LG, Orcutt-Thordarson N, Seigneuret MC, Hansen JA. In vitro regulation of immunoglobulin synthesis by T cell subpopulations defined by a new human T-cell antigen (9.3) Cell Immunol. 1982;72:122–32. [PubMed] [Google Scholar]
9. Damle NK, Engleman EG. Immunoregulatory T cell circuits in man: alloantigen-primed inducer T cells activate alloantigen-specific suppressor T cells in the absence of the initial antigenic stimulus. J Exp Med. 1983;158:159–69. [PMC free article] [PubMed] [Google Scholar]
10. Vinci G, Vernant JP, Nakazawa M, et al. In vitro inhibition of normal human hematopoiesis by marrow CD3+, CD8+, HLA-DR+, HNK1+ lymphocytes. Blood. 1988;72:1616–21. [PubMed] [Google Scholar]
11. Wang EC, Lehner PJ, Graham S, Borysiewicz LK. CD8high (CD57+) T cells in normal, healthy individuals specifically suppress the generation of cytotoxic T lymphocytes to Epstein–Barr virus-transformed B cell lines. Eur J Immunol. 1994;24:2903–9. [PubMed] [Google Scholar]
12. Autran B, Leblond V, Sadat-Sowti B, et al. A soluble factor released by CD8+CD57+ lymphocytes from bone marrow transplanted patients inhibits cell-mediated cytolysis. Blood. 1991;77:2237–41. [PubMed] [Google Scholar]
13. Yamashita N, Nguyen L, Fahey JL, Clement LT. Phenotypic properties and cytotoxic functions of human CD8+ cells expressing the CD57 antigen. Nat Immun. 1993;12:79–91. [PubMed] [Google Scholar]
14. Posnett DN, Sinha R, Kabak S, Russo C. Clonal populations of T cells in normal elderly humans: the T cell equivalent to ‘benign monoclonal gammapathy’ J Exp Med. 1994;179:609–18. [PMC free article] [PubMed] [Google Scholar]
15. Fitzgerald JE, Ricalton NS, Meyer AC, et al. Analysis of clonal CD8+ T cell expansions in normal individuals and patients with rheumatoid arthritis. J Immunol. 1995;154:3538–47. [PubMed] [Google Scholar]
16. Dupuy A, Monier S, Jorgensen C, et al. Increased percentage of CD3+, CD57+ lymphocytes in patients with rheumatoid arthritis. Correlation with duration of disease. Arthritis Rheum. 1993;36:608–12. [PubMed] [Google Scholar]
17. Becker H, Knopp A, Doppl W, et al. Increased activation of CD8-positive lymphocytes in patients with Crohn's disease. Immun Infekt. 1993;21:5–8. [PubMed] [Google Scholar]
18. Choremi-Papadopoulou H, Viglis V, Gargalianos P, Kordossis T, Iniotaki-Theodoraki A, Kosmidis J. Downregulation of CD28 surface antigen on CD4+ and CD8+ T lymphocytes during HIV-1 infection. J Acquir Immune Defic Syndr. 1994;7:245–53. [PubMed] [Google Scholar]
19. Dutra WO, Martins-Filho OA, Cancado JR, et al. Chagasic patients lack CD28 expression on many of their circulating T lymphocytes. Scand J Immunol. 1996;43:88–93. [PubMed] [Google Scholar]
20. Arosa FA, Veltman J, Porto G, de Sousa M. Anomalies of the CD8+ population in genetic hemochromatosis: defective numbers of CD8+CD28+ cells. FASEB J. 1995;9:A3052. (Abstr.). [Google Scholar]
21. Ohteki T, MacDonald HR. Expression of the CD28 costimulatory molecule on subsets of murine intestinal intraepithelial lymphocytes correlates with lineage and responsiveness. Eur J Immunol. 1993;23:1251–5. [PubMed] [Google Scholar]
22. Latthe M, Terry L, MacDonald TT. High frequency of CD8 alpha alpha homodimer-bearing T cells in human fetal intestine. Eur J Immunol. 1994;24:1703–5. [PubMed] [Google Scholar]
23. Boll G, Rudolphi A, Spiess S, Reimann J. Regional specialization of intraepithelial T cells in the murine small and large intestine. Scand J Immunol. 1995;41:103–13. [PubMed] [Google Scholar]
24. Rocha B, Vassalli P, Guy-Grand D. The Vβ repertoire of mouse gut homodimeric alpha CD8+ intraepithelial T cell receptor αβ+ lymphocytes reveals a major extrathymic pathway of T cell differentiation. J Exp Med. 1991;173:483–6. [PMC free article] [PubMed] [Google Scholar]
25. Rocha B, von Boehmer H, Guy-Grand D. Selection of intraepithelial lymphocytes with CD8 alpha/alpha co-receptors by self-antigen in the murine gut. Proc Natl Acad Sci USA. 1992;89:5336–40. [PMC free article] [PubMed] [Google Scholar]
26. Wang EC, Moss PA, Frodsham P, Lehner PJ, Bell JI, Borysiewicz LK. CD8highCD57+ T lymphocytes in normal, healthy individuals are oligoclonal and respond to human cytomegalovirus. J Immunol. 1995;155:5046–56. [PubMed] [Google Scholar]
27. Balk SP, Ebert EC, Blumenthal RL, et al. Oligoclonal expansion and CD1 recognition by human intestinal intraepithelial lymphocytes. Sci. 1991;253:1411–5. [PubMed] [Google Scholar]
28. Panja A, Blumberg RS, Balk SP, Mayer L. CD1d is involved in T cell–intestinal epithelial cell interactions. J Exp Med. 1993;178:1115–9. [PMC free article] [PubMed] [Google Scholar]
29. Shahinian A, Pfeffer K, Lee KP, et al. Differential T cell costimulatory requirements in CD28-deficient mice. Sci. 1993;261:609–12. [PubMed] [Google Scholar]
30. Kawai K, Shahinian A, Mak TW, Ohashi PS. Skin allograft rejection in CD28-deficient mice. Transplantation. 1996;61:352–5. [PubMed] [Google Scholar]
31. Russell GJ, Parker CM, Sood A, et al. p126 (CDw101), a costimulatory molecule preferentially expressed on mucosal T lymphocytes. J Immunol. 1996;157:3366–74. [PubMed] [Google Scholar]
32. Panja A, Barone A, Mayer L. Stimulation of lamina propria lymphocytes by intestinal epithelial cells: evidence for recognition of nonclassical restriction elements. J Exp Med. 1994;179:943–50. [PMC free article] [PubMed] [Google Scholar]
33. Li Y, Yio XY, Mayer L. Human intestinal epithelial cell-induced CD8+ T cell activation is mediated through CD8 and the activation of CD8-associated p56lck. J Exp Med. 1995;182:1079–88. [PMC free article] [PubMed] [Google Scholar]
34. Ebert EC, Roberts AI, O'Connell SM, Robertson FM, Nagase H. Characterization of an immunosuppressive factor derived from colon cancer cells. J Immunol. 1987;138:2161–8. [PubMed] [Google Scholar]
35. Qiao L, Schurmann G, Autschbach F, Wallich R, Meuer SC. Human intestinal mucosa alters T-cell reactivities. Gastroenterol. 1993;105:814–9. [PubMed] [Google Scholar]
36. Cepek KL, Parker CM, Madara JL, Brenner MB. Integrin alpha E beta 7 mediates adhesion of T lymphocytes to epithelial cells. J Immunol. 1993;150:3459–70. [PubMed] [Google Scholar]
37. Cepek KL, Shaw SK, Parker CM, et al. Adhesion between epithelial cells and T lymphocytes mediated by E-cadherin and the alpha E beta 7 integrin. Nature. 1994;372:190–3. [PubMed] [Google Scholar]
38. Cerf-Bensussan N, Jarry A, Brousse N, Lisowska-Grospierre B, Guy-Grand D, Griscelli C. A monoclonal antibody (HML-1) defining a novel membrane molecule present on human intestinal lymphocytes. Eur J Immunol. 1987;17:1279–85. [PubMed] [Google Scholar]
39. Schieferdecker HL, Ullrich R, Weiss-Breckwoldt AN, et al. The HML-1 antigen of intestinal lymphocytes is an activation antigen. J Immunol. 1990;144:2541–9. [PubMed] [Google Scholar]
40. Brew R, West DC, Burthem J, Christmas SE. Expression of the human mucosal lymphocyte antigen, HML-1, by T cells activated with mitogen or specific antigen in vitro. Scand J Immunol. 1995;41:553–62. [PubMed] [Google Scholar]
41. Brandtzaeg P, Bosnes V, Halstensen TS, Scott H, Sollid LM, Valnes KN. T lymphocytes in human gut epithelium preferentially express the alpha/beta antigen receptor and are often CD45/UCHL1-positive [published erratum appears in Scand J Immunol 1989 Nov; 30 (5):653] Scand J Immunol. 1989;30:123–8. [PubMed] [Google Scholar]
42. Dobbins WO. Human intestinal intraepithelial lymphocytes. Gut. 1986;27:972–85. [PMC free article] [PubMed] [Google Scholar]
43. De Maria R, Fais S, Silvestri M, et al. Continuous in vivo activation and transient hyporesponsiveness to TcR/CD3 triggering of human gut lamina propria lymphocytes. Eur J Immunol. 1993;23:3104–8. [PubMed] [Google Scholar]
44. Jarry A, Cerf-Bensussan N, Brousse N, Selz F, Guy-Grand D. Subsets of CD3+ (T cell receptor alpha/beta or gamma/delta) and CD3− lymphocytes isolated from normal human gut epithelium display phenotypical features different from their counterparts in peripheral blood. Eur J Immunol. 1990;20:1097–103. [PubMed] [Google Scholar]
45. Lundqvist C, Baranov V, Hammarstrom S, Athlin L, Hammarstrom ML. Intra-epithelial lymphocytes. Evidence for regional specialization and extrathymic T cell maturation in the human gut epithelium. Int Immunol. 1995;7:1473–87. [PubMed] [Google Scholar]
46. Roberts AI, O'Connell SM, Biancone L, Brolin RE, Ebert EC. Spontaneous cytotoxicity of intestinal intraepithelial lymphocytes: clues to the mechanism. Clin Exp Immunol. 1993;94:527–32. [PMC free article] [PubMed] [Google Scholar]
47. Ebert EC. Proliferative responses of human intraepithelial lymphocytes to various T-cell stimuli. Gastroenterol. 1989;97:1372–81. [PubMed] [Google Scholar]
48. van Kerckhove C, Russell GJ, Deusch K, et al. Oligoclonality of human intestinal intraepithelial T cells. J Exp Med. 1992;175:57–63. [PMC free article] [PubMed] [Google Scholar]
49. Gross GG, Schwartz VL, Stevens C, Ebert EC, Blumberg RS, Balk SP. Distribution of dominant T cell receptor beta chains in human intestinal mucosa. J Exp Med. 1994;180:1337–44. [PMC free article] [PubMed] [Google Scholar]
50. Blumberg RS, Yockey CE, Gross GG, Ebert EC, Balk SP. Human intestinal intraepithelial lymphocytes are derived from a limited number of T cell clones that utilize multiple V beta T cell receptor genes. J Immunol. 1993;150:5144–53. [PubMed] [Google Scholar]
51. Mayer LF, Eisenhardt D, Salomon PDH, Bauer WEM, Plous R, Piccinini L. Expression of class II molecules on intestinal epithelial cells in humans. Gastroenterol. 1991;100:3–12. [PubMed] [Google Scholar]
52. Sollid LM, Brandtzaeg P, Kvale D, Gaudernack G, Scott H, Thorsby E. T cell–epithelium interactions in relation to gut immunity. Monogr Allergy. 1988;24:60–65. [PubMed] [Google Scholar]
53. Liu ZX, Sugawara S, Hiwatashi N, et al. Accessory cell function of a human colonic epithelial cell line HT-29 for bacterial superantigens. Clin Exp Immunol. 1997;108:384–91. [PMC free article] [PubMed] [Google Scholar]
54. Gouttefangeas C, Jacquot S, Meffre E, Schmid M, Boumsell L, Bensussan A. Differential proliferative responses in subsets of human CD28+ cells delineated by BB27 mAb. Int Immunol. 1994;6:423–30. [PubMed] [Google Scholar]
55. Bloom S, Simmons D, Jewell DP. Adhesion molecules intercellular adhesion molecule-1 (ICAM-1), ICAM-3 and B7 are not expressed by epithelium in normal or inflamed colon. Clin Exp Immunol. 1995;101:157–63. [PMC free article] [PubMed] [Google Scholar]
56. Monk T, Spencer J, Cerf-Bensussan N, MacDonald TT. Stimulation of mucosal T cells in situ with anti-CD3 antibody: location of the activated T cells and their distribution within the mucosal micro-environment. Clin Exp Immunol. 1988;74:216–22. [PMC free article] [PubMed] [Google Scholar]
57. McKay DM, Croitoru K, Perdue MH. T cell–monocyte interactions regulate epithelial physiology in a coculture model of inflammation. Am J Physiol. 1996;270:C418–C428. [PubMed] [Google Scholar]
58. McLaren GD, Nathanson MH, Jacobs A, Trevett D, Thomson W. Regulation of intestinal iron absorption and mucosal iron kinetics in hereditary hemochromatosis. J Lab Clin Med. 1991;117:390–401. [PubMed] [Google Scholar]
59. Feder JN, Gnirke A, Thomas W, et al. A novel MHC class I-like gene is mutated in patients with hereditary haemochromatosis. Nat Genet. 1996;13:399–408. [PubMed] [Google Scholar]
60. Feder JN, Tsuchihashi Z, Irrinki A, et al. The hemochromatosis founder mutation in HLA-H disrupts beta2-microglobulin interaction and cell surface expression. J Biol Chem. 1997;272:14025–8. [PubMed] [Google Scholar]
Articles from Clinical and Experimental Immunology are provided here courtesy of British Society for Immunology
-