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CD11b Expression Identifies CD8⁺CD28⁺ T Lymphocytes with Phenotype and Function of Both Naive/Memory and Effector Cells¹

Simona Fiorentini^*, Stefano Licenziati^*, Giulio Alessandri^*, Francesco Castelli, Silvio Caligaris, Monica Bonafede^*, Manuela Grassi^*, Emirena Garrafa^*, Andrea Balsari, Adolfo Turano^* and Arnaldo Caruso^2,*

^* Institute of Microbiology and Department of Infectious Diseases, University of Brescia Medical School, Brescia, Italy; and Chair of Immunology, National Cancer Institute, University of Milan, Milan, Italy

	Abstract

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A previously unreported CD8⁺CD28⁺CD11b⁺ T cell subset occurs in healthy individuals and expands in patients suffering from primary viral infections. In functional terms, these cells share the features of naive/memory CD8⁺CD28⁺CD11b^- and terminally differentiated effector CD8⁺CD28^-CD11b⁺ subpopulations. Like CD28^- cells, CD28⁺CD11b⁺ lymphocytes have the ability to produce IFN-, to express perforin granules in vivo, and to exert a potent cytolytic activity. Moreover, these cells can respond to chemotactic stimuli and can efficiently cross the endothelial barrier. In contrast, like their CD11b^- counterpart, they still produce IL-2 and retain the ability to proliferate following mitogenic stimuli. The same CD28⁺CD11b⁺ subpopulation detected in vivo could be generated by culturing naive CD28⁺CD11b^- cells in the presence of mitogenic stimuli following the acquisition of a CD45RO⁺ memory phenotype. Considering both phenotypic and functional properties, we argue that this subset may therefore constitute an intermediate phenotype in the process of CD8⁺ T cell differentiation and that the CD11b marker expression can distinguish between memory- and effector-type T cells in the human CD8⁺CD28⁺ T cell subset.

	Introduction

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The CD8 T cell subset can be classified according to CD28 surface marker expression as either CD28⁺ or CD28^-, with different biological properties and abilities to produce soluble elements such as cytokines. The CD28⁺ T cell population is predominant in healthy individuals and expands during primary viral infection (1), whereas CD28^- cells are more common in healthy elderly individuals (2) and increase dramatically in HIV patients (3, 4, 5).

CD28⁺ is the only T cell subset present in cord blood (6), which indicates that naive unprimed cells display this phenotype. In adults the population consists of both CD45RA and CD45RO phenotypes, suggesting the presence of both naive and memory cells in the subset (6, 7). Such cells mostly produce IL-2 and exhibit the ability to proliferate in vitro following mitogenic stimulation.

The CD28^- population consists of a subset of memory T cells, and in many respects their phenotype and function are highly suggestive of differentiated armed effector T cells, as they produce IFN- and TNF- and express high levels of granzyme A and perforin. They also contain Ag-specific memory CTL (8, 9) and exert a potent cytolytic activity without requiring deliberate in vitro activation (2, 6). CD28^- are clonally expanded (10), terminally differentiated lymphocytes with little proliferative response to mitogenic stimulation in vitro (11).

The origin of CD28^- cells has long been controversial, but recent data show that they derive from their CD28⁺ counterpart. We were able to generate CD28^- cells in vitro from long term IL-2-stimulated CD28⁺ T cells through an intermediate CD28^dim+ phenotype. The in vitro transformation of CD28⁺ to a stable CD28^- phenotype involved the acquisition of various biological functions (12), thus suggesting a link to the normal pattern of functional CD8⁺ T cell maturation. Labalette et al. (13) confirmed our data, highlighting also the possibility of IL-4 to prevent loss of CD28 expression.

It is well known that CD8⁺CD28⁺ T cells are negative for ₂ integrin -chain CD11b expression, whereas almost all CD28^- cells are CD11b⁺ (14). ₂ integrins mediate the adhesion of lymphocytes to endothelial cells and extravasation (15, 16); they are also required for homing to inflamed tissues (17). Moreover, in the mouse model, the expression of CD11b on CD8⁺ T cells has been associated with acquisition of cytotoxic capacity (18).

We assumed that the functional differentiation of CD8⁺ T cells into mature effector cells, with the disappearance of CD28 and the appearance of CD11b molecules, has to be gradual. It should therefore be possible to observe in vivo, during primary viral infection, the emergence of elements characterized by CD28 and CD11b molecule coexpression. This present article describes the presence of CD8⁺CD28⁺CD11b⁺ T cells in freshly collected PBMCs from healthy donors and, to a greater extent, in PBMCs from virus-infected patients. Such cells have all the functions of effector cells but retain the ability to proliferate in vitro. Finally, we observed that CD8⁺CD28⁺CD11b⁺ emerged from both adult and cord blood IL-2-stimulated CD28⁺CD11b^- cells during their transformation into a CD28^- phenotype.

	Materials and Methods

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Reagents and Abs

PMA, ionomycin, monensin, PBS, BSA, saponin, PHA, paraformaldehyde, L-lysine monohydrochloride, sodium azide (NaN₃), and Hystopaque density gradient were supplied by Sigma (St. Louis, MO). RPMI 1640 medium, FCS, L-glutamine, penicillin, and streptomycin were purchased from Life Technologies (Paisley, U.K.). Endothelial basal medium (EBM),³ heparin, epidermal growth factor, and bovine brain extract were purchased from BioWhittaker (Walkersville, MD). Purified anti-CD3 (OKT3) mAb was supplied by Ortho (Raritan, NJ). FITC- or PE-conjugated anti-CD3, anti-CD4, anti-CD8, anti-CD11b, anti-CD16, anti-CD18, anti-CD28, anti-CD45RA, anti-CD45RO, anti-CD57, anti-perforin, anti-IL-2, PerCP-conjugated anti-CD3, and CyChrome-conjugated anti-CD28 were all supplied by Becton Dickinson (San Jose, CA). mAb IGMB17, an anti-human IFN- mAb, was produced in our laboratory (19). Recombinant human TNF- and recombinant chemokine macrophage inflammatory protein-1 (MIP-1) were purchased from PeproTech (London, U.K.). Recombinant IL-2 came from Roche (Basel, Switzerland). Na⁵¹CrO₄ and [³H]TdR aqueous solutions were supplied by ICN Biomedicals (Irvine, CA).

Patients

Blood samples were obtained from 18 patients suffering from infectious mononucleosis (n = 2), CMV (n = 3), varicella (n = 4), herpes-zoster (n = 4), and measles (n = 5). Blood samples were also obtained from 25 healthy donors and three fetal cords. Mononuclear cells were obtained from heparinized blood by Hystopaque density gradient.

Flow cytometric analysis

Lymphocyte subsets were evaluated on whole fresh blood using different mAb panels. Two- and three-color phenotypic characterizations of lymphocytes were performed as previously described (3). Briefly, 100 µl of heparinized blood was incubated for 30 min on ice with the appropriate amounts of mAb. Cells were then lysed with buffer (FACS lysing solution, Becton Dickinson) and analyzed by flow cytometry (FACScan, Becton Dickinson). The lymphocyte gate was set using the log fluorescence of a two-color Ab panel (Leukogate (anti-CD45 and anti-CD14 mAbs), Becton Dickinson) with linear 90° side scatter. Live gating was used to collect 10,000 events within the lymphocyte gate defined by Leukogate (Becton Dickinson) staining as CD45^bright+ with low side scatter (20). The resulting data were analyzed with CellQuest software (Becton Dickinson).

Purification of CD8⁺ T cell subsets

CD8⁺ cells were purified from lymphocytes by positive selection using anti-CD8 magnetic beads (MiniMACS, Miltenyi Biotec, Bergisch Gladbach, Germany). Sorting of CD8⁺CD28⁺CD11b^-, CD8⁺CD28⁺CD11b⁺, and CD8⁺CD28^-CD11b⁺ T cells was performed from purified CD8⁺ lymphocytes stained with FITC-conjugated anti-CD11b (Immunotech, Marseilles, France), PE-conjugated anti-CD28, and PerCP-conjugated anti-CD3, by flow cytometry (FACSVantage, Becton Dickinson). Only preparations with purity >98% were used for experiments.

Lymphokine production

Purified CD8⁺ T cell subsets (3 x 10⁵ cells/well) were cultured in 24-well plates in complete medium (RPMI 1640 supplemented with 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% of heat-inactivated FCS) and stimulated or not with 10 ng/ml PMA plus 1 µM ionomycin in the presence of 1 µM monensin, which inhibits intracellular traffic pathways leading to protein accumulation (21). Plates were incubated at 37°C in 5% CO₂ in air, and cells were collected for lymphokine staining at 6 and 12 h after the addition of stimulants. The cells were then stained for lymphokine production as previously described (22). Briefly, they were washed twice with PBS, pH 7.2, and suspended using ice-cold periodate-lysine-paraformaldehyde solution for 15 min at -10°C. After two washes in PBS, the cells were resuspended to 1 x 10⁶ in 300 µl of PBS containing 1% BSA (w/v), 0.2% NaN₃ (v/v), and 0.1% saponin (saponin buffer). After 15-min incubation at room temperature, the fixed and permeabilized cells were centrifuged and resuspended in saponin buffer containing 1 µg/ml FITC-conjugated anti-IFN- and 1 µg/ml PE-conjugated anti-IL-2 mAb. The cells were then incubated for 30 min at 4°C and washed with saponin buffer. Stained cells were analyzed by flow cytometry as described above. CD8⁺ T cell subsets were also cultured for 48 h in complete medium containing PMA and ionomycin. Cell-free culture supernatants were collected and assayed for the presence of IFN- or IL-2 by ELISAs (BioSource, Camarillo, CA).

Cell proliferation

Purified CD8⁺ T cell subsets were seeded in triplicate at different concentrations (ranging from 2 x 10⁴ to 6 x 10⁴) in 96-well culture plates in complete medium and stimulated with PHA (5 µg/ml) and IL-2 (10U/ml). On day 3 the cells were pulsed with 1 µCi/ml of [³H]TdR and harvested after 18 h. The SD of the three replicate samples was <10% of the mean in all experiments.

Intracellular perforin detection

Purified CD8⁺ cells were washed twice with PBS, pH 7.2, and fixed in suspension using a 4% paraformaldehyde solution for 5 min at room temperature. After two washes in PBS, cells were resuspended to 1 x 10⁶ in 1 ml of PBS containing 1% BSA (w/v), 0.2% NaN₃ (v/v), and 0.2% saponin (saponin buffer). After a 15-min incubation at room temperature, fixed and permeabilized cells were centrifuged and resuspended in saponin buffer containing FITC-conjugated anti-CD11b, PE-conjugated anti-perforin, and CyChrome-conjugated anti-CD28 mAbs. Cells were then incubated for 30 min at 4°C and washed with saponin buffer. Stained cells were analyzed by flow cytometry as described above.

Cytotoxicity assay

The cytotoxic activity of CD8⁺ T cell subsets was evaluated in an anti-CD3-redirected cytotoxicity assay as described previously (23). Briefly, 5 x 10⁵ Fc receptor-bearing P815 target cells were labeled with 50 µCi of Na⁵¹CrO₄ for 2 h at 37°C. Cells were then washed three times and incubated for 30 min at 4°C in the presence or the absence of 2 µg of anti-CD3 mAb. Purified CD8⁺ T cells were stained for CD11b, CD16, and CD28, but only CD16^- cells were gated and sorted according to their CD11b and CD28 phenotypes. Cells were incubated for 4 h at 37°C with 5 x 10³ P815 target cells at E:T cell ratios ranging from 20:1 to 1:1. Supernatants were then collected and counted. Specific cytotoxicity was calculated as follows: cpm of experimental release - cpm of spontaneous release/cpm of maximum release - cpm of spontaneous release x 100. The SE of the mean percentage lysis never exceeded 5%.

Lymphocyte adhesion to human microvascular endothelial cell cultures

Primary cultures of human adrenal gland capillary endothelial cells (HACECs) were obtained as previously described (24). The endothelial cells were plated onto collagenated 96-well plates at a concentration of 5 x 10³/well in 100 µl of EBM containing 10% FCS, heparin (100 µg/ml), epidermal growth factor (10 ng/ml), and bovine brain extract (15 µg/ml; EBM complete medium). The plates were incubated for 4–5 days to obtain a monolayer. Endothelial cells were activated by adding TNF- (10 ng/ml) for 6 h at 37°C. Cells were then washed with PBS and allowed to interact with purified CD8⁺ T cell subsets (2 x 10⁴ lymphocytes/well in RPMI 1640, containing 0.2% BSA). The plates were incubated for 2 h at 37°C, and unbound lymphocytes were removed by three washes with warm PBS. The lymphocytes attached to endothelial cells were fixed for 5 min with 100 µl of cold methanol, and the cells were stained with Diff-Quick (Merz-Dade, Dudingen, Switzerland) for 30 min at room temperature. Plates were then washed several times with deionized water, and the lymphocytes bound to endothelial cells were counted with a calibrated eyepiece in 15 different fields at x200 magnification. Each test was run in quadruplicate.

Chemotaxis and migration assays

All migration assays were performed in collagen-coated 24-well Trans-well culture inserts (6.5 mm diameter clear polycarbonate membrane with 3-µm pores; Costar, Cambridge, MA). The medium used was RPMI 1640 containing 0.2% BSA. All migration assays were conducted for 4 h at 37°C. Purified CD8⁺ T cell subsets (2 x 10⁵) were placed in the upper chamber in 200 µl; 500 µl of medium containing, or not, MIP-1 was added in the lower well. The optimal chemotactic dose for MIP-1 was 100 ng/ml.

Transendothelial migration experiments were performed simultaneously with chemotaxis. HACECs (5 x 10⁴) were seeded on collagen-coated 24-well plate Trans-well culture inserts and cultured in EBM complete medium until confluence was reached. Under these conditions, HACECs did not cross the membrane and formed a complete monolayer usually after 2 days of culture and only on the upper surface of the filter, as confirmed by staining with Diff-Quick of a batch of Trans-well inserts before use. The HACEC monolayer was treated for 6 h with 10 ng/ml of TNF-. Cells were washed three times, and CD8⁺ T cell subpopulations were added to the insert before Trans-well immersion. The Trans-well inserts were then removed, and migrated cells were collected by centrifugation and counted as described above. Tests were run in triplicate.

Long term culture of CD8⁺ T cells

Purified CD8⁺CD28⁺ T cells (1 x 10⁵) were seeded in 96-well microtiter wells (Nunc, Roskilde, Denmark) in 200 µl of RPMI 1640 complete medium supplemented with PHA (5 µg/ml), IL-2 (100 U/ml), and irradiated (3000 rad) autologous PBMCs (10⁵ cells/well). For cellular expansion, growing cells were split twice a week, and 100 µl of medium was replaced with a fresh aliquot containing 100 U/ml of IL-2. Flow cytometric analysis of growing cells was performed once a week. The viability of the cells at any time of harvesting always exceeded 80% as determined by flow cytometry. Live gating was defined as previously described (5). Cord blood lymphocytes were long term cultured and phenotypically characterized as described above.

Statistical analysis

The data were analyzed by (multivariate) variance analysis. Student’s t test was used to determine significant differences between group averages. When multiple individual groups were compared, p values were corrected with the Bonferroni correction. Significance was defined as p 0.05.

	Results

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CD8⁺CD28⁺CD11b⁺ elements in healthy individuals and in patients suffering from various viral infections

The presence of CD8⁺CD28⁺CD11b⁺ T lymphocytes was evaluated in blood samples from healthy individuals and patients suffering from primary viral infections, characterized by activated cell-mediated immunity. Staining of PBMCs with mAbs to CD8, CD11b, and CD28 revealed three subsets of CD8⁺ T lymphocytes: a CD28⁺CD11b^- subset and two CD11b⁺ subsets (CD28⁺CD11b⁺ and CD28^-CD11b⁺). As shown in Table I, in healthy individuals the CD8⁺CD28⁺CD11b^- subset prevailed over CD28^-CD11b⁺, whereas the CD28⁺CD11b⁺ subset was only barely present. In patients suffering from acute viral infections (n = 18) we observed a significant increase (p < 0.01) in CD11b⁺ cells compared with healthy individuals (n = 25; mean, 41.9 ± 14.2 and 29.0 ± 13.9%, respectively). The increase in CD11b⁺ cells was generally found to be linked to the CD28⁺ subset. In fact, the percentage of CD28⁺CD11b⁺ lymphocytes in patients exhibited an average 3.4-fold increment compared with healthy donors, whereas the percentage of CD28^-CD11b⁺ cells showed no significant difference between the two groups (Table I). The increase in CD28⁺CD11b⁺ cells depended on the virus causing the disease (Fig. 1). At the onset of symptoms, EBV consistently elicited the largest increase in the CD28⁺CD11b⁺ subset (60% of all CD8⁺ T cells) compared with the other viruses considered (range, 9.8–49.7%). All the patients showed a decline in the percentage of CD11b⁺ cells to normal levels as their disease gradually resolved (data not shown).

View larger version (55K): [in this window] [in a new window]   FIGURE 1. Flow cytometric analysis of CD28 and CD11b expression. Blood was obtained from a representative healthy donor (A) and, at the onset of symptoms, from patients suffering from infectious mononucleosis (B), varicella (C), and measles (D). An electronic gate was set on CD8bright+ T cells, and the lymphocytes were evaluated for expression of CD28 and CD11b markers. Data were analyzed using CellQuest software and are displayed as bivariate dot plots. The percentage of cells in each quadrant is given in the upper right corner of each panel.

T cell subsets bearing different surface receptors may also display different functions. The dramatic increase in the percentage of CD28⁺CD11b⁺ T cells during primary viral infections suggests that in humans these may function as primed cells that expand in response to Ag stimulation. Indeed, evaluation of naive and memory cells within the CD28⁺CD11b⁺ subset using CD45RA and CD45RO isoform expression, respectively, revealed that most of these cells had a memory CD45RO⁺ phenotype, whereas considerable heterogeneity was observed within CD28⁺CD11b^- and CD28^-CD11b⁺ populations. The CD28⁺CD11b⁺ population exhibited features of Ag-primed cells, because CD18 was up-regulated (16, 25) compared with CD28⁺CD11b^- (mean fluorescence intensity (MFI), 99 and 65, respectively) but expressed at less intensity than on CD28^-CD11b⁺ cells (MFI, 120). This population, however, did not show a fully differentiated activated effector phenotype. Indeed, while a high percentage of CD28^-CD11b⁺ cells (71%) expressed the CD57 molecule, an effector cell-associated molecule (26), only 14% of CD28⁺CD11b⁺ cells were CD57⁺, a percentage almost comparable to that of CD28⁺CD11b^- lymphocytes (9%). These results suggest that CD28⁺CD11b⁺ cells may be an intermediate phenotype between CD28⁺CD11b^- and CD28^-CD11b⁺ T cells.

The cytokine-producing capacity of CD8⁺ T cell subsets

Unlike naive/memory cells, which mainly synthesize IL-2, the ability to produce IFN- is a typical feature of effector T cells (27, 28). CD8⁺ lymphocytes were purified from freshly collected PBMCs and stained for CD3, CD28, and CD11b. CD8⁺ T cell subsets, namely, CD28⁺CD11b^-, CD28⁺CD11b⁺, and CD28^-CD11b⁺, were then sorted by flow cytometry. The cytokine-producing capacity of CD8⁺ cell subsets was measured after stimulation for 12 h with PMA and ionomycin at the single-cell level. As shown in Fig. 2A, the CD28⁺CD11b^- subset had a higher percentage of IL-2⁺ cells (mean, 18.9 ± 5.7%) than the CD28^- subset (mean, 2.1 ± 1.8%). Conversely, the percentage of IFN--expressing cells was higher in the CD28^-CD11b⁺ than in the CD28⁺CD11b^- subset (mean, 81.3 ± 18.0 and 23.9 ± 11.5%, respectively). Notably, CD28⁺CD11b⁺ cells showed an intermediate pattern of IL-2 and IFN- expression (mean, 6.3 ± 3.1 and 40.6 ± 17.6%, respectively). Results obtained by flow cytometry were confirmed by measurements of cytokines in culture supernatants by ELISA. Again, CD28⁺CD11b^- CD28^-CD11b⁺ T cells mainly produced IL-2 and IFN-, respectively, whereas CD28⁺CD11b⁺ cells were capable of secreting both cytokines (Table II).

View larger version (14K): [in this window] [in a new window]   FIGURE 2. A, Flow cytometric detection of cytokine production. Purified CD8+ lymphocyte subsets from a representative healthy donor were stimulated with PMA (10 ng/ml) plus ionomycin (1 µM) in the presence of monensin (1 µM). Cells were then fixed, permeabilized, and stained for IFN- and IL-2 protein expression as described in Materials and Methods. Histograms show the staining for the intracellular IL-2 (solid line) and IFN- (dotted line). B, Proliferation of CD8+ T cell subsets. CD28+CD11b- (•), CD28+CD11b+ (), and CD28-CD11b+ () T cells from five healthy donors were purified and activated with PHA (5 µg/ml) in the presence of IL-2 (10 U/ml). The proliferative response was measured by [3H]TdR incorporation. Results are the mean ± SD of all the experiments performed.

To test whether the distinction in cytokine production profiles among the three CD8⁺ cell subsets was reflected by their proliferative capacities, the three CD8⁺ subpopulations were stimulated with a combination of PHA and IL-2. As expected, CD28^-CD11b⁺ cells exhibited no proliferative activity in a [³H]TdR incorporation assay (6, 11). Instead, both CD28⁺CD11b^- and CD28⁺CD11b⁺ subsets showed a proliferative response to mitogenic stimulation (Fig. 2B), thus indicating that despite IFN- production the latter group maintains a replicative capacity, which shows that they are nonterminally differentiated effector cells.

CD28⁺CD11b⁺ exhibit perforin expression and cytolytic activity without prior in vitro activation

The protective response in tissues is mediated by mature effector T cells. These lymphocytes are able to produce cytokines such as IFN- and develop in vivo the enzymatic machinery necessary for the exocytic pathways of cytolysis (29, 30). Because the three CD8⁺ subsets differed for cytokine production, we investigated whether there were also differences in intracellular molecules involved in cytolysis, such as perforin. Freshly isolated CD8⁺ cells were stained with anti-CD28, anti-CD11b, and anti-perforin mAbs and were analyzed by flow cytometry. As shown in Fig. 3A, CD28⁺CD11b^- cells did not contain perforin, while high staining was observed in the CD28^- population. As earlier remarked for the analysis of cytokine production, cells with a CD28⁺CD11b⁺ phenotype exhibited an intermediate pattern of perforin expression. We next investigated whether CD28⁺CD11b⁺ cells could exert cytotoxic activity even without previous in vitro activation. The cytotoxic activities of the three CD8⁺ subsets were evaluated using freshly purified lymphocytes as effector cells and the mouse cell line P815 as a nonspecific target in a CD3-redirected cytotoxicity assay. CD28^-CD11b⁺ cells exhibited an efficient cytotoxic response, with >50% specific lysis at an E:T cell ratio of 20:1, whereas CD28⁺CD11b^- cells were unable to efficiently lyse target cells (<10% lysis) at the same E:T cell ratio. Remarkably, the CTL activity of CD28⁺CD11b⁺ was approximately half the cytotoxicity of CD28^- cells. The percentage of lysis reached by CD28^- cells at an E:T cell ratio of 5:1 or 10:1 was achieved with CD28⁺CD11b⁺ cells at E:T cell ratios of 10:1 and 20:1, respectively (Fig. 3B).

View larger version (22K): [in this window] [in a new window]   FIGURE 3. A, Expression of perforin in CD8+ T cell subsets. After purification from freshly collected PBMCs, CD8+ cells were stained with anti-CD28 and anti-CD11b mAbs. Cells were then fixed, permeabilized, and stained for intracellular perforin. The histogram shows the staining for the intracellular protein. The solid line indicates perforin-expressing CD28+CD11b- cells, whereas the bold and dotted lines show perforin-expressing CD28+CD11b+ and CD28-CD11b+ cells, respectively. B, Cytotoxic capacity of CD8+ subpopulations. Purified CD8+ T cell subsets, namely CD28+CD11b- (•), CD28+CD11b+ (), and CD28-CD11b+ (), were directly analyzed for cytotoxicity against P815 target cells in the presence of anti-CD3 mAb in a 4-h 51Cr release assay. Results are the mean of at least three independent experiments.

Effector cells are characterized by further unique biological properties, such as the ability to home to peripheral tissue and secondary lymphoid organs by adhesion to the capillary endothelium and by transendothelial migration (31, 32, 33). All these functions are important in allowing effector cells to exert their protective response in tissues. Integrins mediate the adhesion of lymphocytes to endothelial cells and, accordingly, lymphocyte extravasation (34, 35). We therefore investigated whether the integrin -chain CD11b expression on the surface of CD28⁺ cells renders them capable of adhering to HACECs. As shown in Fig. 4, a high number of CD28^-CD11b⁺ cells adhered to HACECs, whereas only a few CD28⁺CD11b^- cells bound to them. The expression of CD11b molecules on the surface of CD28⁺ cells allowed them to adhere to HACECs with almost the same efficiency showed by CD28^-CD11b⁺ cells.

View larger version (23K): [in this window] [in a new window]   FIGURE 4. Binding of purified CD8+ T cell subsets to HACEC. CD8+ T cell subsets were purified from the blood samples of five healthy donors and allowed to interact with a monolayer of TNF--stimulated HACECs. Lymphocytes bound to HACECs were counted using a calibrated eyepiece in 15 different fields at x200 magnification. Each test was run in quadruplicate. Bars represent the mean number of bound CD28+CD11b- (), CD28+CD11b+ (), and CD28-CD11b+ () lymphocytes per microscopic field ± SD.

View larger version (21K): [in this window] [in a new window]   FIGURE 5. A, Migration of freshly isolated and purified CD8+ T cell subsets. The number of cells migrating into the bottom well in the absence or presence of MIP-1 was evaluated by counting cells from duplicate wells. The net migration of cells was calculated by subtracting the number of cells migrating in the absence of a chemoattractant from the number of cells migrating in response to MIP-1. The experiment is representative of results obtained with cells from three independent healthy donors. B, Transendothelial migration of CD8+ T cell subsets. The number of cells migrated through TNF--activated HACECs in the absence or the presence of MIP-1 was evaluated as described above. The experiment is representative of results obtained with cells from three independent healthy donors.

As evaluated by flow cytometric analysis, no modulation of CD11b or CD28 marker expression was observed on CD28⁺CD11b⁺ and CD28^-CD11b⁺ cells after MIP-1 chemoattraction, whereas a slight decrease in the MFI of CD11b marker expression was observed on both cell subsets after transendothelial migration (data not shown).

Development of CD11b⁺ and CD28^- phenotypes from CD8⁺CD28⁺ T lymphocytes

The stability of the CD28⁺ phenotype and the acquisition of CD11b molecule expression were evaluated in long term cultures of CD8⁺CD28⁺CD11b^- T lymphocytes. When CD28⁺CD11b^- cells were purified and stimulated in vitro, almost all the cells recovered after 10 days were CD45RO⁺. At the same time, we observed a consistent percentage of CD28⁺CD11b⁺ cells, which expanded over time, usually reaching a peak at 3–4 wk after mitogenic stimulation. Following prolonged culture (5–6 wk), the percentage of CD28⁺CD11b⁺ cells decreased concomitantly with an increase in the percentage of CD28^-CD11b⁺ cells. To investigate the likelihood of naive CD8⁺ cells giving rise to CD28⁺CD11b⁺ cells, cord blood T lymphocytes were polyclonally stimulated; all CD8⁺ cord blood lymphocytes showed a CD28⁺CD11b^- phenotype when freshly analyzed but, similarly to peripheral blood CD28⁺ lymphocytes, they acquired a CD45RO⁺ phenotype after 10 days of mitogenic stimulation. Flow cytometric analysis showed that a high percentage of CD28⁺ cells (usually >20%) acquired a CD11b⁺ phenotype after 2-wk stimulation. The expansion of CD28⁺CD11b⁺ cells usually peaked at 3–5 wk and later declined, concomitantly to the emergence of CD28^-CD11b⁺ elements. Data are representatively shown in Fig. 6 and are summarized in Table III.

View larger version (43K): [in this window] [in a new window]   FIGURE 6. Modulation of CD11b and CD28 marker expression on CD8+CD28+CD11b- T cells. Flow cytometrically sorted CD8+CD28+CD11b- T cells from a representative healthy donor (A) were harvested at 14 (A), 21 (B), and 35 (C) days from the beginning of culture and stained for CD3, CD11b, and CD28. Samples were analyzed by three-color flow cytometry. All cells showed a CD3+ phenotype at all test times. Data are given as bivariate dot plots, and the percentages of CD28+CD11b-, CD28+CD11b+, and CD28-CD11b+ T cells are shown in the upper right corner of each panel. Lymphocytes of a representative cord blood sample were harvested at 14 (D), 35 (E), and 52 (F) days from the beginning of culture and stained for CD8, CD11b, and CD28. An electronic gate was set on CD8bright+ T cells, and lymphocytes were then evaluated for the expression of CD28 and CD11b markers. Data are given as described above.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

We found that during the acute phase of a primary viral infection such as infectious mononucleosis most CD8⁺ T cells were CD28⁺CD11b⁺, with a massive presence of CD45RO⁺ cells within the subset. Long term cultures of purified CD8⁺CD28⁺CD11b^- cells, originally containing both CD45RA⁺ and CD45RO⁺ phenotypes, gave rise to an entirely CD45RO⁺ population, which gradually acquired CD11b marker expression. To establish whether CD11b⁺ cells also originate from unprimed naive CD8⁺ T cells, we cultured cord blood lymphocytes and found again that all the cells switched to a CD45RO⁺ phenotype before acquiring the CD11b molecule. These findings suggest that all CD8⁺CD28⁺ T cells regardless of their CD45 isotype expression may acquire an activated CD11b⁺ phenotype, and that acquisition of the CD45RO⁺ phenotype is necessary, at least in vitro, for further development into CD11b⁺ and eventually into CD28^- terminal effector cells. With respect to the parameter analyzed, it appears therefore that the same subsets of memory/effector T cells detected in vivo can also be generated by stimulating cultured naive cells. Hamman et al. (41) have recently suggested a model of human CD8⁺ T cell differentiation in which effector cells may arise from a proliferating memory pool (CD45RA^-CD28⁺) and acquire, during the process of down-regulation of CD28, the features of mature effectors, but the difficulty with this model is that both memory and effector cells, within the CD28⁺ subset, express CD45RO, so there was no way to distinguish between the two. Our data demonstrate that the early stages of CD8⁺ memory differentiation into effector cells are characterized by acquisition of a CD11b⁺ phenotype. Indeed, CD28⁺CD11b^- cells are more similar in behavior to true memory cells, being incapable of cytotoxicity and transendothelial migration, whereas their CD11b⁺ counterpart has all the properties of fully competent effector cells. CD11b expression and CD45RO⁺CD28⁺ phenotype therefore distinguish nonterminally differentiated effector cells from the memory pool. As suggested above, the CD28⁺CD11b⁺ subset provides a model for a critical step in the development of functional CTL, which precedes the process of CD28 down-regulation.

A recent study highlights the relationship between the functional activities of lymphocytes and their migration properties. Cells migrating to lymph nodes lack inflammatory and cytotoxic function, whereas cells migrating to peripheral tissues are endowed with various effector functions (25). CD11b has been described as an important molecule for the extravasation of neutrophils and monocytes to the site of inflammation; it is also involved in adhesion, chemotaxis, and diapedesis (42). Our report demonstrates an increase in the capacity of CD28⁺ cells to migrate in response to MIP-1 at the time that they acquire expression of CD11b, thus supporting the prospect of an effector CD28⁺ subset with tissue-homing properties. Indeed, it was reported that CD11b⁺ cells are present in blood, liver, and spleen, but are absent from tonsil, lymph node, and thymus (43).

In conclusion, the present study demonstrates that the expression level of CD28 and CD11b can discriminate among three subsets of circulating CD8⁺ T cells with different functional properties. The acquisition of CD11b molecules identifies CD28⁺ lymphocytes with effector cell features, which may form an intermediate phenotype in the process of CD8⁺ T cell differentiation. In this respect, it is remarkable that CD28⁺CD11b⁺ cells, unlike their CD28^- counterpart, retain the capacity to proliferate, thus enabling the population to expand greatly both in vivo and in vitro. A better understanding of the mechanisms that govern transition from early (proliferating) to mature (nonproliferating) effector cells will allow researchers to manipulate immunological memory for vaccination and adoptive immunotherapy purposes.

	Acknowledgments

	Footnotes

 
¹ This work was supported by the Istituto Superiore di Sanità, Progetto AIDS (Grant 40B.96).

² Address correspondence and reprint requests to Prof. Arnaldo Caruso, Istituto di Microbiologia, Università degli Studi di Brescia, Piazzale Spedali Civili 1, 25123 Brescia, Italy.

³ Abbreviations used in this paper: EBM, endothelial basal medium; MIP-1, macrophage inflammatory protein-1; HACEC, human adrenal gland capillary endothelial cell; MFI, mean fluorescence intensity.

Received for publication July 5, 2000. Accepted for publication October 23, 2000.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Labalette, M., F. Salez, F. R. Pruvot, C. Noel, J. P. Dessaint. 1994. CD8 lymphocytosis in primary cytomegalovirus (CMV) infection of allograft recipients: expansion of an uncommon CD8⁺ CD57^- subset and its progressive replacement by CD8⁺ CD57⁺ T cells. Clin. Exp. Immunol. 95:465.[Medline]
2. Fagnoni, F. F., R. Vescovini, M. Mazzola, G. Bologna, E. Nigro, G. Lavagetto, C. Franceschi, M. Passeri, P. Sansoni. 1996. Expansion of cytotoxic CD8⁺ CD28^- T cells in healthy ageing people, including centenarians. Immunology 88:501.[Medline]
3. Caruso, A., A. Cantalamessa, S. Licenziati, L. Peroni, E. Prati, F. Martinelli, A. D. Canaris, S. Folghera, R. Gorla, A. Balsari, et al 1994. Expression of CD28 on CD8⁺ and CD4⁺ lymphocytes during HIV infection. Scand. J. Immunol. 40:485.[Medline]
4. Brinckmann, J. E., J. H. Doblough, B. H. Heger, L. L. Haaheim, M. Sannes, T. Egeland. 1994. Expression of costimulatory molecule CD28 on T cells on human immunodeficiency virus type 1 infection: functional and clinic correlations. J. Infect. Dis. 169:730.[Medline]
5. Caruso, A., A. D. Canaris, S. Licenziati, A. Cantalamessa, S. Fiorentini, S. Ausenda, D. Ricotta, F. Dima, F. Malacarne, A. Balsari, et al 1999. Contribution of CD4⁺, CD8⁺CD28⁺, and CD8⁺CD28^- T cells to CD3⁺ lymphocytes homeostasis during the natural course of HIV-1 infection. J. Clin. Invest. 101:137.[Medline]
6. Azuma, M., J. H. Phillips, L. L. Lanier. 1993. CD28^- T lymphocytes. J. Immunol. 150:1147.[Abstract]
7. Wills, M. R., A. J. Carmichael, M. P. Weekes, K. Mynard, G. Okecha, R. Hicks, J. G. P. Sissons. 1999. Human virus-specific CD8⁺ CTL clones revert from CD45RO^high to CD45RA^high in vivo: CD45RA^highCD8⁺ T cells comprise both naïve and memory cells. J. Immunol. 162:7080.[Abstract/Free Full Text]
8. Fiorentino, S., M. Dalod, D. Olive, J. G. Guillet, E. Gomard. 1996. Predominant involvement of CD8⁺CD28^- lymphocytes in human immunodeficiency virus-specific cytotoxic activity. J. Virol. 70:2022.[Abstract]
9. Ho, H. N., L. E. Hultin, R. T. Mitsuyasu, J. L. Matud, M. A. Hausner, D. Bockstoce, C. C. Chou, S. O’Rourke, J. M. Taylor, and J. V. Giorgi. Circulating HIV-specific CD8⁺ cytotoxic T cells express CD38 and HLA-DR antigens. J. Immunol. 150:3070.
10. Weekes, M. P., A. J. Carmichael, M. R. Wills, K. Mynard, J. G. Sissons. 1999. Human CD28^-CD8⁺ T cells contain greatly expanded functional virus-specific memory CTL clones. J. Immunol. 162:7569.[Abstract/Free Full Text]
11. Borthwick, N. J., M. Bofill, W. H. Gombert, A. N. Akbar, E. Medina, K. Sagawa, M. C. Lipman, M. A. Johonson, J. Janossy. 1994. Lymphocyte activation in HIV infection. II. Functional defects of CD28^- T cells. AIDS. 8:431.[Medline]
12. Fiorentini, S., F. Malacarne, D. Ricotta, S. Licenziati, A. Armenta-Solis, S. Ausenda, M. A. De Francesco, E. Garrafa, A. Simonini, L. Imberti, et al 1999. Long term stimulation of CD8⁺CD28^- T cells induces stable disappearance of CD28 molecules: a possible mechanism accounting for the increased number of CD8⁺CD28^- T cells in HIV-1-infected patients. J. Leukocyte Biol. 65:641.[Abstract]
13. Labalette, M., E. Leteurtre, C. Thumerelle, C. Grutzmacher, B. Tourvieille, J. P. Dessaint. 1999. Peripheral human CD8⁺CD28⁺T lymphocytes give rise to CD28^-progeny, but IL-4 prevents loss of CD28 expression. Int. Immunol. 11:1327.[Abstract/Free Full Text]
14. Yamada, H., P. J. Martin, M. A. Bean, M. P. Braun, P. G. Beatty, K. Sadamoto, J. A. Hansen. 1985. Monoclonal antibody 9.3 and anti-CD11 antibodies define reciprocal subsets of lymphocytes. Eur. J. Immunol. 15:1164.[Medline]
15. Springer, T. A.. 1994. Traffic signals for lymphocyte recirculation and leukocyte emigration: the multistep paradigm. Cell. 76:301.[Medline]
16. Hamann, D., P. A. Baars, M. H. Rep, B. Hooibrink, S.R. Kerkhof-Garde, M. R. Klein, R. A. van Lier. 1997. Phenotypic and functional separation of memory and effector human CD8⁺ T cells. J. Exp. Med. 186:1407.[Abstract/Free Full Text]
17. Issekutz, T. B.. 1993. Dual inhibition of VLA-4 and LFA-1 maximally inhibits cutaneous delayed-type hypersensitivity-induced inflammation. Am. J. Pathol. 143:1286.[Abstract]
18. McFarland, H. I., S. R. Nahil, J. W. Maciaszec, R. M. Welsh. 1992. CD11b (Mac-1): a marker for CD8⁺ cytotoxic T cell activation and memory in virus infection. J. Immunol. 149:1326.[Abstract]
19. Caruso, A., S. Licenziati, A. D. Canaris, A. Cantalamessa, M. Corulli, B. Benzoni, L. Peroni, A. Balsari, A. Turano. 1996. Characterization of T cell subsets involved in the production of IFN- in asymptomatic HIV-infected patients. AIDS Res. Hum. Retroviruses 12:135.[Medline]
20. Nicholson, J. K., B. M. Jones, M. Hubbard. 1993. CD4 T-lymphocyte determinations on whole blood specimens using a single-tube three-color assay. Cytometry. 14:685.[Medline]
21. Jung, T., U. Schauer, C. Heusser, C. Neumann, and Rieger. 1993. Detection of intracellular cytokines by flow cytometry. J. Immunol. Methods 159:197.
22. Caruso, A., S. Licenziati, A. D. Canaris, A. Cantalamessa, S. Folghera, M. A. Lonati, G. DePanfilis, G. Garotta, A. Turano. 1995. CD4⁺ and CD8⁺ lymphocytes of AIDS patients synthesize increased amount of IFN-. J. Acquired Immune Defic. Syndr. 10:462.
23. Phillips, J. H., L. L. Lanier. 1986. Dissection of the lymphokine-activated killer phenomenon: relative contribution of peripheral blood natural killer cells and T lymphocytes to cytolysis. J. Exp. Med. 164:814.[Abstract/Free Full Text]
24. Alessandri, G., R. G. Chirivi, P. Castellani, G. Nicolo, R. Giavazzi, L. Zardi. 1998. Isolation and characterization of human tumor-derived capillary endothelial cells: role of oncofetal fibronectin. Lab. Invest. 78:127.[Medline]
25. Sallusto, F., D. Lenig, R. Forster, M. Lipp, A. Lanzavecchia. 1999. Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature 401:708.[Medline]
26. d’Angeac, A. D., S. Monier, D. Pilling, A. Travaglio-Encinoza, T. Reme, and M. Salmon. CD57^{+ T} lymphocytes are derived from CD57^- precursors by differentiation occurring in late immune responses. Eur. J. Immunol. 24:1503.
27. Salmon, M., G. D. Kitas, P. A. Bacon. 1989. Production of lymphokine mRNA by CD45R⁺ and CD45R^- helper T cells from human peripheral blood and by human CD4⁺ T cell clones. J. Immunol. 143:907.[Abstract]
28. Lalvani, A., R. Brookes, S. Hambleton, W. J. Britton, A. V. Hill, A. J. McMichael. 1997. Rapid effector function in CD8⁺ memory T cells. J. Exp. Med. 186:859.[Abstract/Free Full Text]
29. Smyth, M. J., J. A. Trapani. 1995. Granzymes: exogenous proteinases that induce target cell apoptosis. Immunol. Today 16:202.[Medline]
30. Liu, C. C., C. M. Walsh, J. D. Young. 1995. Perforin: structure and function. Immunol. Today 16:194.[Medline]
31. Butcher, E. C., L. J. Picker. 1996. Lymphocyte homing and homeostasis. Science 272:60.[Abstract]
32. Mackay, C. R., W. L. Marston, L. Dudler. 1990. Naive and memory T cells show distinct pathways of lymphocyte recirculation. J. Exp. Med. 171:801.[Abstract/Free Full Text]
33. Baggiolini, M.. 1998. Chemokines and leukocyte traffic. Nature 392:565.[Medline]
34. Kishimoto, T. K., K. O’Connor, A. Lee, T. M. Roberts, T. A. Springer. 1987. Cloning of the subunit of the leukocyte adhesion proteins: homology to an extracellular matrix receptor defines a novel supergene family. Cell 48:681.[Medline]
35. Tedla, N., H. W. Wang, H. P. McNeil, N. Di Girolamo, T. Hampartzoumian, D. Wakefield, A. Lloyd. 1998. Regulation of T lymphocyte trafficking into lymph nodes during an immune response by the chemokines macrophage inflammatory protein (MIP)-1 and MIP-1. J. Immunol. 161:5663.[Abstract/Free Full Text]
36. Kagi, D., B. Ledermann, K. Burki, R. M. Zinkernagel, H. Hengartner. 1996. Molecular mechanisms of lymphocyte-mediated cytotoxicity and their role in immunological protection and pathogenesis in vivo. Annu. Rev. Immunol. 14:207.[Medline]
37. Young, L. H., L. S. Klavinskis, M. B. Oldstone, J. D. Young. 1989. In vivo expression of perforin by CD8⁺ lymphocytes during an acute viral infection. J. Exp. Med. 169:2159.[Abstract/Free Full Text]
38. Joly, P., J. M. Guillon, C. Mayaud, F. Plata, I. Theodorou, M. Denis, P. Debre, B. Autran. 1989. Cell-mediated suppression of HIV-specific cytotoxic T lymphocytes. J. Immunol. 143:2193.[Abstract]
39. Mollet, L., B. Sadat-Sowti, J. Duntze, V. Leblond, F. Bergeron, V. Calvez, C. Katlama, P. Debre, B. Autran. 1998. CD8^hi+CD57⁺ T lymphocytes are enriched in antigen-specific T cells capable of down-modulating cytotoxic activity. Int. Immunol. 10:311.[Abstract/Free Full Text]
40. Callan, M. F., L. Tan, N. Annels, G. S. Ogg, J. D. Wilson, C. A. O’Callaghan, N. Steven, A. J. McMichael, A. B. Rickinson. 1998. Direct visualization of antigen-specific CD8⁺ T cells during the primary immune response to Epstein-Barr virus in vivo. J. Exp. Med. 187:1395.[Abstract/Free Full Text]
41. Hamann, D., M. T. Roos, R.A. van Lier. 1999. Faces and phases of human CD8 T-cell development. Immunol. Today 20:177.[Medline]
42. Butcher, E. C.. 1990. Cellular and molecular mechanisms that direct leukocyte traffic. Am. J. Pathol. 136:3.[Medline]
43. Hoshino, T., A. Yamada, J. Honda, Y. Imai, M. Nakao, M. Inoue, K. Sagawa, M. M. Yokohama, K. Oizumi, K. Itoh. 1993. Tissue-specific distribution and age-dependent increase of human CD11b⁺ T cells. J. Immunol. 151:2237.[Abstract]

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