ICAM-1/LFA-1 interactions are known to enhance T cell/APC interactions and to promote T cell activation and cytokine secretion. We have analyzed the consequences of ICAM-1-mediated signaling on the generation of memory T cell subsets. We report that lack of ICAM-1 on APCs, but not on T cells, leads to poor T cell activation and proliferation in vitro and in vivo, and that the defect can be compensated by Ag dose, exogenous IL-2, additional costimulation, and by increasing responder T cell density on APCs. ICAM-1-null mice do not respond to immunization with OVA peptide, but immunization with OVA or with Salmonella typhimurium leads to good T cell proliferation 7–10 days later, and clearance of a challenge infection is equivalent to that of wild-type mice. However, when followed over time, recall proliferation and antibacterial immunity decay rapidly in ICAM-1-null mice, while recall cytokine responses are unaffected. The decline in immunity is not related to poor survival of T cells activated on ICAM-1-null APCs, or to poor generation of effectors in ICAM-1-null mice. Phenotypic analysis of T cells stimulated on ICAM-1-null APCs reveals preferential generation of CD44highCD62Llow effector memory cells (TEM) over CD44highCD62Lhigh central memory cells (TCM). Further, while the proportion of naive:memory T cells is similar in unmanipulated wild-type and ICAM-1-null mice, there is an accumulation of TEM cells, and a high TEM: TCM ratio in aging ICAM-1-null mice. Together, the data indicate that signaling through LFA-1 during T cell activation may be involved in commitment to a proliferation-competent memory pool.

T cell responses to immunization or infection are characterized by clonal expansion of relatively small numbers of Ag-specific T cells that are present in peripheral lymphoid organs, and their differentiation into effector cells that secrete inflammatory cytokines and other molecules involved in pathogen clearance. A large fraction of T cells responding to primary Ag encounter dies, but some cells survive to form a relatively long-lived memory T cell pool that can respond to a secondary challenge by proliferation and/or cytokine secretion (1, 2, 3, 4, 5, 6, 7, 8, 9). Two types of memory T cells, central memory (TCM)4 and effector memory (TEM), can be identified in vivo based on their differential proliferation capabilities, biochemical properties, recirculation patterns, and persistence, and it is the proliferation-competent TCM cells that may be responsible for long-lived protection against pathogen challenge (10, 11, 12, 13, 14, 15, 16, 17, 18).

The survival of activated T cells and their differentiation into memory cells is related to the magnitude of the primary response and is influenced by the availability of IL-2 and costimulation as well as on the intensity and duration of Ag presentation during priming (19, 20, 21, 22, 23, 24, 25, 26, 27). Previous in vitro studies have shown that ICAM-1/LFA-1 interaction may influence early events in the priming of naive T cells by facilitating T cell/APC conjugate formation and maturation of the immunological synapse, by inducing actin remodeling that contributes to T cell adhesion and movement, and by enhancing T cell activation and proliferation (28, 29, 30, 31, 32, 33, 34, 35, 36). Signaling through LFA-1 on T cells has also been shown to promote apoptosis (37), suggesting that LFA-1/ICAM-1 interactions may be involved in the contraction phase of immune responses.

Using mice that are deficient in ICAM-1 expression, we have attempted to relate these findings to the maturation of T cell responses in vivo. We varied the strength of T cell priming in vivo by using peptide Ags, protein Ags or live bacteria for immunization, and by varying ligand density and responder cell densities in vitro. We looked for T cell-specific and APC-specific effects of ICAM-1 deficiency by using assays in which either T cells or APCs were ICAM-1-null. We followed effector- and memory- T cell responses over time using readouts for T cell proliferation, cytokine secretion and the ability of immunized mice to clear a challenge infection. We also looked at the composition of the primed T cell pool by estimating ratios of TCM and TEM cells. We report that signals provided to T cells by ICAM-1 on APCs promote their differentiation into long-lived, proliferation-competent, TCM cells.

All mouse strains were from The Jackson Laboratory, and were bred and maintained in the Small Animal Facility of the National Institute of Immunology. Mice used for experiments were 8–12 wk old unless specified. Approval from the Institutional Animal Ethics Committee was obtained for all experimental procedures involving animals. Mice were immunized s.c. in the hind foot pad with 10 μg of OVA (Sigma-Aldrich), or 10 μg of OVA peptide (OVA 323–339; Peptron) in CFA (Difco Laboratories). To assess antibacterial immunity, mice were immunized i.p. with 103 CFU of Salmonella typhimurium (38) and bacteria were cleared from internal organs 24 h later by i.p. treatment with ciprofloxacin (39).

S. typhimurium 754 (Stm) is routinely maintained in the laboratory. Bacterial stocks were stored in glycerol broth at −70°C and a fresh aliquot was plated out on Salmonella-Shigella agar (Difco Laboratories) for all experiments. For preparation of bacterial sonicate, an overnight culture of bacteria in Luria Bertani broth (Difco Laboratories) was spun down, washed in PBS, and inactivated by heating the cell suspension in a boiling water bath for 45 min. The suspension was sonicated for 15 min in PBS containing 10 mM PMSF (Sigma-Aldrich) as a protease inhibitor, spun at high speed to reduce LPS, and the supernatant was filtered and used as soluble Ag (39). For bacterial challenge experiments, mice were challenged i.p. with 103 CFU of Stm and 6–8 days later their spleens were harvested and appropriate dilutions of the lysate were plated out on Salmonella-Shigella agar. The number of bacteria was enumerated as CFU/spleen. The limit of detection was 50 CFU/spleen.

Splenocytes from normal mice, or draining lymph node (LN) cells from immunized mice, were cultured at various densities (5 × 105/well to 105/well as indicated) in 96-well flat-bottom plates (Falcon) with titrating doses of anti-CD3 or specific Ag in Click’s medium (Irvine Scientific) supplemented with 10% FBS, 2 mM l-glutamine, 0.1 mM 2-ME, and antibiotics (Invitrogen Life Technologies). Where indicated, 5 μg/ml azide-free anti-CD28 (clone 37.51; BD Biosciences) and 10 μg/ml anti-CD4 (purified GK1.5) or anti-CD8 (purified 53.6.72) was added. Plates were pulsed with 0.5 μCi of [3H]thymidine (NEN) 48–72 h after initiation of culture and harvested 12–16 h later onto glass-fiber filters for scintillation spectroscopy (Betaplate; Wallac).

For enumerating cell divisions, splenocytes from wild-type (WT) or ICAM-1-knockout (KO) mice were labeled with 5–10 μM CFSE (Molecular Probes) before stimulation with anti-CD3, and CFSE dilution on gated CD4 and CD8 cells was estimated by flow cytometry.

LDA for determining clonal frequency of Ag-specific T cells was done essentially as described (40). Briefly, lymphocytes from the draining LNs were titrated from 1.6 × 105/well to 2.5 × 103/well in medium supplemented further with sodium pyruvate and nonessential amino acids (Invitrogen Life Technologies) in the presence or absence of Ag (300 μg/ml OVA or 1 μg/ml OVA peptide 323–339), 105 thymocytes, and 105 irradiated splenocytes (1000 rad) from unimmunized mice (as fillers and APCs, respectively) and the T cell response was read out as [3H]thymidine incorporation 84 h later. For each cell input, 48 wells were plated with Ag and 12 wells without Ag. Wells showing a response at least 3-fold above those of the no-Ag controls for each cell input were scored as positive. The clonal frequency was calculated as the input at which 37% of the wells were negative for proliferation.

For estimating the ability of WT and KO T cells to proliferate in response to allogeneic APCs in vivo, CFSE-labeled splenocytes from WT or KO (H-2b) mice were transferred i.v. into irradiated (600 rad) DBA/2 (H-2d) mice and 72 h later, recipient spleens were harvested, cells stained for Thy1, propidium iodide (PI), and H-2Kd, and proliferation of donor T cells estimated as CFSE dilution of Thy1-positive, H-2d-negative, PI-negative cells (41). Homeostatic proliferation, that may contribute to CFSE dilution of donor cells in irradiated recipients, was assessed by transferring CFSE-labeled WT or KO cells (Thy1.2, H-2b) into irradiated C57BL/6 congenics (Thy1.1, H-2b) and gating on the live Thy1.2+ donor cells. To similarly estimate the ability of WT and KO APCs to induce an allogeneic response in vivo, CFSE-labeled BALB/c splenocytes (H-2d) were transferred into irradiated WT or KO mice (H-2b) and proliferation of donor T cells estimated on Thy1-positive, H-2b-negative, PI-negative cells.

The following reagents were used for various labeling reactions: fluorescein/PE/CyChrome/biotin anti-CD4, CD8, fluorescein/PE/biotin anti-CD90.1, CD90.2, PE/CyChrome anti-CD44, CyChrome streptavidin, PE-Texas Red streptavidin, fluorescein/PE anti-CD2, CD11a, CD25, CD27, CD54, CD69, CD62L, CD122, CD127, CD134, Ly6C, IFN-γ, TCRβ, TCR-Vα2, anti-H-2Kb, anti-H-2Kd (BD Biosciences), PE-streptavidin, fluorescein-streptavidin (Jackson ImmunoResearch Laboratories), PE-Texas Red anti-CD4, CD8 (Caltag Laboratories), and propidium iodide (PI; Sigma-Aldrich).

Cells were incubated with appropriate staining reagents in buffer containing 0.1% sodium azide (Sigma-Aldrich) and 1% FBS for 45 min on ice. Samples were run on an Elite ESP (Beckman Coulter) or a BD LSR (BD Biosciences) flow cytometer and data were analyzed with FlowJo software (Treestar). For visualizing intracellular cytokine levels, splenic or LN cells were stimulated with appropriate Ag, as indicated, for 12 h, stained for surface markers, fixed, and permeabilized with Cytofix/Cytoperm (BD Biosciences) and stained for intracellular IFN-γ. Activation of P14 transgenic cells was estimated by looking at surface TCR levels and CD69 on gated CD8+Vα2+ cells, and differentiation into memory subsets estimated by looking at CD44 and CD62L expression on gated cells. Data were acquired on log scales, except for TCR down-modulation experiments where a linear scale was used for TCR intensity, and the data were plotted as percent decrease of modal intensity over time.

Trophic signal withdrawal death (TSWD) and activation-induced cell death (AICD) were done as described earlier (42). Briefly, splenocytes were stimulated with 160 ng/ml azide-free anti-CD3 (Cedarlane Laboratories) alone for 48 h, supplemented with 5 U/ml IL-2 (Boehringer Mannheim) and cultured for a further 48 h. Viable cells were harvested at 96 h and cultured further in the presence or absence of 5 U/ml IL-2 to induce TSWD, or were cultured in IL-2 with or without plate-coated anti-CD3 to induce AICD. Cells were stained at various intervals from 0 to 48 h with appropriately labeled anti-CD4/CD8, and Hoechst 33342 (5 μg/ml; Molecular Probes) was added just before visualization of nuclear morphology by fluorescent microscopy (TE2000-U; Nikon Eclipse). Between 150–200 cells were counted and results are expressed as the mean number of dying cells.

Activated WT and KO T cells were transferred i.v. into Thy1.1 congenic mice and 18 h later, the efficiency of transfer monitored by estimating the proportion of donor Thy1.2 cells in the PBMC of all recipients. Groups of mice showing equivalent transfer were then euthanized over time, and the survival of donor T cells in spleen and LNs was estimated as a proportion of that seen in PBMC at 18 h.

Bone marrow-derived dendritic cells (BMDCs) were grown from the bone marrow of C57BL/6 or ICAM-1-null mice in medium containing 20 ng/ml GM-CSF (PeproTech) and were activated by addition of 10 ng/ml recombinant TNF-α (PeproTech) 24 h before use (43). Peritoneal exudate cells (PECs) were induced by i.p. injection of thioglycolate broth (Himedia). CD8 cells were purified from total lymphocyte populations by treatment of the cell suspension with biotinylated anti-CD8, followed by Streptavidin magnetic beads (Miltenyi Biotec), and separation on MACS columns (Miltenyi Biotec) according to the manufacturer’s instructions. Purified cells were routinely ≥95% pure by flow cytometric analysis.

In initial experiments, splenocytes from WT and ICAM-1-null mice were stimulated with titrating doses of anti-CD3 in vitro, and the modulation of various activation markers on CD4 and CD8 cells was analyzed 24 or 48 h later. T cells in the WT cultures, but not in the KO cultures, showed significant up-regulation of CD25 at 24 h (Fig. 1, A–D). By 48 h, some up-regulation of CD25 was seen in KO cultures, but only at the higher dose of anti-CD3 used (data not shown). Up-regulation of CD69 expression at 24 h was also compromised in KO cultures (Fig. 1, E–H). Although practically all WT T cells showed enhanced expression of CD69, a relatively large proportion of KO T cells failed to up-regulate CD69, and the pattern was similar at 48 h (data not shown). At 48 h, markers such as CD27, CD44, and CD134 that are associated with secondary T cells (14, 15, 44, 45) were poorly up-regulated on KO T cells (Fig. 2). Modulation of the cytokine receptors CD122 and CD127 that are associated with cell cycling (46, 47, 48) was also different on WT and KO T cells, with KO cells showing poor up-regulation of CD122 and poor down-regulation of CD127 (Fig. 2). Similar differences in activation marker modulation between WT and KO cells were seen with 10 ng/ml anti-CD3 (data not shown).

FIGURE 1.

Poor up-regulation of CD25 and CD69 on KO T cells following polyclonal stimulation. WT (dotted lines) or KO (solid lines) splenocytes were stimulated with 100 or 30 ng/ml anti-CD3 (indicated on right margin) and CD25 (A–D) and CD69 (E–H) expression was assessed on gated CD4 and CD8 cells 24 h later. Staining profiles of unstimulated cells are indicated by shaded histograms. Data are representative of six independent experiments

FIGURE 1.

Poor up-regulation of CD25 and CD69 on KO T cells following polyclonal stimulation. WT (dotted lines) or KO (solid lines) splenocytes were stimulated with 100 or 30 ng/ml anti-CD3 (indicated on right margin) and CD25 (A–D) and CD69 (E–H) expression was assessed on gated CD4 and CD8 cells 24 h later. Staining profiles of unstimulated cells are indicated by shaded histograms. Data are representative of six independent experiments

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FIGURE 2.

Poor modulation of other activation markers on KO T cells following polyclonal stimulation. WT (dotted lines) or KO (solid lines) splenocytes were stimulated with 30 ng/ml anti-CD3 and expression of CD27, CD44, CD122, CD127, and CD134 (indicated on right margin) assessed on gated CD4 (A–E) and CD8 (F–J) cells 48 h later. Staining profiles of unstimulated cells are indicated by shaded histograms. Data are representative of three independent experiments.

FIGURE 2.

Poor modulation of other activation markers on KO T cells following polyclonal stimulation. WT (dotted lines) or KO (solid lines) splenocytes were stimulated with 30 ng/ml anti-CD3 and expression of CD27, CD44, CD122, CD127, and CD134 (indicated on right margin) assessed on gated CD4 (A–E) and CD8 (F–J) cells 48 h later. Staining profiles of unstimulated cells are indicated by shaded histograms. Data are representative of three independent experiments.

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To assess the effect of poor activation of ICAM-1-null T cells on their subsequent proliferation, splenocytes from WT and KO mice were labeled with CFSE, stimulated with titrating doses of anti-CD3, and CFSE dilution was estimated on gated CD4 and CD8 72 h later. In keeping with earlier findings (49), CD8 cells proliferated more than CD4 cells and both WT and KO CD8 cells showed substantial CFSE dilution when stimulated with 100 ng/ml anti-CD3 (Fig. 3,E). However, the proliferation of KO cells decreased substantially at the lower dose of 30 ng/ml (Fig. 3,F) and the response dropped to near background levels at 10 ng/ml (data not shown). KO CD4 cells also showed relatively poor proliferation as compared with WT cells at 72 h (Fig. 3, A, B, and G) and the differences were more pronounced at 96 h (Fig. 3, C and D).

FIGURE 3.

Poor proliferation of KO T cells following polyclonal stimulation. CFSE-labeled splenocytes from WT or KO mice were stimulated with 100 or 30 ng/ml anti-CD3 (as indicated) and CFSE dilution on gated CD4 and CD8 cells was assessed. A and B, Proliferation of CD4 cells at 72 h. C and D, Proliferation of CD4 cells at 96 h. E and F, Proliferation of CD8 cells at 72 h. Data are representative of five independent experiments. G, Division profiles of CD4 cells at 72 h, calculated from the five experiments.

FIGURE 3.

Poor proliferation of KO T cells following polyclonal stimulation. CFSE-labeled splenocytes from WT or KO mice were stimulated with 100 or 30 ng/ml anti-CD3 (as indicated) and CFSE dilution on gated CD4 and CD8 cells was assessed. A and B, Proliferation of CD4 cells at 72 h. C and D, Proliferation of CD4 cells at 96 h. E and F, Proliferation of CD8 cells at 72 h. Data are representative of five independent experiments. G, Division profiles of CD4 cells at 72 h, calculated from the five experiments.

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In parallel experiments, proliferation was also assessed by thymidine incorporation in cultures containing titrating numbers of splenocytes. As seen in Fig. 4,A, WT and KO T cells show equivalent proliferation in high cell density cultures (containing 3 × 105 cells/well), but the proliferation of KO T cells drops precipitously at lower plating densities. The proliferation defect in the low cell density cultures can be rescued partially by addition of nonmitogenic amounts of exogenous IL-2 (Fig. 4,B) and completely by addition of stimulatory anti-CD28 (Fig. 4,C). Together with the data in Figs. 1 and 2, these data indicate that T cell activation in the ICAM-1-null cultures may be of insufficient duration or strength to initiate and support cell division, especially under conditions where costimulation or ligand density are limiting, and when responder T cell frequencies are low.

FIGURE 4.

Partial reversal of the proliferation defect of KO cells by increasing the density of responding cells (A), addition of exogenous IL-2 (B), and increasing costimulation (C). A, WT and KO splenocytes were plated at various densities (105/well to 3 × 105/well) and proliferation was assessed by addition of [3H]thymidine 48 h later. B and C, WT and KO splenocytes were plated at 105/well in the presence or absence of titrating doses of IL-2 (B) or 5 μg/ml anti-CD28 (C), and proliferation was assessed 48 h later. Data are representative of three independent experiments.

FIGURE 4.

Partial reversal of the proliferation defect of KO cells by increasing the density of responding cells (A), addition of exogenous IL-2 (B), and increasing costimulation (C). A, WT and KO splenocytes were plated at various densities (105/well to 3 × 105/well) and proliferation was assessed by addition of [3H]thymidine 48 h later. B and C, WT and KO splenocytes were plated at 105/well in the presence or absence of titrating doses of IL-2 (B) or 5 μg/ml anti-CD28 (C), and proliferation was assessed 48 h later. Data are representative of three independent experiments.

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Because ICAM-1 is expressed on both T cells and APCs, the poor activation of T cells in KO splenocyte cultures may be related to signaling through ICAM-1 or LFA-1 on the T cells. Three different experimental approaches were used to differentiate between the two possibilities. First, we looked at very early T cell activation events by following TCR down-modulation and CD69 up-regulation on transgenic T cells responding to peptide presented by WT or ICAM-1-null APCs. PECs from WT or KO mice were pulsed with titrating amounts of peptide KAVYNFATM (Peptron) of lymphocytic choriomeningitis virus (LCMV) and used as APCs to stimulate purified CD8 cells from P14 mice, which express a transgenic TCR specific for the peptide on H-2Db (50). TCR down-modulation and CD69 up-regulation were then assessed on gated transgenic cells at various times. As seen in Fig. 5, both events occur rapidly when the transgenic T cells are stimulated in the presence of WT APCs. However, no activation by either readout is seen when KO APCs pulsed with low doses of peptide are used. Thus, ICAM-1 on APCs is necessary for inducing very early activation events in T cells stimulated at low ligand densities.

FIGURE 5.

PECs from KO mice support poor activation of purified CD8+ cells from P14 mice. A, TCR down-modulation on transgenic cells assessed over time after addition of 20 ng/ml (circles), 200 ng/ml (triangles), or 2 μg/ml (squares) LCMV peptide. B–E, Up-regulation of CD69 1 h (solid lines), 2 h (dotted lines), and 3 h (thick lines) after addition of titrating doses of peptide (indicated on right margin). Staining profiles of unstimulated cells are indicated by the shaded histograms. Data are representative of four independent experiments.

FIGURE 5.

PECs from KO mice support poor activation of purified CD8+ cells from P14 mice. A, TCR down-modulation on transgenic cells assessed over time after addition of 20 ng/ml (circles), 200 ng/ml (triangles), or 2 μg/ml (squares) LCMV peptide. B–E, Up-regulation of CD69 1 h (solid lines), 2 h (dotted lines), and 3 h (thick lines) after addition of titrating doses of peptide (indicated on right margin). Staining profiles of unstimulated cells are indicated by the shaded histograms. Data are representative of four independent experiments.

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Next, we assessed T cell proliferation in vivo under conditions where either the T cell or the APC was ICAM-1-deficient, using the established method of transferring CFSE-labeled T cells into sublethally irradiated allogeneic hosts, and assessing proliferation by CFSE dilution on gated T cells 60–72 h later (41). To assess the role of ICAM-1 deficiency in T cells, WT and KO splenocytes (H-2b) were transferred into irradiated, ICAM-1-sufficient DBA/2 (H-2d) mice. To assess the role of ICAM-1 deficiency in APCs, CFSE-labeled splenocytes from ICAM-1-sufficient BALB/c mice (H-2d) were transferred into irradiated WT or KO (H-2b) hosts. As seen in Fig. 6,A, APCs in ICAM-1-sufficient mice support equivalent allogeneic proliferation of WT and ICAM-1-null T cells in vivo. In contrast, WT T cells show relatively poor proliferation in ICAM-1-deficient allogeneic hosts (Fig. 6,B). A similar trend was seen at 60 h (data not shown). These results indicate that signaling events associated with ICAM-1 on APCs are required to sustain vigorous T cell proliferation in vivo. To control for homeostatic proliferation of donor T cells in irradiated hosts, CFSE-labeled cells were transferred into irradiated syngeneic hosts, as detailed in Materials and Methods, and no significant proliferation was observed (Fig. 6).

FIGURE 6.

KO APCs support poor allogeneic proliferation of WT T cells in vivo. A, CFSE dilution profile of gated CD4 cells 72 h after transfer of WT (dotted line) or KO (solid line) splenocytes into irradiated DBA/2 mice. B, CFSE dilution profile of gated CD4 cells 72 h after transfer of BALB/c splenocytes into irradiated WT (dotted line) or KO mice (solid line). Homeostatic proliferation of CD4 cells in irradiated syngeneic hosts is shown as shaded histograms. Data are representative of three mice per group and of three independent experiments.

FIGURE 6.

KO APCs support poor allogeneic proliferation of WT T cells in vivo. A, CFSE dilution profile of gated CD4 cells 72 h after transfer of WT (dotted line) or KO (solid line) splenocytes into irradiated DBA/2 mice. B, CFSE dilution profile of gated CD4 cells 72 h after transfer of BALB/c splenocytes into irradiated WT (dotted line) or KO mice (solid line). Homeostatic proliferation of CD4 cells in irradiated syngeneic hosts is shown as shaded histograms. Data are representative of three mice per group and of three independent experiments.

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To confirm that ICAM-1 deficiency on APCs, and not on T cells, was responsible for the poor T cell responses of KO splenocytes in vitro, we stimulated cultures containing equal numbers of splenocytes from KO (Thy1.2) and WT (Thy1.1) mice with titrating amounts of anti-CD3 and assessed T cell activation and proliferation on gated Thy1.1 and Thy1.2 cells in the mixed culture. The responses were then compared with those seen when WT and KO cells were stimulated alone, and the results are shown in Fig. 7. WT cells showed good up-regulation of CD25, CD69, and CD44 when stimulated alone, and the pattern was largely unaffected in the presence of KO cells from Thy1.2 mice (Fig. 7, A, E, I and C, G, K). In contrast, while KO T cells showed poor up-regulation of all markers when stimulated alone (as seen earlier in Fig. 1–2), their activation was significantly enhanced in the mixed cultures (Fig. 7, B, F, J and D, H, L). Similar results were seen when CD25 and CD69 expression was assessed at 48 h (data not shown).

FIGURE 7.

KO T cells respond well when activated in the presence of APCs from WT mice. A–L, Up-regulation of activation markers on T cells following stimulation of WT or KO splenocytes with 30 ng/ml anti-CD3 in independent cultures (dotted lines) or in mixed cultures (solid lines) containing equal proportions of WT (Thy1.1) and KO (Thy1.2) cells. CD25 (A–D) and CD69 expression at 24 h (E–H) and CD44 (I–L) expression at 48 h on gated CD4 and CD8 are shown. For the mixed cultures, CD4 and CD8 gates were applied to Thy1.1 or Thy1.2 gates for WT and KO T cells, respectively. M–T, CFSE dilution on CD4 and CD8 cells, gated as above, 72 h after stimulation of WT or KO splenocytes with 100 or 30 ng/ml anti-CD3, as indicated, in independent cultures (dotted lines) or in mixed cultures (solid lines). Staining profiles of unstimulated cells are shown as shaded histograms. Data are representative of three independent experiments.

FIGURE 7.

KO T cells respond well when activated in the presence of APCs from WT mice. A–L, Up-regulation of activation markers on T cells following stimulation of WT or KO splenocytes with 30 ng/ml anti-CD3 in independent cultures (dotted lines) or in mixed cultures (solid lines) containing equal proportions of WT (Thy1.1) and KO (Thy1.2) cells. CD25 (A–D) and CD69 expression at 24 h (E–H) and CD44 (I–L) expression at 48 h on gated CD4 and CD8 are shown. For the mixed cultures, CD4 and CD8 gates were applied to Thy1.1 or Thy1.2 gates for WT and KO T cells, respectively. M–T, CFSE dilution on CD4 and CD8 cells, gated as above, 72 h after stimulation of WT or KO splenocytes with 100 or 30 ng/ml anti-CD3, as indicated, in independent cultures (dotted lines) or in mixed cultures (solid lines). Staining profiles of unstimulated cells are shown as shaded histograms. Data are representative of three independent experiments.

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CFSE dilution experiments set up in parallel showed that WT cells proliferated well when stimulated alone or in mixed culture (Fig. 7, M, Q, O, and S). In contrast, while KO T cells proliferated very poorly when stimulated alone, they showed excellent CFSE dilution when stimulated in mixed cultures containing WT APCs (Fig. 7, N, R, P, and T). The results were especially striking at the lower dose of anti-CD3, where the response of CD8 cells from WT and KO mice were indistinguishable in the mixed cultures. Together with the data in Fig. 4, these results indicate that signaling through LFA-1 on T cells may be required to augment relatively weak TCR signals that would otherwise be nonstimulatory.

Because the generation of a memory T cell pool in mice depends on the clonal expansion of relatively rare Ag-specific precursors, the inability of ICAM-1-null APCs to support extensive T cell proliferation raised the possibility that ICAM-1-null mice may show compromised T cell responses to immunization. More intriguingly, our data show that increasing plating densities or enhancing costimulation in vitro can compensate for the APC defect, raising the further possibility that the number of T cells responding to a given Ag may determine the vigor of T cell responses in ICAM-1-null mice. We tested these possibilities by tracking T cell recall responses in mice immunized with OVA or with an MHC-II-restricted OVA peptide to prime a relatively large, or a relatively small, number of T cells, respectively. The results are shown in Fig. 8.

FIGURE 8.

Poor generation of proliferation-competent T cells in KO mice following immunization. A–F, Recall proliferation responses of T cells from the draining LN of WT (•) and KO (○) mice to titrating doses of Ag (A–C) and estimation of secondary T cell frequency by LDA (D–F) 7 or 14 days (indicated on the right margin) after immunization with OVA (A and B, D and E) or OVA peptide (C and F). G and H, Intracellular IFN-γ on gated CD4 cells from LN of control mice or from WT/KO mice immunized 7 days earlier with OVA (G) or OVA peptide (H). Cells were stimulated for 16 h with Ag. I, CD8 cells do not contribute to OVA recall. Addition of anti-CD4, but not anti-CD8, inhibits recall proliferation, and CD8 cells show no IFN-γ. Data are representative of four independent experiments (A–C, G, and H), of three independent experiments (D–H), and of two independent experiments (I).

FIGURE 8.

Poor generation of proliferation-competent T cells in KO mice following immunization. A–F, Recall proliferation responses of T cells from the draining LN of WT (•) and KO (○) mice to titrating doses of Ag (A–C) and estimation of secondary T cell frequency by LDA (D–F) 7 or 14 days (indicated on the right margin) after immunization with OVA (A and B, D and E) or OVA peptide (C and F). G and H, Intracellular IFN-γ on gated CD4 cells from LN of control mice or from WT/KO mice immunized 7 days earlier with OVA (G) or OVA peptide (H). Cells were stimulated for 16 h with Ag. I, CD8 cells do not contribute to OVA recall. Addition of anti-CD4, but not anti-CD8, inhibits recall proliferation, and CD8 cells show no IFN-γ. Data are representative of four independent experiments (A–C, G, and H), of three independent experiments (D–H), and of two independent experiments (I).

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One week after immunization with OVA, WT and KO mice showed equivalent T cell proliferation to recall Ag in vitro (Fig. 8,A). However, the response decayed rapidly in KO mice and was barely detectable 2 wk after immunization (Fig. 8,B). WT mice responded even when tested 6 wk after immunization (data not shown). The decline in T cell responses was also reflected in the frequency of secondary T cells, determined by LDA (Fig. 8, D and E). On day 7, a 2-fold difference in frequency, at most, was observed (1/4 × 104 for WT cells and 1/8 × 104 for KO cells, not significantly different) but by day 14, while the frequency of responding WT cells had decreased somewhat (to 1/105) the response of KO cells was too poor to enable calculation of responder frequency. Effector generation, estimated as the proportion of cells containing intracellular IFN-γ following a 16 h stimulation with OVA, was similar in WT and KO mice (Fig. 8,G) and the response declined equivalently in both groups by day 14 (data not shown). Thus, the higher proliferation of cells from WT mice at later time points is not related to an enhanced generation of effector cells which differentiate into proliferation-competent secondary T cells. We have shown previously that recall proliferation and cytokine secretion following immunization of mice with OVA in CFA are mediated by CD4 cells (51) and the primacy of CD4 cells in the recall response was further confirmed here (Fig. 8 I). Addition of anti-CD4, but not anti-CD8 to the recall culture abrogated T cell proliferation, and no IFN-γ was detectable in gated CD8 cells.

Significantly, KO mice did not mount a detectable T cell response when immunized with OVA peptide. No recall T cell proliferation was detectable even 1 wk after immunization, and the frequency of responding T cells was too low to estimate by LDA. Cells from WT mice, in contrast, showed good proliferation to titrating doses of peptide in vitro, and the frequency of responding cells was also high (Fig. 8, C and F). Effector generation in WT mice, estimated by intracellular IFN-γ staining, was similar to that seen after OVA immunization, and was lower in KO mice, in keeping with poor overall priming (Fig. 8 H).

Because early T cell recall responses following OVA immunization are indistinguishable in WT and ICAM-1-null mice, one possible reason for the rapid decay of recall responses in the KO mice is poor survival of T cells primed on ICAM-1-null APCs. To examine this, we stimulated WT and KO splenocytes with anti-CD3 and tested their survival in three independent assays (Fig. 9). Activated T cells were washed and restimulated with anti-CD3 to assess AICD, cultured in the absence of IL-2 to assess TSWD, and transferred into unmanipulated Thy1-congenic mice to assess their survival in vivo in the absence of further stimulation. As seen in Fig. 9, A and B, T cells from KO mice show greater resistance to AICD and TSWD, and their survival is equivalent to that of activated WT cells in vivo (Fig. 9, C and D). Our results are in keeping with earlier observations suggesting that signaling through LFA-1 enhances T cell apoptosis (37) and they indicate that differential survival of activated T cells cannot account for the differential decay of recall responses in WT and KO mice.

FIGURE 9.

Equivalent survival of activated WT and KO T cells in vitro and in vivo. A and B, AICD (A) and TSWD (B) of WT (▪) and KO (▨) T cells following stimulation with anti-CD3 in vitro. Data are representative of five independent experiments. C and D, Survival of activated WT (filled symbols) and KO (open symbols) T cells following transfer into Thy1-congenic mice. Data are plotted as percentage of donor CD4 and CD8 cells recovered from the spleen (circles) and peripheral LN (triangles) of recipient mice over time, normalized to the proportions detected in PBMC of individual recipients 18 h after cell transfer. Data are representative of three mice per group and of two independent experiments.

FIGURE 9.

Equivalent survival of activated WT and KO T cells in vitro and in vivo. A and B, AICD (A) and TSWD (B) of WT (▪) and KO (▨) T cells following stimulation with anti-CD3 in vitro. Data are representative of five independent experiments. C and D, Survival of activated WT (filled symbols) and KO (open symbols) T cells following transfer into Thy1-congenic mice. Data are plotted as percentage of donor CD4 and CD8 cells recovered from the spleen (circles) and peripheral LN (triangles) of recipient mice over time, normalized to the proportions detected in PBMC of individual recipients 18 h after cell transfer. Data are representative of three mice per group and of two independent experiments.

Close modal

Two independent findings have so far been observed in vivo. The first, showing lack of T cell priming to peptide Ags, is in keeping with the generally poor activation of T cells on ICAM-1-null APCs (Figs. 1–3). The second, showing that ICAM-1-null mice respond normally to protein immunization, but only in the short-term, is more curious, especially in conjunction with the data that they are not more susceptible than WT T cells to death after activation (Fig. 9). This suggested to us that T cell priming in the absence of LFA-1/ICAM-interactions might generate a pool of secondary T cells that are biased more toward effector responses (TEM) than toward proliferation (TCM). To examine this, we used a system that allows ready detection of both types of primed cells. Immunization of WT mice with live Stm is known to generate an excellent, short-lived, CD4-mediated effector response that is dominated by Th1 cytokines as well as a long-lasting memory CD4 T cell response that can afford protection against a challenge infection many months after immunization (38, 39). Thus, WT and KO mice were immunized with live Stm, and IFN-γ responses and the ability to clear a challenge infection were scored 2 and 6 wk later. As seen in Fig. 10,A, cells from both groups of mice make substantial amounts of IFN-γ when stimulated for 16 h with bacterial sonicate, and the effector cytokine response declines equivalently between 2 and 6 wk in the two groups. However, while both groups of mice show enhanced clearance of a challenge infection at 2 wk as compared with unimmunized controls (Fig. 10,B), antibacterial immunity declines rapidly in the KO mice, with immunized and control mice showing similar bacterial loads at 6 wk (Fig. 10 C). When tested 8 wk after immunization, effector cytokine responses had declined further, and equivalently, in both groups, but antibacterial immunity was still strong in immunized WT mice (data not shown). These results confirm that T cell priming in the absence of costimulatory signals from ICAM-1 is inadequate for the generation of long-lived, proliferation-competent, TCM cells.

FIGURE 10.

Poor generation of long-lived antibacterial T cell immunity in KO mice. A, Staining for intracellular IFN-γ on gated CD4 cells from spleens of WT and KO mice 2 or 6 wk after immunization with Stm (indicated on right margin). Cells were stimulated with 30 μg/ml sonicate for 16 h or were left unstimulated. B and C, Bacterial loads in spleens of naive mice (□) and mice immunized with Stm (▪) either 2 wk (B) or 6 wk (C) earlier, following a challenge infection.

FIGURE 10.

Poor generation of long-lived antibacterial T cell immunity in KO mice. A, Staining for intracellular IFN-γ on gated CD4 cells from spleens of WT and KO mice 2 or 6 wk after immunization with Stm (indicated on right margin). Cells were stimulated with 30 μg/ml sonicate for 16 h or were left unstimulated. B and C, Bacterial loads in spleens of naive mice (□) and mice immunized with Stm (▪) either 2 wk (B) or 6 wk (C) earlier, following a challenge infection.

Close modal

It may be predicted from the above data that the response of ICAM-1-null mice to environmental Ags will also be dominated by TEM cells. To test this, we looked at the accumulation of TCM and TEM cells in 16- to 20-wk-old mice. Naive T cells can be distinguished phenotypically from previously activated cells by high levels of CD62L and low levels of CD44 (15, 52, 53, 54) and TCM and TEM cells can be differentiated by the relative expression of CD62L on the CD44highLy6Chigh population in the CD8 T cell pool, and on the CD44high population in the CD4 T cell pool (Fig. 11, A–C, Refs. 45 and 55). We used these markers to look at T cell subsets in peripheral lymphoid organs of WT and KO mice, and the results are shown in Table I. It can be seen that while the proportions of total memory-phenotype cells are similar between the two groups, the ratio of TCM:TEM is relatively lower in the KO mice.

FIGURE 11.

Identification of memory T cell subsets in vivo and in vitro. A–C, Gating used to determine the proportions of TCM and TEM cells in spleens and peripheral LNs of mice, for data in Table I. CD8 TCM cells were identified as CD62Lhigh cells in the CD44highLy6Chigh population (A and B) and CD4 TCM cells were identified as CD62Lhigh cells in the CD44high population (C). D, Poor up-regulation of CD62L on activated cells responding to Ag presented on BMDC from KO mice. Purified CD8 cells from P14 transgenic mice were cultured for 5 days with BMDCs from WT or KO mice in the presence or absence of peptide, as indicated. D, Expression of CD44 and CD62L on gated CD8+Vα2+ cells. E, Average proportion of TCM cells generated in such cultures, calculated from four independent experiments.

FIGURE 11.

Identification of memory T cell subsets in vivo and in vitro. A–C, Gating used to determine the proportions of TCM and TEM cells in spleens and peripheral LNs of mice, for data in Table I. CD8 TCM cells were identified as CD62Lhigh cells in the CD44highLy6Chigh population (A and B) and CD4 TCM cells were identified as CD62Lhigh cells in the CD44high population (C). D, Poor up-regulation of CD62L on activated cells responding to Ag presented on BMDC from KO mice. Purified CD8 cells from P14 transgenic mice were cultured for 5 days with BMDCs from WT or KO mice in the presence or absence of peptide, as indicated. D, Expression of CD44 and CD62L on gated CD8+Vα2+ cells. E, Average proportion of TCM cells generated in such cultures, calculated from four independent experiments.

Close modal
Table I.

TEM accumulate in ICAM-1-null mice responding to environmental Ags

Percentage of Cells that are PhenotypicallyWT MiceKO Mice
CD8 memory in spleen (CD44high Ly6Chigh)a 27.8 ± 1.9 22.3 ± 1.9 
CD8 TCM in spleen (CD44highLy6ChighCD62Lhigh)b 37.4 ± 5.9 8.1 ± 2.3 
CD8 memory in PLN (CD44highLy6Chigh)a 17.8 ± 0.9 21.2 ± 0.9 
CD8 TCM in PLN (CD44highLy6ChighCD62Lhigh)b 28.2 ± 4.7 16.2 ± 3.3 
CD4 memory in spleen (CD44high)c 81.1 ± 0.9 82.6 ± 1.2 
CD4 TCM in spleen (CD44highCD62Lhigh)d 34.1 ± 4.2 6.7 ± 1.9 
CD4 memory in PLN (CD44high)c 68.6 ± 1.5 63.3 ± 1.3 
CD4 TCM in PLN (CD44highCD62Lhigh)d 33.5 ± 6.1 9.9 ± 2.1 
Percentage of Cells that are PhenotypicallyWT MiceKO Mice
CD8 memory in spleen (CD44high Ly6Chigh)a 27.8 ± 1.9 22.3 ± 1.9 
CD8 TCM in spleen (CD44highLy6ChighCD62Lhigh)b 37.4 ± 5.9 8.1 ± 2.3 
CD8 memory in PLN (CD44highLy6Chigh)a 17.8 ± 0.9 21.2 ± 0.9 
CD8 TCM in PLN (CD44highLy6ChighCD62Lhigh)b 28.2 ± 4.7 16.2 ± 3.3 
CD4 memory in spleen (CD44high)c 81.1 ± 0.9 82.6 ± 1.2 
CD4 TCM in spleen (CD44highCD62Lhigh)d 34.1 ± 4.2 6.7 ± 1.9 
CD4 memory in PLN (CD44high)c 68.6 ± 1.5 63.3 ± 1.3 
CD4 TCM in PLN (CD44highCD62Lhigh)d 33.5 ± 6.1 9.9 ± 2.1 
a

Percent of CD8 cells. PLN, peripheral LN.

b

Percent of CD8 memory cells.

c

Percent of CD4 cells.

d

Percent of CD4 memory cells.

We also assessed whether T cells stimulated in vitro with ICAM-1-null APCs differentiate preferentially into TEM cells, by culturing purified CD8+ cells from P14 mice with BMDCs from WT or ICAM-1-null mice. The bulk of the CD8+ cells are transgenic, and they exhibit a naive phenotype (>90% also express Vα2, and are CD62Lhigh, data not shown). The recovery of cells left unstimulated in culture for 5 days is poor, but most cells are still CD62Lhigh, CD44low (Fig. 11,D, unpulsed). When stimulated with LCMV peptide, however, all cells up-regulate CD44, whether stimulated with BMDC from WT or KO mice. Ly6C is not expressed on the activated cells 5 days into culture; however, CD62L is up-regulated on a proportion of cells and this proportion is lower when the APCs are ICAM-1-deficient (Fig. 11, D and E, p < 0.05). Similar results were seen with lower doses of peptide and when anti-CD3 stimulated splenocytes from WT and KO mice, gated as shown in Fig. 11, A–C, were analyzed at 96 h (data not shown). Together, our results indicate that ICAM-1 on APCs delivers a signal to responding T cells that is crucial for the generation of proliferation-competent memory cells.

In this study, we undertook a comprehensive analysis of the effect of ICAM-1 deficiency on T cell priming. The approaches included in vitro and in vivo monitoring of early and late events in T cell activation and differentiation, and together our results indicate that during the course of T cell priming, signals provided or modified by ICAM-1 on APCs can have long-term consequences on cell fate determination. In particular, they influence the composition of the memory pool that is generated.

Our in vitro experiments (Figs. 1–3 and 5) indicate a specific role for ICAM-1 on APCs, and not on T cells, in determining both CD4 and CD8 responses. All early T cell activation events tested, including down-regulation of TCRs from the surface, modulation of activation markers, and cell cycling are affected, and the deficiency can be reversed by increasing ligand density or by increasing costimulation through CD28 (Fig. 4). These results are consistent with earlier reports indicating a role for TCR occupancy and costimulatory thresholds in determining T cell responses (29, 32, 33, 35, 56, 57, 58), and they extend them to ex vivo T cells and to a wide variety of readouts. We have also shown that WT T cells proliferate poorly in irradiated allogeneic ICAM-1-deficient hosts, and that no such defect is seen following transfer of T cells from ICAM-1-null mice into irradiated WT hosts (Fig. 6). Together, our results suggest that signals provided by ICAM-1 on APCs are necessary to support T cell proliferation in vivo, and that neither ICAM-2 and ICAM-3 on APCs, nor other integrins on T cells, can compensate.

The diminished proliferation of T cells on ICAM-1-deficient APCs could be compensated by the addition of exogenous IL-2 (Fig. 4). Although it is generally accepted that IL-2 is necessary for T cell proliferation (59), its role in determining the fate of cycling cells is unclear, and the presence of IL-2 has been shown to increase, decrease, or not affect the size of the memory pool (20, 25, 59, 60, 61). There is evidence, however, to link the extent of cell division with cell differentiation; once T cell cycling has been initiated, naive T cells can divide several times and differentiate into effectors and memory cells without further need for Ag (62), but the ultimate fate of the cells appears to depend on the number of cell divisions that occur. Thus, CD8 cells acquire effector potential after a single division cycle but more than five divisions are necessary for the generation of proliferation-competent memory cells (11, 63), and individual T cells that proliferate more in a primary response show greater secondary proliferation capability (64). Our data show that T cells primed on ICAM-1-null APCs differentiate preferentially into a CD44highCD62Llow TEM population (Fig. 11 and data not shown). Thus, they support earlier reports and extend them further by establishing a role for ICAM-1/LFA-1 interactions in the downstream effects.

That a lack of ICAM-1 during T cell priming may have long-term physiological consequences has been indicated by a prior study in which DO11.10 transgenic T cells that were primed on Drosophila APCs in the absence of ICAM-1 failed to cause disease when transferred into RIP-mOVA mice (65). The T cells migrated normally in vivo, but were unable to sustain long-lived expansion and differentiation into inflammation-mediating effectors. One of the significant findings to emerge from our study is that ICAM-1-null mice respond well to immunization with complex replicating and nonreplicating Ags, but that the T cell immunity is relatively short-lived (Figs. 8 and 10). This appears to be the outcome of a combination of events affecting different aspects of T cell priming. ICAM-1-deficent APCs support poor T cell proliferation, and clonal expansion is therefore likely to be more restricted in ICAM-1-null mice. Some compensation for the poor clonal expansion may be afforded by the higher survival of the T cells to AICD- and TSWD-inducing stimuli (Fig. 9), but the surviving T cells tend to differentiate preferentially into effector cells (Figs. 8 and 10). Over time, therefore, an effector population, which may be relatively short-lived, as in the case of OVA immunization (Fig. 8) or relatively long-lived, as in the case of Stm immunization (Fig. 10), accumulates in vivo, and is accompanied by a decline in the proliferation-competent memory pool. Our finding that TEM cells are preferentially generated in vitro following stimulation of T cells on ICAM-1-null APCs and that the TEM:TCM ratio is high in the lymphoid organs of ICAM-1-null mice responding to environmental stimuli lend further support for this model.

A second significant finding is that ICAM-1-null mice respond poorly to immunization with OVA peptide (Fig. 8). Because CFA emulsions were used for peptide and protein immunizations, the presence of inflammatory mediators and nonspecific T cell activation signals are likely to be similar in both situations. The most likely explanation for the difference is that the larger number of T cells responding to various OVA epitopes can augment the responses of individual responders. This is supported by our finding that the proliferation of KO T cells drops rapidly with decreasing cell density, and that exogenous IL-2 and extra costimulation can reverse the effect (Fig. 4).

In summary, our study demonstrates that ICAM-1 expressed on APCs in vivo supports the differentiation of T cells into a proliferation-competent TCM pool, and may be required for the generation of T cell responses to Ags expressed at low ligand density on APCs.

The authors have no financial conflict of interest.

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 grants from the Departments of Science and Technology and Biotechnology, Government of India, the Indian Council of Medical Research (to A.G., S.R., V.B.) and the Wellcome Trust (to V.B.). The National Institute of Immunology is supported by the Department of Biotechnology, Government of India.

4

Abbreviations used in this paper: TCM, central memory T cell; TEM, effector memory T cell; Stm, Salmonella typhimurium 754; LDA, limiting dilution analysis; LN, lymph node; WT, wild type; KO, knockout; PI, propidium iodide; TSWD, trophic signal withdrawal death; AICD, activation-induced cell death; BMDC, bone-marrow derived dendritic cell; PEC, peritoneal exudate cell; LCMV, lymphocytic choriomeningitis virus.

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