Role of CD28 co-stimulation in generation and maintenance of virus-specific T cells

Jeanette E. Christensen1, Jan P. Christensen1, Nanna N. Kristensen1, Nils J. V. Hansen1, Anette Stryhn1 and Allan R. Thomsen1

1 Institute of Medical Microbiology and Immunology, Panum Institute, 3C Blegdamsvej, 2200 Copenhagen, Denmark

Correspondence to: A. Randrup Thomsen; E-mail: A.R.Thomsen{at}immi.ku.dk
Transmitting editor: D. T. Fearon


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Efficient induction of T cell responses is normally assumed to require both TCR-mediated signaling and engagement of co-stimulatory molecules, in particular CD28. However, the importance of CD28 co-stimulation in induction and maintenance of antiviral T cell responses is not clearly established. For this reason antiviral CD4+ and CD8+ T cell responses in CD28-deficient mice were studied using two different viruses [vesicular stomatitis virus and lymphocytic choriomeningitis virus (LCMV)]. Intracellular cytokine staining and/or MHC–peptide tetramers were used to enumerate antigen-specific T cells. In addition, we used DNA constructs encoding viral epitopes to probe the importance of the epitope itself. Our results reveal that while the context of antigen presentation (live virus versus DNA construct) is a critical factor in determining the requirement for CD28 co-stimulation, epitope and virus dose play little if any role. Direct visualization of antigen-specific cells also confirms the notion that CD28 is more critical for the generation of antiviral Th1 cells than for Tc1 cells generated in response to the same virus (LCMV). Most importantly, the present study reveals that CD28 generally is essential for the host to respond optimally over a broad set of conditions, and our results may imply that the relatively CD28 independent activation of LCMV-specific CD8+ T cells may represent an extreme situation related to the non-cytolytic nature of this virus allowing the delivery of a uniquely strong and prolonged signal 1.

Keywords: co-stimulatory molecule, T cell activation, T cell memory, viral infection


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
According to current models of T cell activation, efficient triggering of naive T cells requires two fundamentally different signals (1,2). The first signal derives from interaction of a sufficient number of TCR with appropriate peptide–MHC complexes; without TCR ligation nothing happens, thus assuring the antigenic specificity of T cell activation. The second signal is delivered by a number of co-stimulatory molecules and in recent years it has become abundantly clear that optimal T cell activation is the net result of a complex summation of various positive (and negative) signals, each of which makes some contribution to the setting of the threshold for activation (36). One of the central T cell surface molecules involved in co-stimulation is CD28 which interacts with B7-1 and B7-2 on professional antigen-presenting cells (APC) (715). CD28 is expressed at low density on the majority of both CD4+ and CD8+ cells, and expression is up-regulated upon triggering of the T cell (8,9,16,17). However, CD28–B7 interaction is not pivotal for all T cell responses. Thus the murine virus lymphocytic choriomeningitis virus (LCMV) induces an almost unimpaired cytotoxic T lymphocyte (CTL) response in CD28 knockout mice (18,19). Capacity to bypass CD28 signaling is not, however, a property of all viruses as, for example, vesicular stomatitis virus (VSV) induces a CTL response that is absolutely dependent on CD28 expression (1921). Since studies of TCR transgenic mice indicated that the requirement for co-stimulation through CD28 varied inversely with the strength of signal 1 (3,19), the accepted explanation for the above difference has been that because VSV is not a natural murine pathogen and virus replication is extremely transient (20), this virus provides only a weak and short-lived antigenic stimulus (19). This is in contrast to LCMV, which due to prolonged replication in the secondary lymphoid organs has the capacity to induce a CD28-independent T cell response (19,20). In fact, extrapolating from this, it has been proposed that the generation of potent antiviral CTL responses under normal physiologic conditions do not require co-stimulation delivered through CD28 due to a high level of redundancy generally induced in the context of viral infection (22). However, alternative explanations have not been ruled out. Thus since antiviral CTL responses are often completely dominated by one or two immunodominant epitopes (2326), the above difference in CD28 dependence might simply reflect differences related to the predominant epitopes in the two model systems. Also, since some reports have indicated that the CTL response to VSV may be more CD4+ dependent than the response to LCMV (19,27), the difference in CD28 dependence could entirely reflect a difference in CD4+ dependence in combination with a pivotal role for CD28 co-stimulation in CD4+ T cell activation. The latter assumption is supported by the finding that the Th-dependent IgG antibody response to VSV is markedly reduced in CD28 knockout mice (18).

Similar to the situation for CD8+ T cell activation, which might not invariably require CD28–B7 interaction for activation, results for CD4+ T cells have also been equivocal (18,21,2833). However, the prevailing view is that in general CD4+ T cell responses are more CD28 dependent than CD8+ T cell responses (18). Again this may not be entirely correct (34) since CD4+ responses have mostly been evaluated in the context of relatively weak and non-replicating antigens. In particular, it should be stressed that the CD28 dependence of the CD4+ response to a virus inducing a CD28-independent CD8+ response has not been investigated. Therefore, the apparent difference might reflect different antigenic systems rather than a real biological phenomenon. In this context it is also relevant to note that the methods used to evaluate CD4+ and CD8+ responses up till now have been different and mostly indirect (antibody production versus CTL activity).

Therefore, it was the aim of this study to re-evaluate the importance of CD28 co-stimulation in the triggering of CD4+ and CD8+ responses in vivo. Since the primary objective was to understand the role of co-stimulation in a biologically and evolutionary relevant context, live viruses were used as the main tools to probe both CD4+ and CD8+ responses. Furthermore, to exclude any bias introduced by use of TCR transgenic mice (likely to express TCR with above-average avidity), only polyclonal responses were studied. An essential precondition for undertaking this study was the recent development of techniques to directly visualize antigen-specific T cells, as this made it possible not only to quantify T cell responses more precisely (35,36), but also to evaluate CD4+ and CD8+ responses using the same methodology, thus allowing a direct comparison. Using flow cytometry to detect antigen-specific T cells, we have evaluated T cell responses to two different viruses (VSV and LCMV), differing markedly in their requirement for co-stimulation. Our results confirm the notion that CD28 is more critical for the generation of antiviral Th1 cells than for CD8+ Tc1 cells generated in response to the same virus. More importantly, the present study indicates that CD28 is essential for the host to respond optimally over a broad set of conditions and that the importance of CD28 co-stimulation varies with the context of antigen stimulation, not the epitope as such.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
CD28-deficient (CD28–/–) mice (C57BL/6-Cd28tm1Mak) and CD40 ligand-deficient (CD40L–/–) mice (C57BL/6, 129-Cd40ltm1Imx) were the progeny of breeding pairs obtained from The Jackson Laboratory (Bar Harbor, ME). Female C57BL/6 (B6) mice were purchased from Bomholtgaard (Ry, Denmark). All mice were housed under specific pathogen-free conditions and sentinels were tested regularly for unwanted infections according to FELASA standards; no unwanted infections were detected. Mice from outside sources were always allowed to acclimatize for at least 1 week before entering into an experiment; by that time the animals were ~7–8 weeks old.

Virus infection
Three strains of LCMV were used: slowly replicating wild-type Armstrong virus (clone 53B), a rapidly invasive viscerotropic variant of the same virus (clone 13) and the viscerotropic Traub strain. Wild-type Armstrong virus and clone 13 variant virus were kindly provided by M. B. A. Oldstone (Scripps Clinic and Research Foundation, La Jolla, CA) (37). Traub virus stocks were produced and stored as previously described (38). VSV of the Indiana strain was used in some experiments; VSV was propagated in L929 cells and quantified as previously described (39).

DNA vaccination
Two vaccine constructs were used in this study, named GP33-10-hß2m or NP52-10-hß2m. These plasmids encode the signal sequence of (murine) ß2-microglobulin followed by the peptide of interest, which is tethered to (human) ß2-microglobulin through a 10-amino-acid linker (manuscript in preparation). The expression vector (pcDNA3.1/zeo+) contained the human cytomegalovirus immediate/early promoter and the encoded peptides were either LCMV glycoprotein (GP) 33–41 or VSV nucleoprotein (NP) 52–59. Plasmids were propagated in Escherichia coli XL1 Blue using standard techniques and purified using a Quiagen plasmid purification kit according to the manufacturer’s instructions.

For induction of CTL memory, mice were inoculated intradermally (using a gene gun) with 0.5 mg of plasmid DNA-coated gold particles (2 µg DNA/mg gold). Mice were boosted twice 4 weeks apart; 4 weeks after last immunization, memory cells were re-stimulated in vitro.

Cell preparations
Single-cell suspensions of spleen cells were obtained by pressing the organs through a fine steel mesh and, when required, erythrocytes were lysed by 0.83% NH4Cl treatment.

Re-stimulation
Single-cell suspensions of stimulator splenocytes were pulsed over night with 2 µM of either LCMV GP33–41 or VSV NP52–59. Cells were then washed and irradiated (2000 rad); 5 x 106 stimulator cells in complete medium with 4% IL-2-containing supernatant and 0.5% concanavalin A supernatant were transferred to each cluster well. Splenocytes (5 x 106) from DNA immunized mice in the same medium were subsequently added to each cluster well; cultures were prepared in duplicate. Cells were re-stimulated for 6 days before CTL activity was measured in a standard 51Cr-release assay. Splenocytes from unprimed mice served as controls.

Cytotoxicity assays
Virus-specific CTL activity was evaluated in standard 51Cr-release assays (26,38). Ex vivo virus-specific cytotoxicity was detected using 51Cr-labeled EL-4 cells pulsed with relevant peptides for 1 h. The CTL activity of DNA-immunized mice was assessed using RMA-S target cells pulsed overnight with either NP52–59 or GP33–41. Untreated EL-4 or RMA-S cells served as control targets. Incubation time was 5–6 h at 37°C and four to six different E:T ratios were evaluated in triplicates for each effector cell preparation. Percent specific 51Cr release was calculated as 100 x [(c.p.m. test release – c.p.m. spontaneous release)/(c.p.m. maximum release – c.p.m. spontaneous release)] (38).

Organ virus titers
To determine LCMV virus titers, organs were first gently homogenized in PBS containing 1% FCS to yield a 10% (v/w) organ suspension. Organ suspensions were clarified by centrifugation and serial 10-fold dilutions of the supernatants were prepared in PBS with 1% FCS. Then 0.2 ml of each dilution was transferred in duplicates to flat-bottomed, 24-well plates and MC57G cells in MEM were added. Plates were then incubated for 4–6 h at 37°C in 5% CO2, allowing the cells to settle and adhere to the plastic surface. Subsequently, 0.3 ml of a 1:1 mixture of 2% methylcellulose in double-distilled water and double-strength MEM with 10% FCS, antibiotics and glutamine was added. After 48 h of incubation, cell monolayers were fixed with 4% formaldehyde in PBS over a period of 20–30 min at room temperature and permeabilized by incubation in 0.5% Triton X-100 in HBSS for 20 min. The next day monolayers were labeled with a monoclonal rat anti-LCMV (VL-4) over a period of 60–90 min and, following extensive washing, peroxidase-labeled goat anti-rat antibody was added for 60–90 min. Following another washing, o-phenylendiamine (substrate) was added and the reaction was terminated after 10–30 min by washing the plate with water. The numbers of p.f.u. were counted and organ virus titers were expressed as p.f.u./g tissue (40).

In vivo bromodeoxyuridine (BrdU) labeling
Mice were given BrdU (Sigma, St Louis, MO) at 0.8 mg/ml in their drinking water for a period of 3 days (41,42). BrdU-containing water was protected from light and changed daily.

mAb for flow cytometry
The following mAb were purchased from PharMingen (San Diego, CA) as rat anti-mouse antibodies: phycoerythrin (PE)- and CyChrome-(Cy)-conjugated anti-CD8a (53-6.7, IgG2a), PE- and FITC-conjugated anti-CD4 (H129.19, IgG2a), FITC-conjugated anti-CD49d (common {alpha}4 chain of LPAM-1 and VLA-4) (RI-2, IgG2b), biotin- and FITC-conjugated anti-CD44 (IM7, IgG2b), biotinylated anti-L-selectin (CD62L) (MEL-14, IgG2a ), PE-conjugated anti-IFN-{gamma} (XMG1.2, IgG1), PE-conjugated anti-IL-2 (S4B6, IgG2a) and FITC-conjugated anti-BrdU (B44, IgG1). Rat anti-LCMV (VL-4) was harvested from a B cell hybridoma kindly provided by R. M. Zinkernagel (Institute of Pathology, University Hospital Zurich, Zurich, Switzerland)and peroxidase-conjugated AffiniPure goat anti-rat IgG (code no. 112-035-003) was purchased from Jackson ImmunoResearch (West Grove, PA).

MHC–peptide tetramers for flow cytometry
Relevant tetramers were obtained through the National Institute of Allergy and Infectious Disease Tetramer Facility, and the National Institutes of Health AIDS Research and Reference Reagent Program.

Flow cytometry
Staining of cells for flow cytometry was performed according to standard laboratory procedures (42,43). Briefly, 106 cells were stained with directly labeled mAb for 20 min in the dark at 4°C and washed 2 times in PBS with 1% NaN3. In case of biotin-conjugated antibody, cells were additionally incubated with streptavidin–TriColor (Caltag, San Francisco, CA). Finally, cells were washed and fixed with 1% paraformaldehyde. Tetramer staining was carried as recently described (44).

For BrdU staining (42), cells were stained for surface markers as described above, transferred to cold 0.15 M NaCl solution and fixed by adding cold 96% ethanol. After 30 min incubation on ice, cells were washed once and resuspended in PBS, 0.01% Tween 20 and 1% paraformaldehyde. After 1 h incubation at room temperature, cells were washed and resuspended in PBS, 0.15 M NaCl, 4.2 mM MgCl2, pH 5 containing 50 Kunitz units/ml DNase I (Sigma). After incubation for 15 min at 37°C, cells were washed once in PBS before adding the anti-BrdU antibody. Cells were incubated 30 min at room temperature, washed and analyzed.

To detect intracellular cytokine (42), splenocytes were incubated in vitro at 37°C in 5% CO2 either with or without relevant peptide for 5–6 h in complete RPMI supplemented with monensin (3 µM; Sigma) and murine recombinant IL-2 (50 U/ml; R & D Systems, Abingdon, UK). The peptides used were LCMV GP33–41, NP396–404, GP276–286 (0.1 µg/ml) and GP61–80 (1.0 µg/ml), and VSV NP52–59 (1.0 µg/ml). Following incubation, cells were washed in FACS medium (PBS containing 1% BSA, 0.1% NaN3 and 3 µM monensin), stained with relevant surface antibodies and washed again. Subsequently, cells were fixed with 1% paraformaldehyde for 30 min at 4°C, washed and permeabilized using 0.5% saponin. Cells were then stained with anti-IFN-{gamma} or anti-IL-2 for 20 min at 4°C, washed and analyzed.

Samples were analyzed using a Becton Dickinson (Mountain View, CA) FACSCalibur, and at least 104 live mononuclear cells were gated using a combination of low angle and side scatter to exclude dead cells and debris; data analysis was conducted using CellQuest. CD4+ and CD8+ T cells co-expressing VLA-4 or CD44 (activation markers) and cytokine/tetramers were considered antigen specific.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Requirement for CD28 co-stimulation in antiviral CD8+ T cell responses varies between viruses, but not between epitopes
Previous results have shown that VSV induces little or no CTL response in the absence of CD28 co-stimulation. However, it is not clear to which extent this reflects lack of triggering/clonal expansion or lack of subsequent effector cell differentiation. To resolve this question, CD28–/– and matched wild-type mice were infected with VSV and the virus-induced T cell response was evaluated. Analyzing virus-induced proliferation of splenic CD8+ T cells in VSV-infected mice using in vivo BrdU incorporation as a parameter (Fig. 1A), no difference between CD28–/– and wild-type mice was detected in the initial phase of the response [mice pulsed day 0–3 post-infection (p.i.)]. However, during the period of most extensive CD8+ T cell proliferation in wild-type mice (mice pulsed day 3–6 p.i.) (20), little proliferation was observed in CD28–/– mice, thus separating the virus-induced proliferative response into two distinct phases differing in the requirement for co-stimulatory signals [a similar pattern was observed in CD40L–/– mice (data not shown)]. In comparison, early CD4+ T cell proliferation was marginal in both mouse strains, whereas a distinct proliferative response was detected in wild-type mice, but not in CD28–/– mice, when analyzed on day 6 p.i. Virus-specific Tc1 cells present in the spleen were visualized following short-term stimulation with NP52–59 or by use of NP52–59/H-2Kb tetramers (Fig.1B). As expected from previous analysis of ex vivo CTL activity (19,45), the frequency of cytokine-producing Tc1 cells was markedly reduced in CD28–/– mice. More important, the frequency of tetramer+ cells was also much reduced in CD28–/– mice, demonstrating that CD28 co-stimulation was essential for optimal triggering of virus-specific CD8+ T cells following VSV infection. However, some cells were activated and seemed to differentiate in parallel to wild-type CD8+ T cells. Notably, despite significant and comparable proliferation of CD8+ T cells day 0–3 p.i., no antigen-specific CD8+ T cells were present on day 3 p.i., confirming previous result indicating that most cells proliferating during this phase are memory cells of irrelevant specificities induced to enter the cell cycle by non-specific signals.



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Fig. 1. VSV-induced expansion of CD8+ and CD4+ T cells. CD28–/– and C57BL/6 mice were infected with 106 p.f.u. of VSV i.v. (A) Mice were given BrdU for 3 days prior to analysis; on days 3 and 6 p.i., splenocytes were surface stained with PE-conjugated anti-CD8 or PE-conjugated anti-CD4 and biotin-conjugated anti-L-selectin, permeabilized, and stained with FITC-conjugated anti-BrdU. Gates were set for CD8+ or CD4+ cells, and percentages of BrdU+ cells are shown; BrdU+ cells were predominantly L-selectinhigh on day 3 p.i. and L-selectinlow on day 6 p.i. (B) On the indicated days splenocytes were harvested and NP52-specific CD8+ T cells were quantified using either MHC I tetramers or by intracellular staining as described in Methods. Points represent individual mice; results are representative of two experiments.

 
In contrast to the situation in VSV-infected mice, the frequency of virus-specific Tc1 cells generated in LCMV-infected mice was hardly reduced (Fig. 2), although absolute numbers were reduced somewhat (30–50%, see Fig. 7). In this case we evaluated the responses towards two immunodominant (GP33–41 and NP396–404) and one subdominant (GP276–286) epitope; by using this approach, a broad range of MHC–peptide expression levels on the surface of infected cells (~102–103 copies/infected cell) could be covered (46). As seen in Fig. 2, Tc1 cells directed against either dominant epitopes were detected with almost equal frequency in CD28–/– and wild-type animals, and even the response directed towards the subdominant epitope was only slightly impaired in CD28–/– mice, demonstrating that the ability to raise a LCMV-specific Tc1 response was not limited to a single immunodominant epitope characterized by a high level of surface expression (46). A similar difference in CD28 dependency was observed if the peptide-specific CTL activities induced by VSV and LCMV were compared in standard 51Cr-release assays (data not shown).



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Fig. 2. Frequencies of LCMV-specific CD8+ cells from CD28–/– and C57BL/6 mice. Ten days after infection (104 p.f.u. of LCMV clone 13 i.v.), splenocytes were stimulated in vitro with LCMV GP33–41, NP396–404 or GP276–286 for 5 h. Cells were surface stained with Cy-conjugated anti-CD8 and FITC-conjugated anti-VLA-4, permeabilized, and stained with PE-conjugated anti-IFN-{gamma}. Gates were set for CD8+ cells and percentage of IFN-{gamma}+ cells are shown. Results of individual mice are depicted.

 


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Fig. 7. LCMV-specific CD8+ T cells in CD28–/–, CD40L–/– and wild-type mice infected 10 days earlier with 102, 104 or 106 p.f.u. of LCMV clone 13 i.v. Splenocytes were stimulated in vitro with LCMV GP33–41 or NP396–404 for 5 h, and cells were surface stained with Cy-conjugated anti-CD8 and FITC-conjugated anti-VLA-4, permeabilized, and stained with PE-conjugated anti-IFN-{gamma}. The absolute number of IFN-{gamma}+ cells is shown. Medians and ranges are depicted (n = 2–9/group); the results for 104 p.f.u. are pooled from two independent experiments generating similar results.

 
Absence of CD28 co-stimulation does not select for higher average TCR avidity
Since the requirement for co-stimulation has been found to be inversely related to the strength of signal 1 (3,19), one might expect that LCMV-specific Tc1 cells generated in the absence of CD28 might represent preferential expansion of T cell precursors with TCR of above-average avidity. To test this, we first studied the tetramer-binding capacity of primary GP33-specific CD8+ T cells generated in CD28–/– mice and wild-type controls by titrating the amount of MHC–peptide tetramer needed to obtain half-maximal binding (47); no difference between the strains was observed (Fig. 3A). Second, we measured function-based avidities of splenic CD8+ T cells from LCMV-infected CD28–/– and wild-type mice by peptide titration (47). The percentage of cytokine-producing cells at the highest peptide concentration tested was defined as 100% in either strain and responses at lower peptide concentrations were expressed as a percentage thereof. As can be seen in Fig. 3(B), marginal if any increase in functional avidity was observed for T cells generated in CD28–/– mice and similar results were obtained for NP396-specific CD8+ T cells (data not shown). Together these results suggest that no marked avidity bias exists for LCMV-specific CD8+ T cells triggered in the absence of CD28 and that down-stream signaling pathways operate with similar efficiency.



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Fig. 3. The avidity profiles of GP33-specific CD8+ T cells in LCMV-infected CD28–/– mice and wild-type mice. Mice were infected i.v. with 4800 p.f.u. of LCMV Armstrong. On day 9–10 p.i., cells were incubated with graded concentrations of GP33-specific tetramers or stimulated in vitro with graded doses of GP33–41 peptide for 5 h. Cells were surface stained with Cy-conjugated anti-CD8, FITC-conjugated anti-VLA-4 and tetramers (A) or permeabilized and stained with PE-conjugated anti-IFN-{gamma} (B); gates were set for CD8+ cells. Numbers of tetramer+ and IFN-{gamma}+ cells at each concentration are shown (medians and ranges of n = 2–3 mice/strain) as a percentage of positive cells at the highest concentration; results are representative of two experiments.

 
Absence of CD28 co-stimulation reduces IL-2 production by virus-specific CD8+ T cells
The above results suggest that LCMV-specific CD8+ T cells generated in the presence or absence of CD28 co-stimulation are qualitatively similar. However, we have recently found that a minority subset of virus-specific CD8+ T cells also produce IL-2 and, since it is the dogma that CD28 co-stimulation is essential to induce IL-2 production, we considered it pertinent to test if CD8+ T cells generated in the absence of CD28 co-stimulation would also be able to produce this cytokine. While the frequencies of IFN-{gamma}-producing CD8+ T cells are overlapping, a small but statistically significant decrease in the frequency of IL-2-producing cells is noted in CD28–/– mice (Fig. 4) and a preferential relative decrease in IL-2-producing cells was consistently observed in CD28–/– mice for at least 2 months after infection (data not shown). Thus our findings point to an increased, but not absolute, requirement for CD28 co-stimulation when it comes to IL-2 production by CD8+ T cells.



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Fig. 4. Comparison of frequencies of LCMV-specific CD8+ T cells producing IFN-{gamma} and IL-2 generated in the presence or absence of CD28 co-stimulation. Nine days after infection (103 LD50 of LCMV Traub), splenocytes from CD28–/– and wild-type mice were stimulated in vitro with GP33–41 for 5 h. Cells were surface stained with Cy-conjugated anti-CD8 and FITC-conjugated anti-VLA-4, permeabilized, and stained with either PE-conjugated anti-IFN-{gamma} or anti-IL-2. Gates were set for CD8+ cells and percentages of cytokine+ cells are depicted. Points represent individual mice; data are pooled from two identical experiments. Statistical evaluation was carried out using the Mann–Whitney rank test

 
Requirement for CD28 co-stimulation does not increase with reduction of virus dose
One reason for the ability of LCMV, but not VSV, to largely bypass the requirement for CD28 co-stimulation could be the capacity of the former virus to replicate extensively in the murine host. If this explanation were correct, one would expect CD28 dependency to correlate inversely with the level of virus replication in the secondary lymphoid organs.

To study this, mice were infected i.v. with two doses (low and intermediate) of the slowly invasive LCMV Armstrong strain, and on days 7 and 9 p.i. the Tc1 response against the two major viral epitopes was analyzed. As can be seen in Fig. 5, an impaired Tc1 response was revealed in CD28–/– mice infected with either virus dose 7 days earlier. On day 9 p.i. the difference between knockouts and wild-type mice was smaller although not absent. Most importantly, the overall pattern was not critically dependent on the virus dose and even in CD28–/– mice infected with the lower dose, a clear response was induced. From this we conclude that the number of virus-infected APC per se does not play a major role in determining the requirement for CD28 co-stimulation. However, since the CD8+ T cell response is delayed in mice lacking CD28 expression, it may be inferred that this molecule is critical for an optimal response independent of virus dose.



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Fig. 5. LCMV-specific CD8+ T cells from CD28–/– and wild-type mice infected i.v. with 100 and 4800 p.f.u. of LCMV Armstrong. On days 7 and 9 p.i., splenocytes were stimulated in vitro with LCMV GP33–41 or NP396–404 for 5 h. Cells were surface stained with Cy-conjugated anti-CD8 and FITC-conjugated anti-VLA-4, permeabilized, and stained with PE-conjugated anti-IFN-{gamma}. Gates were set for CD8+ cells and percentage of IFN-{gamma}+ cells is shown. Medians and ranges are presented (n = 3/group); results are representative for two experiments.

 
Requirement for CD28 co-stimulation follows the context of antigen stimulation, not epitope
The above results suggest that LCMV infection provides a nearly optimal stimulating context that allows CD8+ responses to be generated without much need for CD28 co-stimulation. However, formally it still could not be ruled out that the studied LCMV epitopes were somehow unique in their capacity to activate CD8+ T cells.

To directly test whether a difference could lie in the epitope, DNA vectors were generated to include either LCMV GP33–41 or VSV NP52–59 as minimal epitopes. Using this approach, epitopes representing different ends of the CD28 dependency spectrum could be compared under similar priming conditions: would a difference still prevail? The vectors applied were constructed to include the signal sequence of (murine) ß2-microglobulin followed by the relevant epitope linked via a 10-amino-acid linker to (human) ß2-micoglobulin (manuscript in preparation). The aim of this strategy was to enhance (and stratify) the expression of the final peptide–MHC complex on the cell surface. Thus, the gene product was targeted to the endoplasmic reticulum through the signal sequence and the covalent binding to ß2-microglobulin was intended to further increase the likelihood of successful MHC class I association. CD28–/– and wild-type mice were vaccinated twice 4 weeks apart and tested 4 weeks after last priming. Because the frequencies thus induced are too low to be directly visualized, CD8+ T cell priming was evaluated as cytolytic activity following re-stimulation in vitro. Independent of the viral epitope, the CTL response was similarly reduced (~16-fold) in CD28–/– mice (Fig. 6). The increased requirement for co-stimulation regarding the GP33–41 epitope is unlikely to reflect a peripheral versus. a systemic route of antigen presentation as mice infected s.c. with live LCMV have nearly similar frequencies of LCMV-specific memory CD8+ T cells in their spleen at 4 weeks p.i. irrespective of CD28 co-stimulation (data not shown). Therefore, the context of antigen presentation (live virus versus DNA vector) appears to play a greater role in determining the importance of CD28 co-stimulation than does properties directly associated with the individual epitope.



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Fig. 6. Recall CTL responses in DNA immunized CD28–/– and wild-type mice. Mice were primed intradermally (using a gene gun) with 1 µg of plasmid DNA encoding either GP33-10-hß2m or NP52-10-hß2m and boosted twice 4 weeks apart. Four weeks after the last immunization, splenocytes were re-stimulated in vitro for 6 days with irradiated splenocytes pulsed with LCMV GP33–41 or VSV NP52–59. CTL activity was measured using RMA-S target cells pulsed overnight with either GP33–41 or NP52–59; untreated RMA-S cells served as control targets. Results of individual mice are depicted; results are representative of two experiments.

 
Generation of LCMV-specific CD4+ T cells is more dependent on CD28 co-stimulation than are CD8+ T cells
To investigate the role of CD28 in the generation of Th1 cells, IFN-{gamma}- and IL-2-producing CD4+ cells were visualized following in vitro stimulation with the MHC class II-restricted LCMV GP61–80 peptide. As can be seen from Table 1 even under conditions where the reduction in the antiviral CD8+ response is limited (~2-fold), the CD4+ response is substantially reduced (~7-fold). However, a significant antiviral Th1 cell response can be induced in absence of CD28, and, interestingly, we noted that the gap between wild-types and CD28–/– mice tended to decrease with time. Thus, at 2–4 months p.i. the difference was only ~2-fold, albeit still statistically significant (0.96 versus 0.49%, P < 0.05, median of 7 mice/group).


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Table 1. Role of CD28 co-stimulation in generation of cytokine-producing T cell subsets in LCMV-infected micea
 
Different roles of CD28 and CD40 ligand in sustaining LCMV-specific CD8+ T cell responses
Based on the fact that CD28 co-stimulation has been demonstrated to increase cell survival following specific activation (4851), we also wanted to determine whether CD28 co-stimulation could play a significant role in protecting mice against the consequences of infection with high doses of LCMV. Under these conditions most virus-specific CD8+ T cells undergo apoptosis, and the absence of CD4+ T cells shifts the threshold for irreversible exhaustion of the CD8+ T cell response towards lower virus doses (52,53). Previously, CD40L has been found to play a critical role in this context (54,55) and it was therefore of interest to compare these two routes of co-stimulation.

In a previous study we found that mice infected with an intermediate dose of slowly replicating LCMV Armstrong did not require either co-stimulatory system for generation of a primary CTL response (20). However, CD40L but not CD28 was needed to sustain efficient CD8+ immunosurveillance and virus control in the long term. To extend this finding we first infected CD28–/–, CD40L–/– and wild-type mice over a broad range of virus doses using the more invasive clone 13 LCMV strain. When tested at the peak of the primary response (day 10 p.i.), essentially parallel dose–response curves were obtained for CD28 and wild-type mice (Fig. 7); only at the highest dose which is known to induce chronic infection also in wild-type mice (52), the CD8+ response showed evidence of exhaustion in these two mouse strains. This is in marked contrast to CD40L–/– mice, which present an impaired CD8+ T cell response already at 10 days after infection with a low dose of this invasive LCMV variant (Fig. 7).

To study the long-term effects, CD8+ T cell memory in CD28–/– and wild-type mice infected 4–5 months previously with either of the two highest doses of virus were evaluated. While little difference between the two strains was noted at the lower dose, significantly impaired CD8+ T cell memory was observed in CD28–/– mice following infection with 106 p.f.u. (Fig. 8). Thus, while GP33-specific CD8+ T cells in wild-type mice had partially recovered from the initial high dose immune paralysis, neither GP33- nor NP396-specific CD8+ cells were detected in CD28 knockouts and correspondingly high levels of virus were detected in the organs of high-dose infected CD28 mice (Fig. 8). A similar trend towards lower numbers of CD8+ memory cells in CD28 knockouts was noted in mice infected with high doses of another viscerotropic LCMV strain (LCMV Traub, data not shown). Consequently CD28 co-stimulation significantly improves the chances of the host to resist high-dose immune paralysis and control the chronic infection.



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Fig. 8. LCMV-specific memory CD8+ T cells in CD28–/– and wild-type mice infected 4–5 months earlier with 104 or 106 p.f.u. of LCMV clone 13 i.v. Splenocytes were stimulated in vitro with LCMV GP33–41 or NP396–404 for 5 h. Cells were surface stained with Cy-conjugated anti-CD8 and FITC-conjugated anti-VLA-4, permeabilized, and stained with PE-conjugated anti-IFN-{gamma}. Gates were set for CD8+ cells and percentage of IFN-{gamma}+ cells is shown. Medians and ranges of 4–6 mice/group are depicted. *Only three of six wild-type mice had clearly detectable NP396-specific cells. Additionally, kidney and lung virus titers were assayed in those CD28–/– and wild-type mice infected with 106 p.f.u.; ranges of n = 5–6 mice are presented.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
In the present report we have re-evaluated the importance of CD28 co-stimulation for generation of antiviral Tc1 and Th1 responses. We find that some T cells are consistently activated independently of CD28 co-stimulation, but co-stimulation through this molecule is generally required for an optimal T cell response. Furthermore, significant differences exist between different viruses and different T cell subsets.

Thus, in the case of LCMV infection, the reduction in the Tc1 response caused by absence of CD28 is <2-fold, while few CD8+ effector cells are generated in response to VSV. Why this difference exists is not entirely clear; however, several explanations may be considered.

First, unique conditions surrounding the immunodominant epitopes of either viral model infection might play a role. However, both major LCMV epitopes and VSV NP52 bind with high affinity to their presenting class I molecules (46,56), and no difference in the average avidity of the involved TCR could be detected (unpublished observation). Furthermore, all LCMV-derived epitopes seem to induce a CD8+ response in CD28–/– mice despite very different levels of expression and immunodominance. Most importantly, direct comparison of the immunogenicity of a CD28-dependent VSV epitope and a CD28 independent LCMV epitope presented in an identical context (as DNA vector) reveals that there is no intrinsic capacity of the latter causing CD28 co-stimulation to be redundant.

Second, a difference in CD4+ dependency could play a role given that CD4+ T cells seem to rely more on CD28 co-stimulation for an optimal response. However, analyzing the VSV-specific CD8+ response neither we (20) nor others (21) have been able to confirm earlier reports that this response should require CD4+ T cells. Therefore, neither the VSV-specific nor the primary LCMV-specific CD8+ T cell response seem to be critically dependent on CD4+ T cell help, ruling out that a difference in CD4+ dependency constitute the underlying mechanism.

Third, a difference in the capacity of the involved viruses to activate relevant APC might play a role. Recently it has been found that dendritic cells in VSV-infected mice generally express lower levels of critical co-stimulatory molecules than do dendritic cells in LCMV-infected mice (57). Therefore, high expression of other co-stimulatory molecules might render CD28 signaling redundant in the latter mice. This possibility was recently studied by co-infecting mice with VSV and LCMV (20). While co-infection partially rescued the VSV-specific response in CD40L–/– mice, this was not the case in CD28–/– mice, indicating that a general difference in the level of non-specific activation of APC does not suffice as an explanation for the striking difference in requirements for CD28 co-stimulation.

Fourth, the number of virus-presenting APC might be important. It has been suggested that high numbers of APC could provide a more efficient and sustained signal 1 (19) and it is known that the requirement for CD28 co-stimulation is inversely related to the strength of this signal (3). However, this does not suffice to explain the difference between LCMV and VSV responses. Thus, we found little influence of virus dose on the requirement for CD28 co-stimulation in LCMV-infected mice and although it might be argued that even the lowest LCMV dose tested might lead to more extensive antigen presentation than in the case of, for example, VSV infection, this argument carries little weight. Thus, judged from a comparison of the frequency of virus-specific CD8+ T cells generated in VSV-infected and low-dose LCMV-infected wild-type mice, there is no reason to believe that epitope-presenting APC are less abundant following i.v. infection with 106 p.f.u. of VSV than following infection with 102 p.f.u. of LCMV Armstrong. Therefore, we find it unlikely that the number of infected APC per se is the critical determinant. We consider this view to be consistent with recent insight into the T cell–APC interaction (58). Thus, T cells and APC seem to interact in a very intimate fashion (the immunological synapse), and it is difficult to envision how the presence of multiple APC should qualitatively affect T cell activation under these conditions—most TCR are probably already engaged. Therefore, qualities associated with the individual APC are likely to play a much more important role and a stable T cell–APC interaction could be a key factor in bypassing a requirement for CD28 co-stimulation.

Therefore, the last possibility—which to us appears the most likely—is that LCMV-infected APC provide a uniquely stable signal 1 compared to, for example, VSV-infected APC. This could be explained if lytic viruses kill their target cells too rapidly for actively infected cells to provide substantial antigen presentation. Consequently, antigen presentation would have to rely on dendritic cells that are not themselves supporting viral replication (i.e. cross-presentation). In contrast, if a non-cytolytic virus like LCMV infects the dendritic cells, these may directly serve as antigen presenters. Under these conditions there will be a continual feeding of virus-coded proteins into the MHC class I antigen presentation pathway leading to prolonged and stable surface presentation of relevant peptide–MHC complexes. Although a DNA vector might be viewed as being qualitatively comparable to a non-cytolytic virus, this is probably not true in quantitative terms. Thus much more extensive synthesis of relevant proteins is likely to occur in APC infected with live virus compared to vector transfected APC and this may explain the difference in requirement for CD28 co-stimulation.

As regards the role of CD28 co-stimulation in the generation of antiviral Th1 responses, the present analysis allows a direct comparison of the requirement for CD28 in the activation of Tc1 and Th1 cells responding to the same virus. Our data demonstrate that even under conditions that allow almost unimpaired generation of antiviral Tc1 cells, the antiviral Th1 response is significantly impaired. The reason why CD4+ T cells should rely more on CD28 co-stimulation for an optimal response is not entirely clear, but differences in co-stimulatory requirements of CD4+ and CD8+ cells have been reported before (5963), e.g. CD4+ T cells also appear more dependent on CD40–CD40L interaction than are CD8+ T cells (63) whereas the opposite is seen for signaling through 4-1BB (60). Interestingly, despite the reduced frequency of virus-specific CD4+ T cells, Tc1 expansion is normally sustained in CD28–/– mice; only when the mice are infected with very high doses of rapidly invasive virus causing a chronic infection is CD28 important. This is consistent with previous findings demonstrating that CD4+ T cells are not pivotal for the initial expansion of LCMV-specific CD8+ T cells, but Th cells are required to maintain Tc responsiveness in the long term (64). Furthermore previous studies have clearly shown that the requirement for CD4+ help increases when virus clearance is delayed (44,65). Therefore, CD28 co-stimulation appears to be limiting only in the case of a chronic viral infection necessitating CD4+ help for a long-standing CD8+ effector response. The implication of this observation is that except when the immune system is pushed to the limit (as in mice infected with high doses of rapidly invasive virus), CD4+ T cells are present in substantial excess (~5- to 10-fold) of what is actually required.

In conclusion, the present findings underscore an important role of CD28 co-stimulation in the generation of antiviral Tc1 and Th1 responses. While it has previously been claimed to be the rule that viruses do not need CD28 for generation of optimal CD8+ T cell responses (22), the present data points more to a continuous spectrum of CD28 dependency varying with virus and T cell parameter studied. Interestingly, if a relatively CD28-independent response reflects targeting of dendritic cells by non-cytolytic virus—as we suggest—a high degree of functional redundancy of CD28 is likely to be the exception rather than the rule. That CD28 co-stimulation is biologically/evolutionary relevant is underscored by the fact that CD28–/– mice succumb to doses of VSV and influenza virus 50- to 100-fold lower than do matched wild-type animals (unpublished observation).


    Acknowledgements
 
The National Institute of Allergy and Infectious Disease Tetramer Facility, and the NIH AIDS Research and Reference Reagent Program are acknowledged for supplying MHC–peptide tetramers. This study was supported in part by the Danish Medical Research Council, the Biotechnology Center for Cellular Communication and the Novo Nordisk Foundation. J. E. C. is the recipient of a PhD scholarship from the Faculty of Health Sciences, University of Copenhagen, Denmark. J. P. C. is the recipient of a Research Fellowship from the Weimanns Foundation, Denmark. A. S. is the recipient of a Research Fellowship from the Danish Medical Research Council and the Carlsberg Foundation.


    Abbreviations
 
APC—antigen-presenting cell

BrdU—bromodeoxyuridine

CD40L—CD40 ligand

CTL—cytotoxic T lymphocyte

Cy—CyChrome

GP—glycoprotein

LCMV—lymphocytic choriomeningitis virus

NP—nucleoprotein

PE—phycoerythrin

p.i.—post-infection

VSV—vesicular stomatitis virus


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 Top
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 Introduction
 Methods
 Results
 Discussion
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