|Home | About | Journals | Submit | Contact Us | Français|
Immunization in the absence of CD4+ T cell help results in defective CD8+ T cell memory, deficient recall responses and diminished protective immunity. Here we investigated at what stage during the immune response to pathogen CD4+ T cells are essential in the promotion of functional CD8+ T cell memory. Memory CD8+ T cell numbers decreased gradually in the absence of CD4+ T cells despite the presence of similar numbers of memory cell precursors at the peak of the effector phase. Adoptive transfer of effector or memory CD8+ T cells into wild-type or CD4+ T cell–deficient mice demonstrated that the presence of CD4+ T cells was important only after, not during, the early CD8+ T cell programming phase. In the absence of CD4+ T cells, memory CD8+ T cells became functionally impaired and decreased in quantity over time. We conclude that in the context of an acute infection, CD4+ T cells are required only during the maintenance phase of long-lived memory CD8+ T cells.
The rapid and more efficacious response of memory T cells after the second encounter with a pathogen constitutes a hallmark trait of adaptive immunity. The ‘canonical’ CD8+ T cell response to an acute infection consists of three well defined phases: the proliferation of naive cells to produce large numbers of effector cells; the contraction of these effector populations into memory cells once antigen is cleared; and the long-term maintenance of these memory cells. A brief encounter with antigen, as short as 24 h, can initiate ‘programmed differentiation’ in activated CD8+ T cells that leads to many rounds of antigen-independent division and the acquisition of effector function1–4. Effector cell populations at the peak of a CD8 response are also programmed to contract during their priming5. Thus, during an immune response to infection, intrinsic and external signals ‘dictate’ or program the complete CD8+ T cell response6,7.
Although CD4+ T cells are not required for primary CD8+ T cell responses against pathogens, memory CD8+ T cell numbers and secondary responses to bacterial or viral challenge are decreased over time in a CD4+ T cell–deficient animal8–11. In those studies8–11, it was not apparent when the CD4+ T cell help was required to enhance the survival of the antigen-specific memory CD8+ T cells. It has been speculated that CD4+ T cells interacting with antigen-presenting cells also engage in crosstalk with CD8+ T cells during the proliferation or programming phase, delivering the necessary ‘instructive’ and/or survival signals for the generation of a fully functional memory CD8+ T cell pool12–14. It has also been suggested that signals given during the contraction or maintenance phases regulate homeostasis of the memory population14–16. Because activated CD8+ T cell populations undergo a program that dictates their subsequent contraction and eventual memory cell frequencies, we sought to understand at what phase of the immune response to pathogen CD4+ T cells are crucial in determining memory CD8+ T cell numbers and functionality. Are naive CD8+ T cells programmed early during the expansion phase by CD4+ T cell help to become productive memory cells? Or do CD4+ T cells provide an important contribution in the maintenance of memory CD8+ T cell numbers later during the memory phase?
Expression of the interleukin 7 receptor α-chain (IL-7Rα) chain at the peak of the CD8+ T cell primary response marks those effector cells that will differentiate into memory cells17,18. Here we found that effector CD8+ T cells expressing IL-7Rα were present in equal numbers after infection in wild-type and MHC class II–deficient mice, providing evidence that the formation of memory cell precursors does not require CD4+ T cell help. We then ‘dissected apart’ the stages of the CD8+ T cell response to determine precisely when CD4+ T cell help is important for sustained CD8 memory of previously encountered antigen. By generating effector or memory CD8+ T cells in either wild-type or MHC class II–deficient mice and adoptively transferring them into normal or CD4+ T cell–deficient secondary recipients, we found that CD4+ T cell help was not crucial in the generation of functional CD8 memory during the primary expansion phase (days 0–8 after infection), when naive cells are programmed and differentiate to become potent effectors. In the context of an acute infection, CD4+ T cells are crucial to the health and numbers of memory CD8+ T cells only during the memory maintenance phase.
Endogenous memory CD8+ T cells in CD4+ T cell–deficient but not wild-type mice decrease in number over time8. We recapitulated that finding here, tracking adoptively transferred CD8+ T cells specific for lymphocytic choriomeningitis virus (LCMV) glycoprotein amino acids 33–41 (GP(33–41); P14 T cell receptor–transgenic, Thy-1.1+) in wild-type or MHC class II–deficient mice after infection with recombinant Listeria monocytogenes expressing the LCMV GP(33–41) epitope (rLmGP) or after infection with LCMV. We tracked numbers of P14 cells by staining for Thy-1.1, MHC class I tetramer and intracellular interferon-γ (IFN-γ). After either bacterial (Fig. 1a) or viral (Fig. 1b) infection, P14 cell numbers expanded similarly in both hosts to generate large numbers of effectors, which then contracted. Memory CD8+ T cell numbers in the spleen remained almost constant in wild-type mice beyond day 21 after infection, but there was a continuous decrease in memory P14 cell numbers in MHC class II–deficient mice, resulting in a reduction of 85–90% in absolute numbers at 100 d or more after infection, compared with that of wild-type hosts. This decrease in memory cell numbers found in the spleens of MHC class II–deficient mice also occurred in the liver and lymph nodes (Supplementary Fig. 1 online). This concordance between different tissues is not unexpected, given that memory CD8+ T cells in central lymphoid organs and in nonlymphoid tissues are in rapid equilibrium19. Thus, our results do not indicate a specific loss in either ‘effector’ or ‘central’ memory pools in CD4+ T cell–deficient mice. We analyzed the surface phenotype and cytokine production of effector and memory cells in wild-type and MHC class II–deficient mice. Although effector cells in the two environments did not differ, we noted profound differences in CD62L and CD122 (but not CD44) expression on memory cells in wild-type versus MHC class II–deficient mice. We also found an overall reduction in IFN-γ and IL-2 production by the memory cell populations in the MHC class II–deficient mice (Supplementary Fig. 2 online). These findings suggest a diminished functional capacity of memory cells generated in MHC class II–deficient hosts. We obtained similar results in studies in which we adoptively transferred naive ovalbumin (OVA)–specific, OT-1 T cell receptor–transgenic CD8+ T cells into wild-type or MHC class II–deficient mice, followed by infection with recombinant L. monocytogenes expressing chicken OVA (data not shown). In all experiments, the infection was cleared in both wild-type and MHC class II–deficient mice 8 d after infection and was not detected at any subsequent memory time points (data not shown). These findings led us to explore at what stage during the response to an acute infection CD4+ T cells is crucial in sustaining memory CD8+ T cell numbers and functionality.
The subset of effector CD8+ T cells that express IL-7Rα (CD127) 8 d after LCMV infection is predisposed to become long-lived memory cells17. Over 90% of naive T cell receptor–transgenic CD8+ T cells have high expression of IL-7Rα (Fig. 2a). Because we found reduced memory cell numbers in a CD4+ T cell–deficient environment, we compared IL-7Rα expression on effector CD8+ T cells 8 d after LCMV infection in wild-type and MHC class II–deficient mice, with the idea that memory cell precursor generation might depend on the presence of CD4+ T cells. Equal percentages (about 15–20%) of IL-7Rαhi effector CD8+ T cells were present in the two sets of mice at day 8 after infection (Fig. 2b). The similar IL-7Rαhi memory cell precursor numbers at day 8, however, did not ‘translate’ into equal memory cell numbers at later stages in the different host environments (Fig. 1). We compared surface IL-7Rα expression on memory cells from the two environments at day 100 after infection and found that although almost all of the antigen-specific memory CD8+ T cells in wild-type mice expressed IL-7Rα, most of the remaining memory cells in MHC class II–deficient mice did not (Fig. 2b). Therefore, the loss of memory CD8+ T cells in MHC class II–deficient mice correlates with diminished IL-7Rα expression at later time points and not with expression of this receptor at the effector stage.
We sought to more precisely pinpoint the timing of the CD4+ T cell function in determining the fate of the memory CD8+ T cell population. We primed P14 cells in wild-type mice with LCMV and, on day 8 after infection, transferred equal numbers of purified effector cells into wild-type or MHC class II–deficient recipients and tracked donor CD8+ T cell numbers by Thy-1.1 staining in the spleen at 3, 30 and 60 d after transfer (Fig. 3). We analyzed samples at day 3 in this as well as subsequent experiments to ensure that the ‘take’ (engraftment) of donor cells was equivalent in all mice. As expected, effector CD8+ T cell numbers contracted after transfer into wild-type or MHC class II–deficient recipients. However, effector CD8+ T cell numbers underwent a substantial continuing contraction in MHC class II–deficient secondary recipients, resulting in 75–80% lower absolute memory cell numbers by day 60 after transfer compared with those of wild-type recipients (Fig. 3a,c). Similarly, effector CD8+ T cells generated in MHC class II–deficient primary hosts experienced a decrease in total numbers of greater than 80% after transfer into MHC class II–deficient compared with those of wild-type secondary recipients 60 d later (Fig. 3b,d).
Effector cell populations generated in wild-type or MHC class II–deficient primary hosts showed a similar contraction and maintenance of memory cell numbers when transferred into wild-type secondary recipients (Fig. 3a,b). Between days 30 and 60 after transfer, the absolute number of memory cells from both sources remained relatively stable in wild-type secondary hosts. Comparison of the two effector CD8+ T cell populations transferred into MHC class II–deficient recipients showed a similar continuous decrease in memory cell numbers at the later time points. Irrespective of the host environment during programming, both donor cell populations decreased in the MHC class II–deficient recipients. Experiments involving the adoptive transfer of OVA-specific effector CD8+ T cells primed with the recombinant L. monocytogenes expressing chicken OVA gave similar results (data not shown). These findings demonstrate that the presence of CD4+ T cells is important only after the period of expansion of the CD8+ T cell response.
We investigated whether the remaining memory CD8+ T cells in MHC class II–deficient recipients were functionally impaired compared with memory cells in wild-type recipients. We purified effector CD8+ T cells generated in wild-type or MHC class II–deficient primary hosts and transferred them into wild-type or MHC class II–deficient secondary recipients. Using intracellular cytokine staining, we measured the ability of the cells to rapidly produce both IFN-γ and IL-2 in response to peptide stimulation at 3 and 60 d after transfer. By comparing the mean fluorescence intensity of IFN-γ staining, we noted a decreased ability of the transferred memory CD8+ T cells in MHC class II–deficient host mice to produce IFN-γ at 60 d after transfer (Fig. 4a,b). This was accompanied by a 65% decrease in the fraction of memory cells that were capable of IL-2 production (Fig. 4c,d). In accordance with published findings9, memory CD8+ T cells maintained in CD4+ T cell–deficient environments are characterized by this overall decrease in cellular fitness.
To assess the ability of effector CD8+ T cells from wild-type mice to confer protective immunity 60 d after transfer to wild-type or MHC class II–deficient secondary recipients, we challenged recipient mice with a high dose of rLmGP and 3 d later measured bacterial clearance in spleen (Fig. 5a) and liver (Fig. 5b). As a control, we infected naive mice that did not receive transferred effector cells; these mice were unable to control growth of this high dose of listeria. Wild-type mice that had received effector cells 60 d before showed a high degree of protection against the challenge. In contrast, MHC class II–deficient recipients of effector cells showed only minimal protection against the challenge. These results provide further evidence that the maintenance of protective CD8+ T cell memory depends on the presence of CD4+ T cells.
We adoptively transferred memory CD8+ T cells from either wild-type or MHC class II–deficient primary hosts 35 d after infection into wild-type or MHC class II–deficient secondary recipients and tracked memory cell numbers by Thy-1.1 staining. In this set of experiments, we transferred memory cells that had undergone identical expansion and contraction phases into different secondary environments to study memory maintenance. We compared total donor CD8+ T cell numbers in the spleen at 3, 30 and 60 d after transfer (Fig. 6). Memory CD8+ T cells from wild-type primary hosts decreased in number in MHC class II–deficient secondary recipients compared with wild-type secondary recipients, resulting in almost 90% lower absolute numbers by day 60 after transfer (Fig. 6a,c). Similarly, memory CD8+ T cells generated in MHC class II–deficient primary hosts underwent a loss of 75–80% when transferred into MHC class II–deficient compared with that of wild-type secondary recipients (Fig. 6b,d). The unexpected increase in memory cell numbers in the wild-type secondary recipients over the course of the study can be attributed to the young age of the mice (4–5 weeks at the time of transfer), as total spleen cellularity more than doubled by day 60 after transfer in both wild-type and MHC class II–deficient mice (data not shown). These findings demonstrate that CD4+ T cells are crucial for sustaining the memory CD8+ T cell pool only after this stable, long-lived population is generated. In addition, the results in Figure 6b suggest that decreasing memory CD8+ T cells from MHC class II–deficient primary hosts at day 35 after infection can be rescued by transfer to a wild-type environment containing CD4+ T cells.
Even though a primary CD8+ T cell response may seem to be independent of CD4+ T cells, there is a requirement for CD4 help for the subsequent generation of stable, protective CD8 memory8–11,20,21. An important issue arising from those studies8–11,20,21 is when during an immune response CD4+ T cells provide signals that promote the survival of long-lived memory CD8+ T cells. Is CD4+ T cell help needed early during the primary response in a ‘covert’ way that is revealed only at later stages? Or are CD4+ T cells required at later stages to sustain memory CD8+ T cells? In this study, we have shown that during an acute infection, the generation of memory CD8+ T cell precursors is CD4+ T cell independent and have provided evidence that the presence of CD4+ T cells during memory maintenance is crucial to memory CD8+ T cell health and survival.
In the cytotoxic T lymphocyte (CTL) response to noninfectious antigens (such as minor histocompatibility antigens, tumor antigens or protein antigen), which are not nonspecifically inflammatory, CD4+ T cell recognition of antigen is important in activating or ‘licensing’ the antigen-presenting cell such that it can promote a CD8+ T cell response22–24. Whether all the functions provided by CD4+ T cells are mediated via the antigen-presenting cell or via direct CD4-CD8 T cell interactions is not known. Some studies have suggested that in response to noninflammatory antigen, the function of the CD4+ T cell becomes apparent only after the primary effector response; without CD4+ T cell help during the first 3–4 d, effector CD8+ T cells may be generated, but they are unable to mount productive recall responses in vitro11,20. In contrast, other studies have shown that after in vivo priming with noninflammatory antigens, CD4+ T cell help is essential for the stimulation of a measurable primary CD8+ T cell response25–27. The discrepancy between these results probably hinges on the different amounts of endogenous ‘danger’ signals produced by the immunization. Further work is needed to elucidate the contribution of CD4+ T cells to CD8 memory maintenance after priming with noninfectious antigens.
In the CTL response to virulent pathogens, such as L. monocytogenes and LCMV, the situation is very different. In contrast to the suggestion that direct crosstalk between CD4+ and CD8+ T cells via CD40 ligand and CD40 is necessary for the promotion of robust memory generation with noninflammatory antigens20, direct CD40–CD40 ligand interaction between CD4+ and CD8+ T cells is not involved during bacterial or viral infections26,28. In addition, our results presented here show that in these infections, CD4+ T cells do little to promote or program the CD8+ T cell response during the early phase. Effector or memory CD8+ T cells taken from control or CD4+ T cell–deficient mice acted similarly when adoptively transferred to new hosts. Thus, CD8+ T cells generated in a wild-type mouse decrease in number in a CD4+ T cell–deficient secondary host, and CD8+ T cells generated in a CD4+ T cell–deficient mouse recover and are maintained when transferred to a CD4+ T cell–sufficient secondary host. In all scenarios, the environment of the secondary host completely dictates the health and size of the memory CD8+ T cell pool.
Our findings indicate that the CD4+ T cell function in the maintenance of CD8+ T cell memory is not antigen specific simply because antigen is no longer present. Furthermore, because we purified CD8+ T cells before adoptive transfer into secondary recipients, antigen-specific memory CD4+ T cells or other adaptive immune cells were not present. This raises the question of mechanism. What are the noncognate signals CD4+ T cells provide that allow for memory CD8+ T cell health and survival? It is likely that cytokines, which promote survival and basal homeostatic proliferation of memory CD8+ T cells, are involved. CD4+ T cells either directly provide these factors to memory CD8+ T cells or act on an intermediate cell that provides memory CD8+ T cells with such signals.
Studies have focused on the functions of IL-7 and IL-15 in maintaining CD8 memory16,29,30. Because IL-7 is also required for the survival and basal homeostatic proliferation of naive CD8+ T cells31, and normal numbers of these cells are found in MHC class II–deficient mice, it is unlikely that defects in basal IL-7 production account for diminished memory CD8+ T cell numbers. This is in line with our findings that IL-7 mRNA expression in the spleen and lymph nodes of wild-type and MHC class II–deficient mice (measured by real-time PCR) were similar (data not shown). Furthermore, we found that naive CD8+ T cells adoptively transferred into MHC class II–deficient recipients did not decrease in number and were maintained for several months, as with wild-type recipients (data not shown). Studies have shown that IL-7Rαhi effector CD8+ T cells are predisposed to become long-lived memory cells, and as in our study here, those studies found that most memory CD8+ T cells express IL-7Rα in wild-type mice17,18. Here, although MHC class II–deficient mice and wild-type mice had similar numbers of effector cells that were IL-7Rαhi, IL-7Rα expression was not sustained in the absence of CD4+ T cells. How IL-7Rα expression is lost over time in the absence of CD4+ T cells and whether diminished IL-7Rα expression on memory cells is directly responsible for their lack of maintenance remain to be determined.
IL-15 has been linked to the generation and maintenance of memory CD8+ T cells32–39. Il15−/− and Il15ra−/− mice have deficiencies in memory CD8+ T cell numbers. The kinetics of the CD8+ T cell response after infection in these mutant mice seem very similar to those of the memory CD8+ T cell decrease we noted in MHC class II–deficient mice. Expression of the high affinity IL-15Rα on the CD8+ T cells themselves is dispensable for CD8 memory, although binding of IL-15 to the low-affinity receptor composed of the IL-15Rβ and the γc subunits on memory CD8+ T cells is required for their maintenance40,41. Expression of the high-affinity IL-15Rα on other (non-CD8) bone marrow–derived cells is essential for the basal homeostatic proliferation of memory CD8+ T cells40,42, possibly by binding IL-15 and ‘trans-presenting’ it to CD8+ T cells43. We found no difference in IL-15 mRNA expression in the spleens and lymph nodes of control mice and mice lacking CD4+ T cells (data not shown). Whether CD4+ T cells are responsible for producing a factor for memory CD8+ T cell maintenance or for acting on other cells to provide the cytokine signals required for sustaining memory CD8+ T cells remains to be determined. Elucidation of the function of CD4+ T cells during the memory maintenance phase of the CD8+ T cell response will aid in the design of vaccines, which rely heavily on robust secondary CD8+ T cell responses to pathogens, and will also provide insight into the loss of immune function after depletion of CD4+ T cells in certain disease states.
Age-matched C57BL/6 and MHC class II–deficient (C57BL/6-Abβ−/−) mice were purchased from Taconic. A colony of Thy-1.1+ P14 transgenic mice was maintained at our specific pathogen–free animal facility at the University of Washington (Seattle, Washington). Experiments began when mice were approximately 6–12 weeks of age; all experiments were done according to institutional ethical guidelines.
Single-cell suspensions were made from spleen of P14 mice, and 5 × 104 gp33-specific cells were injected intravenously into C57BL/6 or MHC class II–deficient mice before infection. At day 8 or day 35 after infection, CD8+ T cells from spleens of mice were purified with a CD8+ T cell Isolation kit (Miltenyi Biotec) followed by AutoMACS magnetic bead separation. Approximately 5 × 106 to 7 × 106 P14 effector cells and 1 × 106 to 2 × 106 P14 memory cells were injected intravenously into recipient mice. At 3, 30 or 60 d after transfer, total numbers of P14 cells in the spleens of recipient mice were determined by Thy-1.1 staining.
The rLmGP used was provided by H. Shen (University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania)44. Frozen stocks of the rLmGP were grown in brain-heart infusion broth. Bacteria culture samples were grown to mid-log phase, measured by optical density (absorbance at 600 nm) and diluted in PBS for injection. All mice were infected by intravenous injection of the tail vein with a priming dose of 2,000 rLmGP and challenge doses equivalent to 1 × 105 to 2 × 105. Injected bacteria numbers were determined by spreading of bacterial samples on brain-heart infusion plates followed by incubation overnight at 37 °C. LCMV Armstrong 53b was grown on BHK cells and was titered on Vero cells. Mice were infected intraperitoneally with 2 × 105 plaque-forming units of virus.
For determination of the colony-forming units per spleen or liver in the infected mouse, the entire organ was dissociated in PBS with 0.1% Nonidet-P40 (Sigma). Tenfold serial dilutions of spleen or liver suspensions were made in PBS plus 0.1% Nonidet-P40, and each dilution was spread on brain-heart infusion plates. Plates were counted after incubation overnight at 37 °C, and total colony-forming units per organ were determined.
H-2Db tetramers bound to LCMV GP(33–41) were generated as described45,46. Splenocytes (1 × 106) were stained with DbGP33 tetramer or antibody to Thy-1.1 (anti-Thy-1.1; BD PharMingen) along with anti-CD8 (PharMingen), anti-CD44 (PharMingen), anti-CD62L (PharMingen), anti-CD122 (PharMingen), or anti-CD127 (eBioscience). Flow cytometry was done on a FACSCalibur and data were analyzed with CELLQuest software (Becton Dickinson).
Intracellular cytokine staining was done with a kit (PharMingen), following the manufacturer's instructions. In 96-well plates, 2 × 106 splenocytes per well were stimulated for 5 h with media alone or with 10 nM GP(33–41) peptide in the presence of 1 μg/ml of Brefeldin A. Cells were then washed, stained with anti-CD8, resuspended in permeabilization-fixation buffer and stained with anti-IFN-γ and anti-IL-2 (PharMingen). Labeled cells were washed in permeabilization buffer, resuspended in fixation buffer and analyzed on a FACSCalibur.
Supported by the National Institutes of Health (AI19335, AI056809 and CA09537) and the Howard Hughes Medical Institute.