Tissue-Level Regulation of Th1 and Th2 Primary and Memory CD4 T Cells in Response to Listeria Infection

Amanda L. Marzo, Vaiva Vezys, Kristina Williams, David F. Tough and Leo Lefrançois
J Immunol May 1, 2002, 168 (9) 4504-4510; DOI: https://doi.org/10.4049/jimmunol.168.9.4504

Abstract

Ag-specific Th1 and Th2 cytokine-producing CD4 T cells were quantitated in secondary lymphoid and tertiary tissues following oral Listeria monocytogenes infection. Although the response to Listeria was previously believed to be predominately Th1 like, CD4 T cells producing IL-4 or IL-5 comprised a substantial proportion of the overall primary and memory response. The frequency of IFN-γ-, IL-4-, or IL-5-producing primary effector or memory CD4 T cells was significantly higher in lung, liver, and intestinal lamina propria (LP) as compared with spleen and lymph node. However, maximum numbers of IL-4- and IL-5-producing cells were detected in the LP several days after the peak of the Th1 response, and IL-5 production was skewed toward the mucosal tissues. Remarkably, the recall response resulted in sustained Th1 and Th2 responses in tertiary, but not lymphoid tissues and long-term retention of Th1 and Th2 memory cells in equal proportions in the LP. Finally, CD40 ligand was essential for induction of IFN-γ in the spleen and LP, but not in the liver and lung, while the IL-4 response required CD40 ligand only in the spleen. Therefore, the rules governing the effector phenotype, and the overall magnitude of the CD4 response, are regulated at the level of individual tissues.

Upon Ag contact, naive CD4 T cells differentiate into two main subsets, Th1 and Th2, that direct different types of immune responses. These subsets can be distinguished by the secretion pattern of effector cytokines (1) and their expression of chemokine receptors (2). In general, Th1 cells produce IL-2, IFN-γ, and lymphotoxin-β, which have been shown in some systems to be essential in maintaining CD8 T cell effector functions (3). Th2 cells in contrast secrete cytokines such as IL-4, IL-5, and IL-10, which induce class switching to IgE and IgG1 (1, 4). There are likely to be other Th subsets, including Th0 cells, which are considered to be Th1 and Th2 precursors, and produce both Th1- and Th2-type cytokines (5). The balance between these types of Th cells can influence the nature, strength, and duration of systemic immune responses (6).

Although the mechanisms by which CD4 T cells provide help to naive CD8 T cells are not fully understood, several studies have demonstrated that CD40-CD40 ligand (CD40L)3 interactions between dendritic cells (DC) and CD4 T cells activate the DC for effective priming of CD8 T cells (7, 8, 9). Interference with this pathway inhibits formation of germinal centers, Ab isotype switching, production of proinflammatory cytokines, NO, and IL-12, and CTL generation (10, 11). Thus, mice deficient in CD40 or CD40L are impaired in humoral and cell-mediated responses, although this is not absolute (12, 13, 14). Current models suggest that CD40L is induced mainly on CD4 T cells shortly after activation, and that engagement of CD40 expressed by APCs induces up-regulation of costimulatory molecules and production of proinflammatory cytokines (10). CD40 stimulation of DC induces IL-12 production, explaining the ability of this cell type to promote Th1 development (15, 16, 17), although other studies suggest CD40 may also regulate Th2 development (18). CD40L expressed by CD8 T cells also participates in immune responses. For example, optimal activation of CD8 T cells in the intestinal lamina propria (LP) requires CD40L expressed by CD8 T cells (19). Furthermore, the primary CD8 response to Listeria infection also requires CD40L/CD40 interactions, especially in the intestinal mucosa (20). Thus, costimulation and, potentially, CD4 T cell help can be mediated at the tissue level as well as the cellular level.

Recent studies have analyzed in detail the Ag-specific CD4 T cell response to virus infection. In general, the CD4 response to virus infection is numerically ∼10- to 100-fold less than that of the CD8 response. Nevertheless, in the case of lymphocytic choriomeningitis virus (LCMV) infection, long-lived splenic CD4 T cell memory is induced, and although the primary and memory splenic responses are quantitatively skewed toward CD4 T cells producing IFN-γ, IL-4-producing cells are also detected throughout the response (21). Although most studies have examined responses in secondary lymphoid tissues, it has become increasingly clear that CD8 and CD4 T cell responses in nonlymphoid tissues constitute a large portion of the overall response (22, 23, 24). It should also be noted that adoptive transfer studies using in vitro activated CD4 T cells suggest that Th1, but not Th2 cells are able to migrate to tertiary tissues (25, 26). However, infection with Nippostrongylus induces a strong Th2 response in the lung as well as lymphoid tissues (27). In comparison with CD4 responses to viral and parasitic infections (28), relatively little is known with regard to Ag-specific antibacterial CD4 T cell responses. One well-characterized model system used for analyzing cellular immunity is infection with Listeria monocytogenes (LM). Infection of mice with this pathogen induces a strong T cell response that is essential for bacterial clearance from the host. The hallmark of LM infection is a cell-mediated immune response to secreted virulence factors inducing both CD8 and CD4 T cell activation (29, 30, 31, 32, 33, 34, 35). Studies with MHC-deficient mice reveal that both CD4 and CD8 T cells can contribute to protection against listeriosis (36). Although LM has been suggested to induce CD4 T cells that are restricted to the Th1 phenotype (37, 38) and has been proposed as a vaccine vector for immunity induction against HIV and tumors (39, 40), relatively little is known about the generation and function of LM-reactive CD4 T cells in vivo.

In this study, we report the characterization of the Ag-specific CD4 T cell response in secondary lymphoid and tertiary tissues following oral Listeria infection. The results showed that the nonlymphoid memory response outweighed that of the lymphoid response, and included Th1- and Th2-type cells. Furthermore, there were differential activation requirements for Th1 and Th2 CD4 T cells in different tissues.

Materials and Methods

Mice

C57BL/6J and C57BL/6-CD40L−/− mice (41) were purchased from The Jackson Laboratory (Bar Harbor, ME).

Infection with rLM-OVA

A strain of rLM-secreting OVA (rLM-OVA) was produced, as previously described, using a truncated OVA cDNA (20, 39, 42). Mice were infected by gavage with either ∼1 × 109 CFU rLM-OVA for primary infections or ∼1 × 106 CFU for initial priming, followed by infection with ∼1–5 × 109 CFU to induce a recall response. Actual CFU used for infections were calculated by plating dilutions of the inoculum. Bacterial titers within tissues were determined by homogenizing the tissue in PBS containing 1% saponin, plating serial dilutions of homogenate on brain-heart infusion agar plates containing 5 μg/ml erythromycin, and growing for 2 days at 37°C.

Isolation of lymphocyte populations

Single cell suspensions were prepared from lymph nodes and spleens. Lymphocytes were isolated from small intestine LP and liver, as previously described (23, 43). To obtain lymphocytes from lungs, anesthetized mice were perfused with PBS containing 75 U/ml heparin until the lungs were cleared of blood and white in color. The lungs were removed and cut into small pieces and stirred at 37°C for 30 min in HBSS containing 1.3 mM EDTA. Cells were washed in PBS containing 5% FCS twice. Lymphocytes were released from the tissue by digestion with 150 U/ml collagenase (Life Technologies, Rockville, MD) in RPMI containing 1 mM MgCl2, 1 mM CaCl2, and 5% FCS, at 37°C for 30 min. Released cells were pooled and then mashed through a cell strainer (BD Biosciences, Franklin Lakes, NJ). Cells were washed and resuspended in RPMI 1640 medium supplemented with 10% FCS at 1 × 107/ml.

Immunofluorescence analysis

At the indicated times after infection, lymphocytes were isolated, and OVA-specific CD8 T cells were detected using an H-2Kb tetramer containing the OVA-derived peptide SIINFEKL, produced as previously described (44, 45). For staining, lymphocytes were suspended in PBS/0.2% BSA/0.1% NaN3 (PBS/BSA/NaN3) at a concentration of 1 × 106–107 cells/ml, followed by incubation at room temperature for 1 h with OVA-tetramer-allophycocyanin plus the appropriate dilution of anti-CD8-PE (clone 53.6.7; BD PharMingen, San Diego, CA). Cells were washed with PBS/BSA/NaN3, stained with FITC-conjugated anti-CD11a and PerCP-conjugated anti-CD4 (clone RM4-5; BD PharMingen), incubated at 4°C for 20 min, washed, and fixed in 3% paraformaldehyde in PBS. Relative fluorescence intensities were measured with a FACSCalibur (BD Biosciences). Data were analyzed using WinMDI software (J. Trotter, The Scripps Clinic, La Jolla, CA).

ELISPOT assay

Cells secreting IFN-γ, IL-4, and IL-5 in an Ag-specific manner were detected using a standard ELISPOT assay (46, 47). Plates (Multiscreen HA plates; Millipore, Bedford, MA) were coated overnight at 4°C with the cytokine-specific capture Abs specified below, washed with PBS, then incubated with 200 μl RPMI 1640 medium supplemented with 10% FCS for at least 1 h at room temperature. Two-fold dilutions of cells from animals previously inoculated orally with rLM-OVA were added to wells starting at 106–2 × 105 cells/well in the presence of 4 × 105 gamma-irradiated (1200 rad) syngeneic spleen cells. Cells were incubated for 26 h with or without the OVA-derived peptide TEWTSSNVMEERKIKV (OVA265–280) (48) or the listeriolysin O (LLO)-derived peptide NEKYAQAYPNVS (LLO190–201) (49). Peptides were obtained from Research Genetics (Huntsville, AL). Wells were sequentially washed three times each with ddH20, PBS, and PBS containing 0.05% Tween 20, and then incubated for 20 h at 4°C with biotinylated anti-cytokine Abs. Wells were washed and incubated with peroxidase-labeled anti-biotin Ab (2 μg/ml; Vector Laboratories, Burlingame, CA) for 20 h at 4°C. Wells were then washed and spots were developed using freshly prepared substrate: 0.3 mg/ml 3-amino-9-ethyl-carbazole (Sigma-Aldrich, St. Louis, MO), 10% dimethylformamide, 0.015% H2O2 in 0.1 M sodium acetate (pH 4.8). After 30 min, the substrate solution was discarded, and plates were washed under running water and air dried. The following combinations of capture and detection mAbs were used for IFN-γ, IL-4, and IL-5, respectively: R46A2 (10 μg/ml) and XMG1.2-biotin (4 μg/ml), TRFK5 (4 μg/ml) and TRFK4-biotin (2 μg/ml), and BVD4-1D11 (2 μg/ml) and BVD4-24G2-biotin (2 μg/ml) (all Abs were from BD PharMingen). Image analysis of colored spots was performed on an Immunospot Image Analyzer (Cellular Technology, Cleveland, OH) specifically designed for ELISPOT analysis. In all experiments, the background spots (detected using effector cells plus feeders without peptide) were subtracted from wells where spots were generated with the addition of peptide. Statistics were performed using Student’s t test.

Results

Characterization of the primary anti-Listeria CD4 T cell response after oral infection

We undertook a comparison of the CD8 and CD4 T cell responses in tertiary vs lymphoid tissues after oral infection with rLM-OVA. Consistent with our previous findings (20), the CD8 response, as detected using an MHC class I tetramer, peaked at day 9 postinfection, and a substantial population of OVA-specific CD8 T cells was found in all tissues. The liver, lung, and intestinal LP contained a larger population of tetramer+ cells, and the response in these tissues was prolonged as compared with that of secondary lymphoid organs (Fig. 1⇓A). To determine whether the CD4 response followed similar kinetics, Ag-specific CD4 T cells were quantitated from the same tissues using ELISPOT after stimulation with an OVA-derived peptide presented by I-Ab (48). Nine days after oral infection with rLM-OVA (the peak of the response), all tissues contained substantial populations of IFN-γ- and IL-4-producing OVA-specific cells (Fig. 1⇓, B and C). The liver, lung, and LP contained larger populations of IFN-γ-producing CD4 T cells (0.3, 0.4, and 0.4%, respectively) compared with the spleen (0.1%) or mesenteric lymph node (MLN; 0.05%) at the peak of the response (Fig. 1⇓B). Interestingly, at the same time point, the frequency of CD4 T cells producing IL-4 in the LP (0.13%) was greater than in the spleen (0.08%), liver (0.09%), and lung (0.08%). Even more intriguing was the finding that as the IFN-γ response declined, the IL-4 response was sustained or increased in the LP up to 15 days postinfection and did not fall to memory levels until ∼day 35.

FIGURE 1.
FIGURE 1.

High frequencies of Ag-specific T cells are detected in nonlymphoid tissues following oral LM infection. Mice were infected with rLM-OVA by gavage, and lymphocytes were isolated from the indicated organs at the indicated days postinfection. A, Lymphocytes were stained with an allophycocyanin-labeled H-2Kb-SIINFEKL tetramer, anti-CD8, anti-CD4, and anti-CD11a, and analyzed by flow cytometry. Ag-specific CD4 lymphocytes were quantitated by ELISPOT assay to detect cells producing IFN-γ (B) or IL-4 (C). Values represent the means ± SE from four mice per time point.

The total cell numbers producing each cytokine were also determined (Fig. 2⇓). When total numbers of CD4 T cells producing IFN-γ were calculated, the cumulative sum of the cell numbers in the LP, lung, and liver was similar to that in the spleen at the peak of the response (Fig. 2⇓). However, the total number of CD4 T cells producing IL-4 in the spleen was greater than that in the tertiary tissues 9 days postinfection. By day 15, a far greater number of IL-4 producers was present in the tertiary tissues as compared with the spleen, where the numbers had declined to memory levels. This delayed IL-4 increase was attributed to the appearance of significant numbers of IL-4-producing cells in the intestinal LP at day 15. Whether this reflects increased migration of cells to the LP or continued expansion in the LP is unknown. Irrespective of this increase, by 3–5 wk after infection, the total numbers of memory cells producing each cytokine were similar between the tissues and the spleen.

FIGURE 2.
FIGURE 2.

Ag-specific CD4 T cells in nonlymphoid tissues comprise a large proportion of the response to LM infection. The total number of IFN-γ- and IL-4-producing CD4 T cells per tissue was determined by multiplying the number of cytokine-producing CD4 T cells by the total number of lymphocytes isolated from that tissue. Values for “tissues” in A are the total number of either IFN-γ- or IL-4-producing CD4 T cells in the LP, lung, and liver. Values represent means ± SE.

To determine whether the unique kinetics of IL-4 production in the LP was characteristic of the induction of other Th2 cytokines, Ag-specific CD4 T cells producing IL-5 were quantitated in a separate experiment along with IFN-γ-producing cells. Nine days after infection, greater percentages of IFN-γ-producing CD4 T cells were again detected in the nonlymphoid tissues (Fig. 3⇓). By day 15 postinfection, the IFN-γ response had declined in all tissues, although less so in the LP. In the case of IL-5, the largest percentage of IL-5-producing cells was present in the LP at day 9, but, as with IL-4, this value had increased significantly by 15 days after infection, and this was borne out when total cell numbers were evaluated (data not shown). Unlike IL-4 producers, IL-5 producers had also increased in the lung from 9 to 15 days postinfection. In the MLN, spleen, and liver, fewer IL-5-producing cells were detected on day 9, and these numbers had declined by day 15. These data suggested that Listeria-specific Th2 CD4 T cells either preferentially migrated to mucosal sites after primary activation or that differentiation and expansion of Th2 cells occurred in those sites.

FIGURE 3.
FIGURE 3.

Preferential localization and distinct kinetics of IL-5-producing cells in mucosal tissues. Mice were infected with rLM-OVA by gavage, and the frequency of OVA-specific CD4 lymphocytes producing IFN-γ (A) or IL-5 (B) was determined by ELISPOT using lymphocytes from the indicated tissues at 9 or 15 days postinfection. Values represent the means ± SE from four mice per time point.

Preferential localization of CD4 memory cells in tertiary tissues

To determine the localization pattern of CD4 memory cells generated in response to Listeria infection, cytokine-producing OVA-specific CD4 T cells were quantitated 42 days after inoculation (Fig. 4⇓). Interestingly, the frequency of memory cells present in the tertiary tissues as a whole was ∼10-fold greater than that found in the spleen and MLN, implying a preferential retention and/or migration of memory cells in nonlymphoid tissues (Fig. 4⇓, A and B). In the LP and the lung, roughly equal frequencies of IL-4-, IFN-γ-, and IL-5-producing CD4 T cells were detected. Fewer IFN-γ- and IL-4-producing cells were found in the liver, while, as in the primary response, IL-5 producers were barely detectable (Fig. 4⇓A). Analysis of total numbers of cytokine-producing memory cells indicated that the MLN contained very few memory cells, while the spleen contained numbers of memory cells comparable with those of individual nonlymphoid tissues (Fig. 4⇓C). As was observed in the primary response, very few IL-5 producers were detected in the liver (Fig. 4⇓C). Overall, the cumulative sum of Ag-specific memory cells in the tertiary tissues was greater than that present in the secondary lymphoid tissues (Fig. 4⇓D).

FIGURE 4.
FIGURE 4.

The majority of Listeria-specific Th1 and Th2 CD4 memory cells are localized in nonlymphoid tissues. Lymphocytes were isolated from the indicated tissues 42 days postinfection, and ELISPOT assays for detection of cells producing IFN-γ, IL-4, or IL-5 were performed (A and B). Total numbers of cytokine-producing cells from the different tissues were also assessed (C), and the cumulative total cell number from lymphoid (MLN and spleen) vs tertiary tissues (LP, lung, and liver) was determined (D). ∗, Significance of p < 0.002.

Recently, a natural Listeria epitope derived from the LLO protein and presented by I-Ab was described (LLO190–201) (49). Because it was possible that the OVA-specific response was distinct from that directed toward natural epitopes, we quantitated CD4 T cells specific for LLO190–201 after infection with rLM-OVA (Fig. 5⇓). The overall response to the LLO epitope was substantially larger than the OVA-directed response (Fig. 5⇓A, day 9). Nevertheless, the general pattern of cytokine-producing cells generated was qualitatively similar to that observed in the OVA-specific response. Thus, at day 9 after infection, a greater percentage of IFN-γ-producing cells was detected in the LP and the lung (although not liver) than in the spleen and MLN. Th2 cells were again much more prominent in the mucosal tissues at the peak of the response, and this was particularly true for IL-5-producing cells in the lung and LP. Analysis of memory CD4 T cells 108 days postinfection also reiterated the finding that substantially greater frequencies of Ag-specific CD4 T cells producing either of the three cytokines resided in the tertiary tissues compared with secondary lymphoid tissues. Interestingly, in contrast to the IFN-γ response, which had declined ∼10-fold in all tissues by this time, the Th2 response, as a percentage of CD4 T cells, had declined only 2- to 3-fold in the nonlymphoid tissues. These results indicated that, for the most part, the character of the anti-OVA and anti-LLO CD4 responses was qualitatively similar.

FIGURE 5.
FIGURE 5.

LM infection induces a substantial lymphoid and nonlymphoid primary and memory response to a natural listeriolysin-derived epitope. Mice were infected with rLM-OVA by gavage, and lymphocytes were isolated from the indicated organs 9 (left) and 108 (right) days postinfection. ELISPOT assays using the LLO peptide (aa 190–201) were used to determine the frequency of CD4 T cells producing IFN-γ (A), IL-4 (B), and IL-5 (C). Values represent the means ± SE from four mice per time point.

Sustained LP CD4 recall response after oral reinfection

To examine the stability of the Listeria-specific CD4 T cell response following a secondary infection, mice were infected orally with 106 rLM-OVA, and at least 125 days later were orally challenged with 5 × 109 bacteria and the OVA-specific response was measured. The small inoculum used in the primary infection was necessary because oral infection with high doses of Listeria resulted in significant protection against oral reinfection, making it difficult to induce a detectable T cell recall response (data not shown). By day 6 after reinfection, a substantial number of CD4 T cells producing IFN-γ and IL-4 was present in all tissues (Fig. 6⇓), indicating that a secondary, not a primary, response was being detected because few Ag-specific CD4 T cells can be detected at this time point during a primary infection. The Th1 response in all tissues peaked at day 6, with the greatest frequency of IFN-γ-producing CD4 T cells present in the LP (0.6%). While the response in the MLN, spleen, and lung declined to memory levels by day 20, the response in the liver and LP was prolonged at least an additional 2 wk (Fig. 6⇓A). In fact, by day 60, the percentage of IFN-γ producers in the LP was essentially unchanged from day 20, indicating the potential utility of oral boosting for increasing intestinal protection. The peak of IL-4-producing CD4 T cells after secondary infection was day 6 in the MLN, spleen, and lung (0.06, 0.16, and 0.22%, respectively), while in the LP and liver the response peaked at day 20 postinfection (0.32 and 0.35%, respectively) (Fig. 6⇓B). At 60 days postinfection, the frequency of IFN-γ- and IL-4-producing CD4 T cells in the lung and liver was <0.05%, and in the MLN and spleen was <0.02% of CD4 T cells. As was observed with the recall IFN-γ response, the frequency of IL-4-producing CD4 T cells in the LP remained unchanged at ∼0.3% of CD4 T cells from day 20 onward, suggesting that the LP is an important reservoir for LM-specific CD4 memory T cells.

FIGURE 6.
FIGURE 6.

Secondary infection with rLM-OVA induces a sustained Th1 and Th2 recall response in the intestinal mucosa. Mice were initially infected with 3 × 106 rLM-OVA by gavage, then infected with 5 × 109 bacteria by gavage 125 days later. Lymphocytes were isolated from the various tissues at the indicated day after secondary infection and set up in an ELISPOT assay for IFN-γ (A) and IL-4 (B). Values represent means ± SE of values from four mice per time point.

Differential requirement for CD40L in generating primary anti-Listeria CD4 T cell responses

CD40/CD40L interactions are critical for activation of CD4 T cells and play a role in CD8 T cell activation in the intestinal LP (19). Because the anti-Listeria CD4 T cell response was differentially skewed in various tissues, we tested whether CD40L was involved in this regulation by analyzing the response in infected CD40L-deficient mice (Fig. 7⇓). The Listeria-specific primary IFN-γ-producing CD4 T cell response in the spleen was severely reduced in CD40L−/− mice compared with the response in CD40L+/+ mice (Fig. 7⇓, A and C), while the splenic Th2 response was inhibited, but not as dramatically (Fig. 7⇓, B and D). However, although the LP Th1 response was inhibited >80% in CD40L−/− mice, the LP Th2 response was unaffected. In striking contrast to the LP IFN-γ response, CD40L was not required for the generation of Th1 CD4 T cells in the lung or liver, nor was CD40L needed for the induction of Th2 CD4 T cells in these tissues (Fig. 7⇓). Indeed, the Th1 and Th2 responses in the lung were significantly increased in the absence of CD40L, perhaps indicating a compensatory mechanism for decreased responses elsewhere or a redistribution of activated cells.

FIGURE 7.
FIGURE 7.

Differential tissue requirements for CD40-CD40L interactions in the induction of LM-specific CD4 T cells. Nine days after infection of wild-type (B6) or CD40L−/− mice, lymphocytes from the indicated tissues were analyzed by ELISPOT for OVA-specific CD4 T cells producing IFN-γ (A) or IL-4 (B). Total number of IFN-γ (C)- and IL-4 (D)-producing cells from the different tissues was also assessed. Values represent the means and SE from six mice per group and are representative of results from two different experiments. ∗, Significance of p < 0.005.

Another possible explanation for the observed increase in Ag-specific CD4 T cells in the lungs of infected CD40L−/− mice was that the infection was not effectively resolved. To test this possibility, we quantitated the number of viable bacteria in the MLN, spleen, lung, liver, and intestine 4 and 9 days postinfection in control and CD40L−/− mice (Fig. 8⇓). Four days after infection, similar numbers of bacteria were present in all tissues of control and CD40L−/− mice. In the lungs of CD40L−/− mice, ∼50% fewer bacteria were found as compared with controls, perhaps as a result of the heightened response in that tissue. At 9 days after infection, viable bacteria were either not detected or were present in very small numbers in either control or CD40L−/− mice. These results demonstrated that CD40L was not required for bacterial clearance and that IFN-γ responses in spleen and LP were also not critical for resolution of infection in those tissues.

FIGURE 8.
FIGURE 8.

Lack of CD40L does not affect clearance of LM. The number of bacteria in the MLN, spleen, lung, liver, and intestine was determined 4 and 9 days postinfection. Listeria colonies were enumerated after serial dilution of samples and plating on brain-heart infusion agar plates containing erythromycin. Data shown are the mean and SE of three animals per time point and are representative of results from two independent experiments.

Discussion

Kinetic analysis of the CD4 T cell response to LM infection revealed that a major proportion of the Listeria-specific CD4 T cells was present in the tertiary tissues as primary effector and long-term memory cells. These results are in agreement with those of Reinhardt et al. (24), in which adoptively transferred OVA-specific D011.10 cells migrated to many tissues after immunization with soluble Ag and LPS. However, cytokine production was not assessed in each tissue in that study. Our results showed that the kinetics and type of cytokine production were often distinct in different tissues. Thus, the response in the LP was prolonged and the kinetics of the Th2 response in the LP was distinct from that seen in other tertiary or lymphoid tissues (Figs. 1⇑ and 2⇑). Furthermore, in the primary as well as the memory response there was a much greater frequency of IFN-γ-, IL-4-, and IL-5-producing CD4 T cells in the LP, lung, and liver as compared with the MLN and spleen. There are several potential nonmutually exclusive reasons for the preferential localization of effector and memory CD4 T cells in nonlymphoid tissues as compared with secondary lymphoid tissues. The difference may reflect enhanced survival in the nonlymphoid tissues, preferential homing to those sites, selective retention of Ag-specific CD4 T cells, or greater proliferation in the tertiary tissues, particularly in the primary response. Homann et al. (21) recently reported that, while LCMV-specific CD8 T cell memory was stably maintained for life, the numbers of Ag-specific CD4 memory T cells gradually declined in the spleen. However, other reports do not fully support this finding (50, 51). Although we have not yet examined greatly extended time periods, our findings suggested that while CD4 memory T cells may decline in the lymphoid tissues, significant numbers of memory cells remain in tertiary tissues. Such a scenario would imply that the lymphoid pool of memory cells is regulated separately from the tertiary tissue memory pool. Further in-depth migration studies will be needed to address this important question. The substantial population of responding extralymphoid CD4, as well as CD8, T cells should be taken into account when analyzing immune responses.

Our data also demonstrated the capacity to induce both Th1- and Th2-type Listeria-specific effector and memory cells, which was heretofore unappreciated during LM infection. The conclusion that IL-4 was not produced by conventional T cells in LM infection is derived from studies using i.v. or i.p. routes of infection, followed by restimulation of unseparated spleen cells in vitro and measurement of cytokines by ELISA (38, 52), or from PCR analysis of whole tissue lysates (53). Restimulation was achieved using either heat-killed LM (52) or LM culture supernatants (38). IFN-γ, but not IL-4, was readily detected in these systems. It is not clear in these experiments whether sufficient quantities of the relevant antigenic epitopes are available after pulsing APC in vitro with whole killed organisms or culture supernatants. The timing or sensitivity of the assays used or perhaps the use of IL-4 in the cultures may also influence the results. Nonetheless, as we demonstrated in this study, similar numbers of IL-4- and IFN-γ-producing cells are found in the spleen at the peak of the primary response. The relevant quantity of a given cytokine in an immune response in vivo is difficult to determine, but, in any case, the ELISPOT assay using defined MHC class II-presented peptides for restimulation allowed us to quantitate the number of IL-4- and IL-5-producing cells in a number of tissues. The IL-4 producers were conventional MHC class II-restricted CD4 T cells because when CD4 T cells were depleted from lymphocytes before plating or in the absence of peptide or after infection of MHC class II−/− mice, few IL-4-producing cells were detected at the time points tested (data not shown). In contrast, the very early response to LM infection is characterized by rapid production of IL-4 by CD1-restricted NK-T cells in the liver (52, 54). Therefore, it is possible that early IL-4 production plays a role in driving a subset of Ag-specific CD4 T cells toward the Th2 phenotype during the priming phase (55). However, recent work also suggests that early production of IL-4 in an immune response in vivo may direct the response down a Th1 pathway (56). Thus, given the complexity of the innate and adaptive immune components involved in LM infection, it is perhaps not surprising that the CD4 T cell response does not strictly adhere to the conventional Th1/Th2 paradigm. Similarly, following acute infection with LCMV, a mixed CD4 T cell primary and memory response is induced in the lymphoid tissues (50).

While it is clear that IFN-γ plays an important role in protection against LM infection (56), the role of IL-4 or IL-5 derived from conventional T cells has not been determined. Blocking of IL-4 during LM infection results in increased resistance (57), but the source and timing of production of the relevant IL-4 being inhibited are unknown (e.g., IL-4 production by NK/NK-T cells, conventional CD4 T cells, or nonhemopoietic cells). The substantial numbers of IL-4- and IL-5-producing effector and memory cells in the LP and lung could be involved in regulation of Ab production in those tissues or in LN draining those sites. The localization of Th1- and Th2-type memory cells in the nonlymphoid tissues may also be important for a rapid recall response in situ. Indeed, oral reinfection resulted in a sustained increase in memory cells in the LP, but not in the spleen or LN (Fig. 6⇑). The LM-specific primary and recall CD8 response is also sustained in the LP (20). Thus, CD4 T cells in the LM response may also be important for magnification and modulation of the CD8 T cell response in nonlymphoid tissues. This hypothesis is supported by previous studies showing that CD4 and CD8 T cells cooperate in mounting a protective anti-LM immune response (58).

Tissue-level control of Th1/Th2 skewing was exemplified by the requirement for CD40L in differentially regulating the response in different tissues. Remarkably, while the overall splenic CD4 T cell response was inhibited in the absence of CD40L, the induction of IFN-γ producers, and not IL-4 producers, was blocked in the intestinal LP without CD40L. Moreover, both the Th1 and Th2 responses in the lung were in fact augmented in CD40L−/− mice, while the liver and MLN responses were unaffected. In comparison, we previously showed that the splenic anti-LM CD8 T cell response was largely unaffected, while the LP CD8 response was inhibited in the absence of CD40/CD40L interactions (20). Overall, these results demonstrated a conspicuous tissue and cell type-specific regulation and implied that costimulation via CD40L was differentially used depending on the tissue. The influence of CD40/CD40L interactions on helper differentiation is somewhat controversial, although it has been suggested that the CD40 pathway is more important for Th1 development (59). Activation in vitro of CD40L−/− TCR transgenic CD4 T cells results in decreased IFN-γ production, but, interestingly, an increase in IL-4 production, in accordance with what we observed in the lung (60). Consistent with our findings in the spleen, both Th1 and Th2 LCMV-specific splenic responses in CD40L−/− mice were reduced (61). CD40/CD40L interactions are also critical in protection against Leishmania major infection via up-regulation of IL-12 and IFN-γ production (12, 13, 14). In contrast, CD40/CD40L interactions are not essential for Th1-mediated protection against infection with the intracellular pathogens Histoplasma (62) or Mycobacterium (63). Similarly, while IFN-γ is involved in protection against LM infection, the inhibition of IFN-γ in spleen and LP, which we observed in CD40L−/− mice, did not affect clearance of the bacteria. It is interesting to speculate that sufficient levels of IFN-γ were produced in other tissues to afford protection or that other mechanisms can come into play in the absence of IFN-γ.

In summary, the data presented bolster the hypothesis that T cell responses can be compartmentalized at the tissue level, even during a systemic infection. By the same token, the distinct response type in a given tissue is likely to be important in mediating or regulating immunity at that site.

Footnotes

  • ↵1 This work was supported by National Institutes of Health Grants AI41576 and DK45260 (to L.L.), a collaborative grant from the Edward Jenner Institute for Vaccine Research (to D.F.T. and L.L.), C. J. Martin Fellowship 007151 (to A.L.M.), and U.S. Public Health Service Training Grant T32-AI07080 (to V.V.).

  • ↵2 Address correspondence and reprint requests to Dr. Leo Lefrançois, Department of Medicine, University of Connecticut Health Center, M/C 1319, 263 Farmington Avenue, Farmington, CT 06030. E-mail address: llefranc{at}neuron.uchc.edu

  • ↵3 Abbreviations used in this paper: CD40L, CD40 ligand; DC, dendritic cell; LCMV, lymphocytic choriomeningitis virus; LLO, listeriolysin O; LM, Listeria monocytogenes; LP, lamina propria; MLN, mesenteric lymph node.

  • Received January 9, 2002.
  • Accepted March 4, 2002.

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