Systemic C3 Modulates CD8⁺ T Cell Contraction after Listeria monocytogenes Infection

In response to bacterial and viral infections, the number of Ag-specific CD8⁺ T cells expands dramatically. Upon elimination of the infectious agent, the CD8⁺ T cell population decreases to homeostatic levels during a contraction phase (contraction) (1–3). In this scenario, the majority of responding CD8⁺ T cells undergoes apoptosis, whereas 5–10% of the original population survives and becomes long-lived memory cells (1–3). The contraction phase preserves the flexibility of the response to new infections while maintaining enhanced defense against previously encountered pathogens (4). Contraction is programmed early postinfection by cytokines, such as IL-12 and IFN-γ, that regulate differentiation of effector CD8⁺ T cells to KLRG-1^lo (killer-cell lectin-like receptor G1) CD127^hi long-lived memory precursor effector cells (MPECs) that are present at the peak of the response and preferentially survive contraction. In contrast, the CD8⁺ T cell subset KLRG-1^hiCD127^lo is short-lived effector cells (SLECs) that may undergo apoptosis (5–8). Furthermore, cytokines, such as IL-2, IL-7, or IL-15, influence contraction through differential regulation of CD8⁺ T cell subsets (MPECs and SLECs) during the contraction phase (9–11). Nevertheless, the mechanisms that control contraction are not fully understood (12).

A large body of evidence originally established the role of the complement system in the innate immune response against pathogens through its functions of opsonization, lysis, and anaphylatoxin activity (13–15). Subsequent research revealed that the complement system is involved in the induction and regulation of B cell– and T cell–mediated responses (13, 15–18). Complement activation regulates CD8⁺ T cell priming and proliferation in naturally occurring viral infections and in models of transplantation (16, 19, 20). Delayed-type hypersensitivity, a form of a secondary cell-mediated immune response, is suppressed by complement component of C3 (21). Furthermore, a peptide-treated C3-deficient (C3^−/−) mouse induces an increase in Ag-specific CD8⁺ IFN-γ⁺ cells and a higher rate of male graft rejection compared with that for wild-type (WT) mice 10 wk after grafting (22). These studies suggest that complement components may regulate contraction and the induction of memory cells.

C3 plays a central role in the complement cascade (15,). Hepatocytes are the major source of the circulating or systemic pools of C3, although significant, but smaller, amounts are produced locally by cells of the immune system or by other cell types (23, 24). The locally produced complement components contribute to the activation of APCs and T cells and drive CD4⁺ T cell differentiation, expansion, and survival (25–27). Similarly, studies on models of transplantation show that locally produced C3 regulates the expansion of CD8⁺ T cells (20, 28). In contrast, optimal CD8⁺ T cell responses against visceral leishmaniasis depend on circulating complement activation by natural Abs (29). Moreover, C3 depletion by cobra venom factor (CVF) treatment or C3 deficiency lead to weak CD8⁺ T cell responses because fewer blood-borne bacteria are transferred to splenic CD8α⁺ dendritic cells (DCs) postinfection (30). Therefore, it is important to determine the roles of systemic and local C3 production in the regulation of the CD8⁺ T cell response, because they may inform the design of more effective vaccines.

In the current study, using an informative murine model of Listeria monocytogenes infection (31, 32), we demonstrate that systemic C3 plays an important role in regulating CD8⁺ T cell contraction and that regulation of contraction by C3 may correlate with a reduction in the fraction of SLECs at the peak of the response, as well as decreased production of inflammatory cytokines.

C57BL/6 (B6) and BALB/c mice were purchased from the Peking University Animal Center (Beijing, China). C5a receptor-deficient (C5aR^−/−) mice on a BALB/c background, OT-I TCR-transgenic mice, and B6.C3^−/− mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Congenic C57BL/6 mice (Ly5.1) were a gift from Dr. Jinxiong She (Medical College of Georgia, Augusta, GA). Ly5.1-C3^−/− mice were generated by crossing Ly5.1-B6 with B6.C3^−/− mice. All mice were maintained in microisolator cages and housed and treated according to the guidelines of the Animal Care and Use Committee of the Third Military Medical University.

Recombinant L. monocytogenes expressing OVA (rLmOVA) and actA-deficient rLmOVA (△actA rLmOVA) were kindly provided by Dr. John Harty (University of Iowa, Iowa City, IA). Age-matched mice were injected i.v. with a sublethal dose of 4 × 10⁶ CFU △actA rLmOVA or 3 × 10⁴ CFU rLmOVA in 300 μl saline. For secondary infections, mice were injected i.v. with 1 × 10⁷ CFU △actA rLmOVA. Mice were provided 2 mg/ml ampicillin (Dingguo Bio, Beijing, China) in drinking water for 1 wk, starting on day 4 postinfection. The number of bacteria present in each spleen was determined by plating diluted spleen homogenates on brain–heart infusion plates containing erythromycin.

To deplete C3, mice were injected i.p. with 10 μg CVF (Quidel, San Diego, CA) in sterile PBS (300 μl) on day −1 before infection with △actA rLmOVA and then on days 1, 2, 3, 5, 7, 9, 11, and 13 postinfection. To determine the effects of complement during the early priming/expansion phase or the later contraction phase, mice were treated with CVF on days −1, +1, +2, +4, and +6 or were treated with CVF on days +7, +8, +10, and +12 or were treated with sterile PBS (control group) on the same day as CVF treatment.

Murine C3 was measured as described (33). Capture and detection Abs were from Cappel (ICN Pharmaceuticals, Costa Mesa, CA). Serum levels of IL-12, IL-23, IFN-γ, and TNF-α were determined using ELISA kits (BioLegend), according to the manufacturer’s protocol.

A small cyclic hexapeptide AcF-(OPdChaWR), which is a selective C5a receptor antagonist (C5aR-A) (34), was synthesized by GL Biochem (Shanghai, China). C5aR-A was administered by i.p. injection (1 μg/g mouse body weight) on days −1, +1, +2, +3, +5, +7, +9, +11, and +13 postinfection.

Anti-CD8α (clone 53-6.7), anti-Ly5.1 (clone A20), anti-Ly5.2 (clone 104), anti-CD127 (IL-7Rα; clone A7R34), anti–KLRG-1 (clone 2F1), anti-CD44 (clone IM7), anti–IFN-γ (clone XMG1.2), rat IgG2a and IgG2b isotype controls and mouse IgG1 isotype controls, anti–IFN-γ (clone XMG1.2), and anti–IL-2 (clone JES6-5H4) were purchased from eBioscience; anti-CD8α (clone 53-6.7) and anti-CD62L (clone MEL-14) were purchased from BioLegend; and anti-TCR Vβ4 (clone KT4), anti-TCR Vβ5.1, 5.2 (clone MR9-4), anti-TCR Vβ6 (clone RR4-7), anti-TCR Vβ7 (clone TR310), and anti-TCR Vβ8.1, 8.2 (clone MR5-2) were purchased from BD Biosciences. Splenocytes or PBMCs were prepared and stained with mAbs specific for proteins expressed on the cell surface. To detect intracellular cytokines, cells were cultured at 37°C for 4–5 h in complete medium supplemented with 10% calf bovine serum and 1 μl/ml brefeldin A (eBioscience), in the absence or presence of OVA_257–264 peptide (SIINFEKL; 1.0 μg/ml). Listeriolysin O 91–99 (LLO_91–99) peptide (GYKDGNEYI; 1.0 μg/ml) was used to detect the CD8⁺ T cell response of mice with a BALB/c background. These peptides were synthesized by Chinese Peptide (Hangzhou, Zhejiang, China). After culture, cells were harvested and stained using a Foxp3 Staining Buffer Set (eBioscience), following the manufacturer’s protocol. OVA-specific CD8⁺ T cell responses were also determined by staining with H2-K^b/OVA_257–264 pentamers (Proimmune, Oxford, U.K.), according to the manufacturer’s instructions. All FACS data were acquired using a BD FACSCanto II and analyzed using FlowJo software (TreeStar).

Single-cell suspensions of spleen and lymph node cells were split and pulsed with OVA_257–264 peptide (5 μg/ml) or buffer for 1 h in an incubator (37°C, 5% CO₂). The cells were washed with PBS and labeled with the fluorescent dye CFSE (Invitrogen, Carlsbad, CA) for 10 min at 37°C. The cells pulsed with OVA were labeled with CFSE^hi (5 μM) and used as targets, whereas those pulsed without peptide were labeled with CFSE^lo (0.5 μM) and served as internal controls. The cells were washed, counted, mixed 1:1, and injected i.v. into B6 and C3^−/− mice immunized with 4 × 10⁶ CFU △actA rLmOVA. The unimmunized mice were used as control. Spleens were removed 48 h later, and the ratio of CFSE^lo/CFSE^hi cells was determined using flow cytometry.

CD8⁺ T cells were enriched (∼90%) from splenocytes by immunodepletion of CD8⁻ cells using a CD8α⁺ T Cell Isolation Kit II (Miltenyi Biotec), according to the manufacturer’s instructions. To further purify CD8⁺ T cell subpopulations, magnetic bead–purified CD8⁺ T cells were stained with appropriate mAbs on ice in PBS containing 1% FCS and were further sorted using a FACSAria (Becton Dickinson). The purity of FACS-sorted samples was ∼98%. Groups of congenic recipient mice were injected i.v. with equivalent numbers of the respective CD8⁺ T cell subpopulations.

B6 WT and C3^−/− mice were irradiated (1100 cGy) and injected i.v. with 10 × 10⁶ bone marrow (BM) cells from congenic B6 or C3^−/− mice, respectively. After 2 mo, the animals were bled, and CD8⁺ T cells were analyzed by surface staining to confirm chimerism. The mice were immunized with △actA rLmOVA, and analyzed 7, 14, and 21 d after immunization.

Total RNA was extracted using TRIzol reagent (Invitrogen) from the FACS-sorted populations of CD8⁺ T cells, according to the manufacturer’s instructions. RNA purity and concentration were determined using a NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE). Total RNA (1 μg) was reverse transcribed using the PrimeScript RT Reagent Kit with gDNA Eraser (TAKARA BIO, Otsu, Japan), according to the manufacturer’s instructions. PCR conditions were as follows: denaturation at 95°C for 30 s, followed by 45 cycles of amplification at 95°C for 5 s, 60°C for 30 s, and 72°C for 30 s. Quantitative PCR was performed using a Stratagene Mx3000P system (Agilent Technologies, Santa Clara, CA) and a SYBR Premix Ex Taq (Perfect Real Time) Kit (TaKaRa), according to the manufacturer’s instructions. Each reaction mixture, consisting of 7.0 μl water, 1 μl forward primer (10 μM), 1 μl reverse primer (10 μM), and 10 μl 2× Master mix, was added to 1.0 μl cDNA. Data are expressed relative to those of the controls. The average threshold-cycle differences among the samples were normalized to the β-actin (Actb) (control) in the corresponding cDNA preparation. Primers used were 5′-ACC TTA CCT CGG CAA GTT TCT-3′ and 5′-TTG TAG AGC TGC TGG TCA GG-3′ for C3 and 5′-CAC TAT CGG CAA TGA GCG GTT CC-3′ and 5′-CAG CAC TGT GTT GGC ATA GAG GTC-3′ for Actb.

Statistical analysis was performed using an unpaired two-tailed Student t test by Prism GraphPad Software (La Jolla, CA). The p values < 0.05 were considered statistically significant.

C3^−/− mice were immunized with △actA rLmOVA, and changes in the numbers of OVA-specific CD8⁺ T cells were determined by staining for intracellular IFN-γ from days 7 to 50. On day 7 following immunization, effector CD8⁺ T cell expansion was approximately two-thirds lower in C3^−/− mice compared with WT mice (Fig. 1A, 1B), which is consistent with the results of another study showing that CD8⁺ T cell responses to L. monocytogenes infection were downregulated in the absence of C3 (35). Interestingly, compared with control mice, the contraction (days 7–16) in C3^−/− mice was much slower, whereas the numbers of OVA-specific effector CD8⁺ T cells in C3^−/− mice were greater (∼2–3-fold) on day 50 postinfection (Fig. 1B). K^b/OVA pentamer analysis also revealed decreased contraction in C3^−/− mice (Fig. 1C, 1D). In addition, we used rLmOVA to immunize mice and found slower contraction in C3^−/− mice than in WT mice (Fig. 1E). These results suggest that C3 is required for CD8⁺ T cell contraction after L. monocytogenes infection.

CVF treatment leads to complement depletion by activating the alternative pathway of C3 activation (36). To verify the role of C3 in regulating contraction, we depleted circulating C3 in WT mice by ∼90% by continuous administration of CVF from days −1 to +13 (Fig. 2A). The primary response of the CVF-treated mice was ∼50% of that of the controls on day 7 postinfection. Contraction was significantly slower, whereas the percentage of OVA-specific CD8⁺ T cells on day 50 was higher in CVF-treated mice than in the controls (Fig. 2B, 2C).

To further determine the effect of systemic C3 during the priming/expansion or contraction phases, we treated mice with CVF from days −1 to +6 or from days +7 to +12. Only the mice treated with CVF from days −1 to +6 showed decreased contraction postinfection (Fig. 2D), indicating that systemic C3 was involved in the regulation of contraction during the early priming/expansion phase.

C3 activation generates downstream pathway components, including anaphylatoxin C5a (15, 37,). Because C5a/C5aR signaling enhances the CD8⁺ T cell response (38, 39), we hypothesized that C3 regulated CD8⁺ T cell contraction through C5a. Therefore, we used a peptide antagonist (C5aR-A) specific for C5aR to test this possibility. Contraction in mice treated with C5aR-A did not differ significantly compared with controls (Supplemental Fig. 1). We also assessed the LLO_91–99-specific CD8⁺ T cell response in C5aR^−/− mice on a BALB/c background and found that CD8⁺ T cell contraction was not significantly different from that in WT mice (Fig. 2E). These results suggest that the regulation of CD8⁺ T cell contraction under conditions of C3 deficiency is independent of C5aR signaling.

To further address the role of systemic C3 on contraction, we generated chimeras by infusing WT BM cells into WT or C3-deficient recipients. Thus, in the presence or absence of serum C3, BM cells produced local C3. C3^−/− BM cells also were infused into WT or C3-deficient recipients. Thus, BM cells do not produce local C3, regardless of whether serum C3 is present or absent. The presence of chimerism after 2 mo was determined using flow cytometry (Fig. 3A). The chimeric animals were immunized with △actA rLmOVA, and the changes in CD8⁺ T cell responses were analyzed using PBLs on days 7, 14, and 21 postinfection. We found that the C3^−/− recipients exhibited slower contraction compared with WT recipients when engrafted with cells from the same donor BM. In contrast, CD8⁺ T cell contraction in the same recipient mice, when engrafted with BM from C3^−/− or WT donors, respectively, showed no significant difference (Fig. 3B, 3C). These findings support the conclusion that systemic C3 regulates CD8⁺ T cell contraction.

Because the unique phenotype of memory CD8⁺ T cells contributes to their protective effect (5, 8, 40), we tested whether the phenotype of OVA-specific CD8⁺ T memory cells from C3^−/− mice was similar to those from WT mice (5, 8, 40). The expression of CD62L, KLRG-1, CD127, and CD44 of OVA-specific CD8⁺ T cells, as well as the production of IL-2, was not significantly different between WT and C3^−/− mice (Fig. 4).

Memory CD8⁺ T cells undergo a vigorous expansion to protect against a challenge infection (1, 41). Therefore, we next investigated whether memory T cells harvested from C3^−/− mice functioned normally. Because complement is involved in phagocytosis and killing of C3-bound L. monocytogenes by activated macrophages (42) and is required for the efficient generation of CD8⁺ T cell responses (35), it is reasonable to hypothesize that C3^−/− mice acquire greater bacterial loads after L. monocytogenes infections. Surprisingly, the bacterial load was significantly lower in the spleen of C3^−/− mice than in WT mice postinfection because of reduced blood-borne bacterial shuttling to splenic CD8α⁺ DCs (30). We speculated that bacterial loads might not correlate with the protective function of effector CD8⁺ T cells in the model of L. monocytogenes infection. Therefore, we performed an in vivo killing assay on day 34 and found that approximately twice as many OVA-specific target cells were killed in C3^−/− mice compared with WT mice (Fig. 5A, 5B). Because the amount of memory T cells in C3^−/− mice was about twice that in WT mice (Fig. 1), these results indicate that the cytotoxic potential of memory cells in C3^−/− mice was comparable to that in WT mice.

We next determined whether the expansion abilities of the memory cells of C3^−/− and WT mice were the same. After rechallenge at 31 d after primary infection, higher numbers of CD8⁺ T cells were detected in C3-depleted mice, and the slopes of the CD8⁺ T cell response curves for C3-depleted and control mice were not significantly different (Fig. 5C). This result suggests that memory cells in C3-depleted and control mice underwent equivalent response after challenge. Unexpectedly, the population of OVA-specific CD8⁺ T cells in WT mice expanded faster and to a greater extent than did that in C3^−/− mice after rechallenge, although more memory CD8⁺ T cells were detected in C3^−/− mice than in WT mice (Supplemental Fig. 2).

The recall response of memory CD8⁺ T cells of C3^−/− mice differs from that of C3-depleted mice. We hypothesized that the difference may be attributed to the direct effect of C3 on CD8⁺ T cell proliferation (35), as well as on contraction and memory induction. Therefore, we asked whether CD8⁺ memory T cells generated in C3^−/− mice mounted a full response in mice with normal levels of C3. For this purpose, CD8⁺ memory T cells from C3^−/− and WT mice were transferred into WT naive mice 40 d postinfection (Fig. 5D). One day later, recipient mice were infected with △actA rLmOVA. The fold increase in memory CD8⁺ T cells in the recall response was not significantly different between C3^−/− and WT donors (Fig. 5E, 5F), suggesting that the recall capability of memory CD8⁺ T cells in C3-deficient and C3-depleted mice were similar compared with WT mice. Taken together, these data indicate that the phenotypes of memory cells generated in C3-deficient and C3-depleted mice were comparable to those in WT mice and that the memory cells protected against infection.

Next, we sought to identify factors that participated during the priming/expansion phase. Curtailing prolonged stimulation of CD8⁺ T cells toward the end (days 6–7) of an acute infection decreases the differentiation potential of terminal effectors (7, 8). To test the possibility that curtailed bacterial persistence in C3^−/− mice inhibited CD8⁺ T cell contraction, C3^−/− and WT mice were treated with ampicillin from day 4 postinfection to eliminate persistent bacteria; on day 6 postinfection, the bacteria were eliminated from C3^−/− and WT mice (Fig. 6A). Moreover, the numbers of OVA-specific CD8⁺ T cells in C3^−/− mice were less than half of those in WT mice on day 6, whereas this population contracted more slowly, which resulted in ∼2-fold more OVA-specific CD8⁺ T cells on day 50 (Fig. 6B, 6C). These results indicate that the reduced CD8⁺ T cell contraction is not due to the duration of Ag persistence.

Functional avidity of effector CD8⁺ T cells may contribute to their initial fate decision and to the generation of SLECs (43). The functional avidity of effector CD8⁺ T cells is determined by their sensitivity to peptide Ag or their distribution of TCR Vβ (44).

We attempted to determine whether the functional avidity of effector CD8⁺ T cells in C3^−/− mice differed from that in WT mice and correlated with reduced CD8⁺ T cell contraction. We found that the distribution of TCR Vβ expression among OVA-specific CD8⁺ T cells in C3^−/− mice was not significantly different from that in WT mice on day 7 postinfection (Fig. 7A). To determine whether the sensitivity to peptide of effector CD8⁺ T cells differed between C3^−/− and WT mice, splenocytes were isolated on day 7 postinfection and stimulated with different concentrations of OVA peptide. There was no significant difference in the peptide sensitivity between C3^−/− and WT mice (Fig. 7B). Together, these data suggest that altered contraction of effector CD8⁺ T cells is not associated with changes in the functional avidity of effector CD8⁺ T cells.

Postinfection or postimmunization, early effector CD8⁺ T cells were proposed to differentiate into at least two major subsets that can be distinguished by their expression of cell surface markers, such as CD127 and KLRG-1 (5, 7, 8). KLRG-1^hiCD127^lo SLECs are more likely to die after the peak response, whereas KLRG-1^loCD127^hi MPECs preferentially survive contraction and slowly develop into memory cells (7, 8). Therefore, CD8⁺ T cell contraction correlates with the proportions of these two effector CD8⁺ T cell subsets after differentiation (7).

We hypothesized that decreased contraction in C3^−/− mice may be due to diminished differentiation of SLECs. Indeed, there were fewer SLECs in C3^−/− mice than in WT mice (p < 0.01; Fig. 8A, 8B). In contrast, more MPECs were generated in C3^−/− mice than in WT mice (p < 0.01; Fig. 8A, 8B). This result indicates the generation of more MPECs during the effector T cell differentiation phase and is consistent with our findings that more memory cells were present in C3^−/− mice (Fig. 1). We also found fewer KLRG-1^hi subsets in C3-depleted mice compared with the controls on day 7 postinfection (Fig. 8C, 8D).

We further investigated whether C3 deficiency influenced the contraction of MPECs or SLECs adoptively transferred into infection-matched mice. Because KLRG-1^loCD127^hi MPECs represented a very small proportion, we were unable to obtain a sufficient number of cells to perform the transfer experiments; only KLRG-1^hiCD127^lo SLECs subpopulations were transferred. Their contraction was not affected by C3 during the later contraction phase (Supplemental Fig. 3), which was consistent with the result of the CVF-treatment experiment (Fig. 2D). These results suggest that C3 facilitates early CD8⁺ T cell differentiation toward SLECs, which promotes CD8⁺ T cell contraction.

Inflammatory cytokines regulate CD8⁺ T cell contraction through their influence on early priming and differentiation (6, 7). For example, IL-12 promotes the preferential differentiation of CD8⁺ T cells toward the SLEC phenotype (7). Because Ag-specific CD8⁺ T cells differentiated into fewer SLECs in C3^−/− mice (Fig. 8A, 8B), we asked whether cytokines secreted by DCs in C3^−/− mice differed from those secreted in control mice and contributed to reduced contraction. WT or C3^−/− mice were infected with △actA rLmOVA, and the levels of IL-12, IL-23, IFN-γ (6), and TNF-α (45) in sera were detected at 24 h postinfection. The levels of IL-12, IL-23, and IFN-γ in the sera of C3^−/− mice were significantly lower compared with those of WT mice (Fig. 9). These data suggest that the decreased inflammatory cytokines in C3^−/− mice may correlate with the inhibition of differentiation to SLECs and decreased contraction.

During an immune response against acute infection, CD8⁺ T cell contraction is a critical homeostatic process that occurs following the peak of Ag-specific T cell responses. Accumulating evidence shows that the complement system plays a very important role in the regulation of T cell immunity (16, 17, 46), particularly in the regulation of primary CD8⁺ T cell responses in certain models of bacterial and viral infection (19, 29, 35, 39). The present study was undertaken to determine the role of C3 in regulating CD8⁺ T cell contraction and memory induction. Our findings demonstrate that CD8⁺ T cell contraction was less in C3^−/− mice than in WT mice, as well as upon C3 depletion induced by CVF treatment. The contraction also was diminished in C3^−/− recipient mice to a greater extent than in WT recipient mice when each was engrafted with the same BM cells. Moreover, our findings suggested that decreased CD8⁺ T cell contraction in C3^−/− mice may correlate with the generation of fewer SLECs and reduced levels of serum inflammatory cytokines.

Marked increases in primary viral Ag–specific CD8⁺ T cell expansion and numbers of memory CD8⁺ T cells occur in decay-accelerating factor (Cd55/Daf1)-knockout mice after acute infection with lymphocytic choriomeningitis virus compared with those in WT mice (39). Deletion of C3 or C5aR in Cd55^−/− mice reverses the enhanced CD8⁺ T cell response but does not influence contraction (39). In this study, we show reduced contraction in C3-deficient or C3-depleted mice compared with the controls. Moreover, CD8⁺ T cell contraction was not influenced in C5aR^−/− mice or by treatment with C5aR-A. Therefore, the effect of C3 activation on the regulation of CD8⁺ T cell responses may be complex and variable, depending on the pathogen.

Although understanding how the complement regulates T cell responses is incomplete and varies in different models of infection, recent data generated by transplantation and infection models suggest that local production of C3 determines the responses of CD8⁺ T cells (28, 35). For example, in a model of L. monocytogenes infection, C3 directly promotes CD8⁺ T cell expansion in vitro (35), which suggests that locally produced C3 acts on CD8⁺ T cells. We detected increased C3 mRNA expression in activated CD8⁺ T cells from WT mice (Supplemental Fig. 4). However, there was no significant difference in contraction in the WT recipient mice when respectively engrafted with BM from WT or C3^−/− donors; also the contraction was not significantly different in the C3^−/− recipient mice with BM adoptive transfer from WT or C3^−/− donors, suggesting that local C3 might not play an important role in regulating CD8⁺ T cell contraction.

How does systemic C3 regulate CD8⁺ T cell contraction? We show in this study that C3 exerted an effect early postinfection but not during later contraction. DCs are essential for the induction of antilisterial CD8⁺ T cell immunity early postinfection (47, 48), and multiple signals integrated (e.g., inflammatory cytokines) by DCs program the CD8⁺ T cell response (2, 49). Therefore, we hypothesize that systemic C3 depletion or C3 deficiency may alter the characteristics of DCs to regulate contraction. Evidence indicates that the strength or duration of an antigenic signal influences contraction through altering SLEC differentiation (8, 43). Nevertheless, we found that CD8⁺ T cell contraction was independent of prolonged duration of infection or functional avidity of effector CD8⁺ T cells. The present results suggest that antigenic signals were not involved in regulating contraction, consistent with findings that the dose and duration of infection do not influence contraction (4, 6). Moreover CD8⁺ T cells do not undergo cell division in response to residual Ag (50), and C3 deficiency does not affect the ability of DCs to present Ag in vitro or during the maturation of DCs in vivo after L. monocytogenes infection (35). In contrast, we found that the levels of critical cytokines, such as IL-12 and IFN-γ, were reduced 24 h postinfection. Because IL-12 and IFN-γ program contraction (6, 7), it is reasonable to conclude that these cytokines may link C3 to CD8⁺ T cell contraction.

In summary, our results reveal an important role for systemic C3 in influencing CD8⁺ T cell contraction following an infection with intracellular bacteria and provide insights into the mechanisms by which complement regulates the CD8⁺ T cell response. Furthermore, because depletion of C3 by CVF treatment reduced CD8⁺ T cell contraction and increased the generation of CD8⁺ memory T cells with enhanced protective function, our findings suggest that strategies that alter the functions of the complement pathway may enhance vaccine efficacy.

We thank Dr. Liyun Zou for critical reading and comments on the manuscript.

The authors have no financial conflicts of interest.