Human C-reactive protein (CRP) is a serum-soluble pattern recognition receptor that serves as a marker of inflammation and directly contributes to innate immunity. In this study, we show that human CRP also directly contributes to adaptive immunity, that is, native CRP binds specifically to human Jurkat T cells and to mouse naive CD4+ T cells and modulates their Th1 and Th2 responses. In vitro both exogenously added (purified) and endogenously expressed (via transfection) human CRP inhibited Th1 differentiation and augmented Th2 differentiation of naive CD4+ T cells. In vivo for human CRP transgenic compared with wild-type mice, a lesser proportion of the T cells recovered from the spleens of healthy animals were Th1 cells. Moreover, in both CRP transgenic mice and in wild-type mice treated with human CRP, during myelin oligodendrocyte glycoprotein peptide–induced experimental autoimmune encephalomyelitis both the Th1 cell response and disease severity were inhibited. These pattern recognition–independent actions of CRP directly on T cells highlights the potential for this soluble pattern recognition receptor to act as a tonic regulator of immunity, shaping global adaptive immune responses during both homeostasis and disease.
The C-reactive protein (CRP) is evolutionarily conserved and there is no known natural deficiency in humans (1). CRP is a major human acute phase reactant, and thus the plasma concentration of CRP can increase up to 1000-fold upon tissue injury or infection (1). Although a rise in circulating CRP is widely used as a nonspecific clinical marker of inflammation and several of its properties have been well defined in vitro, there is still no consensus about the exact function of CRP in vivo (1, 2). The recognized capacity of CRP to bind Fc receptors, to activate the classical pathway of complement, and to opsonize both apoptotic cells and microbes supports the proposition that CRP acts as a soluble pattern recognition receptor (PRR) in vivo and thereby directly contributes to innate host defense (3, 4). Additional studies done using human CRP transgenic (CRPtg) mice indicate that CRP might also regulate autoimmunity (5–8), and our recent identification of highly recurrent promoter mutations in the CRP gene in multiple types of cancers suggests that CRP might also play a critical role therein (9, 10).
CD4+ effector T cells are key components of adaptive immunity and they play a major role in controlling infections and the development of autoimmunity and cancer (11–16). The propagation of effector CD4+ T cells begins when TCRs on naive CD4+ T cells are engaged by cognate Ags in the context of MHC class II and costimulation provided by APCs. Thus activated and depending on the nature of cytokines produced by cells of the innate immune system, naive T cells differentiate into multiple kinds of effectors, including IFN-γ–secreting Th1 cells, IL-4–secreting Th2 cells, and IL-17–secreting Th17 cells (17, 18). PRRs were originally thought to regulate T cell differentiation and effector responses indirectly via their actions on APCs and other kinds of innate immune cells. However, recent evidence indicates that TLRs, the representative membrane PRRs, are themselves expressed by T cells and hence can directly modulate T cell responses following TLR ligation by their cognate ligands (19–21).
In the mid-1970s, it was initially reported that CRP could bind T cells and thereby modulate their effector functions (22–24). Subsequently, however, that observation could not be reproduced by the same group (25). The paradoxical outcomes were attributed to differences in CRP purity (25). Nevertheless, because T cell heterogeneity was not fully appreciated at the time, its likely contribution to the observed variance in CRP binding and actions was not explored. Importantly, although FcRs were identified as major receptors for CRP (26, 27), there is little evidence that T cells express FcRs (28). Thus, whether purified CRP is able to directly interact with T cells still remains equivocal.
In the present study, we rigorously characterized both the CRP preparations and T cells that we used and revisited the question of CRP binding by T cells. We demonstrate that human CRP in its native pentameric conformation does indeed bind to both primary mouse naive T cells and to human leukemic Jurkat T cells. This binding is independent of calcium or the classic CRP ligand phosphorylcholine, and it does not require either FcR or LOX-1, another recently identified CRP receptor (29). CRP binding to T cells is abrogated by pretreatment of cells with proteases, however, indicating a requirement for an as yet unidentified receptor. Importantly, to our knowledge, we show for the first time that CRP binds preferentially to the naive T cell subset and thereby modulates their differentiation, favoring the Th2 effector program while inhibiting the Th1 program both in vitro and in vivo. Although the identity of the CRP-binding receptor on naive T cells has yet to be determined, these new results suggest that CRP might play a direct role in regulating the adaptive immune response. Given the inducible nature of CRP and its systemic presence, it remains plausible that CRP is an important tonic regulator of adaptive immunity (8).
Materials and Methods
Wild-type mice (strain C57BL/6) were from the Animal Center of Lanzhou University. Human CRPtg mice have been fully described elsewhere (30). Human CRP is present in the blood of CRPtg mice at concentrations manifested in humans, that is, low levels under steady-state conditions (<1–10 μg/ml) and high levels during the acute phase response (∼30–500 μg/ml). Mice were housed at constant humidity (60 ± 5%) and temperature (24 ± 1°C) with a 12-h light cycle (6 am to 6 pm) and maintained ad libitum on sterile bottled water and regular chow (Harlan Teklad). Eight- to 12-wk-old mice were used unless specifically noted otherwise. All animal use protocols were approved by the Institutional Animal Care and Use Committees at the University of Alabama at Birmingham and Lanzhou University and were consistent with the Guide for the Care and Use of Laboratory Animals, 8th Ed. (2010).
Native human CRP purified (>99% purity) from ascites was purchased from Binding Site (Birmingham, U.K.). To ensure that calcium and ligand binding ability was retained, CRPs were repurified with phosphorylcholine (PC)-agarose beads (Thermo Fisher Scientific, Rockford, IL), and dialyzed extensively to remove any residual NaN3. Finally, CRP was passed through Detoxi-Gel columns (Thermo Fisher Scientific) to remove endotoxin. The functional integrity of the CRP molecule was then directly verified by SDS-PAGE (0.5% of the normal concentration) (31) (Supplemental Fig. 1A), electron microscopy (Supplemental Fig. 1B), conformation-specific ELISA (32) (Supplemental Fig. 1C), and calcium-dependent binding to the classic CRP ligand PC (1, 4) (Supplemental Fig. 1D). As a further safeguard, polymyxin B (20 μg/ml) was included in cell response experiments to exclude possible confounding effects from residual endotoxin in any of the culture media used. Anti-CD3 mAb (2C11, catalog no. 555273), anti-CD28 mAb (37.51, catalog no. 553295), anti–IL-4 mAb (catalog no. 554432), anti–IFN-γ mAb (catalog no. 554408), anti–IL-12 mAb (catalog no. 554475), mouse (m)IL-2 (catalog no. 550069), mIL-12p70 (catalog no. 554592), mIL-4 (catalog no. 550067), ELISA kits for IL-4 (catalog no. 555232), and IFN-γ (catalog no. 555138) were from BD Biosciences (San Jose, CA). Anti-Ly6G mAbs were from BioLegend (San Diego, CA, catalog no. 127611) and other reagents were from Sigma-Aldrich (St. Louis, MO). Anti-CRP mAbs 1D6, 2C10, and 3H12 were prepared as described (33).
T cell differentiation
CD4+ and CD62L+ naive T cells were purified from the spleens of male C57BL/6 mice using MACS kits (Miltenyi Biotec, Bergisch Gladbach, Germany, catalog no. 130-093-227) according to the manufacturer’s instructions. T cells (2 × 105) were cultured in 300 μl medium for 3 d with plate-bound anti-CD3 (2 μg/ml, immobilized overnight at 4°C) and fluid-phase anti-CD28 mAbs (2 μg/ml), in the presence or absence of CRP under Th1-polarizing conditions (10 ng/ml mIL-2, 20 ng/ml mIL-12p70, 10 μg/ml anti–IL-4 mAb) or Th2-polarizing conditions (10 ng/ml mIL-2, 20 ng/ml mIL-4, 10 μg/ml anti–IL-12 mAb, 10 μg/ml anti–IFN-γ mAb). In some experiments, cells were transfected with control or CRP-expressing pcDNA 3.1 plasmids via electroporation with a Multiporator apparatus (Eppendorf, Hamburg, Germany) or by using the X-tremeGENE HP DNA transfection reagent (Roche, Basel, Switzerland, catalog no. 550067). The cells were cultured under resting condition for an additional 2 d followed by treatment with PMA (20 ng/ml), calcium ionophore (1 μg/ml), and monensin (BD Biosciences, catalog no. 550069) for 5 h. Cells were washed with PBS and FcRs were blocked with anti-mouse CD16/32 mAb (Abcam, Cambridge, U.K., catalog no. AF1460) for 10 min on ice. After staining with FITC-rat anti-mouse CD4 (BD Biosciences, catalog no. 553046) or rat IgG2a κ isotype control (BD Biosciences, catalog no. 553929) for 30 min at 4°C, cells were washed twice and treated with Cytofix/Cytoperm fixation/permeabilization buffer (BD Biosciences, catalog no. 554714). Intracellular IL-4 and IFN-γ were stained with PE-rat anti-mouse IL-4 (BD Biosciences, catalog no. 554435) and PerCP-Cy5.5–rat anti-mouse IFN-γ (BD Biosciences, catalog no. 560660) or corresponding isotype controls (BD Biosciences, catalog nos. 554685 and 560537) for 30 min at 4°C followed by washing and flow cytometry detection. Negligible cell staining was observed with all isotype control Abs.
The mRNA levels of cytokines and transcription factors were measured by quantitative PCR using the SYBR Premix Ex Taq II kit (Takara, Shiga, Japan, catalog no. DRR091C) and a CFX96 real-time thermal cycler (Bio-Rad, Hercules, CA). Cytokine and transcription factor mRNA levels were normalized to β-actin (ACTB) or GAPDH. The primer sequences used are: human IFN-γ forward, 5′-GAATGTCCAACGCAAAGCAAT-3′, reverse, 5′-GACCTCGAAACAGCATCTGACTCCT-3′; human IL-4 forward, 5′-CTGTGCACCGAGTTGACCGTA-3′, reverse, 5′-GTCCTTCTCATGGTGGCTGTAGAAC-3′; human ACTB forward, 5′-GCAAAGACCTGTACGCCAACA-3′, reverse, 5′-ACACGGAGTACTTGCGCTCAG-3′; mouse IFN-γ forward, 5′-CCATCAGCAACAACATAAGCGTC-3′, reverse, 5′-TTGACCTCAAACTTGGCAATACTCA-3′; mouse IL-4 forward, 5′-TTCCAAGGTGCTTCGCATA-3′, reverse, 5′-TGCAGCTTATCGATGAATCCA-3′; mouse IL-12Rβ2 forward, 5′-TCTGCGAAATTCAGTACCGAC-3′, reverse, 5′-GCCCACCGTGATGATAGC-3′; mouse IL-4Ra forward, 5′-GGGCATGGAGGCTACAAG-3′, reverse, 5′-CTCCGTGTCTAGTCCGAAAGT-3′; mouse T-bet forward, 5′-CCATTCCTGTCCTTCACCG-3′, reverse, 5′-CTGCCTTCTGCCTTTCCAC-3′; mouse GATA-3 forward, 5′-AGTCCTCATCTCTTCACC-3′, reverse, 5′-CACTCTTTCTCATCTTGC-3′; mouse GAPDH forward, 5′-GGAGAAACCTGCCAAGTATGA-3′, reverse, 5′-GTGGGTGCAGCGAACTTTA-3′.
CRP was incubated with 5 × 105 Jurkat or mouse T cells for 1 h at 4°C in PAB buffer (PBS, 0.1% BSA, 0.05% NaN3, 1 mM Ca). After washing twice with PAB, cells were incubated sequentially with anti-CRP mAbs 2C10, 1D6, or 3H12 (33) and DyLight 650–labeled anti-mouse Ab (Abcam, catalog no. ab96874). Cells were then incubated with propidium iodide (PI) for 15 min at room temperature to allow identification of live/dead cells by flow cytometry. In some binding experiments, allophycocyanin-labeled streptavidin (BioLegend, San Diego, CA, catalog no. 405207) was used to detect the binding of biotin-labeled CRP.
Mouse naive T cells stimulated in vitro as indicated above were lysed in the presence of protease/phosphatase inhibitor mixture (Cell Signaling Technology, Danvers, MA, catalog no. 5872) for 30 min on ice. The lysates were centrifuged at 12000 × g for 30 min at 4°C and the supernatants were subjected to standard SDS-PAGE followed by transferring to polyvinylidene difluoride membranes for detection of STAT, T-bet, or GATA-3 proteins. Rabbit anti-STAT1 (catalog no. 9172S), anti–phospho-(Tyr701) STAT 1 (catalog no. 9167S), anti-STAT4 (catalog no. 2653S), and anti–phospho-(Tyr693) STAT 4 (catalog no. 4134S) were from Cell Signaling Technology. Rabbit anti-STAT6 (catalog no. ab32520) and anti–phospho-(Tyr641) STAT6 (catalog no. ab54461) were from Abcam. Anti–T-bet mAb 4B10 (catalog no. sc-21749) and anti–GATA-3 HG3-31 (catalog no. sc-268) were from Santa Cruz Biotechnology (Dallas, TX).
Experimental autoimmune encephalomyelitis
Female C57BL/6 mice housed in conventional conditions were immunized s.c. with 200 μg myelin oligodendrocyte glycoprotein (MOG) peptide 35–55 (>99% purity; Shanghai Science Peptide Biological Technology, Shanghai, China) in CFA containing 10 mg Mycobacterium tuberculosis strain H37Ra (Chondrex, Redmond, WA, catalog no. 7027). On days 0 and 2, immunized mice were injected i.p. with 200 ng pertussis toxin (Enzo Life Sciences, Farmingdale, NY, catalog no. BML-G101). On day 2, immunized mice received s.c. a single injection of 200 μg human CRP or buffer control. The clinical severity of experimental autoimmune encephalomyelitis (EAE) was scored using the standard grading scale: 0, asymptomatic; 1, limp tail; 2, limp tail and weakness of hindlimb; 3, limp tail and partial hindlimb paralysis; 4, limp tail, complete hindlimb and partial foreleg paralysis; 5, moribund. The splenocytes were isolated on day 7 or at the peak of EAE symptoms and restimulated ex vivo with 100 μg/ml MOG peptide 35–55. Intracellular and secreted cytokines were then determined by flow cytometry and ELISA, respectively. In experiments comparing wild-type and CRPtg splenocytes, flow cytometry was done without prior MOG restimulation in vitro.
Assessing spinal cord cell infiltration
To examine spinal cord–infiltrating cells, spinal cords were removed from mice at the peak of EAE. After perfusion with PBS, the spinal cords were ground through a cell strainer, washed in buffer, resuspended in 30% Percoll, and layered onto 70% Percoll. After centrifugation at 2500 rpm (room temperature, 30 min), cells at the interface were carefully removed and washed in PBS. Cells were plated in complete medium using round-bottom 96-well plates and incubated at 37°C for 4 h in the presence of GolgiStop (BD Biosciences). After treatment with fixing/permeabilization buffer the cells were washed and stained with cell marker–specific mAbs and processed for flow cytometry.
Data are presented as means ± SEM. Statistical analysis was performed using two-tailed Student t tests or one-way ANOVA with Tukey post hoc comparisons. The p values <0.05 were considered significant.
Specific binding of CRP to naive CD4+ T cells
Human Jurkat clone E6-1 is a widely used human T cell line. We found that highly purified human CRP bound to live E6-1 Jurkat T cells in a concentration-dependent manner with >70% of cells staining positive for CRP at saturation (Fig. 1A). The calculated KD of this interaction was 163.3 μg/ml, but substantial CRP binding could still be observed at lower concentrations. This binding was not attributable to CRP with an altered conformation, as the native structure of our CRP preparations were thoroughly validated by documenting the integrity of CRP’s pentameric assembly (Supplemental Fig. 1A, 1B), the exclusive expression of its native antigenicity (Supplemental Fig. 1C), and the calcium dependence of its interaction with PC (Supplemental Fig. 1D). Moreover, CRP bound to E6-1 cells could be detected with an mAb specific to native CRP (1D6) but not with an mAb (3H12) that recognizes a CRP epitope exposed in the altered CRP conformation (Fig. 1B).
Because CRP binds avidly to necrotic or apoptotic cells (34), this might also account for the observed binding of CRP to E6-1 cells. However, counterstaining cells with PI or annexin V revealed only negligible proportions of dead or dying cells in our cultures (Fig. 1B–E, Supplemental Fig. 2). Moreover, the binding of CRP to live E6-1 Jurkat T cells was both calcium- and PC-independent (i.e., not inhibited with 5 mM EDTA and 5 mM PC, respectively) (Fig. 1C, top panels). In stark contrast, interaction of CRP with necrotic cells (generated by exposure to boiling buffer) was both EDTA and PC inhibitable (Fig. 1C, bottom panels). Surface receptors were likely required for the association of CRP with live E6-1 cells, as pretreating the cells with proteinase K or trypsin abrogated CRP binding (Fig. 1D). Nonetheless, CRP binding to live E6-1 cells was not FcγR-dependent because these cells did not express FcγRs (Supplemental Fig. 3A) and because neither monomeric nor aggregated IgG was able to prevent CRP binding (Fig. 1E). LOX-1 has recently been identified as a potential receptor for CRP on endothelial cells (29). However, the presence of polyinosinic acid, an inhibitor of LOX-1 (35), also failed to reduce the binding of CRP to live Jurkat cells (Supplemental Fig. 3B).
To test whether CRP binding to T cells extended beyond the special case of E6-1 Jurkat cells, we also examined the interaction of CRP with primary mouse CD4+ T cells. Unexpectedly, only a small percentage of live CD4+ T cells (isolated from the spleens of healthy adult mice) bound CRP under the conditions we tested (Fig. 2A). As Jurkat T cells more closely resemble the naive phenotype (36), we purified primary mouse naive T cells based on their high expression of the homing receptor CD62L (l-selectin) (37). On this CD4+CD62L+ T cell subpopulation we detected appreciable binding of CRP (Fig. 2B). Confocal microscopy was used to confirm prominent binding of CRP to CD4+CD62L+ T cells but not to CD4+CD62L− T cells (Fig. 2C). Similar to binding to human Jurkat cells, CRP binding to mouse primary CD4+CD62L+ naive T cells did not require calcium or PC, but it did require surface (proteinaceous) receptors (Fig. 2D, 2E) other than FcγR and LOX-1 (Supplemental Fig. 3C). Taken together, these results establish that human CRP binds preferentially to the naive CD4+ T cell subset and that this action requires proteinaceous surface receptors.
Binding of CRP to naive CD4+ T cells alters their Th1/Th2 responses
Having established that CRP binds to human Jurkat and mouse naive CD4+ T cells, we asked whether this was sufficient to drive functional consequences. The effector functions of T cells are predicated on their activation and differentiation consequent to TCR recognition of cognate Ag and costimulation by APCs (17, 18). To investigate the impact of CRP binding on this process, we first measured expression of IL-4 and IFN-γ, two key effector T cell cytokines, produced by Jurkat T cells activated with immobilized anti-CD3 and soluble anti-CD28 mAbs, a combination that drives polyclonal T cell activation. To ensure that any effect we observed was not due to NaN3 or endotoxin contamination, we removed these from our CRP preparations by extensive dialysis, passage through Detoxi-Gel columns, and inclusion of polymyxin B to neutralize any residual endotoxin. Even after these exhaustive measures, CRP significantly enhanced IL-4 expression and inhibited IFN-γ expression by CD3/CD28-activated Jurkat cells (Fig. 3A). These effects were attributable to native CRP, as both boiling the protein and eliminating it by filtration through a 10-kDa cut-off membrane abrogated these actions (Fig. 3A). Importantly, the IL-4–augmenting and IFN-γ–suppressing effects were also recapitulated in E6-1 cells transfected with a human CRP expression vector (Fig. 3B).
Next, we examined the effects of CRP on primary mouse naive CD4+CD62L+ T cells and obtained comparable results. Thus, CD3/CD28-activated primary mouse naive T cells treated with purified human CRP expressed increased IL-4 and decreased IFN-γ at both the mRNA (Fig. 3C) and protein levels (Fig. 3D). Additionally, IL-5 was found to be also significantly upregulated by CRP treatment (data not shown). Without CD3/CD28 activation, however, naive T cells barely secreted IL-4 and IFN-γ, regardless of the presence or absence of CRP (not shown), indicating that CRP alone is incapable of evoking effector programs in naive T cells, which instead require Ag and costimulatory signals. Further experiments revealed little impact of CRP on the proliferation of CD3/CD28-activated naive T cells (data not shown). These findings thus demonstrate that CRP is able to directly regulate the cytokine responses of activated naive mouse CD4+ T cells.
Because IFN-γ and IL-4 production is influenced by CRP and because these are Th1 and Th2 cell signature cytokines (36, 37), respectively, we reasoned that CRP binding may also influence Th1/Th2 differentiation of naive CD4+ T cells. The fate of Ag-activated naive T cells is dictated primarily by specific master cytokines, such as Th1-inducing IL-12 (38) and Th2-inducing IL-4 (39). Accordingly, we found that CRP treatment suppressed phosphorylation of STAT4 and STAT1 downstream of IL-12 signaling (38) whereas it augmented phosphorylation of STAT6 evoked by IL-4 (39) (Fig. 3E). Moreover, the inclusion of CRP nearly halved the number of IFN-γ+IL-4− cells generated under Th1-polarizing conditions (41.2 ± 6.6% decrease for CRP early treatment compared with vehicle controls, p < 0.001; n = 7) whereas it increased the number of IFN-γ−IL-4+ cells by ∼40% under Th2-polarizing conditions (40.8 ± 6.0% increase for CRP early treatment compared with vehicle controls, p < 0.001; n = 7) (Fig. 4A). Parallel and consistent changes were also observed in the expression of lineage-specific transcription factors (Fig. 4B) and STAT phosphorylation (Fig. 4C).
The results shown above demonstrate that in vitro, CRP is able to directly bind primary mouse naive T cells and thereby regulates their effector responses. To determine whether human CRP alters the Th1/Th2 balance in vivo during homeostasis, we analyzed splenocytes harvested from healthy adult wild-type versus CRPtg mice (30) housed under conventional conditions. Splenocytes recovered from CRPtg versus wild-type mice contained nearly equal proportions of CD3+CD4+ double-positive cells and CD3+CD4+CD62L+ triple-positive cells (Fig. 4D). Importantly, however, following PMA/ionomycin activation, fewer T cells from CRPtg mice expressed the Th1 signature cytokine IFN-γ whereas there was no effect of genotype on the number of cells producing the Th2 signature cytokine IL-4 (Fig. 4D). Although the observed difference between genotypes did not achieve statistical significance, in light of the in vitro data described above the results do support a role for CRP in the direct modulation of Th1/Th2 differentiation in vivo.
CRP-mediated suppression of the Th1 response in MOG-induced EAE
In the aforementioned in vitro studies CRP was present in cell cultures throughout the Th1/Th2 polarization process. This “early treatment” should mimic the homeostatic state. In the context of disease, however, because any rise in CRP serum levels necessarily lags behind the onset of inflammation, prominent actions of CRP on T cell differentiation might be manifest at a later stage. We modeled this scenario in vitro by adding CRP to naive T cells 24 h after the initiation of Th1 or Th2 polarization, an approach we term “late treatment.” Interestingly, we found that the effect of late treatment was more pronounced than that of early treatment, with the former leading to a maximum of ∼67% inhibition of Th1 differentiation (p < 0.01; n = 4) and ∼71% enhancement of Th2 differentiation (p < 0.05; n = 5) at the doses we tested (Fig. 4A, top and bottom panels, respectively, Supplemental Fig. 4A). Importantly, with the late treatment regimen the effect of CRP on both Th1 and Th2 cell differentiation were dose-dependent, and an effect on Th1 polarization was obvious even with as little as 1 μg/ml CRP (Supplemental Fig. 4A). In stark contrast to the early and late treatment scenarios, “pre-treatment,” wherein CRP was incubated with naive T cells for 24 h under resting conditions but was absent during the following Th1 or Th2 polarization (a nonphysiological scenario), had little effect (Fig. 4A).
To extend our findings to an in vivo setting wherein T cells are known to be integral and wherein CRP has been shown to play a beneficial role, we immunized C57BL/6 mice with MOG peptide to induce EAE, followed 2 d later by a single s.c. injection of CRP (6, 7). Splenocytes were collected 7 d later to assess Th1/Th2 balance. We found that splenocytes from MOG-immunized and CRP-treated mice secreted 40% less IFN-γ and ∼5-fold more IL-4 than did splenocytes from MOG-immunized/vehicle-treated mice (Fig. 5A). Moreover, the proportion of MOG-reactive CD4+IFN-γ+ T cells recovered from the spleens was reduced by 60% for MOG-immunized and CRP-treated mice (p < 0.001; n = 3) (Fig. 5B). Only a few MOG-reactive CD4+IL-4+ T cells were recovered, likely because of the strong Th1-polarizing milieu created by MOG immunization (40). Importantly, the weaker Th1 response seen early after s.c. treatment with purified human CRP was associated with a significant reduction in severity of ensuing EAE (Fig. 6A). At the peak of the disease, a reduction in the proportion of MOG-reactive splenic Th1 cells (Fig. 6B) and their impaired ability to secret IFN-γ upon activation ex vivo (Fig. 6C) were also noted. Corroborating these results, we further found that CRP treatment decreased T-bet whereas it increased GATA-3 levels in splenocytes of mice with EAE (Fig. 6D). We did not directly assess Th17 cells in these experiments but because the encephalitogenicity of Th17 cells has been shown to be dependent on T-bet (41), weakening of the Th17 response might also contribute to the protection of CRP against EAE. Finally, the number of cells infiltrating the spinal cord and the extent of spinal cord demyelination were dramatically less in CRP-treated mice compared with vehicle-treated mice (not shown). In CRPtg mice with EAE, wherein disease is less severe compared with wild-type mice (not shown and Refs. 6, 7), significantly fewer CD3+CD4+IFN-γ+ T cells (p < 0.05) were recovered from the spinal cord (Fig. 6E). Collectively, these data suggest that the observed capacity of CRP to suppress Th1 responses in vitro is recapitulated in vivo, and that CRP’s ability to alleviate EAE likely involves inhibition of the pathological Th1 response.
CRP is an ancient soluble PRR whose emergence predates that of the adaptive immune system. Therefore, it should not be surprising if this protein is found to have an impact on adaptive immunity. Unlike the cell type–restricted expression of membrane PRRs such as TLRs, the secretion of CRP by hepatocytes into the blood and lymph should, in principle, allow it direct access to a variety of leukocytes in both the circulation and in inflamed tissues. Although the direct interaction of CRP with T cells was investigated previously, these studies yielded conflicting results (22–25). Indeed, little evidence supports the expression by T cells of established CRP receptors, including FcRs (28) and LOX-1 (42). Consequently, it seems more likely that CRP modulation of T cell responsiveness is indirect and mediated by innate immune cells that do express one or more of the receptors for CRP (4). However, in the present study we provide compelling evidence that CRP in its native conformation binds directly to human Jurkat and primary mouse naive T cells via an as yet unidentified receptor, and it thereby alters T cell differentiation. The failure to detect CRP binding to Jurkat T cells in previous work (34) might have been due to the use of an inappropriate Ab for CRP detection, the epitope of which might have been masked when CRP was bound to the cell. Indeed, we found that mAb 1D6 was much more efficient than mAb 2C10 at revealing T cell–bound CRP (see Fig. 1B). Moreover, apparent binding of CRP to live Jurkat T cells has previously been noted, although it was interpreted as nonspecific (43). Based on our new finding that binding of human CRP to CD4+ T cells is largely restricted to the naive CD62L+ subset (see Fig. 2B, 2C), we propose that the inconsistent findings reported about binding of human CRP to T cells (22–25) is most likely due to phenotypic heterogeneity of the T cells used.
Most importantly, to our knowledge, we demonstrate in this study for the first time that CRP is able to directly modulate Th1/Th2 balance, favoring the Th2 effector program while inhibiting Th1 differentiation of TCR-activated naive T cells in vitro (see Figs. 3, 4). In contrast to the rather conditional and localized actions of membrane-bound TLR on T cells, which requires the concurrent presence of its cognate ligands (19–21), the effects of CRP are likely to be tonic and global in vivo because CRP circulates in the blood and lymph and because its interaction with naive T cells does not require additional ligand. Of course CRP undoubtedly can also regulate T cell responses indirectly via its actions on FcR-expressing APCs for example. Indeed, CRP has been shown to promote DC maturation (44), to enhance the uptake of Ags by DCs (45), and to induce cytokine release from monocytes or macrophages (46, 47). Nevertheless, our results show that accessory cells are not an essential requirement. As such, both direct and indirect actions of CRP on T cells could ultimately lead to the suppression of Th1 responses in health and disease. Consistent with this notion, we provide preliminary evidence that the proportion of Th1 cells in healthy CRPtg mice is lower than that in wild-type mice (see Fig. 4D) and that a reduction of pathological Th1 responses in mice with EAE can be achieved by CRP injection or by its transgenic expression (see Figs. 5, 6). Notably, transgenic expression of CRP has also been shown to be protective in mouse models of other autoimmune diseases wherein Th1 cells play a detrimental role (12–14), including spontaneous lupus (5) and collagen-induced arthritis (8). Additionally, the prolonged effect of a single dose of administered CRP observed in this study and previous ones (6, 48) strongly suggests that CRP is able to prominently and permanently remodel the adaptive immune response. A direct action of CRP on naive T cells may be the explanation that ties these observations together, because CRP treatment 2 or 9 d following disease induction is still very effective (see Figs. 5, 6) (6, 44). Notably, in preliminary experiments we also observed a moderate influence of CRP on the response of already differentiated Th1 and Th2 cells (data not shown). Given the recognized importance of Th17 cells in EAE as well as in other autoimmune diseases, future studies will be needed to determine whether CRP influences the generation of Th17 cells.
CRP is not the panacea for all T cell–mediated diseases. In fact, under certain circumstances CRP’s ability to modulate T effector cell balance might prove to be detrimental. For example, in asthma and cancer, CRP-mediated augmentation of Th2 responses would be predicted to promote disease (15, 49). Notwithstanding these caveats, there is growing evidence that CRP is a tonic regulator of the adaptive immune response and this will need to be considered if either purified CRP or drugs designed to modulate the level of CRP (50, 51) are to be used in patients. In conclusion, CRP can directly affect both innate and adaptive immunity and these capacities are fine-tuned by the protein’s conformation (2, 32, 52, 53), location (54), and concentration (1, 4). This diversity of traits, although no doubt accounting for much of the seemingly contradictory effects of CRP reported in the literature, underlies the role of CRP as a central node in the regulatory network of inflammation and immunity (9, 10).
We thank Liang Peng, Li Xie, and Jing Zhao for excellent technical assistance.
This work was supported by Ministry of Science and Technology of China Grant 2011CB910500; National Natural Science Foundation of China Grants 31470718, 31222015, 31270813, 31170696, and 30930024; Ministry of Education of China Grant PCSIRT: IRT1137; and by National Institutes of Health Grants F31NS081903 and R01DK099092.
The online version of this article contains supplemental material.
Abbreviations used in this article:
experimental autoimmune encephalomyelitis
myelin oligodendrocyte glycoprotein
pattern recognition receptor.
The authors have no financial conflicts of interest.