Our human CRPtg have been fully described elsewhere (13, 14). In brief, CRPtg (C57BL/6 background) carry a 31-kb human DNA fragment encoding the CRP gene and all the cis-acting elements required for tissue specificity and acute phase inducibility, while the trans-acting factors required for its human-like pattern of regulation are conserved from mouse to man (13, 14). Consequently, unlike WT, CRPtg exhibit a robust human CRP acute phase response during inflammation. FcγRIIB deficient mice (FcγRIIB^−/−; B6.129S4-Fcgr2b^tm1TtK N12) (15) were purchased from Taconic Farms (Germantown, NY). 2D2 mice [C57BL/6-Tg(Tcra 2D2, Tcrb 2D2) 1Kuch/J] (16) are transgenic for a T cell receptor (TCR) that recognizes residues 35–55 of myelin oligodendrocyte glycoprotein (MOG_35–55) and were purchased from Jackson Laboratories (Bar Harbor, ME, USA; JAX 006912). OT-II mice [B6.Cg-Tg(Tcra Tcrb)425Cbn/J] (17) are transgenic for a TCR that recognizes residues 323–339 of ovalbumin (OVA_323–339) and were purchased from Jackson Laboratories (Bar Harbor, ME, USA; JAX 004194). FcγRIIB^−/− mice expressing a human FCGR2B transgene driven by a mouse CD11c minimal promoter (_cd11cFcγRIIB^hu) were generated herein and are fully described in the Section “Results.” To date, no embryonic lethality or unusual phenotype has been observed for _cd11cFcγRIIB^hu. C57BL/6 mice (WT) were obtained from the Jackson Laboratories (Bar Harbor, ME, USA; JAX 000664). All mice were housed in the same vivarium at constant humidity (60 ± 5%) and temperature (24 ± 1°C) with a 12-h light cycle (6:00 a.m. to 6:00 p.m.), and maintained ad libitum on sterile water and regular chow (Harlan Teklad). Mice were 8–12 weeks old when used and both sexes were combined unless specifically noted. All animal use protocols were approved by the Institutional Animal Care and Use Committees at the University of Alabama at Birmingham and were consistent with the Guide for the Care and Use of Laboratory Animals; Eighth Edition (NIH Academies Press, 2011).

Bone marrow progenitors were grown under conditions known to drive DC generation and expansion (18, 19). Briefly, BM was harvested from femurs, the red blood cells lysed (Hybri-Max Red Blood Cell Lysing Buffer; Sigma, Salem, MA, USA), and the marrow passed through a 70 µM cell strainer and brought to single-cell suspension in RPMI 1640 (Gibco, Grand Island, NY) containing 5% fetal bovine serum (Gibco), 1% Penicillin/Streptomycin (Gibco), 2 mM GlutaMAX™ (Invitrogen), non-essential amino acids (Gibco), 55 µM β-mercaptoethanol (Gibco), and 20 ng/ml granulocyte macrophage-colony stimulating factor (Shenandoah Biotechnology, Warwick, PA, USA). BM progenitors were then added to 12-well tissue culture plates (1 × 10⁶ cells in 1 ml per well) that were incubated at 37°C, 5% CO₂ for 7 days. The culture medium was replaced on days 3 and 5. On day 5, cells were exposed to 50 µg/ml of highly purified human CRP (endotoxin and azide-free CRP from US Biological; Salem, MA, USA), purified chicken OVA_323–339 peptide (MISC-011; CPC Scientific, San Jose, CA, USA), or purified MOG_35–55 peptide (12668-01; Biosynthesis Inc., Lewisville, TX, USA). OVA_323–339 and MOG_35–55 loaded BMDCs were subsequently used in BMDC:T cell co-cultures with OT-II and 2D2 T cells, respectively, as described below. To trigger BMDC maturation in some experiments LPS from Escherichia coli, serotype 055:B5 (Sigma Aldrich) was added (1 µg/ml) on day 6. Alternatively, culture medium was supplemented with 100 ng/ml interleukin-4 (IL-4; Shenandoah Biotechnology, Warwick, PA, USA). IL-10 and IL-12p70 production was assessed by ELISA (88-7105-22 and 88-7121-22; Invitrogen, Eugene, OR, USA) according to the manufacturer protocol. Flow cytometry was performed on a BD LSR-II cytometer (described below) and, after excluding dead cells and aggregated cells, BMDCs were identified as CD11b⁺ CD11c⁺ cells. For detailed analysis of cell death, cells were stained with Annexin V and 7-AAD and were defined as early apoptotic (Annexin V⁺ 7-AAD⁻), late apoptotic (Annexin V⁺ 7-AAD⁺), necrotic (Annexin V⁻ 7-AAD⁺), or live (Annexin V⁻ 7-AAD⁻).

From OT-II and 2D2 mice, the spleens and lymph nodes (axillary, brachial, inguinal) were harvested and mechanically homogenized, the red blood cells lysed, and the homogenate passed through a 70 µM cell strainer, and brought to single-cell suspension in media at 1 × 10⁸ cells/ml. CD4⁺ T cells were enriched by negative selection according to the manufacturer’s guidelines using a kit from StemCell Technologies (Vancouver, BC, Canada). Enriched CD4⁺ T cells were then stained for 20 min with 1 µM CellTrace™ carboxyfluorescein succinimidyl ester (CFSE; Invitrogen, Eugene, OR, USA). BMDCs, cultured as described above, were treated with MOG_35–55 or OVA_323–339 peptide on day 6. On day 7, the peptide-loaded BMDCs were mixed with the freshly isolated and CFSE-stained CD4⁺ T cells (1:5 ratio in triplicate), placed into 96-well round bottom plates, and incubated for 3 days before analysis of CD4⁺ T cell proliferation (dilution of CFSE). BMDC:T cell co-cultures exposed to plate-bound anti-CD3ε and soluble anti-CD28 antibodies (both from Biolegend, San Diego, CA, USA) served as positive controls.

Cells were washed with PBS, spun down at 300 × g for 5 min at 4°C, stained with the viability dye eFluor 780 (eBioscience, San Diego, CA, USA) for 30 min at room temperature, fixed in Fixation Buffer (Biolegend, San Diego, CA, USA) for 10 min at room temperature, blocked with anti-mouse CD16/32 (clone 93; eBioscience) at 4°C for 15 min, and stained with specific antibodies at 4°C for 30 min. For BMDCs, we used anti-mouse CD11c (clone N418), MHC class II IA/IE (clone M5/114.15.2), CD40 (clone HM40-3), CD80 (clone 16–10 A1), CD86 (clone GL-1) (all from Biolegend), and FcγRIIB (clone AT 130-5, Bio Rad, Hercules, CA, USA), and anti-human FcγRIIB (clone AT 10, AbD Serotec, Raleigh, NC, USA). For T cells we used anti-mouse CD4 (clone RM4-5) (Biolegend). Stained and labeled cells were run on a BD LSR-II cytometer and the acquired data analyzed using BD FACSDiva version 6.1.3 and FlowJo version 10.3. For all gating analyses, debris was gated out using a FSC by SSC dot plot, followed by selection of single cells using a SSC-A by SSC-H dot plot, and live cells were selected based on the viability dye eFluor 780 dot plot. For assessment of T cell proliferation (CFSE dilution), the bounds for the CD4⁺ CFSE⁺ “parents” gate was determined using unstimulated T cells and the bounds for the “progeny peaks” were based on anti-CD3ε/anti-CD28 stimulated T cells (see Figure 3A). As T cells divide, the progeny:parent ratio increases.

Experimental autoimmune encephalomyelitis was induced as we described previously (5, 6, 20). Briefly, 10–12-week-old mice were immunized subcutaneously with 150 µg MOG_35–55 emulsified in Freund’s complete adjuvant plus 400 µg heat-killed Mycobacterium tuberculosis (Difco, Detroit, MI, USA). On days 0 and 2, mice received an intraperitoneal injection of 200 ng pertussis toxin (List Biological Laboratories, Campbell, CA). For 30 days thereafter the development of EAE was monitored daily. EAE symptoms were scored on a clinical scale ranging from 0 to 6 as follows: 0, asymptomatic; 1, loss of tail tone; 2, flaccid tail; 3, incomplete paralysis of one or two hind limbs; 4, complete hind limb paralysis; 5, moribund (at which case the mouse was humanely euthanized); 6, dead. For mice that developed EAE, the day of onset was defined as the first of two consecutive days, wherein the clinical score was ≥2.

Raw data were pooled and are expressed graphically as the mean ± SEM or SD, as noted. Group comparisons were done using one-way analysis of variance (ANOVA) followed by post hoc Bonferroni’s and Tukey’s multiple comparison tests, or using linear trends tests. Differences were considered significant when p was <0.05. For EAE, the maximum clinical score achieved by each animal during the 30-day observation period was used to calculate the average maximum clinical score (a measure of severity). To study the time-course of disease, average clinical scores were calculated and plotted daily for each group of mice, and cumulative disease index (CDI) was calculated by area under the curve analysis. Statistical analyses were done using GraphPad Prism version 7.00.

We first examined the influence of human CRP on the generation of DCs from BM progenitors. On day 7 of culture, nearly 90% of all cells were viable (dashed horizontal lines in Figure 1A) and the cultures routinely achieved a yield of nearly 75% BMDCs (dashed horizontal line in Figure 1B). Whether CRP at 10 or 100 µg/ml was added on day 0 or 6 of culture it had no significant effect on cell viability (Figure 1A). CRP treatment also had no effect on the proportion of early apoptotic, late apoptotic, and necrotic BM cells (data not shown). However, CRP treatment did significantly decrease (by 10–15%) the proportion of CD11b⁺ CD11c⁺ BMDCs that developed (Figure 1B). Notably, when CRP was added at the initiation of culture, the inhibitory effect on the final yield of BMDCs was strongest (Figure 1B) and was dose-dependent (Figure 1C). These results show that while CRP has no significant influence on the viability of cultured BM progenitors, it does significantly impede the generation of CD11b⁺ CD11c⁺ BMDCs in both a temporal and dose-dependent manner.

Next, we assessed the influence of CRP on activation/maturation of BMDCs. Treating immature BMDCs with CRP (50 µg/ml) had no effect on their surface expression of MHC class II, CD86, CD40, and CD80 (Figure 2A), whereas treatment of immature BMDCs with LPS (1 µg/ml) significantly upregulated these markers (Figure 2A), indicative of BMDC maturation. While CRP did not trigger BMDC maturation, CRP did significantly inhibit the LPS-triggered increase in surface expression of MHC class II and the co-stimulatory markers, CD86 and CD40 (Figure 2A). Also, the suppressive effect of CRP on LPS-triggered BMDC maturation was dose-dependent, as evidenced by a stepwise reduction of MHC class II, CD86, and CD40 (Figure 2B). This suppressive effect was specific as CRP had no effect on the expression of CD80, CD11b, or CD172a (Figure 2B). Finally, BMDCs treated with LPS (1 µg/ml) robustly produced both the T cell suppressive cytokine IL-10 and the T cell stimulatory cytokine IL-12p70 (225.7 ± 8.8 and 1245.8 ± 191.0 ng/ml, respectively), but the production of both cytokines was significantly suppressed by CRP (no detectable IL-10 and 773.2 ± 13.2 ng/ml IL-12p70; p < 0.05, t-tests). These data demonstrate that CRP dose-dependently prohibits LPS-triggered activation/maturation of BMDCs and limits their production of IL-10 and IL-12p70, cytokines with pleiotropic effects in immunoregulation.

We next sought to determine if the observed effects of CRP on BMDC activation/maturation phenotype and cytokine production affects their T cell stimulatory function. We found that CRP (1–100 µg/ml) had no significant effect on the proliferation of OT-II T cells co-cultured with BMDCs in the absence of any stimulus (Figure 3B; nil) or in the presence of T cell activating antibodies (Figure 3B; CD3/CD28). Importantly, however, when BMDCs loaded with OVA_323–339 peptide were used as APCs, the addition of CRP caused a dose-dependent inhibition of OT-II T cell proliferation (Figure 3B; OVA). Using the MOG TCR-transgenic model (2D2) we obtained similar results, i.e., CRP (50 µg/ml) significantly inhibited the proliferation of 2D2 T cells co-cultured with BMDCs loaded with MOG_35–55 peptide (Figure 3C). These data confirm that CRP’s prohibition of BMDC activation/maturation and cytokine production reduces their ability to stimulate antigen-specific T cell proliferation. The fact that in both model systems, CRP had no effect on T cells directly stimulated with anti-CD3ε/anti-CD28 antibodies shows that CRP’s influence on T cell proliferation must be via its actions on BMDCs.

C-reactive protein binds to both activating and inhibitory Fc receptors, thereby triggering a diversity of cellular responses in vitro (2, 21) and many of the in vivo biological actions of human CRP in CRPtg are fully supported by FcγRIIB (6, 22). Since FcγRs per se, and FcγRIIB in particular, are widely expressed by both human and mouse DCs (7), we generated DCs using FcγRIIB^−/− BM to test if CRP’s influence on BMDC phenotype and function required FcγRIIB. Like the expression on immature WT BMDCs (Figure 2A), expression of MHC class II, CD80, CD40, and CD86 on immature FcγRIIB^−/− BMDCs was unaffected by CRP alone (50 µg/ml), and LPS triggered their increase (Figure 4A). However, in stark contrast to its effect on LPS-matured WT BMDCs (Figure 2A), CRP did not impair the LPS-triggered upregulation of MHC class II, CD86, and CD40 by FcγRIIB^−/− BMDCs (Figure 4A). Like for WT BMDCs, expression of IL-10 by LPS-treated FcγRIIB^−/− BMDCs (308.5 ± 12.5 ng/ml) was lowered by CRP (69.6 ± 8.7 ng/ml). However, unlike for WT BMDCs, for FcγRIIB^−/− BMDCs treated with LPS the amount of IL-12p70 produced (948.9 ± 25.3 ng/ml) was not reduced by CRP (1017.1 ± 51.6 ng/ml). These findings strongly suggest that FcγRIIB expression is required for CRP to prohibit LPS-induced activation/maturation of BMDCs and to suppress production of the T cell stimulatory cytokine IL-12p70. As expected, when MOG_33–55 peptide-loaded FcγRIIB^−/− BMDCs were used as APCs, CRP (50 µg/ml) did not impair their proliferation (Figure 4B). In our hands, FcγRIIB^−/− BMDCs did not stimulate OT-II T cell proliferation even when loaded with OVA_323–339 (data not shown), precluding us from assessing if CRP requires FcγRIIB in the OT-II model system. Nevertheless, the results from the 2D2 model confirmed that CRP’s ability to prohibit BMDC stimulation of an antigen-specific T cell response is facilitated by FcγRIIB expressed on BMDCs.

The apparent requirement of mouse FcγRIIB for human CRP-mediated prohibition of BMDC activation/maturation and 2D2 T cell proliferation prompted us to investigate this biology further. Accordingly, we generated FcγRIIB^−/− mice that express a human FCGR2B transgene. Expression of the human FcγRIIB receptor was restricted to DCs by using a vector that contains the CD11c minimal promoter (kindly provided by Dr. Thomas Brocker, Institute for Immunology, LMU Munich Goethestr. 31, D-80336 Munich, Germany) (8). Briefly, a full-length cDNA clone encoding human FCGR2B (23) was inserted into the vector (Figure 5A) to drive FCGR2B expression on CD11c⁺ DCs in all mouse tissues. Transgenic mice (_cd11cFcγRIIB^hu) were then established by injecting the construct directly into fertilized FcγRIIB^−/− eggs in the UAB Transgenic & Genetically Engineered Models Core. Offspring were screened for presence of the human transgene by PCR and flow cytometry was used to detect surface expression of human FcγRIIB on peripheral blood mononuclear cells (Figure 5B, left). Of the three potential founders identified (M27-1, F6-5, and F6-4; Figure 5B, left), only one (F6-5) showed germline transmission of the transgene. Transgenic descendants of F6-5 showed uniform expression of human FcγRIIB (Figure 5B, right) and were used for all further experiments.

We generated _cd11cFcγRIIB^hu BMDCs and confirmed that they upregulated expression of MHC class II, CD86, CD40, and CD80 after LPS-triggered activation/maturation (Figure 6A) and that CRP alone had no effect on expression of these markers (Figure 6A). Expression of human FcγRIIB partly reconstituted the CRP prohibitory effect on BMDC maturation, i.e., upon LPS-stimulation, CRP prohibited the expression of MHC class II and CD40 (Figure 6A). CRP inhibited IL-10 production by LPS-stimulated _cd11cFcγRIIB^hu BMDCs (459.4 ± 3.1 ng/ml without CRP and no detectable amounts with CRP), but not IL-12p70 production (485.9 ± 94.8 ng/ml and 689.9 ± 235.9 ng/ml without or with CRP, respectively). Although the effect was not significant (ns), when MOG_35–55 peptide-loaded _cd11cFcγRIIB^hu BMDCs were used as APCs, their ability to stimulate the proliferation of 2D2 T cells was reduced by CRP (Figure 6B). These data generally support the premise that CRP’s influence on DCs requires their expression of FcγRIIB, since some of the effects of CRP on FcγRIIB^−/− BMDCs are recovered by expression of human FcγRIIB. Interestingly, although reconstitution of FcγRIIB^−/− BMDCs with human FcγRIIB restored their ability to promote OVA_323–339-driven OT-II T cell proliferation, CRP (50 µg/ml) did not have a significant effect (data not shown).

We had previously shown that CRPtg undergoing EAE have delayed onset and reduced severity of disease compared to WT and that this beneficial effect of CRP is FcγRIIB-dependent (5, 6, 20, 25), and herein we provide new evidence that this FcγRIIB-dependency extends to BMDCs in vitro. Moreover, although not all the observed effects of human CRP on BMDCs were supported by human FcγRIIB, CD11c-specific expression of human FcγRIIB was sufficient to fully reconstitute human CRP’s beneficial actions in EAE (Figure 7; Table 1). Given that human CRP can utilize human FcγRIIB expressed by CD11c⁺ cells in transgenic mice, it is possible that the same or a similar CRP→FcγRIIB pathway operates in humans to regulate tolerance and prevent autoimmunity.