Hepatitis B Virus Blocks the CRE/CREB Complex and Prevents TLR9 Transcription and Function in Human B Cells

This article is featured in In This Issue, p.2191

Infection with human hepatitis B virus (HBV) in immunocompetent adults results in transient liver disease, leading to resolution of hepatitis B infection. However, 5–10% of adults and 80–90% of children develop a chronic infection (1). Over 240 million people worldwide are chronically infected with HBV and are at high risk to develop liver cirrhosis and hepatocellular carcinoma (2). Chronic HBV infection (CHB) occurs because the antiviral immune response is insufficient. Although ineffective T cell and Ab responses have been demonstrated, the precise events that may contribute to insufficient B cell responses remain to be determined (3). B cells play an essential role in humoral immunity as well as capturing Ags for presentation, producing immunomodulatory cytokines, and influencing T cell and dendritic cell (DCs) responses (4). B cell function, expansion, and differentiation are controlled by three signals: the first two are BCR cross-linking and CD40/CD40L signaling, and the third signal is delivered by TLR7 or TLR9 in response to infection. Combination of the three signals allows for optimal Ab responses toward T cell–dependent Ags (5).

TLR9 recognizes DNA sequences from bacteria or viruses in the form of unmethylated CpG motifs. In humans, TLR9 expression is restricted to plasmacytoid DCs (pDCs) and B lymphocytes (6). Upon TLR9 stimulation by distinct classes of CpG oligonucleotides (ODN), pDCs are activated to produce IFN-α/β and various chemokines, and B lymphocytes proliferate, differentiate, and secrete cytokines and Abs. The B cell population in the peripheral blood is composed of naive, memory, and plasma cells. Naive and memory B cells produce cytokines such as IL-2, IL-4, TNF-α, and IL-6 in response to TLR9 stimulation. Following development, B cells migrate to peripheral lymphoid organs and recirculate in blood where they require external signals for survival and constitute IgD⁺ naive mature B cells reservoir. Cells that recognize their cognate Ag are activated and proliferate and differentiate into CD27⁺ memory B cells or effector cells [i.e., CD38^hi Ab-producing plasma cells (7)].

HBV is primarily hepatotropic, yet accumulating evidence strongly supports its extrahepatotropic nature. In fact, the dsDNA genome of HBV as well as viral transcripts and proteins have been found in the PBMCs, including B cells from CHB carriers (8). The hepatitis B surface Ag (HBsAg) is the most abundant HBV protein in the liver and peripheral blood of patients with CHB. Numerous studies have shown direct interactions between HBV-encoded proteins and innate immune responses, suggesting that HBV has developed multiple mechanisms to counteract the host cellular defense. Antiviral immune responses by pDCs, Kupffer, NK, and T cells are reduced because of HBV infection (9–11). Furthermore, several reports have shown specific interactions with viral proteins from HIV, EBV, and human papillomavirus (HPV) with regulatory molecules that impair TLR9 expression and/or signaling (12, 13). HBV has been shown to specifically impair TLR9, but not TLR7, function in pDCs by decreasing the amount of IFN-α upon TLR9 stimulation (14). By contrast, very little is known about the impact of HBV or HBV components on B cell physiology. Therefore, we tested the effect of HBV infection on TLR9 cell expression and function on human B cells using complementary in vitro systems and cohorts of CHB patient samples.

Human blood from healthy subjects was obtained from the French blood agency (Établissement français du sang, Lyon, France). Human PBMCs were separated from peripheral blood of healthy donors (HD) by gradient centrifugation on Ficoll (Eurobio Laboratoires et AbCys) at room temperature (RT). For the clinical study, cells were suspended in heat-inactivated FCS with 20% DMSO, progressively cooled down to −80°C, and stored in cryotubes in liquid nitrogen. B cells were negatively selected using human B Cell Isolation Kit II (Miltenyi Biotec). Purity of sorted B cells (95–99%) was analyzed by staining with anti-CD19 mAb (BD Biosciences) by flow cytometry (LSR II and LSRFortessa; BD Biosciences). PBMCs from CHB carriers (Limoges samples) were obtained with informed consent from each patient with the procedure approved by the local ethics committee (Comité de Protection des Personnes, Centre Hospitalier Universitaire Limoges, Limoges, France). Patients were recruited according to several inclusion and exclusion criteria detailed in Table I. Demographic characteristics and clinical features of recruited patients are described in Table II. The majority of CHB patients were HBeAg negative (ratio HBeAg < 1).

Flow cytometry determined the expression of distinct surface molecules on PBMCs. B cell subsets markers were used to differentiate between the different B cell populations. Cells were simultaneously immunostained using the following anti-human Abs: CD19-PE-CF594 (BD Biosciences), IgD-PerCP-Cy5.5, CD27-APC-Cy7, and CD38-PE-Cy7 (all from BioLegend). In brief, cells were stained for 30 min at 4°C in the dark. After two additional washing steps, cells were stained with LIVE/DEAD Fixable Blue Dead Cell Stain Kit (Life Technologies) for 30 min at 4°C in the dark to assess viability. After two additional washing steps, cells were incubated with 100 μl of BD Cytofix/Cytoperm solution (BD Biosciences) for 20 min at 4°C in the dark and then washed twice with BD Perm/Wash Buffer prior to being exposed to human TLR9-PE Ab (Affymetrix eBioscience) for assessment of TLR9 expression. Cells were resuspended in FACS buffer and analyzed on a BD LSR II. Fluorescence Minus One controls were used to set the gates. FlowJo 7.4.5 software (Tree Star) was used for analysis.

HBV inoculum was a 100-fold–concentrated culture supernatant from HepG2.2.15 cells or HBV-infected HepaRG cells. Briefly, supernatants were clarified, layered onto a sucrose cushion (10 and 20% in 20 mM Tris [pH 8]) and centrifuged at 4°C for 16 h at 25,000 rpm using a SW41 Rotor (Beckman Coulter). HBV inoculum was also obtained following concentration using Centricon Plus-70 (Biomax-100; MilliporeSigma). Similar procedure was applied to HepG2 or HepaRG cells supernatants to generate the mock lysate. HBV stock titer (genome equivalent per milliliter) was assessed using real-time PCR. After DNA extraction (QIAamp UltraSens Virus Kit; Qiagen), HBV inoculum was titrated by quantitative PCR (qPCR) with forward 5′-GCTGACGCAACCCCCACT-3′ and reverse 5′-AGGAGTTCCGCAGTATGG-3′ probes using a standard curve from a quantified HBV-encoded plasmid. All preparations were tested for the absence of endotoxin (Lonza Verviers).

Recombinant HBsAg (adw subtype) and HBeAg were purchased from Jena Bioscience and were used at 40 μg/ml. Viral particles and HBV virions (Dane particles) were obtained from David Durantel’s laboratory (U1052; CRCL). HBsAg productions were used at 5 μg/ml, HBeAg at 4.5 μg/ml, and 400 viral genome equivalents (v.g.e.) of Dane particles. Endotoxin levels in HBsAg and HBeAg productions were <0.01 EU/ml within HBV stocks (Limulus Amebocyte Lysate Assay; BioWhittaker).

PBMCs, primary B cells and RPMI 8226 cells, were cultured in RPMI 1640 medium (Life Technologies; Invitrogen Life Technologies) supplemented with 10% of FBS, 2 mM l-glutamine, 10 μM of ciprofloxacin or penicillin/streptomycin (HCL Technologies), and 1 mM sodium pyruvate (PAA Laboratories).

CpG 2006, CpG 2006 FITC, and imiquimod were purchased from InvivoGen and were used at the concentration of 5 μM according to the manufacturer’s recommendations.

Cells were seeded at 2 × 10⁵ per 200 μl per well and stimulated for 24 h. The supernatant was harvested, and IL-6, TNF-α, and IL-10 (R&D Systems) were measured following the manufacturer’s protocol. Nunc-Immuno Plate MaxiSorp plates (Thermo Fisher Scientific) were coated with the correspondent capture Ab and incubated overnight at 4°C. After blocking with 1% BSA in PBS, the plates were incubated with serially diluted standard samples and then with HRP-conjugated detection Abs. After colorimetric reaction (tetramethylbenzidine; Moss), the absorbance at 450 nm was measured using an ELISA plate reader. The IgG secretion was evaluated in supernatants harvested 120 h after CpG 2006 and imiquimod stimulation using the Human IgG ELISA Quantitation kit (Bethyl Laboratories).

B cells were fixed at 37°C in 4% paraformaldehyde complemented with 0.2% glutaraldehyde. Cells were washed three times in cacodylate (0.2 M)/saccharose (0.4 M) for 1 h at 4°C, dehydrated through a series of washes with 30, 50, and 70% ethanol maintained previously at 4°C for 5 min, and infiltrated with London Resin White (LRWhite) using 1:1 LRWhite and 4°C absolute ethanol for 60 min, followed by pure LRWhite at 4°C for three periods of 60 min each, then embedded in pure LRWhite in gelatin capsules for polymerization at 50°C for 48 h. Ultrathin sections (∼70-nm thick) were cut on a Reichert UltraCut E (Leica Microsystems) ultramicrotome, mounted on 200 mesh nickel grids coated with 1:1000 polylysine, and stabilized for 1 d at RT. Immunogold labeling was performed by flotation of the grids on drops of reactive media. Nonspecific sites were coated with 1% BSA and 1% normal goat serum in 50 mM Tris-HCl (pH 7.4) for 20 min at RT. Thereafter, incubation was carried out overnight at 4°C in a wet chamber with a rabbit anti-HBsAg Ab (M.A. Petit, INSERM U1052; CRCL). Sections were successively washed three times in 50 mM Tris-HCl (pH 7.4 and pH 8.2) at RT. They were incubated in a wet chamber for 45 min at RT in 1% BSA, 50 mM Tris-HCl (pH 8.2) for 20 min at RT and labeled with gold-conjugated secondary Ab (Aurion Resources). Sections were successively washed three times in 50 Mm Tris-HCl (pH 8.2 and pH 7.4) and three times with infiltrated distilled water. The immunocomplex was fixed by a wash in glutaraldehyde 4% for 3 min. Sections were stained with 0.5% uranyl acetate in ethanol 50% for 5 min in darkness and observed with a JEOL JEM-1400 Transmission Electron Microscope operating at 80 kV equipped with a camera Orius 600 Gatan DigitalMicrograph. Analysis was done using Olympus Soft Imaging System.

Total RNA was extracted from cells (RPMI 8226 cells, PBMCs, and B cells) using the RNA/Protein Extraction Kit (MACHEREY–NAGEL) and was reverse transcribed using the First Strand cDNA Synthesis Kit according to the manufacturer’s protocol (Fermentas).

qPCR was performed using the Mx3000P Real-Time PCR System (StrateGene) with MESA GREEN qPCR MasterMix Plus (Eurogentec) and TLR9- and β2-microglobulin–specific primers for relative quantification. PCR assays were conducted for each sample using the primers specific for TLR9, forward: 5′-CGT CTT GAA GGC CTG GTG TTG A-3′, reverse: 5′-CTG GAA GGC CTT GGT TTT AGT GA-3′. Housekeeping gene β2-microglobulin was used. The sequence of the BCL-2 primers was forward: 5′-TGG CCA GGG TCA GAG TTA AA-3′, reverse: 5′-TGG CCT CTC TTG CGG AGT A-3′and for cyclin D1, forward: 5′-AAC TAC CTG GAC CGC TTC CT-3′, reverse: 5′-CCA CTT GAG CTT GTT CAC CA-3′. Relative quantification was performed using standard curve analysis. TLR9, BCL-2, and cyclin D1 expression data were normalized with β2-microglobulin.

Total protein extracts were obtained with the RNA/Protein Extraction Kit (MACHEREY–NAGEL) following manufacturer’s instructions. Protein extracts were quantified with the protein quantification assay (MACHEREY–NAGEL). Abs used for immunoblotting were TLR9, β-tubulin, CREB, p-CREB, p-PKA, protein kinase A (PKA)–RIα/β, and histone H3 (Cell Signaling). Western blots were developed using the Intelligent Dark Box (Fujifilm).

Densitometries were performed on TIFF images of radiographical films exposed to Western blots. We used ImageJ software analysis and assigned values to each lane by integrating the area under the curve.

RPMI 8226 cells have been transiently transfected (1 μg per well) in triplicate using FuGENE HD Transfection Reagent (Promega). Cells were harvested after 48 h, and the activities of both Firefly and Renilla luciferase were measured with a luminometer (AB-2200; ATTO) using a Dual-Luciferase Reporter System (Promega). Firefly luciferase activity was normalized to that of Renilla luciferase.

The pGL3-TLR9 luciferase promoter and the deleted TLR9 promoters were obtained from F. Takeshita (Division of Cellular and Molecular Medicine, National Cancer Center Research Institute, Tokyo, Japan). The cAMP-responsive element (CRE) site in TLR9 promoter (-686TGACGTGG-679) was mutated to (-686TGTGATGG-679) using the QuickChange Lightning Site-Directed Mutagenesis Kit (StrataGene) according to manufacturer’s instructions. All mutations were confirmed by direct sequencing.

EMSA assay was performed using the CREB (1) EMSA kit (Affymetrix eBioscience). For each binding reaction, 10 μg of nuclear extracts was used. Proteins or protein–DNA complexes were detected using ECL kit (GE Healthcare).

The wild-type (WT) (CRE) and mutated CRE ODN used were melted at 95°C then annealed to their respective complementary ODN at RT for 1 h. The ODN sequences were as follows: TLR9pWT-F, 5′-biotin-ATCTGGAGTGACGTGGTGTGTG-3′; TLR9pWT-R, 5′-CACACACCACGTCACTCCAGAT-3′; TLR9pMut-F, 5′-biotin-ATCTGGAGTGGTATGGTGTGTG-3′; TLR9pMut-R, 5′-CACACACCATACCACTCCAGAT-3′. Nuclear extracts prepared from RPMI 8226 cells, previously treated with 400 v.g.e. of HBV for 24 h, were incubated with streptavidin-coupled agarose beads and 500 pmol of each dsODN for 2 h at RT with gentle rocking. The protein–DNA–streptavidin–agarose complex was washed four times with PBS containing protease inhibitors. The precipitates were subjected to immunoblotting using the indicated Abs.

2 × 10⁵ PBMCs per well were stimulated for 7 d with F(ab′)2 Anti-Human IgM⁺IgG (Affymetrix eBioscience), CD40L (soluble human recombinant) (Enzo Life Sciences), IL-4 (Affymetrix eBioscience), and either CpG 2006 or imiquimod (InvivoGen) in flat-bottom 96-well culture plates. Multiscreen plates (Millipore MultiScreen) were precoated with unlabeled Goat Anti-Human Ig (SouthernBiotech). Cultured PBMCs were seeded onto anti-Ig–coated plates at 2 × 105 cells per well and diluted successively to achieve the best dilution factor and incubated overnight. Alkaline phosphatase–conjugated goat anti-human IgG, IgA, and IgM Abs (SouthernBiotech) were used as secondary Abs. Spots were developed using BCIP/NBT (Bio-Rad Laboratories) and counted using the CTL ImmunoSpot Analyzer (Cellular Technologies). The background was accounted for by subtracting the average number of spots in noncoated wells from the spots in Ab-coated wells.

PBMCs (1.5 × 105 per well) were stained with CellTrace Violet (CTV) (Molecular Probes, Invitrogen) fluorescent dye (5 μM) at 37°C for 8 min and were then cultured in completed RPMI 1640 medium in 96 well flat-bottom plates (Nunc). Stimulation involved the use of a mixture consisting of F(ab′)2 Anti-Human IgM⁺IgG (Affymetrix eBioscience) (10 μM), CD40L soluble human recombinant plus enhancer (Enzo Life Sciences) (1 μM), IL-4 (Affymetrix eBioscience) (10 nM), and CpG 2006 (InvivoGen) (5 μM). Analytical flow cytometry was performed with a BD LSRFortessa (BD Biosciences).

Nuclear and cytosolic protein extracts from RPMI 8226 cells were obtained using the Nuclear/Cytosolic Fractionation Kit (BioVision) according to the manufacturer’s protocol.

Prism software (GraphPad) was used for two-tailed paired Student t tests. The p values were *p < 0.05, **p < 0.01, and ***p < 0.001. For the clinical study, statistical analyses were carried out using R software version 3.2.3. Statistical significance was determined by an unpaired, two-tailed Student t test, with p values of *p < 0.05, **p < 0.01, and ***p < 0.001 that were considered statistically significant.

We used a human B cell line (RPMI 8226) as well as freshly isolated primary human B cells to study the effect of HBV on TLR9 expression. Cells were exposed to HBV at 200 and 400 v.g.e. per cell for 24 h, and TLR9 mRNA and protein levels were examined. The HBV inoculum produced in our study contained viral proteins HBsAg, HBcAg, and HBeAg as well as Dane particles (HBV virions). As shown in Fig. 1A, HBV significantly downregulated TLR9 mRNA levels in RPMI 8226 cells at 200 and 400 v.g.e. We next corroborated our findings in primary human blood B cells and observed that HBV inhibited TLR9 mRNA levels (Fig. 1B). Accordingly, the inhibition of TLR9 mRNA levels correlated with a massive decrease in TLR9 protein expression in both RPMI 8226 cells and primary human B cells treated with HBV (Fig. 1C, 1D, respectively). We confirmed the loss of TLR9 expression by HBV in human primary B cells using flow cytometry. Fresh PBMCs were cultured for 24 h with/without HBV; cells were stained for CD19 (a surface total B cell marker) and intracellular TLR9. We observed that the TLR9 mean fluorescence intensity decreased when cells were treated with HBV (Fig. 1E, 1F; p < 0.01). This effect was reversed using a blocking anti-HBs Ab (Fig. 1G) and TLR9 expression was restored, which shows a specific effect of HBV. We next assessed the effect of HBV on TLR9 expression on the different peripheral B cell subpopulations (see gating strategy in Supplemental Fig. 1A). We observed that TLR9 was expressed by the majority of naive B cells, CD27⁺ memory B cells, and plasma cells, but the latter subset displayed the highest expression intensity (Fig. 1H, 1I). This observation is concordant with a previous study that has shown that plasma cells express the highest level of TLR9 (15). Reverse transcription–qPCR also confirmed these results on sorted B cell subsets, showing that TLR9 mRNA was the most abundant in plasma cells and CD27⁺ memory B cells (data not shown). TLR9 expression was blocked in all three subsets when exposed to HBV, predominantly on CD27⁺ memory and plasma B cells (Fig. 1J). No effect on cell viability was observed between HBV- and non-HBV–treated cells (Supplemental Fig. 1B). Our results show that HBV downregulates TLR9 expression in all B cell subsets found in peripheral blood.

HBV covalently closed circular DNA as well as viral transcripts and proteins have been found in PBMCs, including B cells from chronically infected carriers (8). To assess the ability of B cells to internalize HBV virions, primary B cells were exposed to HBV for 24 h, and cells were analyzed by electron microscopy. Internalization of HBV virions was determined by immunogold labeling using anti-HBsAg Abs. Forty-two– to forty-five–nanometer spheres that represent the size of HBV virions (red arrows) were identified within the cytoplasm of B cells (Fig. 2). We also noted that HBsAg (blue arrows) were present in human B cells. These results show that HBV and its HBsAg particles can be efficiently internalized in peripheral blood B cells.

To assess whether HBV alters TLR9 function by HBV in primary B cells, we stimulated freshly isolated B cells with CpG ODN 2006 (henceforth simply referred to as CpG) with/without HBV for 24 h, and IL-6 and IL-8 secretion was measured. CpG-induced IL-6 and IL-8 secretion was significantly reduced in the presence of HBV inoculum, whereas the mock lysate (a control for HBV production) exerted no effect (Fig. 3A, 3B). Human B cells also express TLR7 (16, 17), yet HBV exerted no effect on TLR7-mediated IL-6 secretion (Fig. 3C). As already reported for pDCs, these results showed that HBV specifically targets TLR9 without affecting TLR7 (18). Additionally, we stimulated the TLR9 pathway using HSV-2 and observed that HBV also blocked IL-6 secretion (Supplemental Fig. 2A). Furthermore, the drop in CpG 2006 internalization correlated with the decrease of TLR9 expression when exposed to HBV (Supplemental Fig. 2B, 2C).

We next investigated the effect of HBV on TLR9-induced B cell proliferation. PBMCs were treated with anti-BCR Abs (signal 1) and CD40L (signal 2) with/without CpG (signal 3) with/without HBV for 5 d. B cell proliferation was assessed by following dilution of the CTV dye proliferation marker. TLR9 engagement boosted B cell proliferation induced by BCR and CD40 engagement. However, this effect was virtually abolished in the presence of HBV (Fig. 3D). In fact, the percentage of proliferating B cells upon TLR9 stimulation decreased by 50% following HBV treatment. In contrast, the HBV inoculum did not alter B cell proliferation, induced by the sole triggering of BCR and CD40 (Fig. 3D, 3E). Next, we tested the effect of HBV on Ig secretion by B cells upon TLR9 stimulation. By ELISPOT analysis upon CpG stimulation, we observed that HBV significantly increased the frequency of IgG- and IgA-producing cells upon 7-d culture of B cells with CpG (Fig. 3F, 3G). Increased production of IgG by HBV-stimulated cells was indeed confirmed by an ELISA assay on supernatants of cultured human B cells (Fig. 3H). In summary, we observed that loss of TLR9 expression by HBV leads to reduced proliferation and cytokine secretion but increased Ig production in human B cells.

Previous reports showed that oncoviruses such as HPV16, EBV, and hepatitis C virus (HCV) downregulate TLR9 transcription using NF-κB, which is a negative regulator of TLR9 transcription (12, 13, 19). We hypothesized that HBV used the same mechanism to block TLR9 transcription in human B cells. By transient transfection into RPMI 8226 B cell line, we evaluated TLR9 promoter activity with/without HBV inoculum and observed that HBV blocked TLR9 luciferase activity (Fig. 4A). To investigate which region of the promoter was implicated in TLR9 deregulation by HBV, we used several deletions of the TLR9 promoter (Fig. 4B). We found that deleting the CRE cis site (-686’/TGACGTGG/-679’) on TLR9 promoter eliminated the inhibitory effect exerted by HBV (Fig. 4C).

To further investigate the implication of the CRE region, we performed site-directed mutagenesis of the CRE site on the full-length construct of the TLR9 promoter as well as the del-1 deleted construct (also containing the CRE site). The mutated site is shown in Fig. 4D. As expected, HBV decreased TLR9 WT–promoter activity; however, when the CRE site was mutated, HBV could no longer exert a suppressive effect on TLR9 transcription (Fig. 4E). CRE is a cis element responsible for recruiting the CREB to activate the transcription of target genes. CREB at steady state is expressed in the nucleus of cells. To bind to CRE sites, CREB must be phosphorylated at Ser¹³³. We sought to see if HBV affects CREB or p-CREB levels in B cells, which may affect TLR9 promoter activity. RPMI 8226 B cells were exposed to HBV, and cell fractionation of nuclear and cytoplasmic proteins was performed. As expected, HBV suppressed TLR9 expression in the cytoplasm (Fig. 4F). In the nuclear extracts, we observed that total CREB, as well as p-CREB proteins, were reduced in B cells exposed to HBV (Fig. 4F). Furthermore, EMSA revealed a reduction in the CRE/CREB complex in HBV-treated cells versus nontreated cells (Fig. 4G). Our data were corroborated by an ODN pull-down assay using ODN for the specific CRE site found on the TLR9 promoter (Fig. 4H). In this study, we concluded that HBV affects the TLR9 promoter by abrogating TLR9 transcription specifically via the CRE/CREB pathway. HBV disrupts CREB phosphorylation to prevent binding on the CRE region within the TLR9 promoter.

Several HBV proteins have been shown to alter or suppress innate immune responses (20–22). HBV components include HBsAg (which is expressed on the surface of virions or secreted into the blood of HBV-infected patients), HBcAg (which is the core protein of the virus, enveloping the partially dsDNA of the virus), and HBeAg (which is derived from the precore protein). As the HBV inoculum contains viral components, we decided to determine which viral component downregulates TLR9. Primary B cells were exposed to HBV viral components (HBsAg and HBeAg) as well as Dane particles (HBV virions). We observed that both HBsAg and HBeAg suppressed TLR9 at the protein level, whereas Dane particles (purified virions) had a minor effect on TLR9 levels (Fig. 5A). Both HBsAg and HBeAg also decreased TLR9 promoter activity comparably to HBV inoculum (Fig. 5B). These results showed that HBsAg and HBeAg are involved in HBV-mediated suppression of TLR9 expression in B cells. Indeed, HBsAg, but not HBeAg, lost its inhibitory effect on the TLR9 promoter when the CRE site was mutated (Fig. 5C, Supplemental Fig. 3A). We also observed that HBsAg inhibited CREB phosphorylation similarly to HBV (Fig. 5D). Furthermore, EMSA assay revealed a reduction in the CRE/CREB complex in HBs-treated cells versus nontreated cells (Fig. 5E). As CREB is phosphorylated by PKA upstream of the CRE/CREB pathway, we wanted to assess the effect of HBsAg on the PKA. Human B cells were exposed to HBsAg, and total PKA regulatory subunits α and β, as well as p-PKA protein levels, were examined. We show in Fig. 4F that PKA regulatory subunits α and β, as well as p-PKA levels, were decreased in the presence of HBsAg, which correlated with the reduction in p-CREB and TLR9 levels.

To assess whether HBV altered other CRE/CREB target genes in B cells, we analyzed the mRNA expression of cyclin D1 implicated in the cell cycle (23) and BCL-2 involved in B cell survival and apoptosis control (24). Regulation of both these targets could explain the altering effect of HBV on B cell proliferation as shown in Fig. 3C. HBV suppressed the expression of cyclin D1 and BCL-2, thus confirming the involvement of the CRE/CREB pathway (Supplemental Fig. 3B). In summary, we have shown a new mechanism by which HBV blocks TLR9 transcription. HBV uses HBsAg to alter the phosphorylation of PKA and thus the activation of CREB to prevent it from binding to the CRE site on the TLR9 promoter.

Our results demonstrate TLR9 inhibition by HBV using in vitro models. Next, we wanted to corroborate these results in a clinical cohort; we collected blood samples from CHB patients without any evidence of liver disease and a family history of hepatocellular carcinoma or cirrhosis (Table I). Demographic characteristics and clinical features of recruited patients are described in Table II.

PBMCs were isolated from CHB patients as well as from HD. TLR9 expression was analyzed by flow cytometry on the different B cell subsets. Concordant with our in vitro data, we observed that CHB patients had a significant reduction of TLR9 expression in total B cells (Fig. 6A) as well as on each individual B cell subset (Fig. 6B–D). Next, we investigated if TLR9-mediated functions were also suppressed in CHB patients. TLR9 stimulation with CpG motifs led to the secretion of IL-6 and TNF-α in HD, whereas CpG had virtually no effect on cytokine secretion in CHB patients (Fig. 7A, 7B). We noticed that even the basal secretions of IL-6 and TNF-α in CHB patients were reduced in comparison with controls (Fig. 7A, 7B). IL-10 has been described to be upregulated in the plasma of CHB patients (25), preserving an immune-suppressive state. Interestingly, IL-10 secretion was not affected by HBV infection as we observed comparable amounts of IL-10 between control and CHB groups in the unstimulated cells. However, we observed that when PBMCs of CHB patients are stimulated with CpG, they produce less IL-10 than of the HD. (Fig. 7C).

Next, we wanted to analyze the ability of B cells to proliferate in CHB patients following TLR9 stimulation. To this aim, PBMCs were activated with anti-BCR Abs, CD40L, and CpG for 5 d, and B cell proliferation was analyzed by flow cytometry by gating on CD19⁺ cells. TLR9-mediated B cell proliferation was strongly reduced in CHB patients in comparison with the control group (Fig. 7D). Indeed, chronically infected patients displayed a significant decrease in the percentage of proliferating B cells by nearly 70% (decrease in the percentage of divided B cells). Furthermore, PBMCs from CHB patients displayed an overall reduction of TLR9 mRNA expression in comparison with controls (Fig. 8A). TLR7 mRNA expression, however, was similar between HD and CHB patients (Supplemental Fig. 4A). We did not find any significant disruption in naive and CD27⁺ memory percentages between HD and CHB patients (Supplemental Fig. 4B–D), confirming other studies (26); however, CHB patients had significantly more plasma B cells (Supplemental Fig. 4E).

To assess the implication of CRE/CREB pathway observed in in vitro data, we also analyzed the mRNA expression of cyclin D1 and BCL-2 (27). Although cyclin D1 mRNA levels were not significantly affected by the HBV infection in patients, BCL-2 levels were downregulated in CHB patients compared with the control group (Fig. 8B, 8C). This result suggests the involvement of CRE/CREB pathway in the TLR9 downregulation observed in CHB patients. In summary, these results show that in CHB patients, TLR9 expression and function are significantly decreased in PBMCs and specifically in human B cells.

Several oncoviruses can escape immune recognition by deregulating TLR9 expression and function (13, 28–30). Reported is the ability of each virus to use a unique strategy to target transcription factors that regulate the TLR9 promoter or mRNA stability. LMP-1 from EBV, as well as E7 from HPV16, activated NF-κB, leading to the suppression of TLR9 transcription in B cells and keratinocytes, respectively (13, 28). The T Ag locus of Merkel cell polyomavirus blocked the C/EBPβ transactivator, which also regulated TLR9 transcription (29). Recently, Fischer et al. (19) showed that TLR9 promoter SNPs associated with natural clearance of HCV infection showed higher transcriptional activities. We previously showed that TLR9 expression on pDCs, as well as TLR9-mediated secretion of IFN, was impaired by HBV (14). Furthermore, we showed that TLR9 promoter activity was deregulated in RPMI 8226 cells. However, the effect and mechanism of HBV on TLR9 expression and function in peripheral blood B cell subsets is still unknown.

Our data suggest a novel mechanism by which HBV can block TLR9 transcription and function in human B cells. We demonstrated that HBsAg inhibited p-CREB levels and function, thereby reducing its ability to bind to the CRE site located in the TLR9 promoter. Furthermore, HBV via HBsAg could no longer suppress TLR9 promoter activity when we introduced a mutation on the CRE site. CREB has a well-documented role in neuronal plasticity and long-term memory formation, although emerging evidence has revealed specific functions of CREB in immune cells by inducing macrophage survival as well as promoting the proliferation, survival, and regulation of T and B cells (31). BCL-2 is also regulated by CREB and is implicated in B cell proliferation and inhibiting the actions of proapoptotic proteins (32). BCL-2 mRNA levels in B cells were also reduced by HBV, indicating that the loss of CREB function could also contribute to a decrease in B cell proliferation, an effect we observed in this study. Furthermore, several studies have shown that HBsAg can suppress the release of cytokines by interfering with TLR signaling pathways in monocytes/macrophages, which may contribute to the establishment of chronic infections (10, 33, 34). We show that HBsAg can abrogate PKA levels upstream of CREB. However, further investigation is needed to elucidate how HBsAg can affect PKA as well as CREB phosphorylation in human B cells. HBV has been shown to use CREB/PKA signal transduction pathways in hepatocytes to enhance HBsAg expression during homeostasis and hepatic inflammation (35, 36). However, to our knowledge, this is the first time that CRE/CREB pathway has been shown to downregulate the expression of TLR9 to enhance immune evasion by HBV. Therefore, HBV may use the CRE/CREB pathway to 1) expand its infection in hepatocytes and 2) to abrogate TLR9-mediated B cell functions.

In this study, we showed by electron microscopy that HBV virions, as well as HBsAg, can enter B cells. The nature of the cellular site of attachment involved in virus binding to B cells is under investigation. Although sodium taurocholate cotransporting polypeptide is a potential candidate, as shown for hepatocytes (37), unpublished observations (I. Chemin) indicate that B cells do not express this receptor, and therefore, another cell receptor might be implicated. We found that upon stimulation of the TLR7 pathway, IL-6 secretion is not affected by HBV compared with the control, showing that in B cells, HBV specifically targets TLR9. TLR7 has been shown in several studies not to be affected by HBV or HCV infection. Woltman et al. (38) demonstrated that HBV and HBsAg were unable to suppress loxoribine/TLR7–induced immune responses. We also showed that HBV particle internalization by pDCs did not lead to suppression of TLR7-mediated secretion of IFN-α (14). Moreover, other groups have shown that TLR9 expression and function are altered in CHB patients. Martinet et al. (11) found that pDCs from patients infected with CHB displayed poor responses to stimulation with TLR9 ligands and were unable to induce cytolytic activity of NK cells. Furthermore, mRNA levels of TLR9 were negatively correlated with serum levels of liver enzymes while being positively associated with HBV viral load in patients with CHB (39). IL-6 and TNF-α play antiviral roles and are implicated in limiting HBV replication (40–43). We observed that HBV blocked TLR9-mediated IL-6 as well as TNF-α secretion in PBMCs and B cells, which could contribute to the loss of the antiviral response. Along with the abrogation of IFN observed for pDCs and recently shown immunosuppressive monocytes (44), these data show that several innate immune responses are abrogated in CHB patients.

We also studied the effect of HBV on the expression of TLR9 in different peripheral blood B cell subsets from CHB patients. Consistent with our in vitro data, TLR9 expression was downregulated in naive, CD27⁺ memory B cells, and plasma cells from CHB patients. Whether TLR9 downregulation is also observed on other B cell subsets found in secondary lymphoid tissues in the germinal center and marginal zone would require further studies. Our results show that TLR9 inhibition is not restricted to a particular peripheral B cell subset. HBV infection did not significantly affect the percentages of naive or CD27⁺ memory B cells, but interestingly, the percentage of plasma B cells is increased in CHB patients (Supplemental Fig. 4E), a finding in accordance with other clinical studies (26, 45). We also observed that HBV does not affect TLR9-mediated IgM production but enhanced both IgA and IgG plasma cell numbers and IgG levels. IgA and IgG are mainly produced by plasma B cells subsets; in fact, we did observe a tendency of increased percentages of plasma cells when PBMCs were exposed to HBV for 7 d (Supplemental Fig. 4F). This stimulatory effect of HBV, which supports increased survival of pre-existing plasma cells and/or enhanced differentiation of naive and memory B cells into plasma cells, is surprising given that TLR9 expression is diminished on the three B cell subsets. This may indicate that HBV can increase plasma cell numbers independently of TLR9 engagement or by a mechanism involving a functional TLR9 on non–B cells. Another hypothesis is that the plasma B cells exposed to HBV produced more IgA and IgG than nontreated cells.

Our data highlight a key role for TLR9-mediated B cell responses against hepatitis B. Our results suggest that TLR9 ligands can be considered a powerful objective for adjuvants to use in HBV vaccination, especially in patients who fail to respond to conventional vaccination. Along with antiviral treatment, future studies can be directed to discover new ligands to restore TLR9 expression and thus improve immune responses against HBV in patients with prolonged chronic infection.

We thank the Flow Cytometry Core Facility, SFR BioSciences Gerland UMS3444, and the French blood agency (EFS) for contribution in the clinical studies, especially Sébastien Dussurgey. We also thank the Quantitative Imagery Center Lyon East and Elisabeth Erraduriz-Cerda for the electron microscopy photos.

The authors have no financial conflicts of interest.