1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Chronic hepatitis C virus (HCV) infection is a leading cause of cirrhosis and hepatocellular carcinoma (HCC). Both advanced solid tumors and HCV have previously been associated with memory B-cell dysfunction. In this study, we sought to dissect the effect of viral infection, cirrhosis, and liver cancer on memory B-cell frequency and function in the spectrum of HCV disease. Peripheral blood from healthy donors, HCV-infected patients with F1-F2 liver fibrosis, HCV-infected patients with cirrhosis, patients with HCV-related HCC, and non-HCV-infected cirrhotics were assessed for B-cell phenotype by flow cytometry. Isolated B cells were stimulated with anti–cluster of differentiation (CD)40 antibodies and Toll-like receptor (TLR)9 agonist for assessment of costimulation marker expression, cytokine production, immunoglobulin (Ig) production, and CD4+ T-cell allostimulatory capacity. CD27+ memory B cells and, more specifically, CD27+IgM+ B cells were markedly less frequent in cirrhotic patients independent of HCV infection. Circulating B cells in cirrhotics were hyporesponsive to CD40/TLR9 activation, as characterized by CD70 up-regulation, tumor necrosis factor beta secretion, IgG production, and T-cell allostimulation. Last, blockade of TLR4 and TLR9 signaling abrogated the activation of healthy donor B cells by cirrhotic plasma, suggesting a role for bacterial translocation in driving B-cell changes in cirrhosis. Conclusion: Profound abnormalities in B-cell phenotype and function occur in cirrhosis independent of HCV infection. These B-cell defects may explain, in part, the vaccine hyporesponsiveness and susceptibility to bacterial infection in this population. (HEPATOLOGY 2012)

A complex interaction of hepatitis C virus (HCV) infection and B cells evolves during the natural history of HCV infection. Upon initial infection, virus-specific neutralizing antibody responses develop weeks after initial viremia target hypervariable regions of the HCV envelope proteins, continuously selecting antibody escape variants, an evolution that continues throughout the chronic phase of infection.1, 2 In addition to chronic stimulation of virus-specific B cells, chronic HCV infection is often characterized by a nonspecific polyclonal activation of B cells,3 which has been attributed to interactions between the HCV E2 envelope protein and cluster of differentiation (CD)81, an activating tetraspannin coreceptor that colocalizes with the B-cell receptor complex.4 Despite the activation of virus-specific and non-virus-specific B cells, which could result in the proliferation and accumulation of memory B cells, several studies have demonstrated that the frequency of CD27+ memory B cells is either unchanged5 or modestly reduced in chronic HCV infection.6, 7 Controversy persists as to the fate of memory B cells, with the reduced frequency attributed to the following: (1) increased activation-induced apoptosis,6 a theory that has been contradicted by recent data showing relative resistance to apoptosis of memory B cells in HCV8, 9; (2) increased conversion of B cells into short-lived plasmablasts7; or (3) increased intrahepatic compartmentalization.7, 10

Cirrhosis ultimately evolves in 20%-30% of chronically HCV-infected patients. In cirrhotics, hepatic decompensation eventually develops as a result of progressive portal hypertension, hepatic synthetic insufficiency, and/or neoplastic transformation. Particularly after decompensation, cirrhotic patients are at high risk of invasive bacterial infections, such as spontaneous bacterial peritonitis and bacteremia, likely mediated by reduced production or increased consumption of complement, altered neutrophil function,11 increased intestinal permeability,12 and bacterial translocation.13 B-cell dysregulation might also contribute to this immunocompromised state. Cirrhotic patients exhibit suboptimal seroconversion rates after vaccination with recombinant hepatitis B virus (HBV) vaccine14 and impaired immunoglobulin (Ig)G production after pneumococcal vaccination.15 Despite poor response to vaccination, cirrhosis has been associated with abnormally increased serum levels of pathogen-specific Igs.16–19 Despite these observations, the impact of cirrhosis on B cells has not been thoroughly investigated.

We recently reported that advanced solid tumors, such as melanoma and breast cancer, were associated with marked reductions of peripheral memory B-cell populations and related B-cell hypofunction.20 Unpublished pilot data revealed a similar phenotype in HCV-infected cirrhotics with hepatocellular carcinoma (HCC). In this study, we sought to dissect the contribution of these three potential precipitating factors—HCV infection, cirrhosis, and cancer—to the observed phenotype and to further characterize the functional capacity of B cells in early and advanced liver disease.

Patients and Methods

  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information


Subjects and controls were recruited from the Gastroenterology Clinics at the Philadelphia Veterans Affairs Medical Center under an institutional review board–approved protocol. Patients were assessed for baseline demographics, hepatitis viral serologies, alcohol-use history, and radiological findings. Healthy donors (HDs) had no evidence of liver disease or malignancy. Study subjects with HCV infection confirmed twice by commercial polymerase chain reaction assays were classified in this study as having the following: (1) early fibrosis (EF) based upon a liver biopsy within 3 years of the bleed date showing ≤ METAVIR F2 fibrosis and/or FibroTest ≤ F1-F2 testing within 6 months; (2) cirrhosis (CIR) based upon clinical decompensation (e.g., ascites, jaundice, encephalopathy, or thrombocytopenia), radiological finding (e.g., splenomegaly, nodular liver, varices, or ascites), liver biopsy within 5 years, and/or Fibrotest F4; or (3) HCC based on standard American Association for the Study of Liver Diseases diagnostic guidelines.21

Cell Isolation.

Peripheral blood mononuclear cells (PBMCs) were isolated using Ficoll-Histopaque (Sigma-Aldrich, St. Louis, MO) density centrifugation. Surface phenotyping for CD27 expression was performed on freshly thawed cryopreserved PBMCs, but the remainder of experiments were performed with fresh PBMCs. CD19+ B cells were negatively selected using a B-cell isolation kit II (Miltenyi Biotec, Auburn, CA) on an AutoMACS platform. Purity of isolated B cells was greater than 95%. CD4+ T cells were isolated from cryopreserved PBMCs via a negative selection bead cocktail (Miltenyi), with purity greater than 95%. Isolated lymphocytes were resuspended in Roswell Park Memorial Institute 1640 medium with L-glutamine (Invitrogen, Carlsbad, CA), supplemented with 10% heat-inactivated human AB serum (Sigma), 1.5% HEPES (Invitrogen), and 1% penicillin/streptomycin (Invitrogen).22

Flow Cytometry.

Surface phenotyping was performed using antibodies against CD3 (PerCP, SK7), CD14 (PerCP, MϕP9), CD19 (allophycocyanin [APC]-H7, SJ25C1), CD21 (APC, B-ly4), CD27 (phycoerythrin [PE] and V450; M-T271), CD38 (fluorescein isothiocyanate [FITC], HIT2), and FcRL4 (PE, 413D12; BioLegend, San Diego, CA) with Live/Dead Aqua. A subset of fresh PBMCs were also stained with IgD (AlexaFluor 700, IA6-2), IgG (V450, G18-145), and IgM (FITC, G20-127). Isolated B cells were stained with CD40 (FITC, LOB7/6), CD70 (PE, Ki-24), CD86 (V450, 2331 (FUN-1)), and human leukocyte antigen (HLA)-DR (APC, G46-6). Responder CD4+ T-cells were carboxyfluorescein succinimidyl ester (CFSE)-labeled (Invitrogen) and stained for CD3 (PerCP, UCHT-1) and CD4 (APC, RPA-T4). All monoclonal antibodies (mAbs) were purchased from BD Biosciences (Franklin Lakes, NJ), except for anti-CD40 (AbD Serotec, Raleigh, NC), anti–Fc-receptor-like protein 4 (anti-FcRL4; BioLegend), and a fixable Live/Dead Aqua Staining kit (Invitrogen). All data were acquired on FACSCanto (BD) and analyzed using FlowJo (Tree Star Inc., Ashland, OR) using cutoffs based on isotype antibodies.

B-cell Culture and Activation.

B cells were activated using anti-CD40 mAb and TLR9 ligation, as previously described.23 Briefly, 2 × 105 freshly isolated B cells were incubated with both CP-870,893 (kindly provided by Pfizer, New London, CT) plus CpG oligodeoxynucleotide (ODN) 2006 (Invitrogen) or dual control (human IgG2κ; Chemicon International, Temecula, CA) and ODN2006 control (Invitrogen). After 48 hours, B cells were washed, stained for activation markers, and utilized for mixed lymphocyte reaction (MLR) experiments.


MLR was performed as described previously.23 Briefly, after 48 hours of stimulation, 6 × 104 dual-activated or dual-control B cells were irradiated (3,000 rad) and cocultured with CFSE-labeled CD4+ T cells (B:T ratio = 1:2) from a normal donor. CFSE-labeled CD4+ T cells were also coincubated with media alone or with anti-CD3/CD28 beads (kindly provided by Dr. Carl June). After 5 days, CD4+ T-cell proliferation was assessed by flow cytometry. To compare B-cell allostimulatory capacity across dates, we normalized CFSE dilution results according to the positive and negative control in each experiment. The percent maximal CFSE dilution for each test condition was thus obtained by the following formula: [(log10 geometric MFI of media exposed CD4+ T-cells) − (log10 geometric MFI of dual-activated or dual-control B-cell-exposed CD4+ T-cells)/(log10 geometric MFI of media exposed CD4+ T-cells) − (log10 geometric MFI of anti-CD3/CD28-stimulated CD4+ T-cells)], controlling for background in dual-control conditions. Simple comparisons of geometric mean fluorescence intensity (MFI) of dual-activated cells divided by geometric mean MFI of dual-control cells yielded similar results.

Cytokine/Ig Detection.

Undiluted culture supernatant from 48-hour B-cell activation and 5-day T-cell cocultures were collected and stored at −80°C. Cytokine (interferon-gamma, interleukin [IL]-2, IL-4, IL-6, IL-8, IL-10, IL-12, IL-17A, TNF-α, TNF-β, and IL-21) or Ig levels (IgG1, IgG2, IgG3, IgG4, IgA, and IgM) were quantified using Milliplex MAP Kit (Millipore, Billerica, MA) on a Luminex 200 system (Luminex Corporation, Austin, TX) using Masterplex QT software (Hitachi/MiraiBio, South San Francisco, CA).

Enzyme-Linked Immunosorbent Assay.

Freshly isolated plasma from whole blood were aliquoted and stored at −80°C for enzyme-linked immunosorbent assay (ELISA) analysis of soluble CD14 (sCD14; R&D Systems, Minneapolis, MN), according to the manufacturer's instructions.

TLR4 and TLR9 Blockade.

B cells from healthy donors were negatively isolated, as described above. First, 5 × 104 B-cells were cultured in 50% plasma from CIR donors, 50% plasma from HDs, 10% human AB serum alone or supplemented with IgG/A/M (Jackson Immunoresearch, West Grove, PA), 1 μg/mL of lipopolysaccharide (LPS; Sigma), or 1 μg/mL of CpG oligodeoxynucleotides (ODN) 2006 (Invitrogen). Plasma wells were cultured in the presence or absence of the TLR4 antagonist, Rhodobacter sphaeroides/LPS (LPS-RS; Invitrogen), anti-CD14 mAb (61D3; eBioscience), anti-TLR4 mAb (HTA125; Thermo Fisher, Rockford, IL), or the TLR9 antagonist TTAGGG (Invitrogen). After 72 hours, B cells were stained for Live/Dead Aqua, HLA-DR, and CD38 and were acquired on a FACSCanto.

Statistical Analysis.

The median values for clinical and immunologic parameters were compared using analysis of variance (ANOVA) (for normally distributed values), matched-pair comparisons, nonparametric Kruskal-Wallis ANOVA, Wilcoxon rank sum, or the Mann-Whitney U test, as appropriate. Spearman rank correlation was used for bivariate correlation of variables. Multivariate regression was performed using JMP 9 (SAS Institute, Inc., Cary, NC). P < 0.05 was considered significant with Bonferroni's correction, where required.


  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Patient Characteristics.

Samples from 18 HDs, 25 HCV-infected patients with F1-F2 fibrosis (EF), 19 with CIR, 30 HCC patients, and 5 non-HCV cirrhotics were studied (Table 1). Median age for HCC patients was approximately 6 years older than cirrhotic subjects, consistent with the natural history of HCC in HCV-related cirrhosis, but there were no other significant demographic differences among these groups. Expected differences in total bilirubin, serum albumin, platelet count, and International Normalized Ratio (INR) were observed in patients with cirrhosis. Absolute lymphocyte counts were slightly reduced in patients with cirrhosis or HCC (P = 0.039); therefore, phenotypic differences were evaluated both as percentage per lymphoid population and as absolute population numbers.24

Table 1. Baseline Patient Characteristics
  Healthy DonorsEarly FibrosisCirrhosisHCCNon HCV-CIRStatistical Analysis
Patient number I82519305OverallCIR/HCC versus EFCIR versus HCC
  1. HCC, hepatocellular carcinoma; HCV, hepatitis C virus; CIR, cirrhotic group; EF, early fibrosis; M, male; F, female; As, Asian; H, Hispanic; ALT, alanine aminotransferase; INR, International Normalized Ratio; WBC, white blood cell count; ALC, absolute lymphocyte count; AFP, alpha-fetoprotein.

GenderM/F17/125/019/030/05/0 11
ALT (g/dL)Median (25th-75th)23.9 (20.3-26.3)52.6 (36.0-57.0)81.5 (55.0-92.5)75.9 (45.3-87.5)39.4 (23.0-53.0)0.00010.0140.37
Albumin (g/dL)Median (25th-75th)4.4 (4.3-4.6)4.3 (4.1-4.5)4.1 (3.8-4.4)3.3 (3.0-3.8)4.0 (3.7-4.4)0.0001<0.00010.0001
Total bilirubin (g/dL)Median (25th-75th)0.6 (0.5-0.8)0.8 (0.6-0.9)1.2 (0.8-1.5)1.3 (0.8-1.7)0.9 (0.7-1.1)0.000I0.00030.61
INRMedian (25th-75th)1.1 (1.0-1.1)1.0 (1.0-1.0)1.2 (1.1-1.2)1.2 (1.1-1.3)1.2 (1.1-1.2)<0.0001<0.00010.21
Platelets (K/mm3)Median (25th-75th)241.3 (215.0-269.5)228.0 (180.0-265.0)136.7(100.0-169.5)118.0 (62.0-146.0)163.8 (109.0-191.0)<0.0001<0.00010.18
WBC (K/mm3)Median (25th-75th)6.4 (5.2-7.9)6.8 (5.6-7.715.7 (4.5-7.6)6.0 (4.3-7.2)6.4 (5.0-7.0)0.520.120.98
ALC (K/mm3)Median (25th-75th)2.2 (1.5-2.712.0 (1.7-2.4)1.8 (1.3-2.4)1.5 (0.8-2.212.3 (2.0-2.2)
AFP (ng/mL)Median (25th-75th)  15.1 (2.6-14.0)636.4 (23.1-409.5)3.6 (2.5-4.2)  0.0056

Cirrhosis Is Associated With Relative and Absolute Reduction of CD27+ Memory B Cells Independent of HCV Infection.

B-lymphocytes were defined using lymphoid gating, excluding nonviable cells, CD3+ T-cells, CD14+ monocytes, and then gating on CD19+ cells (Fig. 1A). Across the four patient groups, no significant differences were observed in the relative frequency and absolute number of CD19+ B cells among lymphoid cells (Fig. 1C). Though the frequency of CD27+ memory B cells among CD19+ cells was not significantly altered in HCV-infected patients with F1-F2 fibrosis, there were strongly significant reductions in relative and absolute CD27+ memory B-cell frequency in cirrhotic patients with or without HCC (Fig. 1D). The frequency of CD27+ B-cells among CD19+ B cells was not significantly different between fresh and cryopreserved samples (Supporting Fig. 1), and the intragroup differences remained significant when limiting analysis to cryopreserved samples (data not shown). Reduced CD27+ B-cell frequency was also found in patients with non-HCV-related cirrhosis (e.g., alcohol, HBV, nonalcoholic steatohepatitis) (Fig. 1E). The reduction of CD27 expression was B-cell specific, and the expression of CD27 on T cells was not different across the patient groups (data not shown). Unlike CD27+IgG+ B-cell frequency that was preserved in cirrhotics, CD27+IgM+ B cells were strikingly reduced (cirrhotic 16.3% versus noncirrhotic 32.4%; P = 0.021; Fig. 1F). A significant increase in CD27+CD38hi plasmablasts among cirrhotic patients was also observed (Supporting Fig. 2). FcRL4, an inhibitory coreceptor on B cells potentially identifying “exhausted” B cells, was not found to be expressed in CD27+, CD27-CD21+, or CD27CD21 B-cell subsets in any patient group (data not shown). The frequency of CD27+/CD19+ B cells was strongly correlated with several parameters related to progressive liver disease, including total bilirubin, hypoalbuminemia, thrombocytopenia, and INR (Fig. 2A-D; all P ≤ 0.0001). In summary, reductions in CD27+ memory B-cell frequency, particularly CD27+IgM+ B cells, are associated with cirrhosis independent of HCV infection, possibly because of increased peripheral conversion to short-lived plasmablasts.

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Figure 1. Reduction of CD27+ memory B cells in cirrhosis. (A) Gating strategy for identification of CD19+ B cells. (B) Representative histograms of CD27 expression of CD19+ B cells in HDs, HCV with early fibrosis (F1-2) (EF), HCV cirrhotics (CIR), and HCV cirrhotics with HCC. (C) Relative and absolute number of CD19+ B cells in HD, HCV+ EF, CIR, and HCC. (D) Relative and absolute number of CD27+ B cells in HD, HCV+ EF, CIR, and HCC. (E) Frequency of CD27+CD19+ B cells in non-HCV cirrhotic patients, relative to other patient groups. (F) Distribution of CD27+IgM+ and CD27+IgG+ B cells across patient groups. All statistical comparisons made using Kruskal-Wallis and pairwise Wilcoxon rank sum tests.

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Figure 2. Correlation of CD27+ B-cell frequency and hepatic dysfunction. Spearman rank correlation of CD27+ memory B-cell frequency and (A) total bilirubin, (B) serum albumin, (C) platelet count, and (D) INR.

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Cirrhotic B Cells Are Hyporesponsive to Activation by Ligation of CD40 Plus TLR9.

In our earlier work, peripheral B-cell CD27 expression was directly related to the capacity of B cells to be activated by CD40 plus TLR9 ligation.23 To determine the effect of CD27+ B-cell reduction and B-cell function in cirrhosis, we stimulated isolated B cells with anti-CD40 mAb combined with CpG ODN or appropriate controls for 48 hours, then assessed the expression of the activation markers, CD40, CD70, CD86, and HLA-DR. We detected a slight increase in the up-regulation of the activation/costimulation markers, CD86 and HLA-DR, among CIR relative to EF patients, but no difference in CD40 up-regulation (Fig. 3A-C). By contrast, up-regulation of CD70 was significantly reduced in cirrhotic patients (with and without HCC), relative to normal donors (Fig. 3D). The up-regulation of CD70 was strongly associated with baseline CD27 expression (R2 = 0.36, P < 0.001; Fig. 3E). We noted no significant intragroup differences in the production of IL-4, IL-6, IL-8, IL-10, IL-12, or TNF-α by activated B cells (Table 2A). However, cirrhotic B cells tended to secrete less TNF-β, relative to HD B cells (70.2 versus 27.9 pg/mL, P = 0.01; Table 2A; Fig. 4A). TNF-β production was strongly correlated with baseline CD27 expression (R2 = 0.34, P = 0.005; Fig. 4B). Interestingly, strong associations were also observed between baseline CD27 expression and IL-6, IL-12, and TNF-α production, although no significant intragroup differences were observed (Fig. 4C,D). Furthermore, cirrhotic B cells also produced less total IgG (but not IgA or IgM) than normal donor B cells (Fig. 4E). Thus, cirrhotic B cells are hyporesponsive to strong activating stimuli, as manifested by impaired up-regulation of CD70, TNF-β, and IgG production.

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Figure 3. Impaired activation of cirrhotic B cells by CD40/TLR9 ligation. Difference in geometric MFI of (A) HLA-DR, (B) CD86, (C) CD40, and (D) frequency of CD70+ B cells in HD, EF, and CIR/HCC patients, compared by Kruskal-Wallis and pairwise Wilcoxon rank sum tests. (E) Spearman rank correlation between up-regulation of CD70 expression upon CD40/TLR9 stimulation and baseline CD27+ B-cell frequency.

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Figure 4. Impaired cytokine production by cirrhotic B cells after CD40/TLR9 stimulation. (A) TNF-β secretion upon CD40/TLR9 stimulation in HD, EF, and CIR/HCC patients. Spearman rank correlations of (B) TNF-β, (C). TNF-α, and (D) IL-6 and CD27+ B-cell frequency. (E) IgG, IgM, and IgA titers in CD40/TLR9 across patient groups, compared by Kruskal-Wallis and pairwise Wilcoxon rank sum tests.

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Table 2. Activated B-cell and T-cell Cytokine Production
N = 8N = 9N = 18ANOVAHD versus CIR
  1. HD, healthy donor; EF, early fibrosis; CIR, cirrhotic group; ANOVA, analysis of variance; IFN, interferon; IL, interleukin; TNF, tumor necrosis factor.

A. B-cell (48 Hours)
IFN-γ (pg/mL) Median (25th-75th)3.7 (0.0-8.2)0.0 (0.0-1.5)0.0 (0.0-4.5)0.16 
IL-4 (pg/mL) Median (25th-75th)0.0 (0.0-2.2)15.2 (0.0-43.9)1.0 (0.0-2.7)0.10 
IL-6 (pg/mL) Median (25th-75th)2,503.7 (733.3-5692.0)1,163.5 (595.1-5842.9)2,481.3 (637.8-5000.5)0.99 
IL-8 (pg/mL) Median (25th-75th)767.2 (222.8-1694.6)2,545.7 (191.6-7746.3)2,990.9 (288.5-6425.0)0.42 
IL-10 (pg/mL) Median (25th-75th)324.1 (58.6-487.1)190.7(44.7-566.4)67.3 (17.1-444.0)0.49 
IL-12p40 (pg/mL) Median (25th-75th)45.5 (28.5-106.9)21.7(6.1-100.4)19.0 (7.6-82.0)0.39 
IL-17 (pg/mL) Median (25th-75th)0.0 (0.0-.0)0.0 (0.0-.0)0.0 (0.0-.0)0.99 
TNF-α (pg/mL) Median (25th-75th)187.7 (123.8-275.2)133.3 (112.8-294.9)243.9 (142.0-399.2)0.47 
TNF-β (pg/mL) Median (25th-75th)66.8 (34.5-114.3)31.3 (19.6-70.4)27.4 (3.8-53.1)0.0190.0082
B. T-cell (120 Hours)
IFN-γ (pg/mL) Median (25lh-751h)470.0 (252.3-1505.9)398.8 (74.0-838.8)249.2 (15.5-814.5)0.39 
IL-2 (pg/mL) Median (25th-75th)194.2 (153.4-289.9)186.9 (84.7-238.8)138.1 (29.7-357.5)0.56 
IL-4 (pg/mL) Median (25th-75th)30.5 (25.3-35.6)26.9 (14.7-29.7)16.8 (3.2-35.1)0.42 
IL-6 (pg/mL) Median (25th-75th)112.8 (78.1-206.2)82.0 (53.9-167.5)126.0 (60.0-182.0)0.89 
IL-8 (pg/mL) Median (25th-75th)215.9 (155.7-332.9)636.5 (136.5-2284.3)306.8 (134.7-671.3)0.21 
IL-10 (pg/mL) Median (25th-75th)0(0-0)0 (0-5.5)0(0-9.3)0.34 
IL-17 (pg/mL) Median (25th-75th)0 (0-0)0 (0-0)0 (0-0)0.99 
TNF-α (pg/mL) Median (25th-75th)214.7 (172.6-233.1)265.5 (58.1-437.9)65.8 (26.9-91.8)0.00740.003
TNF-β (pg/mL) Median (25th-75th)163.4(96.0-199.5)168.9 (9.4-259.4)70.5 (9.1-95.0)0.050.013

Cirrhotic B-Cell Antigen-Presentation Capacity Is Impaired Relative to HD B Cells.

To test the allostimulatory capacity of cirrhotic B cells, relative to HD B cells, we performed a mixed lymphocyte reaction using 48-hour–activated and control B cells to stimulate normal donor CD4+ T cells. Cirrhotic B cells (with or without HCC) were less capable of stimulating alloreactive CD4+ T-cell proliferation than noncirrhotic HCV patient or HD B cells (Fig. 5A). Interestingly, B-cell allostimulatory capacity was not correlated with memory B-cell frequency, CD86, or HLA-DR (Fig. 5B-E), but did correlate strongly with the degree of up-regulation of CD40 upon activated B cells (R2 = 0.37, P = 0.002; Fig. 5F). B-cell allostimulatory capacity did not significantly correlate with B-cell cytokine production (data not shown). T-cells stimulated by cirrhotic B cells were impaired in their capacity to produce TNF-α and TNF-β (Table 2B). In multivariable logistic regression analysis, only CD40 expression and percent CD70+ B cells were the only independent predictors of B-cell allostimulatory capacity (data not shown). Thus, cirrhotic B cells are impaired in their capacity to stimulate CD4+ T cells, an effect that appears to correlate with impaired up-regulation of costimulation markers after CD40/TLR9 activation.

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Figure 5. Reduced allostimulatory capacity of cirrhotic B cells. (A) Maximal CFSE dilution (calculated as geometric MFI of CFSE in T cells cocultured with activated B cells minus geometric MFI of media-control T cells divided by geometric MFI of anti-CD3/CD28 bead-stimulated T cells minus geometric MFI of media-control T cells) across patient groups. Correlation of maximal CFSE dilution and (B) baseline CD27+ B-cell frequency, (C) frequency of CD70+ on post-CD40/TLR9 activation B cells, (D) postactivation B-cell HLA-DR expression, (E) postactivation B-cell CD86 expression, and (F) postactivation B-cell CD40 expression.

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Circulating Factors in Cirrhotic Plasma Hyperactivate B Cells.

By ELISA, levels of sCD14, a soluble LPS adaptor protein produced and shed by monocytes after LPS exposure,25 were significantly increased in cirrhotic plasma (Fig. 6A). sCD14 concentrations were strongly inversely associated with CD27+ B-cell frequencies (R2 = 0.40, P < 0.001; Fig. 6B). B cells do not express membrane-bound cluster of differentiation 14 (mCD14), but sCD14 can directly transfer LPS to myeloid differentiation-2 (MD-2), activating the TLR4 pathway.26 It has also previously been shown that bacterial DNA, a potential TLR9 ligand, can often be detected in cirrhotic plasma.27 We, therefore, investigated the potential role of TLR4 and TLR9 ligands in cirrhotic plasma in activating B cells. HD B cells were cultured with 50% plasma from noncirrhotic (non-CIR; n = 8) or cirrhotic (CIR; n = 8) patients for 72 hours for the measurement of activation (i.e., HLA-DR, CD38, CD27, and CD19). Cirrhotic plasma induced a significant up-regulation of the expression of HLA-DR, up-regulation of CD38, and down-regulation of CD19 (Fig. 6C). HLA-DR expression was associated with serum sCD14 levels (R2 = 0.29, P = 0.033) (data not shown). Up-regulation of HLA-DR expression by cirrhotic plasma, which could also be induced by exposure to LPS or CpG, was abrogated by the antagonism of either TLR4 or TLR9 (Fig. 6D). Abrogation of HLA-DR up-regulation was detected with three different approaches to TLR4 blockade: LPS-RS, which inhibits LPS binding to LPS-binding protein (LBP), neutralization of sCD14, and direct blockade of TLR4 (Fig. 6E). Last, similar to LPS and CpG, cirrhotic plasma protected B cells from apoptosis in 72-hour culture, an effect that was abrogated by TLR4 and/or TLR9 blockade (Fig. 6F). Thus, soluble factors associated with bacterial translocation, such as LPS and CpG motifs, that are elevated in cirrhotic plasma are capable of activating B cells in vitro. Though the long-term effects of such activation cannot be modeled ex vivo, these data suggest a possible mechanism underlying the phenotypic and functional perturbations of peripheral blood B cells in cirrhosis.

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Figure 6. TLR4 and TLR9 activation of B cells in cirrhosis. (A) sCD14 plasma concentration by ELISA across patient groups. (B) Inverse correlation of sCD14 concentrations and CD27+ B-cell frequency. (C) Effect of cirrhotic versus healthy donor plasma on expression of HLA-DR, CD38, and CD19 on normal donor B cells after 72 hours of culture. Representative data from three separate experiments with different B-cell donors are shown. (D) Impact of TLR4 (LPS-RS) and/or TLR9 (TTAGGG) antagonism of HD and CIR plasma on HD B cells after 72 hours of culture. Representative data from three separate experiments with different B-cell donors are shown. (E) Impact of three approaches to block TLR4 activation (LPS-RS, anti-CD14, and anti-TLR4) on the expression of HLA-DR on normal donor B cells cocultured with HD and CIR plasma. (F) Preservation of B-cell viability (lack of Live/Dead Aqua staining) by cirrhotic plasma and effect of TLR4 (LPS-RS) and/or TLR9 (TTAGGG) antagonism.

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  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

In our study, we have uniquely found that among patients with chronic HCV, only those that have progressed to cirrhosis display a loss of CD27+ memory B cells with associated functional abnormalities. The noncirrhotic and cirrhotic HCV-infected patients we studied were similar in age, gender, ethnicity, viral genotype, and duration of infection, making viral or demographic factors very unlikely to explain the observed differences. Furthermore, this phenotype was also identified in patients with non-HCV-related cirrhosis, strongly implicating hepatic fibrosis and/or portal hypertension in the development of this phenotype. The loss of CD27+ memory B cells appears to be a phenomenon common to several immunocompromised states, such as advanced solid tumors,23 human immunodeficiency virus (HIV) infection,28 and common variable immunodeficiency (CVID).29 Though HIV and cirrhosis are both associated with bacterial translocation, a common underlying pathophysiology with CVID and advanced malignancy is not immediately obvious, but perhaps may be related to splenic dysfunction.

The loss of CD27+ memory B cells in cirrhosis was associated with several functional consequences, including impaired activation, impaired TNF-β and IgG production, and impaired allostimulatory capacity. This impaired activation and reduced capacity to recruit T-cell help may explain the observed vaccine hyporesponsiveness in cirrhotic patients.14, 15 Paradoxically, overall Ig levels are elevated in cirrhotics because of increased levels of pathogen-specific Igs, such as antibodies against Saccharomyces cerevisiae and against Galα1-3Galβ1-3GlcNAc, a glycan epitope found in bacterial cell walls.16, 17 Quite strikingly, we have shown that cirrhosis is associated with profound reductions of CD27+IgM+ B cells, a subset of memory B cells thought to be generated in response to T-independent antigens.30 Based on these observations, further investigation is warranted to determine the specific effect of cirrhosis on T-dependent and T-independent antigen responses, as well as on optimal adjuvants that may improve vaccine efficacy in cirrhotics.

Our findings indicate that TLR ligands associated with bacterial translocation circulating in cirrhotic patients directly activate B cells in vitro, an effect that can be attenuated with TLR4 and/or TLR9 blockade. TLR9 is constitutively expressed on B cells,31 and it has been suggested that TLR9 agonists might affect the nature of B-cell Ig responses in cirrhosis.18 Human B cells express minimal basal levels of TLR4, but up-regulate TLR4 expression on exposure to various stimuli.32 LPS-LBP bound to sCD14 can directly bind MD-2 on mCD14-negative cells.26 Consistent with earlier studies, we found that sCD14 levels were elevated in cirrhotic plasma,33 and that sCD14 levels correlated with in vitro B-cell activation. Elevated sCD14 levels have previously been found in systemic lupus erythematosus34 and HIV infection,35 both of which are also associated with CD27+ B-cell reductions. In particular, HIV, which infects gastrointestinal lymphoid tissue early in infection and compromises intestinal integrity, leads to increased bacterial translocation, nonspecific immune activation,36 and, ultimately, is associated with memory B-cell loss.37–39 Our data suggest a similar pathogenesis of memory B-cell loss in cirrhosis, albeit within the limitations of what can be demonstrated in ex vivo human B cells. In vivo animal studies will be critical to determine the complex interaction of portal hypertension, bacterial translocation, hypersplenism, and hepatic microenvironmental factors on B-cell memory generation and maintenance.

The fate of “lost” CD27+ B-cells in cirrhosis remains incompletely defined. One potential fate is the evolution of an “exhausted” phenotype similar to that described in HIV disease, in which an increased frequency of hypoproliferative CD27CD21 B cells with elevated expression of an inhibitory molecule, such as FcRL4 and other inhibitory molecules, disproportionately consisting of HIV-specific B cells has been identified.39 Though we did identify an increase in CD27CD21 B-cells in cirrhotic patients with HCC, we did not identify an increase of FcRL4-expressing cells in any group of patients or cell subset (data not shown). An alternative explanation for the reduction of CD27+ B cells in chronic HCV patients is an increased conversion of activated CD27+ B-cells to short-lived plasmablasts.6, 7 Our data, showing an increase in CD27+CD38hi in cirrhotics, provide modest support for this hypothesis for the cirrhotic patient subset. HCV E2-CD81 interactions40 also have been postulated to drive activation-induced apoptosis in chronic HCV. In vitro studies support an activating role of CD81 ligation in B cells from chronic HCV patients.4, 41 However, E2-CD81 interactions cannot explain the loss of CD27+ memory B-cells we identified in non-HCV cirrhotics or alterations of B-cell memory that have been identified in some HBV patients.7 Further challenging the activation-induced apoptosis hypothesis are data from Sugalski et al. and Mizuochi et al., which demonstrate that HCV-infected patient B cells manifest increased survival in vitro, relative to HD B-cells.8, 9 Our in vitro data do suggest that soluble factors in plasma from cirrhotic patients promote B-cell survival.

A third explanation for peripheral memory B-cell loss could be compartmentalization of activated CD27+ memory B cells to the intrahepatic or lymphoid compartments resulting from up-regulation of homing markers, such as CXCR3,8, 10, 42 a possible mechanism that was not explored in this study. In the intrahepatic compartment, a profibrotic role of B-cells has been suggested by work in the B-cell-deficient mice treated with carbon tetrachloride,43 by association of plasma cells and activated stellate cells in autoimmune liver disease,44 and by anecdotal regression of cirrhosis associated with rituximab in case reports.45 The intrahepatic compartment in cirrhotics does appear to be enriched for CD27+ memory B-cells (Supporting Fig. 3), but study of animal models will be critical to precisely define the fate of CD27+ memory B cells in cirrhosis and will be helpful in determining whether or not intrahepatic B-cells may play a pathological role in chronic liver injury/fibrosis.

Independent of chronic HCV infection, memory CD27+ and, more specifically, CD27+IgM+ B-cells are profoundly reduced in the peripheral blood of patients with cirrhosis with or without HCC in direct relationship with parameters associated with hepatic metabolic dysfunction and portal hypertension. The remaining B-cells are hyporesponsive to activation via CD40 and TLR9, with impaired up-regulation of costimulation markers, production of TNF-β, and production of IgG. The remaining B cells, upon activation, are also less effective at stimulating CD4+ T-cell responses. The presence of elevated levels of sCD14 and attenuation of B-cell activation by TLR4 and TLR9 blockade in vitro suggest that the loss of peripheral memory B-cells may be a consequence of chronic B-cell activation as a result of increased gut permeability caused by portal hypertension. These findings shed light on vaccine hyporesponsiveness and increased susceptibility to bacterial infection in cirrhotic patients, which might be ameliorated by therapies designed to reduce microbial translocation or block chronic pathogen-induced B-cell activation.


  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

The authors thank Mary E. Valiga, R.N., for her support of the study. The authors also thank the patients and volunteers who contributed samples.


  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Patients and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Additional Supporting Information may be found in the online version of this article.

HEP_24689_sm_SuppFig1.tif1419KSupporting Information Figure 1. Comparison of CD27+ memory B-cell frequency in fresh and cryopreserved lymphocyte samples. (A) Representative dot plots of fresh and frozen CD27+ B-cell frequency. (B) Matched-pair comparison of CD27+/CD19+ B-cell frequency.
HEP_24689_sm_SuppFig2.tif1268KSupporting Information Figure 2. Frequency of CD27+CD38hi plasmablasts in patient cohorts. (A) Representative fluorescence-activated cell-sorting plots for CD27+CD38hi populations. (B) Comparison of CD38hi/CD27+ B-cell across patient groups.
HEP_24689_sm_SuppFig3.tif1960KSupporting Information Figure 3. Peripheral versus intrahepatic memory B-cell frequency in cirrhosis. Data from 2 patients with HCC, who underwent hepatic resection, are shown.

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.