Immunoregulation of dendritic and T cells by alpha-fetoprotein in patients with hepatocellular carcinoma

Journal of Hepatology 41 (2004) 999–1007 www.elsevier.com/locate/jhep

Immunoregulation of dendritic and T cells by alpha-fetoprotein in patients with hepatocellular carcinoma Marcus Ritter1,†, Mona Y. Ali1,†, Christian F. Grimm1,†, Robert Weth1, Leonhard Mohr1, Wulf O. Bocher2, Katja Endrulat1, Heiner Wedemeyer3, Hubert E. Blum1, Michael Geissler1,* 1

Department of Medicine II, University Hospital Freiburg, Hugstetter Strasse 55, D-79106, Freiburg, Germany 2 Department of Medicine, University Hospital Mainz, Mainz, Germany 3 Department of Gastroenterology, Hepatology and Endocrinology, Medizinische Hochschule Hannover, Germany

Background/Aims: Novel immunotherapeutic and other strategies are being explored for the treatment of hepatocellular carcinoma (HCC). Alpha-fetoprotein (AFP) may be a target antigen for immunotherapy. Little is known, however, about the immunobiology of AFP. Therefore, the impact of AFP on dendritic cells (DC), CD4C and CD8C T cells was studied in detail. Methods: Immune cells from peripheral blood of 27 HCC patients were studied using FACS, ELISPOT, and proliferation assays. Results: The in vitro generation, maturation, and T cell stimulatory capacity of DCs were not altered by AFP up to concentrations of 20 mg/ml. Higher AFP concentrations (O20 mg/ml) resulted in phenotypic changes on DCs without impairing their capacity to stimulate CD4C T cells. Frequencies and function of DCs and AFP specific T cells were not reduced in HCC patients independent on serum AFP levels. Finally, T lymphocytic infiltrations in the liver were not dependent on AFP serum levels. Conclusions: These studies clearly demonstrate that (i) DC-based immunotherapeutic approaches are a promising approach for HCC treatment and (ii) AFP-reactive T cell clones have not been deleted from the human T cell repertoire establishing AFP as a potential target for T cell based immunotherapy of HCC. q 2004 European Association for the Study of the Liver. Published by Elsevier B.V. All rights reserved. Keywords: Alpha-fetoprotein; Cytotoxic T lymphocytes; Dendritic cells; Hepatocellular carcinoma; Liver

1. Introduction Hepatocellular carcinoma (HCC) is one of the major malignancies worldwide [1,2]. Most of the HCC patients are inoperable at the time of diagnosis [3]. Despite

Received 23 March 2004; received in revised form 25 July 2004; accepted 3 August 2004; available online 28 August 2004 * Corresponding author. Tel.: C49-761-2703260; fax: C49-7612703610. E-mail address: [email protected] (M. Geissler). Abbreviations: AFP, a-fetoprotein; CTL, cytotoxic T lymphocyte; DC, dendritic cell; FCS, fetal calf serum; HCC, hepatocellular carcinoma. † These authors contributed equally to this study.

several palliative therapeutic options [4–6] there is a high rate of recurrence or of intra-hepatic metastases. Therefore, novel treatment strategies such as immuno-gene therapy [7,8] are necessary to be developed to lower the frequency of tumor recurrence. Recent reports have demonstrated that alpha-fetoprotein (AFP) may be used as a target for active immunotherapy of HCCs [9–12]. However, until now the immunobiological function of AFP remains unclear. Some in vitro studies postulated an immunomodulating and immunosuppressing function of AFP [13–24]. The impact of serum AFP levels on immune cell functions in HCC patients is not known. Therefore, we analyzed different antitumoral immune effectors such as dendritic

0168-8278/$30.00 q 2004 European Association for the Study of the Liver. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.jhep.2004.08.013

1000

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

cells, CD4C and CD8C T cells in 27 HCC patients with different AFP serum levels and five healthy volunteers.

2. Materials and methods

Maturation of immature DCs was induced by incubation for 48 h in the presence of maturation medium: RPMI 1640 (Invitrogen GmbH, Karlsruhe, Germany) medium supplemented with 5% human AB serum, 1000 U/ml IL-4, 800 U/ml GM-CSF, 10 ng/ml IL-1b (R and D Systems, Wiesbaden, Germany), 1000 U/ml IL-6, 10 ng/ml TNF-a (both CellGenix Inc.), and 1 mg/ml PGE2 (Sigma, Taufkirchen, Germany).

2.4. Loading of immature DC

2.1. Patients and PBMC isolation Institutional Review Board approval and informed consent were obtained before drawing blood. All healthy volunteers (nZ5) and HCC patients (nZ27) enrolled in this study were from the University Hospital Freiburg, Department of Medicine II. The patients were included in the order of appearance in the hospital independent on AFP level or etiology of liver cirrhosis. The primary inclusion criteria were (a) histology proven HCC, (b) absence of systemic or transarterial chemotherapy/chemoembolization prior or during this study, and (c) hemoglobin O10 g/dl, leukocytes O3500 per ml. Exclusion criteria of the study were (a) presence of a second malignancy, (b) intake of immunosuppressive or chemotherapeutic drugs, and (c) anemia (Hb !10 g/dl) or leukopenia !3500 per ml. HCC etiology was alcoholic (nZ11), HCV (nZ8), HBV (nZ7), or cryptogenic (nZ1) cirrhosis (Table 1). Nine HCC patients had normal (0–20 ng/ml), 18 had elevated (O20 ng/ml) AFP serum levels (Table 1). Blood (180–200 ml/ patient) was processed within 1–2 h after drawing.

2.2. Separation of blood dendritic cells (BDC) and T cells PBMCs were isolated from heparin blood by Ficoll (Biochrom, Berlin, Germany) centrifugation. The different cell types were magnetically separated (Miltenyi Biotec, Bergisch Gladbach, Germany). Shortly, PBMCs were depleted of CD19C(clone SJ25-C1) cells. Then, MDC were isolated with anti-BDCA-1 mAb (AD5-8E7) and anti-biotin microbeads, PDC were directly isolated with anti-BDCA-4 mAb (AD5-17F6)conjugated microbeads. Finally, CD8C and CD4C cells were isolated using anti-CD8-(BW135/80) and anti-CD4-Mircobeads (M-T321), respectively. Purity of isolated cell types was O95% (Fig. 1A). The remaining cells were used for in vitro generation of Mo-DC.

For ELISpot and proliferation assays immature DC were loaded either with 20 mg/ml AFP derived form human cord blood (ICN, Eschwege, Germany, purity O98%), 10 mg/ml tetanus toxoid (TT) (List Biological Laboratories, Campbell, CA, USA) or 7.5 mg/ml ConA (Sigma, Taufkirchen, Germany) as a positive control for 2 h followed by maturation.

2.5. FACS analysis of different cell types The following mAb (clone names) were used to analyze the maturation status DCs: CD80 FITC (L307.4), CD83 PE (HB15e), CD86 FITC (2331, (FUN1)), CD 86 PE (IT2.2), CD40 FITC and PE (5C3), MHC II FITC ¨ 36), and MHC II PE (L243) from Becton Dickinson (BD), Heidelberg, (TU Germany. To analyze the frequency of MDC and PDC, mAbs were added to the whole blood and incubated for 15 min on ice, followed by addition of Optilyse B (Immunotech, Marseille, France) for 15 min at room temperature and FACS analysis. The following mAbs were used: CD1c FITC (B-B5, Biosource, Camarillo, CA, USA), BDCA-2 FITC (AC144), BDCA-3 FITC (AD-14H12), BDCA-4 PE (AD5-17F6) from Miltenyi Biotec, Bergisch Gladbach; CD4 FITC (RPA-T4) and PE, CD19 PE (HIB19), HLA-DR II PE, CD123 PE (9F5) from BD and streptavidinFITC/-PE from Biosource, Camarillo, CA, USA. Isolated T-cells were stained for CD4, CD8, CD25, CD45RA, CD54, CD62L, CCR7 (BD).

2.6. Quantitation of endocytosis using FITC dextran and luciferin yellow capture

2.3. Generation of Mo-DC

DCs were resuspended and FITC dextran or lucifer yellow (both Molecular Probes, Molecular Probes Europe, Leiden, Netherlands) was added at a final concentration of 0.1 mg/ml for 15 min at 37 8C. Cells were analyzed by FACS after fixation.

Mo-DCs were generated from CD 14C monocytes using CellGro DC medium (CellGenix Inc., Freiburg, Germany) supplemented with 1% human AB serum, 1000 U/ml IL-4 (CellGenix Inc.; Freiburg, Germany) and 800 U/ml GM-CSF (Leukomaxw, Novartis, Nu¨rnberg, Germany).

2.7. T-cell proliferation assay

Table 1 Patient data No. Average age and range (years) M/F Cirrhosis † HBsAg positive † anti-HCV positive † alcoholic † cryptogenic † Child Pugh A/B/C Serum AFP † Average and range (ng/ml) † !20 ng/ml † 20–999 ng/ml † 1000–19,999 ng/ml † O20,000 ng/ml

27 66 (47–80) 21/6 7 8 11 1 18/5/4 12,002 (2.7–308,900) 9 (mean 7 ng/ml, range 2.9–19.1 ng/ml) 13 (mean 149 ng/ml, range 21–468 ng/ml) 4 (mean 3030 ng/ml, range 1019–5754 ng/ml) 1 (308,900 ng/ml)

Triplicates of 2!105 PBMCs or 5!104 CD4C T cells per well were pulsed with 20 mg AFP/ml, 10 mg/ml tetanus toxoid (TT), or 7.5 mg ConA/ml for 4 days, respectively. As negative controls, unpulsed PBMCs were used. For the last 18 h the cells were labelled with 4 mCi/ml [methyl-3H]-thymidine (Amersham, Freiburg, Germany) and incorporation into DNA was measured after harvesting. Results were corrected for background activity (D cpm).

2.8. Allogenic mixed lymphocyte reaction (MLR) Isolated MDC and PDC were incubated at different T cell: DC ratios, using 2!105 PBMCs from a healthy donor. 2!105 PBMCs were added to MDCs and PDCs. As a positive control Con A (Sigma), was added to the PBMCs in a concentration of 7.5 mg/ml. MLR was allowed to incubate for 4 days in a humidified atmosphere (37 8C, 5% CO2) and [methyl-3H]thymidine incorporation into DNA was measured after harvesting.

2.9. ELISpot Plates were coated with 1 mg anti-IFN-g (MabTech, Nacka, Sweden) or anti-IL-5 (BD, Heidelberg, Germany) antibodies. 5!103 DC per well and 5!104 CD4C and CD8C per well were incubated for 40 h. For ELISpot with PBMCs, 2!105 PBMCs per well were incubated with 20 mg AFP/ml,

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

1001

Fig. 1. Immune cell separation and phenotypic characterization of Mo-DCs. A. Purity of immunomagnetically purified MDC, PDC, CD4C and CD8C T-cells using FACS analyses. Purified MDC, PDC, CD4C, and CD8C T cells were stained with antibodies against CD1c, BDCA-2, CD4, and CD8, respectively. B–C. Representative FACS analyses of Mo-DCs derived from a healthy volunteer. B. FACS analyses of lucifer yellow uptake of immature Mo-DCs generated in medium without or with AFP 20 mg/ml). Background staining was measured after incubation of the cells with lucifer yellow at 4 8C. C. Phenotype of Mo-DC and impact of AFP on maturation of Mo-DCs. Mo-DC phenotype was examined using FACS analysis of CD80, CD83, CD86, CD40, and HLA-DR expression. Immature Mo-DC were maturated in maturation medium alone (/) or supplemented with 20 mg/ml AFP or 10 mg/ml TT. D. Measurement of FITC dextran uptake was analyzed by FACS. im4 and im37 8C, immature Mo-DC cultured at 4 and 37 8C, respectively; m4 and m37 8C, mature Mo-DC cultured at 4 and 37 8C, respectively; m/AFP37 8C and m/TT37 8C, mature Mo-DC cultured in maturation medium at 37 8C supplemented with 20 mg/ml AFP and 10 mg/ml TT, respectively. [This figure appears in colour on the web]. 10 mg TT/ml, or 7.5 mg ConA/ml. Secreted cytokines were detected with either biotinylated anti-IFN-g (MabTech), or biotinylated anti-IL-5 antibodies (BD), both in a concentration of 1 mg/ml followed by Streptavidin-AP (MabTech) and substrate (BioRad, Munich, Germany). Plates were analyzed with a Bioreaderw 3000 Pro (BioSys, Karben, Germany).

3. Results

2.10. Immunohistochemical studies

Since an AFP-specific receptor has been identified on human monocytes [25], we analyzed the impact of AFP on generation, maturation, and function of Mo-DC. Mo-DCs from healthy volunteers (nZ3) were generated both in the absence and presence of 20 mg/ml AFP. The numbers and the lucifer yellow uptake of immature Mo-DCs (Fig. 1B), the expression of MHC class II antigens, CD86, CD83, CD80, and CD40 (Fig. 1C), the Mo-DC maturation (Fig. 1C), and the FITC-dextran uptake of mature Mo-DCs (Fig. 1D) was comparable between the groups. These analyses were expanded in two additional healthy individuals (controls) and 15 patients with a HCC (Table 2). Expression of HLA-DR, CD80, CD83, CD86, and CD40 on immature Mo-DCs and upregulation of CD80, CD83, and CD86 expression following Mo-DC maturation was

Distributions of CD3C, CD8C, and CD57C cells in the livers were analyzed by immunohistochemistry. Paraffin-embedded liver sections were stained using mAbs against CD3, CD8, and CD57 (all Dako, Hamburg, Germany) followed by the application of biotinylated antibodies and alkaline phosphatase-conjugated streptavidin and substrate (*DAKO ChemMatee Detection Kit, alkaline phosphatase/RED, Rabbit/mouse). Sections were counterstained with hematoxylin. The number of CD3C, CD8C, and CD57C cells in the portal tracts and the intralobular areas was scored in equivalent fields (!400).

2.11. Statistical analysis One way analysis of variance and the Wilcoxon signed rank test were used for statistical comparison of means (GSD) and proportions between groups, respectively. A P value of less than 0.05 was considered significant.

3.1. Impact of alpha–fetoprotein on generation and function of Mo-DC in vitro

1002

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

Table 2 Comparison of the phenotypic characteristics of Mo-DCs derived from controls and HCC patients at both immature and mature state dependent on AFP pulsing Immature day 7

Mature day 8 unpulsed

Controls (nZ2) % CD80 CD83 CD86 CD40 HLA-DR

HCC (nZ14) MFI

30G0.1 18G1 13G2 30G3 91G3 136G18 94G4 72G39 100 864G55 Mature day 8 AFP pulsed (48 h) Controls (nZ2)

CD80 CD83 CD86 CD40 HLA-DR

% 96G1 97G1 98G1 98G1 100

Controls (nZ2) MFI

%

33G24 31G14 95G3 98G1 99G0.5

19G3 48G10 151G68 75G41 1097G773

96G1 42G0.4 97G1 136G3 98G2 335G64 98G2 97G19 100 2416G239 Mature day 8 TT pulsed (48 h)

MFI 37G16 121G52 203G64 90G29 1835G853

% 96G2 97G2 98G2 97G2 100

HCC (nZ15) MFI 41G0.7 112G27 309G66 90G11 1998G22

HCC (nZ15)

%

% 89G7 92G13 98G2 97G2 99G2

MFI

Controls (nZ2)

%

MFI

88G9 92G11 98G1 97G2 100

34G12 121G55 225G65 95G31 2027G896

HCC (nZ12) MFI 42G0.4 98G11 334G60 91G15 1877G53

% 90G9 92G15 97G6 98G2 100G1

MFI 38G16 125G62 213G70 97G48 1526G679

Both the proportion of positive cells (%) and the mean fluorescence intensity (MFI) are presented for each marker as the medianGSD. Mo-DC were pulsed with 20 mg/ml AFP or 10 mg/ml tetanus toxoid (TT).

comparable between HCC patients and controls (PO0.5 for all surface markers). AFP or TT did not alter the percentage and mean fluorescence intensity of Mo-DC surface markers (PO0.5 for all comparisons). Increasing the AFP concentrations to 100 mg/ml during maturation resulted in a reduction of the mean expression levels of CD83, CD86, CD40, and HLA-DR compared to maturation w/o AFP (Table 3). However, these phenotypic changes did not result in an impaired allogeneic stimulatory capacity of Mo-DCs (Fig. 2A). Finally, we examined the effect of AFP on the capacity of Mo-DC to induce antigen specific CD4C T cell proliferative responses. The addition of AFP or PSA to TT during Mo-DCs pulsing did not affect CD4C T cell proliferation (Fig. 2B). Similar results were obtained after pulsing Table 3 Comparison of the phenotypic characteristics of Mo-DCs derived from two healthy controls (c1 and c2) at both immature and mature state dependent on pulsing with 100 mg/ml AFP Mo-DC treatment

CD80 (%/MFI)

CD83 (%/MFI)

CD86 (%/MFI)

CD40 (%/MFI)

HLA-DR (%/MFI)

66/23 94/57

25/29 80/66

93/149 94/302

62/62 94/74

97/1000 98/1536

94/50

73/48

95/229

94/55

98/1218

54/23 93/48

13/44 92/75

94/143 97/248

90/45 97/66

97/601 99/1392

91/46

85/65

96/210

94/55

98/935

c1 Immature Mature unloaded MatureC 100 mg/ml AFP

Mo-DCs with recombinant hepatitis-B-virus surface antigen (HBsAg) for stimulation of CD4C T cells derived from three anti-HBs positive vaccinated individuals (Fig. 2C). The number of IFN-g secreting CD4C T cells was not reduced after co-stimulation with HBsAg and AFP or HBsAg and CEA compared to HBsAg alone. Taken together, these results indicate that the presence of AFP in concentrations observed in patients with HCCs does not change the in vitro generation or function of Mo-DCs. 3.2. Phenotype and function of blood dendritic cells Pooled frequencies of MDCs (0.41G0.1%) and PDCs (0.37G0.08%) derived from all HCC patients were comparable to controls (Table 4). MDC and PDC frequencies were independent on AFP serum levels. Even a subgroup analysis of patients with AFP serum concentration ranging from 1000–308,900 ng/ml (nZ5) did not reveal significantly reduced MDC and PDC frequencies. MDC and PDC frequencies in HCC patients with HBV (nZ7) versus HCV (nZ8) cirrhosis were 0.36G0.25% and 0.51G0.31% versus Table 4 Absolute numbersa of circulating CD11cC and CD123C DC subsets in healthy controls and HCC patients dependent on serum AFP levels DC subsets

Controls (nZ5)

HCC (nZ9) AFP 0–20 ng/ml

HCC (nZ18) AFP O20 ng/ml

HCC (nZ4) AFP O 1000 ng/ml

MDC (CD1cC, CD11cC) PDC (CD11cK, CD123C)

0.39G0.09

0.39G0.22

0.36G0.23

0.47G0.07

0.32G0.17

0.41G0.28

0.34G0.22

0.36G0.15

c2 Immature Mature unloaded MatureC 100 mg/ml AFP

a Values represent percentage of positive cells and are expressed as meanG standard deviation.

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

Fig. 2. Functional characterization of Mo-DCs. A. Mo-DC from two healthy individuals (c1 and c2) were generated and subsequently maturated in maturation medium without or with AFP at concentrations up to 100 mg/ml for 48 h. All four Mo-DCs of each control were analyzed in an allogeneic T cell proliferation assay. Allogeneic responder CD4C T cells, derived from one buffy coat batch, were cultured for 5 days in the presence of increasing numbers of stimulator Mo-DCs followed by analysis of [methyl-3H]-thymidine incorporation into DNA. No relevant proliferation of CD4C T cells alone was detected. B. Antigen specific recall CD4C T cell response in three healthy controls. 5!103 Mo-DC were pulsed with the helper T cell recall antigen TT (10 mg/ml), AFP (20 mg/ml), TTCAFP, or, as a control, TTCPSA (20 mg/ml). Subsequently, they were cocultured with 5!104 autologous CD4C T cells. As negative control (background activity) unpulsed Mo-DCs and CD4C T cells were cocultured. Furthermore, proliferation of TT pulsed MDCs without CD4C T cells and CD4C T cells alone was measured. Results were corrected for background activity (D cpm). C. Frequencies of IFN-g secreting AFP and HBsAg specific CD4C and CD8C T cells derived from three HBV vaccinated control persons were determined by ELISPOT. 5!103 Mo-DCs were pulsed with the helper T cell recall antigen HBsAg (10 mg/ml), AFP (20 mg/ml), HBsAgCAFP, or, as a control, HBsAgCCEA (20 mg/ml). Subsequently, they were cocultured with 5!104 autologous isolated CD4C and CD8C T cells for 40 h and spots were counted automatically.

0.44G0.13% and 0.31G0.1%, while they were lower in patients with alcoholic cirrhosis (0.28G0.16% and 0.28G 0.18%, respectively). Patients with alcoholic cirrhosis had a more advanced Child–Pugh Score B and C (nZ7/11) as compared to patients with HBV or HCV cirrhosis (Child–Pugh Score B and C: nZ2/15) which may be

1003

the reason for the statistically non-significant lower numbers of MDC and PDC in the patients with alcoholic cirrhosis. MDCs and PDCs derived from peripheral blood displayed the characteristic phenotype of immature DCs: expression of moderate to high levels of MHC class II antigens and CD86, low levels of CD80, CD80, and CD40 (Table 5). Mean fluorescence intensity (MFI) values of HLA-DR were significantly higher on MDCs compared to PDCs (PZ0.001). No significant differences (PO0.1 for all comparisons) in the percentage or mean fluorescence intensity of HLA-DR, CD80, CD83, CD86, or CD40 was observed in MDCs or PDCs with respect to normal and elevated AFP levels in the HCC patients (Table 5). Proliferation of allogeneic T cells upon stimulation with MDCs or PDCs (MLR) was much weaker compared to stimulation with Mo-DCs. The allostimulatory function of MDCs (Fig. 3A) and PDCs (Fig. 3B) was similar (PO0.1) between HCC patients with elevated or normal AFP serum levels. Moreover, no significant difference was observed in the MLR assays between HCC patients and healthy individuals. Therefore, our results demonstrate that number and function of MDCs and PDCs from HCC patients are not affected by AFP. 3.3. Hepatocellular carcinoma specific CD4C and CD8C T cell responses Pilot experiments in healthy individuals and AFPpositive HCC patients demonstrated that pulsing Mo-DCs with 2–20 mg/ml of fetal cord blood derived AFP results in presentation of AFP derived peptides in the context of both MHC class I and II, leading to activation of autologous AFP specific CD4G and CD8G T cells, consistent with efficient cross-priming of CD8G T cells (Fig. 4A). In fact, in 59% of HCC patients CD4C TH1 and CD8C Tc1 cells producing IFN-g could be detected directly ex vivo at frequencies between 0.01 and 0.02% (Fig. 4B). Significantly lower frequencies were detected with respect to IL-5 producing CD4C TH2 (PZ0.01) and CD8C Tc2 cells (PZ0.001) in 93% of patients. Interestingly, CD8C T cells secreting IFN-g and/or IL-5 could be detected in a significantly higher number of HCC patients (87.5%)

Table 5 Percentages (%) and mean fluorescence intensity (MFI) of CD11cC and CD123C DC surface markers in HCC patients (nZ27)

CD1cC CD11cC total CD1cC CD11cC AFP!20 ng/ml CD1cC CD11cC AFPO20 ng/ml CD1cC CD123C total CD1cC CD123C AFP!20 ng/ml CD1cC CD123C AFPO20 ng/ml

HLA-DR (%/MFI)

CD80 (%/MFI)

CD83 (%/MFI)

CD86 (%/MFI)

CD40 (%/MFI)

94G19/633G97 87G29/315G192 99G1/855G552 93G21/191G130 85G32/151G79 99G0.6/214G148

7G4/11G10 3G2/12G8 11G8/11G8 3G2/7G3 2G2/5G1 3G3/8G3

54G31/30G17 45G25/24G7 63G32/35G21 79G23/54G42 75G33/36G20 82G12/63G47

91G16/86G42 86G21/76G27 95G8/94G49 98G2/63G26 98G3/58G25 98G1/66G27

24G22/16G11 12G6/10G4 33G23/20G13 33G24/14G6 24G21/11G5 38G22/16G5

Values are presented for each DC subset as a total derived from all HCC patients and with respect to normal (!20 ng/ml) or elevated (O20 ng/ml) serum AFP levels.

1004

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

Fig. 3. Allogenic CD4C T cell response upon stimulation with MDC and PDC. Isolated MDCs (A) and PDCs (B) from three healthy individuals and HCC patients with normal (!20 ng/ml) and elevated AFP levels (O20 ng/ml) were incubated at different T cell: DC ratios, using 2!105 isolated CD4C T cells from a healthy individual as effectors and the indicated numbers of DC as stimulators. After 72 h 4 mCi/ml of [methyl-3H]-thymidine was added for the remaining 18 h followed by analysis of [methyl-3H]-thymidine incorporation into DNA. As a negative control (background activity), proliferation of MDCs, PDCs, and CD4C T cells alone was measured. Results were corrected for background activity (D cpm).

compared to CD4C TH1 or TH2 cells (50%, PZ0.001) suggesting that activation of AFP specific CD8C T cells does not necessarily require the presence of AFP specific CD4C T cell help. In fact, in 20% of HCC patients with IFN-g producing CD8C T cells neither IFN-g nor IL-5 producing CD4C T cells could be detected. Similarly, 40% of patients with IL-5 secreting CD8C T cells lacked TH1 and TH2 cells. HCC patients with chronic HBV or HCV infection harbored significantly higher numbers of IFN-g compared to IL-5 secreting AFP specific PBMCs (Fig. 5, PZ0.02). In HCC patients with alcoholic cirrhosis there were no differences between the presence of IL-5 or IFN-g secreting AFP specific PBMCs (PO0.05). HCC patients with elevated AFP levels were characterized by statistically non-significant (PO0.1) reduced

Fig. 4. AFP specific CD4C and CD8C T cell responses.A: 5!103 MoDCs pulsed with AFP (20 mg/ml) were incubated for 40 h with 5!104 CD4C or CD8C T cells derived from two healthy volunteers (HV) and two patients with HCC (HCC) and secreted IFN-g was detected using ELISpot. As a control unloaded Mo-DC as stimulators were used (w/o AFP). B: 5!103 Mo-DCs derived from HCC patients (nZ27) and pulsed with AFP (20 mg/ml) were incubated for 40 h with 5!104 CD4C or CD8C T cells. Secreted IL-5 and IFN-g was detected using ELISpot. Results are corrected for background activity (unloaded Mo-DC as stimulators, T cells alone).

numbers of IFN-g producing CD8C T lymphocytes (Fig. 6). AFP levels had no significant impact on the presence of AFP specific TH1, TH2 polarized CD4C T cells or Tc2 CD8C T cells.

Fig. 5. Frequency and polarization of AFP specific PBMC and HCC etiology. Triplicates of 2!105 PBMCs per well were pulsed with 20 mg AFP/ml for 40 h followed by ELISpot. As negative controls, unpulsed PBMCs were used. Results were corrected for background activity (specific spots). Only patients with measurable spots are displayed.

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

1005

4. Discussion

Fig. 6. Frequencies of AFP specific IL-5 or IFN-g producing CD4C and CD8C T cells at different AFP levels. 5!103 Mo-DC pulsed with AFP (20 mg/ml) were incubated for 40 h with 5!104 CD4C or CD8C T cells. Secreted IL-5 and IFN-g was detected using ELISpot. Results are corrected for background activity (unloaded Mo-DC as stimulators, T cells alone).

By FACS analyses (CD45RA, CCR7, CD25, HLA-DR, CD54, CD62L) we could not observe significant differences in the pool of naive, central memory, effector memory, and effector CD4C and CD8C T cells between HCC patients with normal or elevated AFP levels (data not shown) excluding a systemic inflammation at the time of drawing blood that could have affected the analysis of AFP specific T cell responses. 3.4. Immunohistochemical staining for CD3, CD8, CD68, CD57 in liver tissues In patients with ALT elevation and histological evidence of inflammation, a large number of CD8C T cells was present in the portal areas (Table 6). By contrast, subjects with normal ALT had relatively low numbers of infiltrating CD8C T cells, preferentially localized within the lobules between hepatocytes. There was no correlation between serum AFP levels and infiltration of the liver by CD3C and CD8C T cells (Table 6), suggesting that production of AFP by tumor cells or regenerating normal hepatocytes does not impair T cell recruitment to the liver. CD57C cells could be only rarely detected.

Immunotherapy may be a promising approach to prevent metastasis and recurrence after HCC resection and/or local ablation because intra-hepatic metastatic lesions seem to be indistinguishable from the primary tumors at the genomic level, regardless of tumor size, encapsulation, and age of the patient [26]. The concept of HCC immunotherapy, albeit not AFP specific, is supported by two reports [30,31], and further studies have clearly demonstrated that AFPreactive T cell clones have not been deleted from the human or murine T cell repertoire establishing AFP as a potential target for T cell based immunotherapy of HCCs [9–11,27–29]. Before using AFP as a potential tumor rejection antigen in clinical studies, we examined its role on immune effector cells because immunosuppressive effects of AFP on immune cell function such as inhibition of antibody synthesis and T cell proliferation, induction of suppressor cells, as well as modulation of MHC class II expression and phagocytosis on macrophages has been postulated in the past [14–17,32–34]. Furthermore, an AFP specific receptor has been identified on human monocytes [25] and AFP was found to reduce the prostaglandin mediated production of TNF-a and IL-1b by monocytes [35]. In the presence of high AFP concentrations up to 20 mg/ml we could not find any evidence that AFP affects the in vitro generation, maturation, and function of MoDCs. Such AFP concentrations are rarely observed in HCC patients. By contrast, pulsing Mo-DCs with 100 mg/ml AFP resulted in a downregulation of CD83, CD86, CD40, and HLA-DR expression which, however, did not result in an impaired T cell stimulatory capacity of Mo-DC. Therefore, the previously described defective functions of in vitro generated Mo-DC in HCC patients [36,37] may not be due to AFP but rather related to HCV [38] or HBV [39] infection. Similarly, the frequency and phenotype of MDCs and PDCs in HCC patients with normal and elevated AFP levels were comparable excluding an AFP-mediated systemic immunosuppression since these circulating DC subpopulations may reflect the state of the systemic immunity. Kunitani et al. [40] recently described reduced numbers of

Table 6 Analysis of liver infiltrating CD3C, CD8C, and CD57C cells dependent on AFP serum and ALT level CD3

AFP!20 ng/ml (nZ9) AFP O20 ng/ml (nZ14) ALT normal (nZ6) ALT elevated (nZ17)

CD8

CD57 Total

Portal

Intralobular

Total

Portal

Intralobular

Total

114G26 127G31 22G7 171G42

22G7 29G11 43G6 14G5

136G29 156G37 62G11 185G49

82G16 91G19 15G3 125G22

15G3 22G6 31G5 9G3

97G20 103G25 46G7 134G29

7G2 8G1 3G1 10G3

Liver biopsies were stained with anti-CD3, anti-CD8, and anti-CD57 mAbs. The distribution of CD3C, CD8C, and CD57C cells in the liver was visualized by immunostaining in formalin-fixed, paraffin-embedded liver specimens. The number of cells in the portal tracts and intralobular areas was scored in equivalent histological fields (original magnification !400).

1006

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

CD11cC MDCs and CD123C PDCs in patients with various liver diseases incl. HCC. The lowest numbers were observed in patients with cirrhosis suggesting that the Child Pugh Score, not assessed by Kunitani et al. [40], is a major factor in DC immunoregulation. In our patients with liver cirrhosis, the majority was Child Pugh A (Table 1) that may explain the differences in MDC and PDC numbers between Kunitani et al. [40] and our study. Serum AFP levels had no significant impact on the numbers of liver infiltrating CD3C, CD8C, and CD57C cells or the strength and frequency of AFP specific CD4C or CD8C T cell responses in HCC patients. We detected AFP specific CD8C T lymphocytes in peripheral blood of a high percentage of HCC patients, while AFP specific CD4C T lymphocytes were present at low frequencies in few HCC patients only suggesting that CD4C T lymphocytes are either present at very low frequencies below the detection limit of our ELISPOT assay or specifically deleted/anergized. It is unlikely that the reduction of AFP specific CD4C T lymphocytes was due to a general down-regulation of immune cell function since TT specific immune responses could be easily detected in most patients. More importantly, AFP specific CD4C and CD8C T cells, preferentially secreting INF-g were detected in HCC patients which is an important finding, because IFN-g is an important antitumor effector in vivo [41]. The median survival of HCC patients with detectable CD4C and CD8C T cell responses against AFP was 30.4 months compared to 26.6 months of patients with no detectable AFP specific T cell responses (PZ0.5) suggesting that liver function and tumor treatment rather than the pure presence of AFP specific T cells influences the outcome of HCC patients. Nevertheless, it has been demonstrated that patients with HCCs and a high density of tumor infiltrating lymphocytes have a significantly better 5-year survival and lower recurrence rates after HCC resection compared to HCC patients without lymphocyte infiltrations [42]. Therefore, a better prognosis of HCC patients with preexisting T cell responses against HCC antigens might be obtained by immunotherapeutic interventions that selectively expand and induce cytotoxic T lymphocytes (CTL) cytokines in HCC tissues. Taken together, the results of our study demonstrate that AFP may be used as tumor rejection antigen in most of patients with HCCs overexpressing AFP.

Acknowledgements M.G. is supported by Grant Ge824/7-1 and SFB364 from the Deutsche Forschungsgemeinschaft, and from the BMBF-HepNet network.

References [1] El-Serag HB, Mason AC. Rising incidence of hepatocellular carcinoma in the United States. N Engl J Med 1999;340:745. [2] Bosch X, Ribes J, Borras J. Epidemiology of primary liver cancer. Semin Liver Dis 1999;19:271. [3] Llovet JM, Bourroughs A, Bruix J. Hepatocellular carcinoma. Lancet 2003;362:1907. [4] Bruix J, Sherman M, Llovet JM, Beaugrand M, Lencioni R, Burroughs AK, et al. Clinical management of hepatocellular carcinoma. Conclusions of the Barcelona-2000 EASL Conference. European Association for the Study of the Liver. J Hepatol 2001;35: 421. [5] Allgaier HP, Galandi D, Zuber I, Blum HE. Radiofrequency thermal ablation of hepatocellular carcinoma. Dig Dis 2001;19:301. [6] Allgaier HP, Deibert P, Zuber I, Olschewski M, Blum HE. Percutaneous radiofrequency interstitial thermal ablation of small hepatocellular carcinoma. Lancet 1999;353:1676. [7] Butterfield LH, Ribas A. Immunotherapy of hepatocellular carcinoma. Expert Opin Biol Ther 2002;2:123. [8] Mohr L, Geissler M, Blum HE. Gene therapy for malignant liver disease. Expert Opin Biol Ther 2002;2:163. [9] Butterfield LH, Meng WS, Koh A, Vollmer CM, Ribas A, Dissette VB, et al. T cell responses to HLA-A*0201-restricted peptides derived from human alpha fetoprotein. J Immunol 2001;166:5300. [10] Butterfield LH, Koh A, Meng W, Vollmer CM, Ribas A, Dissette V, et al. Generation of human T-cell responses to an HLA-A2.1restricted peptide epitope derived from alpha-fetoprotein. Cancer Res 1999;59:3134. [11] Grimm CF, Ortmann D, Mohr L, Michalak S, Krohne TU, Meckel S, et al. Mouse alpha-fetoprotein-specific DNA-based immunotherapy of hepatocellular carcinoma leads to tumor regression in mice. Gastroenterology 2000;119:1104. [12] Geissler M, Mohr L, Ali MY, Grimm CF, Ritter M, Blum HE. Immunobiology and gene-based immunotherapy of hepatocellular carcinoma. Z Gastroenterol 2003;41:1101–1110. [13] Abramsky O, Brenner T, Mizrachi R, Soffer D. Alpha-fetoprotein suppresses experimental allergic encephalomyelitis. J Neuroimmunol 1982;2:1. [14] Auer IO, Kress HG. Suppression of the primary cell-mediated immune response by human alpha1-fetoprotein in vitro. Cell Immunol 1977;30:173. [15] Murgita RA, Tomasi Jr TB. Suppression of the immune response by alpha-fetoprotein on the primary and secondary antibody response. J Exp Med 1975;141:269. [16] Murgita RA, Wigzell H. The effects of mouse alpha-fetoprotein on Tcell-dependent and T-cell-independent immune responses in vitro. Scand J Immunol 1976;5:1215. [17] Murgita RA. The immunosuppressive role of alpha-fetoprotein during pregnancy. Scand J Immunol 1976;5:1003. [18] Neumann U. Inhibition of in vitro lymphocyte blastogenesis by chicken alpha-fetoprotein. Comp Immunol Microbiol Infect Dis 1984; 7:43. [19] Ohashi K, Mikami T, Kodama H, Izawa H. Suppression of NK activity of spleen cells by chicken fetal antigen present on Marek’s disease lymphoblastoid cell line cells. Int J Cancer 1987;40:378. [20] Olinescu A, Laky M, Pofescu DE, Dumitrescu A, Ganea D. The effect of alpha-fetoprotein on the immune response. III. Diminution of the phagocytic capacity of macrophages cultures in vitro in the presence of mouse amniotic fluid or alpha-fetoprotein. Arch Roum Pathol Exp Microbiol 1977;36:247. [21] Olinescu A, Popescu DE, Babes S, Dumitrescu A. Does alphafetoprotein influence the immune response? Arch Roum Pathol Exp Microbiol 1978;37:303. [22] Olinescu A, Popescu DE, Babes S, Dumitrescu A, Ganea D, Andreescu V, et al. The effect of alpha-fetoprotein on the immune

M. Ritter et al. / Journal of Hepatology 41 (2004) 999–1007

[23]

[24] [25]

[26]

[27]

[28]

[29]

[30]

[31]

response. IV. Effect of treatment with alpha-fetoprotein on mice infected with bacteria in various experimental conditions. Arch Roum Pathol Exp Microbiol 1978;37:137. Peck AB, Murgita RA, Wigzell H. Cellular and genetic restrictions in the immunoregulatory activity of alpha-fetoprotein. III. Role of the MLC-stimulating cell population in alpha-fetoprotein-induced suppression of T cell-mediated cytotoxicity. J Immunol 1982;128:1134. Sell S, Sheppard Jr. HW, Poler M. Effects of alpha-fetoprotein on murine immune responses. II. Studies on rats. J Immunol 1977;119:98. Suzuki Y, Zeng CQ, Alpert E. Isolation and partial characterization of a specific alpha-fetoprotein receptor on human monocytes. J Clin Invest 1992;90:1530. Ye QH, Qin LX, Forgues M, He P, Kim JW, Peng AC, et al. Predicting hepatitis B virus-positive metastatic hepatocellular carcinomas using gene expression profiling and supervised machine learning. Nat Med 2003;9:416. Hanke P, Rabe C, Serwe M, Bohm S, Pagenstecher C, Sauerbruch T, Caselmann WH. Cirrhotic patients with or without hepatocellular carcinoma harbour AFP-specific T-lymphocytes that can be activated in vitro by human alpha-fetoprotein. Scand J Gastroenterol 2002;37: 949. Geissler M, Mohr L, Weth R, Kohler G, Grimm CF, Krohne TU, et al. Immunotherapy directed against alpha-fetoprotein results in autoimmune liver disease during liver regeneration in mice. Gastroenterology 2001;121:931. Vollmer Jr CM, Eilber FC, Butterfield LH, Ribas A, Dissette VB, Koh A, et al. Alpha-fetoprotein-specific genetic immunotherapy for hepatocellular carcinoma. Cancer Res 1999;59:3064. Iwashita Y, Tahara K, Goto S, Sasaki A, Kai S, Seike M, et al. A phase I study of autologous dendritic cell-based immunotherapy for patients with unresectable primary liver cancer. Cancer Immunol Immunother 2003;52:155. Takayama T, Sekine T, Makuuchi M, Yamasaki S, Kosuge T, Yamamoto J, et al. Adoptive immunotherapy to lower postsurgical recurrence rates of hepatocellular carcinoma: a randomised trial. Lancet 2000;356:802.

1007

[32] Yamashita T, Nakane A, Watanabe T, Miyoshi I, Kasai N. Evidence that alpha-fetoprotein suppresses the immunological function in transgenic mice. Biochem Biophys Res Commun 1994;201:1154. [33] Toder V, Blank M, Nebel L. Immunoregulatory mechanisms in pregnancy. 1. Evidence for the alpha-fetoprotein-induced generation of suppressor cells in vitro. Transplantation 1982;33:41. [34] Brenner T, Zielinski A, Argov Z, Abramsky O. Prevention of experimental autoimmune myasthenia gravis in rats by fetal alphafetoprotein-rich fractions. Tumour Biol 1984;5:263. [35] Wang W, Alpert E. Downregulation of phorbol 12-myristate 13acetate-induced tumor necrosis factor-alpha and interleukin-1 beta production and gene expression in human monocytic cells by human alpha-fetoprotein. Hepatology 1995;22:921. [36] Ninomiya T, Akbar SM, Masumoto T, Horiike N, Onji M. Dendritic cells with immature phenotype and defective function in the peripheral blood from patients with hepatocellular carcinoma. J Hepatol 1999;31:323. [37] Kakumu S, Ito S, Ishikawa T, Mita Y, Tagaya T, Fukuzawa Y, Yoshioka K. Decreased function of peripheral blood dendritic cells in patients with hepatocellular carcinoma with hepatitis B and C virus infection. J Gastroenterol Hepatol 2000;15:431. [38] Bain CFA, Zoulim F, Zarski JP, Trepo C, Inchauspe G. Impaired allostimulatory function of dendritic cells in chronic hepatitis C infection. Gastroenterology 2001;120:512. [39] Beckebaum S, Cicinnati VR, Zhang X, Ferencik S, Frilling A, GrosseWilde H, et al. Hepatitis B virus-induced defect of monocyte-derived dendritic cells leads to impaired T helper type 1 response in vitro: mechanisms for viral immune escape. Immunology 2003;109:487. [40] Kunitani H, Shimizu Y, Murata H, Higuchi K, Watanabe A. Phenotypic analysis of circulating and intrahepatic dendritic cell subsets in patients with chronic liver diseases. J Hepatol 2002;36: 734. [41] Kemp RA, Ronchese F. Tumor-specific Tc1, but not Tc2, cells deliver protective antitumor immunity. J Immunol 2001;167:6497. [42] Wada Y, Nakashima O, Kutami R, Yamamoto O, Kojiro M. Clinicopathological study on hepatocellular carcinoma with lymphocytic infiltration. Hepatology 1998;27:407–414.