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. Author manuscript; available in PMC: 2015 Dec 1.
Published in final edited form as: J Immunol. 2014 Oct 29;193(11):5723–5732. doi: 10.4049/jimmunol.1400725

Tumor-derived alpha-fetoprotein impairs the differentiation and T cell stimulatory activity of human dendritic cells

Angela D Pardee *, Jian Shi *, Lisa H Butterfield *,†,‡,§
PMCID: PMC4239186  NIHMSID: NIHMS633765  PMID: 25355916

Abstract

Several tumor-derived factors have been implicated in DC dysfunction in cancer patients. Alpha-fetoprotein (AFP) is an oncofetal antigen that is highly expressed in abnormalities of prenatal development and several epithelial cancers, including hepatocellular carcinoma (HCC). In HCC patients exhibiting high levels of serum AFP, we have observed a lower ratio of myeloid-to-plasmacytoid circulating DC compared to patients with low serum AFP levels and healthy donors. To test the effect of AFP on DC differentiation in vitro, peripheral blood monocytes from healthy donors were cultured in the presence of cord blood-derived normal AFP (nAFP) or HCC tumor-derived AFP (tAFP), and DC phenotype and function was assessed. Although the nAFP and tAFP isoforms only differ at one carbohydrate group, low (physiological) levels of tAFP, but not nAFP, significantly inhibited DC differentiation. tAFP-conditioned DC expressed diminished levels of DC maturation markers, retained a monocyte-like morphology, exhibited limited production of inflammatory mediators, and failed to induce robust T cell proliferative responses. Mechanistic studies revealed that the suppressive activity of tAFP is dependent on the presence of low molecular weight (LMW) species that i) co-purify with tAFP, and ii) function equivalently to the LMW fractions of both tumor and non-tumor cell lysates. These data reveal the unique ability of tAFP to serve as a chaperone protein for LMW molecules, both endogenous and ubiquitous in nature, which function cooperatively to impair DC differentiation and function. Therefore, novel therapeutic approaches that antagonize the regulatory properties of tAFP will be critical to enhance immunity and improve clinical outcomes.

Keywords: alpha-fetoprotein, Dendritic Cells, Hepatocellular Carcinoma, immune suppression, T cells, cytokines

INTRODUCTION

Alpha-fetoprotein (AFP), an oncofetal protein synthesized in the yolk sac and fetal liver, is the most abundant serum protein in the fetus, reaching a maximum level of 3 mg/ml in the third month of gestation (1). Neural tube defects, chromosomal abnormalities, and other fetal abnormalities can be identified by elevated levels of AFP in the amniotic fluid. AFP is transcriptionally repressed shortly after birth, and normal adult levels typically range between 1–40 ng/ml. Reappearance of AFP in the circulation of adults is associated with liver regeneration, hepatitis, chronic liver diseases and malignant growth, particularly hepatomas, teratomas, and gastrointestinal cancers of endodermal origin (2). Various glycoforms of AFP have been identified in the serum of HCC patients (3, 4). The fucosylated variant AFP-L3 is the major glycoform found in individuals with HCC and is associated with poor prognosis. While cord serum-derived AFP contains <5% of the fucosylated variant, >80% of the fucosylated variant has been observed in HCC patient serum.

Several functions of AFP have been described. With its structural similarity to albumin, it has been hypothesized to play a role in the transport of serum components, including fatty acids, steroids, and heavy metals, and may serve as an embryonic analogue of albumin (2). There have also been reports of AFP interfering with intracellular signaling, including both caspase 3 and PI3K/AKT pathways (5, 6). An immunoregulatory role for AFP has also been proposed. Early studies revealed an inhibitory effect of cord blood-derived human AFP on lymphocyte function, while more recent reports suggest that AFP exerts its immunosuppressive activity through the induction of DC dysfunction (7–12). There is currently little consensus, however, on which cell subsets and/or signaling pathways are the primary targets of AFP-mediated immunosuppression. Furthermore, because the majority of these studies have solely used cord blood AFP preparations, the immunoregulatory nature of tumor-derived AFP remains unknown.

Given its specificity to HCC tumors, AFP represents an attractive target for immunotherapy. Several MHC class I and II restricted epitopes have subsequently been identified (13). Our group observed that CD8+ T cells from healthy donors can recognize 4 dominant and 10 subdominant HLA-A*0201-restricted peptides in vitro (14). Two clinical trials were initiated to test the following vaccine regimens: i) 4 immunodominant AFP peptides emulsified in Montanide adjuvant, and ii) AFP peptide-pulsed autologous DC (15, 16). Although no objective clinical responses were observed in the small numbers of vaccinated patients, AFP-specific T cell responses were either generated or enhanced in the majority of patients, providing proof-of-principle for AFP-targeted immunotherapeutic approaches.

In order to more fully characterize the immunosuppressive activity of AFP, as well as to inform the rational design of AFP protein-based vaccines, we examined the effects of cord blood-derived normal AFP (nAFP) or HCC tumor-derived AFP (tAFP) on DC differentiation and function. Here we show that tAFP has profound negative effects on the generation, maturation, and function of monocyte-derived DC, and that this activity is dependent on the presence of low molecular weight (LMW) molecules that co-purify with tAFP. Moreover, we further report that elevated serum levels of AFP are associated with alterations in the frequency of DC subsets in HCC patient peripheral blood.

MATERIALS AND METHODS

Antibodies and reagents

Purified ovalbumin (OVA; Fisher Scientific), human cord serum AFP (Cell Sciences; purity >95% by SDS-PAGE), and HCC cell line culture-derived AFP (Bio-Rad; purity >95% by SDS-PAGE) were added to cultures at 10 µg/ml, unless otherwise indicated. AFP concentrations were routinely confirmed by clinical laboratory tests (University of Pittsburgh Medical Center), and the degree of fucosylation (AFP-L3 percentage) was determined by Quest Diagnostics (both CLIA-certified assays). As expected, the tAFP preparations used in our experiments exhibited a high degree of fucosylation (61.7–84.6%) compared to nAFP (3.6–5.2%). Lot-to-lot variability in the preparations’ inhibitory activity was negligible (data not shown). AFP proteins were examined by stained SDS PAGE gels and Western blot to confirm identity (Santa Cruz anti-AFP SC-8399) (Supplementary Fig. S1A). AFP and OVA preparations were divided into high and low molecular weight (HMW & LMW, respectively) fractions using Amicon Ultra 3kDa MWCO centrifugal filters (Millipore). DC, T cell, and MDSC phenotypes were examined using fluorochrome-conjugated antibodies against the following cell-surface markers: HLA-ABC (BioLegend), CD206 (mannose receptor), CD40, CD80, CD83, IL-2, IFN-γ, Lineage Cocktail 1 (Lin1; CD3, CD14, CD16, CD19, CD20, CD56), CD33, (BD Biosciences), CD1c/BDCA-1, CD303a/BDCA-2 (eBioscience) CD4, CD8, TNF-α, HLA-DR, CD14, and CD11b (Beckman Coulter). DC apoptosis was detected using Annexin V-FITC and 7AAD staining (BD Pharmingen).

Cell lines

T2 (HLA-A2+; ATCC), T2-DR4 (HLA-DR4+; kindly provided by Dr. Janice Blum (Indiana University School of Medicine, Indianapolis, IN)), and the HepG2 hepatoma cell line (ATCC) were all cultured in RPMI 1640 medium, supplemented with 10% fetal bovine serum, 1% penicillin-streptomycin, and 1% L-glutamine (all reagents from Life Technologies). Cultures were maintained in a humidified 37°C incubator under 5% CO2 tension.

Isolation of PBMCs

Peripheral blood mononuclear cells (PBMC) were obtained from healthy donors (HD) and from HCC patients enrolled in an AFP DNA plasmid prime/adenovirus boost vaccine or a peptide-pulsed DC vaccine (UPCI #04-001; UPCI #04–101, IND BB13653; UCLA IRB #00-01-026, IND BB9395; informed consent was obtained from all patients and donors). Limited patient data is listed in Supplementary Table S1. PBMC were separated from blood using gradient centrifugation (Ficoll-Paque, GE Healthcare).

DC preparation

CD14+ monocytes were isolated from PBMC using magnetic cell sorting (Miltenyi Biotec) and cultured for 5 days in 800 IU/ml rGM-CSF (Sargramostim; Genzyme) and 500 IU/ml rIL-4 (R&D Systems). In some cases, DC were matured with IFN-γ (1000 IU/ml; PeproTech) and lipopolysaccharide (LPS, 250 ng/ml; Sigma-Aldrich) for an additional 24 hours prior to collection. DC culture supernatants were tested by multiplex Luminex assay (Life Technologies) for 30 analytes (cytokines, chemokines and growth factors), as previously published (17).

Confocal immunofluorescence staining and imaging

DC cultured in 8-well chamber slides (Nunc) were fixed (with 4% paraformaldehyde), permeabilized (0.1% Triton X-100), and stained with rhodamine phalloidin (Life Technologies) and DRAQ5 (eBioscience), to label F-actin and nuclei, respectively. Images were acquired using a Leica TCS-SL confocal microscope.

T cell proliferation, multimer staining, and cytokine production assays

For mixed lymphocyte reaction (MLR) assays, 2 × 104 mature DC were co-cultured with 2 × 105 allogeneic CFSE-labeled CD8+ or CD4+ T cells in T cell media (RPMI 1640 medium, supplemented with 10% human AB serum in the presence of rhIL-2 (30 IU/ml; PeproTech)). Cell proliferation was tested by measuring CFSE dilution on day 6. For intracellular cytokine staining, T cells were stimulated with PMA (0.2 ng/ml) and ionomycin (0.2 µM) in the presence of brefeldin A. Six hours later, T cells were stained for surface markers, fixed, permeabilized, and stained for intracellular cytokines (IL-2, TNF-α, and IFN-γ).

For Flu peptide-specific assays, immature DC from HLA-A2+ or HLA-DR4+ donors were cultured overnight with 10 µg/ml FluM158–66 (University of Pittsburgh Peptide Synthesis Facility) or HA307–319 (Mimotopes), respectively, in the presence of IFN-γ and LPS. The next day, 2 × 105 FluM158–66 or HA307–319 loaded mature DC were co-cultured with 1 × 106 autologous CD8+ or CD4+ T cells, respectively, in T cell media plus 30 IU/ml IL-2. Cultures were supplemented with IL-2 at 30 U/ml every 3–4 days. After 11 days, CD8+ T cells were collected and stained with PE-labeled FluM158–66 pentamer (ProImmune) plus CD8-APC. For intracellular cytokine staining, CD8+ or CD4+ T cells were restimulated with FluM158–66 pulsed T2 cells or HA307–319 pulsed T2-DR4 cells in the presence of brefeldin A. Six hours later, T cells were stained for surface markers, fixed, permeabilized, and stained for intracellular cytokines (IL-2, TNF-α, and IFN-γ). Data were acquired with an Accuri C6 cytometer (BD Biosciences) and analyzed using Accuri C6 software.

Enzymatic treatment of AFP

For all enzymatic treatments, AFP and OVA preparations were first heat denatured at 95°C for 15 min. Proteinase K (Ambion) was added at 0.5 µg enzyme per 1 µg AFP (or OVA control) at 56°C for 4hr. AFP degradation was confirmed by clinical laboratory test, which detected 0 ng/ml AFP. PNGase F (New England BioLabs) was added to AFP and OVA preparations according to the manufacturer’s instructions, and incubated 2hr at 37°C. α-L-Fucosidase (Sigma-Aldrich) was added to AFP and OVA preparations at 0.5 U/ml and incubated overnight at 37°C. Enzymes were used at concentrations recommended by the manufacturers based on functional testing by lot. After all treatments, enzymes were heat-inactivated at 95°C for 15 min before AFP and OVA preparations were added to culture.

Preparation of cell lysates

HepG2 hepatoma cells and healthy donor PBMC were resuspended in 0.9% saline and lysed by 3 freeze (on methanol and dry ice)–thaw (37°C) cycles. Larger particles were removed by centrifugation (13,000 rpm, 20 min, 4°C), and supernatants were either left whole or separated into HMW and LMW fractions using Amicon Ultra 3kDa MWCO centrifugal filters (Millipore). Lysates were added to cultures on day 0 at a lysed cell-to-monocyte ratio of 1:1, and DC were collected and analyzed on day 5.

Gene regulation and signaling analysis

DC cultured with nAFP, tAFP and OVA as described above were lysed and total mRNA was isolated and quantified. Transcriptome analysis was performed by Affymetrix HG–U133A array and analyzed by Significance Analysis of Microarrays (SAM) and Ingenuity software. Selected immune-related gene regulation by nAFP and tAFP was confirmed by real time PCR using MicroFast 96 well qPCR plates and primers (Life Technologies) and qPCR SuperMix kit (VWR), and HPRT1 as a housekeeping gene for normalization. To confirm NF-κB pathway changes, total NF-κB p65, phospho-NF-κB, total IκBα and phospho-IκBα were examined by Western blot (Cell Signaling NF-κB sampler) with β-actin as a housekeeping protein for densitometric calculations. Expression of selected NF-κB pathway immune-related genes were further tested by real time PCR by TaqMan Gene Expression Master Mix and Fast 96 well plates (Life Technologies) with HPRT1 as the housekeeping gene, according to manufacturer’s instructions.

Statistical Analysis

All comparisons of inter-group means were performed using a Student’s t test, with P values < 0.05 considered significant.

RESULTS

Elevated serum AFP levels are associated with a lower mDC-to-pDC ratio in the peripheral blood of HCC patients

Two independent studies have shown that i) the frequency of circulating myeloid DC is decreased in HCC patients (18), and ii) the frequency of circulating plasmacytoid DC is significantly increased in the context of advanced liver disease (19). We tested banked PBMC samples from HCC patients (Supplementary Table S1) for levels of myeloid DC (mDC; Lin1negCD1c+) and plasmacytoid DC (pDC; Lin1negCD303a+) (Fig. 1A). When patients were stratified into AFPlow (serum AFP < 1 µg/ml) and AFPhigh (serum AFP > 1 µg/ml) groups, the mean ratio of mDC-to-pDC was significantly reduced for AFPhigh HCC patients versus AFPlow HCC patients and healthy donors (Fig. 1B).

Figure 1. Alteration of circulating DC subsets in the peripheral blood of AFPhigh HCC patients.

Figure 1

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(A) Circulating DC gating strategy. Percentages of mDC (Lin1negCD1c+) and pDC (Lin1negCD303a+) among total PBMC are indicated. Shown is representative data from HCC patient C3. (B) Myeloid-to-plasmacytoid DC ratio in the peripheral blood of healthy donor (HD; n = 6) or HCC (n = 12) patients was determined by flow cytometry. HCC patients were divided into AFPlow (serum AFP < 1 µg/ml) and AFPhigh (serum AFP > 1 µg/ml) groups. *, P < 0.05.

DC differentiation is inhibited by physiologic levels of tumor-derived AFP

To investigate the effect of AFP on DC differentiation in vitro, monocytes were isolated from the PBMC of healthy donors and cultured for 5 days in the presence of GM-CSF + IL-4 and either cord blood-derived nAFP or HCC tumor-derived tAFP. Because median AFP serum levels of 9 µg/ml have been observed in HCC patients (2), nAFP and tAFP were added to the culture at a biologically-relevant dose range (1–20 µg/ml), with ovalbumin (a member of the albuminoid gene superfamily) used as a control. Both nAFP- and tAFP-treated DC expressed diminished levels of the DC markers HLA-ABC, CD206 (mannose receptor), CD40, CD80, and CD83, while elevated levels of the monocyte marker CD14 was observed (Fig. 2A). Expression levels of HLA-DR, CCR7, and CD86 were not significantly altered (data not shown). A high dose of nAFP (10–20 µg/ml), however, was necessary to suppress DC differentiation, consistent with a previous report (9). Conversely, tAFP demonstrated suppressive activity at lower doses (5–10 µg/ml), suggesting that DC are particularly sensitive to the tumor-derived isoform of AFP. As depicted in Figure 2B, when cells were treated at a concentration of 10 µg/ml (used for all subsequent experiments), only CD83 expression was significantly diminished in nAFP-conditioned DC, whereas expression levels of all five DC markers were significantly reduced on tAFP-conditioned DC compared to both nAFP- and OVA-treated cells. CD14 expression was significantly elevated by nAFP treatment (P < 0.05 vs. OVA), and enhanced to an even greater extent by tAFP treatment (P < 0.01 vs. OVA), suggesting that AFP (and tAFP in particular) blocks the differentiation of DC from monocyte precursors.

Figure 2. DC fail to acquire a fully-differentiated phenotype when cultured in the presence of tAFP.

Figure 2

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(A) CD14+ monocytes isolated from the peripheral blood of healthy donors were cultured with GM-CSF and IL-4 for 5 days either alone (“no protein”) or in the presence of increasing doses of ovalbumin (OVA), nAFP, or tAFP (1, 5, 10, 20 µg/ml). Mean fluorescence intensity (MFI) of HLA-ABC, CD206 (mannose receptor), CD40, CD80, CD83, and CD14 is indicated. Data from one representative healthy donor (of three total HD tested) is shown. (B) Monocytes from HDs (n = 5–6) were differentiated into DC (as in panel A) in the presence of OVA, nAFP, or tAFP (all at 10 µg/ml). The solid line represents the mean value. *, P < 0.05; **, P < 0.01.

It has been documented that the differentiation program of DC precursors can be skewed towards myeloid-derived suppressor cell (MDSC) generation by various tumor-associated factors (20). We therefore assessed our AFP-treated, GM-CSF- and IL-4-cultured DC for an MDSC phenotype (Lin1negHLA-DRlowCD11b+CD33+), but no evidence of MDSC-skewing was observed (Supplementary Fig. S1B).

Consistent with their phenotype, tAFP-conditioned DC also displayed a poorly-differentiated morphology as determined by fluorescent microscopy (Fig. 3A). tAFP-treated DC lacked prominent dendrites, which were present in their nAFP- and OVA-treated counterparts, and in general retained a monocyte-like morphology. Along with the surface marker analysis above, these data indicate that both DC phenotype and morphology are negatively impacted by physiologic levels of tAFP. Although cell recovery was modestly reduced in the tAFP-treated group (Fig. 3B), the percentage of apoptotic cells after five days of treatment did not exceed 14% (Fig. 3C), suggesting that tAFP does not impair DC differentiation primarily through the induction of cell death.

Figure 3. tAFP impairs DC morphology, viability, and the production of inflammatory cytokines and chemokines.

Figure 3

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Purified monocytes from healthy donors (n = 4) were differentiated into DC for 5 days in the presence of OVA, nAFP, or tAFP (all at 10 µg/ml). (A) Representative confocal analysis of OVA-, nAFP-, and tAFP-conditioned DC. Actin (red), nuclei (blue). Images are representative of three independent experiments performed. (B) Percent DC recovery (from total plated monocytes) on day 5 of culture is shown for four HD. ns, P ≥ 0.05. (C) DC apoptosis was assessed with Annexin V and 7AAD staining on day 5 of cultures, with percent apoptotic cells calculated as the sum of % Annexin V+/7AADneg and % Annexin V+/7AAD+ cells. Data from six HD are shown. *, P < 0.05. (D) Cell-free supernatants were collected from the above cultures and tested for IFNα, IL-8, IL-10, MCP-1/CCL2, MIP-1α/CCL3, and MIP-1β/CCL4 by Luminex assay. ns, P ≥ 0.05; *, P < 0.05; **, P < 0.01.

The production of inflammatory cytokines and chemokines is compromised in tAFP-conditioned DC

To further characterize the effect of AFP on DC generation and function, cell-free supernatants from the previous cultures (Fig. 2B) were assessed by a 30-plex Luminex assay. Production of several cytokines, including IL-1β, IL-6, TNF-α, IL-7, and IL-15, were not significantly changed by nAFP or tAFP treatment, and IL-12 was undetectable in these samples (data not shown). Although not statistically significant, there was a trend towards lower levels of IFN-α secreted by tAFP-DC (Fig. 3D). However, we observed a highly-significant (P < 0.01) decrease in IL-8 production by tAFP-conditioned DC compared to both nAFP- and OVA-treated cells. We also observed elevated levels of IL-10 in the culture supernatant of both nAFP- and tAFP-treated DC. The chemokines MCP-1/CCL2, MIP-1α/CCL3, and MIP-1β/CCL4, which are known to recruit innate cells (neutrophils, monocytes, NK cells) to sites of infection/inflammation and enhance the cytolytic activity of NK cells, were produced at markedly lower levels in tAFP-DC versus nAFP- and OVA-treated cells (21, 22).

To examine the molecular mechanism involved in the phenotypic and secreted molecule changes induced by tAFP and nAFP, DC from five different donors were tested for gene expression changes by gene array. Several immune-related genes were significantly down-regulated in tAFP-treated DC, including CCR7, CXCL1, and MIP-1α/CCL3 while IL-6 and CD14 mRNAs were upregulated (Supplementary Fig. S2A). Because the NF-κB pathway is known to be critical for DC maturation and inflammatory cytokine regulation (23), and several of the specific gene products we identified by microarray, we examined whether this signaling pathway was modulated by tAFP. We tested this first by examining gene products specifically regulated by the NF-κB pathway by real time PCR. In immature DC, tAFP increased IL-6 mRNA levels and decreased expression of CD40, IL-12A, IL-12B and CD83 (Supplementary Fig. S2B). In mDC, tAFP increased IL-10 mRNA levels and decreased IL-12A, IL-12B and CD83 (Supplementary Fig. S2B). To further confirm the modulation of the NF-κB pathway by tAFP, Western blotting showed that in both iDC and mDC, tAFP-treated DC have less total NF-κB protein. As expected, maturation with IFN-γ and LPS increases phospho-NF-κB in both OVA and tAFP-DC. IκBα phosphorylation was reduced by 51–63% in both iDC and mDC exposed to tAFP (Supplementary Fig. S2C). Together, these tAFP-induced changes in gene expression, protein expression and phosphorylation support tAFP modulation of the NF-κB pathway.

Allo-reactive T cell proliferation, but not cytokine production, is impaired by tAFP-conditioned DC

To enhance their allostimulatory activity, OVA-, nAFP-, and tAFP-conditioned DC were cultured overnight with LPS and IFN-γ to generate mature DC (mDC). While this maturation step enhanced the expression of HLA-ABC, CD206, and CD40 by tAFP-DC to levels similar to those expressed by OVA-DC, CD80 and CD83 expression remained significantly lower (and CD14 significantly higher) in tAFP-treated mDC compared to their nAFP- and OVA-treated counterparts (Supplementary Fig. S3A). The production of inflammatory cytokines and chemokines (TNF-α, IL-12, MCP-1, and MIP-1α) was also significantly reduced in tAFP-treated mDC (Supplementary Fig. S3B). Similar to immature DC, the recovery and viability of tAFP-treated mDC was slightly impaired, although the percentage of apoptotic cells after five days of treatment did not exceed 10% (Supplementary Fig. S3C–D).

When these OVA-, nAFP-, and tAFP-conditioned mDC were used to stimulate allogeneic T cells in a mixed lymphocyte reaction (MLR), considerable variability between donor-pair responses was observed, with OVA-mDC-induced proliferation at day 6 ranging from 34–72%. To standardize the donor-pair responses, nAFP-mDC- and tAFP-mDC-induced proliferation was normalized to control OVA-mDC-induced proliferation (set at 100%). This showed that both nAFP-mDC and tAFP-mDC, but particularly tAFP-mDC, fail to induce a robust allogeneic T cell proliferative response (Fig. 4A). CD8+ T cell proliferation was significantly impaired by both nAFP-mDC and tAFP-mDC, compared to OVA-mDC control (P < 0.01 for both). Meanwhile, proliferation of the CD4+ T cell subset was only inhibited by tAFP-mDC in comparison to nAFP-mDC (P < 0.05) and OVA-mDC control (P < 0.01).

Figure 4. tAFP-conditioned DC fail to induce robust allogeneic T cell proliferation.

Figure 4

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Monocytes from healthy donors (n = 5) were differentiated into DC for 5 days in the presence of OVA, nAFP, or tAFP (all at 10 µg/ml). DC were then matured for an additional 24 hours prior to collection. (A) Conditioned mDCs were cultured with allogeneic CFSE-labeled CD8+ or CD4+ T cells for 6 days. T cell proliferation was assessed by measuring CFSE dilution. The solid line represents the mean value. For each allogeneic donor pair, nAFP-mDC- and tAFP-mDC-induced proliferation was normalized to control OVA-mDC-induced proliferation (set at 100%). (B) Conditioned mDCs were cultured with allogeneic CD8+ or CD4+ T cells for 7 days. After stimulation with PMA and ionomycin, the cells were surface stained for CD8 or CD4 expression, and intracellular cytokine production was assayed by flow cytometry. Data are reported as the mean ± SD. ns, P ≥ 0.05; *, P < 0.05; **, P < 0.01.

Alloreactive T cells were also analyzed for the ability to produce inflammatory cytokines. After a brief restimulation, the percentage of IL-2+, TNF-α+, or IFN-γ+ T cells was unaltered between the OVA-, nAFP-, and tAFP-mDC groups (Fig. 4B). This was true for both CD8+ and CD4+ T cell subsets.

tAFP-conditioned DC are poor stimulators of antigen-specific T cell expansion

We next employed a more biologically relevant model by using autologous AFP-conditioned DC to stimulate Flu peptide-specific T cell activation. Autologous CD8+ T cells were stimulated for 11 days by FluM158–66 peptide-pulsed OVA-, nAFP-, and tAFP-mDC. Multimer staining revealed that the expansion of FluM158–66 peptide-specific CD8+ T cells was impaired in both nAFP- and tAFP-mDC groups (Fig. 5A). To standardize the variability that was observed between donors (multimer-positive cells in the OVA-mDC group ranged from 2.4–7.5%), nAFP-mDC and tAFP-mDC groups were normalized to control OVA-mDC (set at 1.0% for each individual donor). As depicted in the right panel of Figure 5A, expansion of FluM158–66 peptide-specific CD8+ T cells from three healthy donors was significantly dimished by nAFP-mDC (P < 0.05), and reduced still more by tAFP-mDC (P < 0.01), compared to OVA-mDC control.

Figure 5. Antigen-specific T cell expansion, but not effector function, is blunted by tAFP-conditioned DC.

Figure 5

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(A and B) Monocytes from healthy HLA-A2+ donors were differentiated into DC in the presence of OVA, nAFP, or tAFP, then matured overnight prior to collection. Conditioned mDCs were cultured with autologous CD8+ T cells in the presence of peptide FluM158 for 11 days. (A) Representative FluM158–66 pentamer staining (left) and from all donors (right) is shown. For each donor (n = 3), nAFP-mDC and tAFP-mDC groups were normalized to control OVA-mDC group, which was set at 1.0%.*, P < 0.05; **, P < 0.01. (B) After a 6 hour stimulation of collected CD8+ T cells with FluM158–66, intracellular cytokine production was assayed by flow cytometry. (C) Monocytes from healthy HLA-DR4+ donors were differentiated into DC in the presence of OVA, nAFP, or tAFP, then matured overnight prior to collection. Conditioned mDCs were cultured with autologous CD8+ T cells in the presence of peptide HA307 for 11 days. After a 6 hour stimulation of collected CD4+ T cells with HA307–319, intracellular cytokine production was assayed by flow cytometry. (B and C) For each donor (n = 4), nAFP-mDC- and tAFP-mDC-induced cytokine production was normalized to control OVA-mDC-induced cytokine production, which was set at 1.0%. ns, P ≥ 0.05.

CD8+ T cells from this FluM158–66 in vitro stimulation were further studied for cytokine production following a brief peptide-specific restimulation. Again, there was notable donor-to-donor variability. In the OVA-mDC control group, TNF-α+ cells ranged from 0.2–1.1%, and IFN-γ+ from 0.2–2.1%. nAFP-mDC and tAFP-mDC groups were normalized to control OVA-mDC (set at 1.0%). No alterations in TNF-α or IFN-γ production were observed, however, between the different DC treatment groups (Fig. 5B).

Next, autologous CD4+ T cells were stimulated with HA307–319 peptide-pulsed OVA-, nAFP-, and tAFP-mDC. After 11 days, cells were briefly restimulated and intracellular cytokine production was determined by flow cytometry. Donor variability (OVA-mDC control, TNF-α: 0.5–2.3%; IFN-γ: 0.3–1.2%) was standardized by setting OVA-mDC-induced cytokine production at 1.0%. Again, HA307–319 peptide-specific cytokine production by CD4+ T cells was not impaired by nAFP- or tAFP-mDC (Fig. 5C). Along with the results from the MLR, these data demonstrate that although effector function is unaltered, the ability of T cells to proliferate in response to both alloantigen and cognate antigen was significantly impaired by AFP-conditioned DC.

A low molecular weight AFP co-purifying ligand is required for tAFP-induced DC dysfunction

Previous reports have suggested that protein conformation (24), the degree of glycosylation (25), and fucosylation status (26) play important roles in the function of AFP. However, when OVA, nAFP, and tAFP preparations were heat denaturated or treated with the enzymes PNGase F or α-L-fucosidase (to remove N-linked glycan or fucose residues, respectively), tAFP retained suppressive activity, as evidenced by diminished levels on DC of the surface markers CD206 (Fig. 6A), as well as HLA-ABC, CD40, CD80, and CD83 (data not shown). To evaluate whether the inhibitory activity of tAFP is dependent on full length protein, we next pretreated OVA, nAFP, and tAFP with a serine protease (proteinase K). As shown in Figure 6A, CD206 expression in DC treated with digested tAFP was restored to levels observed in digested OVA- or nAFP-treated cells, indicating that intact primary structure is necessary for tAFP activity. Similarly, expression of HLA-ABC, CD40, CD80, and CD83 (data not shown) was almost completely restored to control levels with protease-digested tAFP.

Figure 6. The inhibitory activity of tAFP requires intact whole protein and low molecular weight co-purification products.

Figure 6

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(A) DCs were differentiated for 5 days in the presence of native OVA, nAFP, or tAFP (all at 10 µg/ml), or the same preparations prepared with heat denaturation or enzymatic treatment with PNGase, fucosidase, or protease (as described in Materials & Methods). (B) OVA, nAFP, and tAFP (all at 10 µg/ml) were separated into high or low molecular weight (HMW & LMW, respectively) fractions with a 3kDa molecular weight cut off (MWCO). DCs were differentiated for 5 days in the presence of intact native preparations, HMW fraction, LMW fraction, or both HMW & LMW fractions together, as indicated. (A and B) MFI of CD206 is shown. Similar patterns were observed for HLA-ABC, CD40, CD80, and CD83 (data not shown). Columns, mean of four HD; bars, standard deviation. ns, P ≥ 0.05; *, P < 0.05; **, P < 0.01.

Because one of the main functions of AFP (69kDa) is to bind and transport low molecular weight (LMW) hydrophobic ligands (4), we divided our OVA, nAFP, and tAFP preparations into high molecular weight (HMW) and LMW fractions (>3kDa and <3kDa, respectively). Compared to the native tAFP preparation, neither the isolated HMW nor LMW fractions of tAFP inhibited DC expression of CD206 (Fig. 6B), HLA-ABC, CD40, CD80, and CD83 (data not shown). However, when both HMW and LMW fractions of tAFP were added to cultures together, we observed suppressive activity to the extent of the native tAFP preparation. We next dialyzed tAFP against PBS using membranes of various molecular weight cutoffs (MWCO). Even using a MWCO of 0.5kDa, the HMW fraction of tAFP lacked suppressive activity (data not shown), indicating that the co-purifying ligands in the LMW fraction of tAFP are <0.5kDa in size. Extensive analysis of our native AFP preparations and isolated HMW and LMW fractions via mass spectrometry revealed no unexpected protein or peptide species (other than AFP itself) or glycosylation pattern that correlated with the immunosuppressive activity of AFP (data not shown), confirming the expected identity of these preparations, and suggesting that the LMW ligands are not peptide-based molecules.

Finally, to further characterize the LMW species that function cooperatively with tAFP, cell lysates were prepared from HepG2 hepatoma cells and healthy donor PBMC (representing tumor and non-tumor tissue, respectively), and either left whole or separated into HMW and LMW fractions. DCs were then differentiated for 5 days in the presence of lysates with or without the purified (HMW) fractions of OVA, nAFP, or tAFP. As illustrated in Figure 7, CD206 expression was unaltered by HepG2 and PBMC lysates (both whole and fractionated) alone. No change was also observed when LMW lysates were added to cultures in combination with HMW OVA or nAFP. However, when the LMW lysates of either HepG2 cells or PBMC were added to cultures together with HMW tAFP, expression levels of CD206 (Fig. 7), HLA-ABC, CD40, CD80, and CD83 (data not shown) were significantly reduced, effectively reproducing the tAFP-DC phenotype. These data suggest that the immunoregulatory LMW co-purifying species in our tAFP preparations is a ubiquitous self molecule that is present in both tumor and healthy tissue.

Figure 7. LMW cell lysates in combination with purified tAFP fully recapitulate the tAFP-DC phenotype.

Figure 7

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Lysates were prepared from HepG2 hepatoma cells and healthy donor PBMC, and either left whole (“whole lys”) or separated into HMW and LMW fractions (“HMW lys” and “LMW lys”, respectively). OVA, nAFP, and tAFP (all at 10 µg/ml) were either left intact or separated into HMW and LMW fractions. DCs were differentiated for 5 days in the presence of these preparations, as indicated. MFI of CD206 is shown. Similar patterns were observed for HLA-ABC, CD40, CD80, and CD83 (data not shown). Columns show mean of three HD; bars indicate standard deviation. *, P < 0.05; **, P < 0.01.

Cumulatively, we have observed that the ability of tAFP to impair DC differentiation and function is not dependent on native protein conformation or glycosylation status, but rather full-length AFP protein, along with the presence of LMW co-purifying species, are absolutely essential for tAFP to exert its immunosuppressive activity.

DISCUSSION

Compared with healthy donors, reduced frequencies of circulating myeloid DC are observed in the peripheral blood of HCC patients, while DC generated in vitro from HCC patient PBMC exhibit impaired IL-12 production and limited allo-stimulatory activity (18, 27). In this report, we show a significant relationship between AFP serum levels and the ratio of distinct DC subsets in the peripheral blood of HCC patients. Several tumor-derived factors have been implicated in DC dysfunction in cancer patients and tumor-bearing mice. These include VEGF, gangliosides, prostaglandins, TGF-β, and indoleamine 2,3-dioxygenase (IDO) (28). Our current data suggests that AFP, which can be found at high levels in the serum of patients with gastrointestinal (i.e. HCC, stomach, pancreatic) and reproductive (i.e. yolk sac, teratoma) cancers, serves as a key regulator of DC differentiation.

Consistent with a previous study, nAFP exerted inhibitory activity in our DC cultures at higher concentrations (>10 µg/ml) (9). However, tumor-derived AFP preparations induced aberrant DC differentiation at much lower doses. These tAFP-DC retain a monocyte-like phenotype and morphology, down-regulate MHC and costimulatory molecule expression, and produce limited levels of inflammatory cytokines and chemokines, including IL-8, MCP-1, MIP-1α, and MIP-1β. This reduced stimulatory phenotype involves NF-κB pathway signaling. We have recently identified DC-derived IL-8 as a crucial factor in NK cell chemotaxis (22). Moreover, the chemokines MCP-1/CCL2, MIP-1α/CCL3, and MIP-1β/CCL4 are known to recruit NK cells to sites of inflammation and enhance their IFN-γ production and cytolytic activity (21, 29). Notably, limited numbers of NK cells have been reported in the peripheral blood and tumor lesions of HCC patients (30), and the density of intratumoral NK cells correlates with patient survival (31). Our data suggest that this paucity of NK cells in HCC patients, which serves as a significant barrier to anti-cancer immunity, may be a result of tAFP-mediated mechanisms that reduce inflammatory chemokine production. This is a new area we are currently investigating.

Based on the poorly-differentiated phenotype and limited cytokine/chemokine profile of tAFP-DC, we hypothesized that these cells would be unable to induce a robust T cell response. Indeed, in both allogeneic and autologous T cell stimulation models, tAFP-DC, and nAFP-DC to a lesser extent, drove limited T cell proliferation compared to OVA-DC control. Conversely, cytokine production by activated T cells was not impaired by nAFP- and tAFP-DC. This dichotomy may be explained by a recent study, where it was reported that while a low density of T cell receptor (TCR)-CD3 complexes was sufficient to induce cytokine secretion, the threshold for T cell proliferation was significantly higher (32). Therefore, while tAFP-DC may be capable of engaging a sufficient level of TCR-CD3 complexes to allow for cytokine production by T cells, they are unable to surpass the threshold necessary to drive proliferation.

In order to identify the molecular basis of tAFP’s immune suppressive activity, nAFP and tAFP preparations were tested under several conditions to identify the impact of glycosylation, structure, and conformation. Currently, the only reported variation between nAFP and tAFP isolates is altered glycosylation, namely the predominance of the fucosylated variant in tAFP preparations (33). However, when we pretreated OVA, nAFP, and tAFP with fucosidase, the immunosuppressive activity of tAFP remained intact. Similarly, when these preparations were pretreated with the glycosidase PNGase F, the immunosuppressive activity of tAFP was unaltered. This is consistent with an earlier study, where it was shown that several different AFP isolates exerted immunoregulatory activity, irrespective of their carbohydrate content (34).

An alternative hypothesis is that only specific conformational states of AFP protein possess immune suppressive activity. AFP can assume three different conformations: i) the natural compact conformation found during circulation in the blood, ii) a slightly denatured state, termed the molten globule form (MGF), typically present when AFP is localized in the cytoplasmic compartment of cells, and iii) a completely denatured state induced by extreme, non-physiologic environments (4, 24). Heat denaturation of our AFP preparations failed to abrogate tAFP-mediated DC dysfunction, indicating that tAFP in both its compact (natural) and completely denatured states is immunosuppressive.

One of the main functions of AFP is to bind and transport LMW hydrophobic ligands such as fatty acids (35), neopterin (G. Mizejewski, personal communication, 2013), and bilirubin (36). Immunologically, it has been shown that high levels of these serum factors are capable of inducing immune dysfunction (37–39). Setiyono et al. report that domains 2 and 3, but not domain 1, of the AFP protein possess immunoregulatory activity (12). Notably, domain 2 contains the major fatty acid-binding site of the protein, providing further evidence of a potential role for fatty acids in tAFP-based DC dysfunction (40). These observations, along with our current data, support a model in which LMW species that i) co-purify with tAFP, and ii) are abundant in the LMW fractions of both tumor and non-tumor cell lysates are efficiently chaperoned into monocytes/DC by tAFP, where they interfere with molecular programming pathways and inhibit full DC differentiation. We are currently using mass spectrometry and biochemical methods to further uncover the identity of this molecule(s).

Purified AFP is being explored as a therapeutic in several clinical settings. Based on its immunosuppressive properties, recombinant AFP is under development for the treatment of autoimmune diseases (41). In the setting of AFP-expressing cancers, namely HCC, AFP serves as an attractive target for immunotherapy, and several groups have investigated the anti-cancer potential of HCC tumor lysate- or AFP-based vaccines (16, 42, 43). Our data suggest that caution must be applied in the formulation of these therapeutics. In the setting of autoimmune therapy, it will be essential to maintain AFP co-purifying, LMW immunoregulatory species, while for AFP-based cancer vaccines, considerable effort must be taken to eliminate this component.

Supplementary Material

1

ACKNOWLEDGMENTS

The authors wish to thank Drs. Jerry Mizejewski and Tim Greten for their constructive comments used in developing this manuscript, and Drs. T. Clark Gamblin and David A. Geller for providing several HCC patient blood samples.

Financial Support: This study was supported by research funding from the University of Pittsburgh Cancer Institute and NCI RO1 CA 138635 (LHB). This project used the UPCI Immunologic Monitoring and Cellular Products Laboratory (Lisa H. Butterfield, PhD, Director) and the UPCI Cancer Biomarkers Facility (mass spectrometry services: Nathan Yates, PhD; genomics services: William La Framboise, PhD) that are supported in part by award P30CA047904.

List of abbreviations

AFP

alpha-fetoprotein

nAFP

cord blood-derived normal AFP

tAFP

tumor-derived AFP

OVA

ovalbumin

HCC

hepatocellular carcinoma

DC

dendritic cell

mDC

mature DC

LMW

low molecular weight

HMW

high molecular weight

MWCO

molecular weight cutoff

PBMC

peripheral blood mononuclear cells

MLR

mixed lymphocyte reaction

MDSC

myeloid-derived suppressor cell

Footnotes

Disclosure of Conflicts of Interest: ADP and JS: none to declare. LHB is co-inventor of patents covering aspects of AFP as a target for T cell-mediated anti-HCC immunity.

Microarray Data: The data discussed in this publication have been deposited in NCBI’s Gene Expression Omnibus (Edgar et al., 2002) and are accessible through GEO Series accession number GSE62005 (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE62005).

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