Dendritic cells (DC) are professional APCs that regulate innate and adaptive immunity. The role of fatty-acid synthesis in DC development and function is uncertain. We found that blockade of fatty-acid synthesis markedly decreases dendropoiesis in the liver and in primary and secondary lymphoid organs in mice. Human DC development from PBMC precursors was also diminished by blockade of fatty-acid synthesis. This was associated with higher rates of apoptosis in precursor cells and increased expression of cleaved caspase-3 and BCL-xL and downregulation of cyclin B1. Further, blockade of fatty-acid synthesis decreased DC expression of MHC class II, ICAM-1, B7-1, and B7-2 but increased their production of selected proinflammatory cytokines including IL-12 and MCP-1. Accordingly, inhibition of fatty-acid synthesis enhanced DC capacity to activate allogeneic as well as Ag-restricted CD4+ and CD8+ T cells and induce CTL responses. Further, blockade of fatty-acid synthesis increased DC expression of Notch ligands and enhanced their ability to activate NK cell immune phenotype and IFN-γ production. Because endoplasmic reticulum (ER) stress can augment the immunogenic function of APC, we postulated that this may account for the higher DC immunogenicity. We found that inhibition of fatty-acid synthesis resulted in elevated expression of numerous markers of ER stress in humans and mice and was associated with increased MAPK and Akt signaling. Further, lowering ER stress by 4-phenylbutyrate mitigated the enhanced immune stimulation associated with fatty-acid synthesis blockade. Our findings elucidate the role of fatty-acid synthesis in DC development and function and have implications to the design of DC vaccines for immunotherapy.

Dendritic cells (DC) have emerged over the past two decades as the most specialized professional APCs, which initiate adaptive and innate immune responses (1). As a consequence, DC have an important role in immune surveillance against developing cancer or invading pathogens and have potential to serve as vehicles for immunotherapy. Hence, elucidating the cellular biochemistry of DC has implications both for understanding immunity and for the design of immunotherapy regimens.

Fatty-acid synthesis is an essential element of cellular metabolism. However, its role in DC development and function is uncertain. A study by Zeyda et al. (2) investigated the effects of exogenous administration of polyunsaturated fatty acids (PUFA) to DC and found that PUFA block DC immunogenic function independent of NF-κB activation. In particular, DC capacity for T cell activation was markedly inhibited in DC treated with PUFA. Similarly, a more recent report by Herber et al. (3) found that DC acquire exogenous lipids within the tumor microenvironment in both mice and humans, which renders them poorly functional, accounting for their inability to generate a potent antitumor immune response. The diminished DC immunogenicity facilitates the cancer’s capacity to evade immune recognition. Nevertheless, whereas exogenous fatty-acids, either directly administered or accumulated in tumor bearing hosts, appear to lessen the immunogenic potential of DC, the role of endogenous fatty-acid synthesis on dendropoiesis in vitro and in vivo and on DC functional properties is uncertain. In this study, we found that blocking fatty-acid synthesis using either inhibitors of acetyl CoA carboxylase or fatty-acid synthase diminished dendropoiesis from bone marrow or PBMC precursors. However, surprisingly, inhibition of fatty-acid synthesis upregulated DC expression of TLRs and markedly augmented DC capacity to stimulate Ag-restricted CD4+ and CD8+ T cells, induce CTL, and activate innate immune effector cells. Our mechanistic studies revealed that blockade of fatty-acid synthesis enhances MAPK, PI3K/Akt, and Notch signaling in DC and leads to higher endoplasmic reticulum (ER) stress. These findings suggest an important role for fatty-acids synthesis in modulating basic DC biology and have implications for the design of more effective immunotherapy regimens.

Male C57BL/6 (H-2kb), BALB/c (H-2kd), OT-I (B6.Cg-RAG2tm1Fwa-TgN), and OT-II (B6.Cg-RAG2tm1Alt-TgN) mice were purchased from Taconic Farms (Germantown, NY). Age-matched 6–8-wk-old mice were used in experiments. Animals were housed in a clean vivarium and fed standard mouse chow. In selected experiments, mice were injected three times weekly for 4 wk with saline or C75 (250 μg, i.p.; Sigma-Aldrich, St. Louis, MO), an inhibitor of fatty-acid synthase. Animal procedures were approved by the New York University School of Medicine Institutional Animal Care and Use Committee.

Bone marrow–derived DC (BMDC) were generated as described (4). Briefly, bone marrow aspirates were cultured for 8 d in complete RPMI (RPMI 1640 with 10% heat-inactivated FBS, 2 mM L-glutamine, and 0.05 mM 2-ME) supplemented with GM-CSF (20 ng/ml). To generate human monocyte-derived DC (moDC), leukocyte-enriched buffy coats were obtained from the New York Blood Center. PBMCs were separated by density gradient centrifugation on Ficoll-Hypaque (GE Healthcare, Piscataway, NJ). Cells were cultured for 5–7 d in complete RPMI supplemented 10% human serum, 800 U/ml GM-CSF, and 1000 U/ml IL-4 (R&D Systems, Minneapolis, MN). In selected experiments, acetyl CoA carboxylase was inhibited in murine BMDC or human moDC cellular suspensions using tall oil fatty acid (TOFA; 5 μg/dl; Cayman Chemical, Ann Arbor, MI) beginning on day 2 of culture (57). In selected experiments, a lower dose of TOFA was used (1 μg/dl). Ethanol (0.5%) was used a solvent for TOFA. In additional experiments, staurosporine (10 μM) was employed to induce DC apoptosis (8, 9).

Murine hepatic nonparenchymal cells were isolated as described (10). Briefly, the portal vein was infused with Collagenase IV (Sigma-Aldrich) followed by hepatectomy and mechanical digestion. Hepatocytes were excluded by serial low-speed (300 rpm) centrifugation. Nonparenchymal cells were further enriched over an Optiprep (Sigma-Aldrich) gradient. Splenocytes were isolated by manual disruption of whole spleen. In selected experiments, splenic T cells and NK cells were purified by FACS or using anti-CD90, anti-CD4, anti-CD8, or anti-DX5 immunomagnetic beads, respectively, and passage through positive selection columns (Miltenyi Biotec, Bergisch-Gladbach, Germany).

Flow cytometry was performed using the FACSCalibur (BD Biosciences, Franklin Lakes, NJ) after incubating 5 × 105 cells/tube with 1 μg anti-FcγRIII/II Ab (2.4G2, Fc block; mAb Core, Sloan-Kettering Institute, New York, NY) and then labeling with 1 μg fluorescently conjugated mAb against MHC class II (MHC II; I-Ab), CD4 (RM4-5) CD8α (53-6.7), CD11b (M1/70), CD11c (HL3), CD19 (1D3), CD25 (3C7), CD40 (HM40-3), CD45 (30-F11), CD54 (YN1/1.7.4), B7-1 (16-10A1), B7-2 (GL1), Foxp3 (FJK-16s; all from eBioscience, San Diego, CA), CD3ε (145-2C11; BioLegend, San Diego, CA), and TLR2 (T2.5) (Imgenex, San Diego, CA). Alternatively, cells were labeled with unconjugated Abs against Jagged-1 (Santa Cruz Biotechnology, Santa Cruz, CA), TLR4, TLR7, and TLR9 (all from Imgenex) and subsequently stained with fluorescently labeled secondary Abs. Human moDC were tested using mAbs directed against HLA-DR and CD11c (BD Biosciences). For cytokine analysis, cell suspensions were cultured in complete RPMI at a concentration of 1 × 106 cells/ml for 24 h before supernatant harvest and analysis either using either a cytometric bead array (BD Biosciences) or the Milliplex Immunoassay (Millipore, Billerica, MA).

For CD4+ or CD8+ T cell proliferation assays, BMDC were pulsed with the appropriate Ova peptide (10 μg/ml; Abcam, Cambridge, Massachusetts) for 90 min before washing and plating, respectively, with CD4+OT-II TCR-transgenic T cells (1 × 105) specific for Ova323–339 peptide or CD8+OT-I TCR-transgenic T cells specific for Ova257–264 peptide for 72 h in 96-well plates as described (10). In selected experiments, non–peptide-pulsed DC were used to stimulate allogeneic BALB/c T cells in an MLR as described (10). For cross-presentation experiments, DC were loaded with OVA (1 mg/ml; Sigma-Aldrich) and used to stimulate OT-I T cells as described (10). For the last 24 h, 1 μCi [3H]thymidine was added to wells and proliferation measured using a MicroBeta counter (PerkinElmer, Waltham, MA). Alternatively, T cell activation was assessed by measuring Th1, Th2, and Th17 cytokine production using a cytometric bead array or by examination of T cell surface phenotype. In selected experiments, soluble inhibitors of PI3K (50 μM; LY294002) and MAPK (100 μM; PD98059; both Invivogen, San Diego, CA) signaling were employed. To assess BMDC induction of regulatory T cells, DC were cultured with equal numbers of splenocytes before determination of CD4+ T cell coexpression of CD25 and Foxp3 at 96 h. In some experiments, BMDC were incubated overnight with the chemical chaperone 4-phenylbutyrate (10 mM; Sigma-Aldrich) before washing, peptide loading, and coculture with T cells. To determine the ability of DC to produce a CTL response in vivo, naive mice were immunized i.p. twice at weekly intervals with DC.Ova257–264 (3 × 105) or mock immunized. One week later, splenocytes were harvested from immunized mice, restimulated in vitro with Ova257–264, and cell-culture supernatant assayed for IFN-γ and IL-10 as described (4).

DC–NK cell cocultures were performed as described with slight modifications (4). Briefly, splenic NK cells (1 × 105) were plated with BMDC (1 × 105) in a 1:1 ratio in 96-well plates for 24 h. IFN-γ was measured in cell-culture supernatant using a cytometric bead assay (BD Biosciences). In addition, NK cell expression of CD25 was analyzed by flow cytometry.

To assess DC capacity for Ag uptake, BMDC were incubated with FITC-dextran, FITC-albumin, or FITC-mannose albumin (1 mg/ml; all from Sigma-Aldrich) at 37°C for various time intervals. Ag uptake was determined by flow cytometry. For in vivo Ag uptake, control or C75-treated mice were injected i.p. with 1 mg FITC-albumin. Mice were then sacrificed at 30 min and splenic DC fluorescence determined by flow cytometry.

For light and fluorescent microscopic analysis, cells were spun onto slides and stained with H&E, Giemsa, HCS LipidTOX Red specific for neutral lipids, and HCS LipidTOX Green specific for phospholipids (Invitrogen, Grand Island, NY). Light microscopic images were captured using an Axiovert 40 microscope (Zeiss, Thornwood, NY). Fluorescent images were captured on an Axiovert 200M (Zeiss). Cells were also tested by flow cytometry using BODIPY (Invitrogen) as described (11). For electron microscopic analysis, BMDC suspensions were first incubated with 4% formaldehyde and 2% glutaraldehyde in 0.1 M (pH 7.5) PIPES buffer (PB) for 30 min at room temperature. Cells were further fixed with 4% glutaraldehyde and 0.4% tannic acid in PB for 30 min followed by 2% osmium tetroxide in PB for 1 h. The samples were then counterstained with 2% uranylacetate for 12 h at 4°C before exchanging the solution with ethanol. Samples were then infused with epoxy resin and polymerized at 60°C. The sample blocks were sectioned to a thickness of 50–70 μm, collected on electron microscope grids, and stained with uranylacetate and Sato Lead Stain. The samples were then imaged using a Philips CM12 microscope (Philips, Eindhoven, The Netherlands). For each section, the cellular, nuclear, and cytosolic areas were estimated by assuming the cell and nucleus sections are ellipses and measuring the long and short axes. To measure the rate of intracellular fatty-acid synthesis in BMDC, C-14–labeled acetate was added to BMDC cultures (2 μCi/well) for 6 h. More than 40 cells from each treatment group were analyzed. Intracellular lipids were isolated by the Folsch extraction method, and C-14 uptake was determined by scintigraphy as described (11).

Western blotting was performed as described (11, 12). Briefly, BMDC were homogenized in RIPA buffer, and proteins were separated from larger fragments by centrifugation at 14000 × g. Samples were equiloaded onto 10% polyacrylamide gels (NuPage; Invitrogen), electrophoresed at 200 V, electrotransferred to polyvinylidene difluoride membranes, and probed with mAbs to GRP-78, eukaryotic translation initiation factor 2α (eIF2α), p-eIF2α, XBP-1, peroxisome proliferator–activated receptor γ (PPAR-γ), caspase-3, BCL-xL, cyclin B1, Jagged-1, Δ-4, Akt, p-Akt, Erk1, p-Erk1, NF-κB, p–NF-κB, phosphatase and tensin homolog, p38MAPK, p-p38MAPK, p70S6 kinase, and β-actin (all from Santa Cruz Biotechnology). Blots were developed by ECL (Thermo Scientific, Asheville, NC).

Statistics were calculated using GraphPad Prism V5.00 (GraphPad Software, San Diego, CA). Data are presented as mean ± SEM. Statistical significance (p < 0.05) was determined using the Student t test and the log-rank test.

To determine whether blockade of fatty-acid synthesis in vivo affects dendropoiesis in lymphoid and nonlymphoid organs, mice were serially administered C75, an inhibitor of fatty-acid synthase (13, 14), and the number of CD11c+ cells was measured in the bone marrow, spleen, and liver. Treatment for 4 wk resulted in an 80% reduction in the fraction and total number of CD11c+ cells in the liver (Fig. 1A, 1B) and an ∼20% reduction in the spleen and bone marrow (Fig. 1B). Other cell types, including B cells, T cells, neutrophils, and macrophages, were not affected (Fig. 1C).

FIGURE 1.

Blockade of fatty-acid synthesis inhibits dendropoiesis in mice and humans. (AC) Mice were treated for 4 wk with C75 or saline. (A) Live CD45+ liver leukocytes were gated using flow cytometry, and the subfraction of hepatic CD11c+ cells was determined. (B) The percentage decrease in the number of liver, spleen, and bone marrow DC was calculated. (C) The fraction of splenocytes expressing CD3, CD19, and CD11b in saline- or C75-treated mice was tested. (DG) BMDC were grown alone or with TOFA. (D) The fraction of propidium iodide (PI)+ cells was calculated on day 8 of culture. (E) Day 8 BMDC and T-BMDC were also tested for expression of caspase-3, cleaved caspase-3, BCL-xL, cyclin B1, and β-actin by Western blotting. (F) In addition, the total number and fraction of CD11c+ cells was calculated in day 8 BMDC and T-BMDC cultures. (G) Cellular proliferation was compared in day 8 BMDC and T-BMDC by pulsing with [3H]thymidine. (H) moDC grown in control media and TOFA-enriched media were tested for HLA-DR and CD11c expression. Median fluorescence intensity is indicated for each respective histogram (*p < 0.05, **p < 0.01, ***p < 0.001).

FIGURE 1.

Blockade of fatty-acid synthesis inhibits dendropoiesis in mice and humans. (AC) Mice were treated for 4 wk with C75 or saline. (A) Live CD45+ liver leukocytes were gated using flow cytometry, and the subfraction of hepatic CD11c+ cells was determined. (B) The percentage decrease in the number of liver, spleen, and bone marrow DC was calculated. (C) The fraction of splenocytes expressing CD3, CD19, and CD11b in saline- or C75-treated mice was tested. (DG) BMDC were grown alone or with TOFA. (D) The fraction of propidium iodide (PI)+ cells was calculated on day 8 of culture. (E) Day 8 BMDC and T-BMDC were also tested for expression of caspase-3, cleaved caspase-3, BCL-xL, cyclin B1, and β-actin by Western blotting. (F) In addition, the total number and fraction of CD11c+ cells was calculated in day 8 BMDC and T-BMDC cultures. (G) Cellular proliferation was compared in day 8 BMDC and T-BMDC by pulsing with [3H]thymidine. (H) moDC grown in control media and TOFA-enriched media were tested for HLA-DR and CD11c expression. Median fluorescence intensity is indicated for each respective histogram (*p < 0.05, **p < 0.01, ***p < 0.001).

Close modal

To investigate the effects of inhibition of fatty-acid synthesis on DC generation in vitro from bone marrow precursors, we isolated bone marrow cells and cultured them in GM-CSF–supplemented media for 8 d to drive dendropoiesis, as described (4). In parallel, for the duration of in vitro culture, bone marrow cells were coincubated with TOFA, which inhibits acetyl CoA corboxylase (15, 16). The number of nonviable propidium iodide+ cells was increased on day 8 of culture (Fig. 1D) as well as at earlier time points (not shown) in cellular suspensions incubated with TOFA. Further, there was increased expression of cleaved caspase-3 and BCL-xL in TOFA-treated BMDC (T-BMDC), consistent with increased rates of apoptosis (Fig. 1E). Accordingly, cyclin B1, an anti-apoptotic gene, was downregulated in T-BMDC (Fig. 1E). The total number and fraction of CD11c+ cells produced per mouse femur (Fig. 1F) and BMDC cellular proliferation (Fig. 1G) were also lower in TOFA-treated bone marrow cultures. Generation of human moDC was similarly hindered by TOFA (Fig. 1H). Furthermore, serial in vivo administration of C75 resulted in less-efficient generation of BMDC after bone marrow harvest (Supplemental Fig. 1A). Taken together, these data show that blockade of fatty acid synthesis inhibits dendropoiesis in vitro and in vivo and in both mice and humans.

As anticipated, bone marrow–derived cells grown in TOFA exhibited a decreased rate of fatty-acid synthesis (Fig. 2A). Accordingly, on both electron microscopy and light microscopy, T-BMDC exhibited decreased vacuolization and numbers of lipid droplets (Fig. 2B, 2C, Supplemental Fig. 1B). Similarly, HCS LipidTOX Red staining revealed a substantial reduction in total neutral lipids (Fig. 2D, Supplemental Fig. 1C), and HCS LipidTOX Green staining revealed decreased phospholipid levels in T-BMDC (Fig. 2E, Supplemental Fig. 1D). Further, T-BMDC had diminished staining for BODIPY, which binds total neutral lipids (Supplemental Fig. 1E).

FIGURE 2.

Blockade of fatty-acid synthesis alters DC phenotype. (A) C14 acetate uptake was compared in day 8 BMDC and T-BMDC cultures. Day 8 BMDC and T-BMDC were examined by electron microscopy (B), H&E and Giemsa staining (C), or immunofluorescence using HCS LipidTOX Red (D), which binds neutral lipids, and HCS LipidTOX Green (E), which binds phospholipids. Original magnification ×40. CD11c+ cells in BMDC or T-BMDC cultures were gated and analyzed for expression of MHC II, adhesion, and costimulatory molecules (F) as well as TLR2, TLR4, TLR7, and TLR9 (G). Median fluorescence intensities are indicated for each respective histogram. ***p < 0.001.

FIGURE 2.

Blockade of fatty-acid synthesis alters DC phenotype. (A) C14 acetate uptake was compared in day 8 BMDC and T-BMDC cultures. Day 8 BMDC and T-BMDC were examined by electron microscopy (B), H&E and Giemsa staining (C), or immunofluorescence using HCS LipidTOX Red (D), which binds neutral lipids, and HCS LipidTOX Green (E), which binds phospholipids. Original magnification ×40. CD11c+ cells in BMDC or T-BMDC cultures were gated and analyzed for expression of MHC II, adhesion, and costimulatory molecules (F) as well as TLR2, TLR4, TLR7, and TLR9 (G). Median fluorescence intensities are indicated for each respective histogram. ***p < 0.001.

Close modal

Because we found that inhibition of fatty-acid synthesis prevents dendropoiesis, we postulated that it may also affect BMDC maturation. To test this, bone marrow–derived CD11c+ cells were analyzed for expression of MHC II, costimulatory, and adhesion molecules. As anticipated, T-BMDC exhibited decreased expression of MHC II, ICAM-1, B7-1, and B7-2 (Fig. 2F). However, CD40 and CD11b were consistently upregulated in BMDC grown in TOFA (Fig. 2F). Similar phenotypic differences between T-BMDC and controls were seen when gated exclusively on CD11c+MHC II+ cells (not shown). Surprisingly, despite a diminished maturational phenotype, blockade of fatty-acid synthesis upregulated DC surface expression of TLR2 and TLR4 and intracellular expression of TLR7 and TLR9 (Fig. 2G). Conversely, in contrast to the effects of TOFA, staurosporine, which also induced BMDC apoptosis (Supplemental Fig. 2A), upregulated MHC II expression on BMDC (Supplemental Fig. 2B), and did not increase BMDC TLR expression (Supplemental Fig. 2C), suggesting that effects of TOFA are specific to fatty-acid synthesis inhibition.

ER stress can have marked affects on the immune-stimulatory capacity of APCs (1719). Because inhibition of fatty-acid synthesis induces ER stress in neoplastic cells (20), we postulated that TOFA-grown BMDC would exhibit high ER stress. In consort with our hypothesis, we found that GRP-78, eIF2α, p-eIF2α, and XBP-1, all markers of ER stress (21), were more highly expressed in T-BMDC compared with controls (Fig. 3A). Human moDC generated in TOFA also expressed markedly elevated p-eIF2α (Fig. 3B). Higher PPAR-γ expression has been linked to increased ER stress and is associated with enhanced DC capacity to present Ag (2225). Accordingly, we found substantial upregulation of PPAR-γ expression in murine T-BMDC at both the protein (Fig. 3C) and mRNA levels (Fig. 3D). TOFA-treated human moDC also expressed higher PPAR-γ (Fig. 3E).

FIGURE 3.

Inhibition of fatty-acid synthesis during DC development increases ER stress and alters DC production of inflammatory mediators. (A) Expression of markers of ER stress (GRP-78, eIF2α, p-eIF2α, XBP-1, and β-actin) was tested in day 8 BMDC and T-BMDC by Western blotting. (B) Human moDC generated in control or TOFA-enriched media were tested for expression of selected ER stress markers. Control or TOFA-treated murine BMDC were tested for expression of PPAR-γ by Western blotting (C) and PCR (D). (E) Human moDC generated in control media or TOFA-supplemented media were tested for expression of PPAR-γ by Western blotting. β-actin was used as a loading control. (FH) Twenty-four–hour cell-culture supernatants from day 8 BMDC and T-BMDC plated at equal densities were tested for the presence of numerous cytokines and chemokines. (I) Control BMDC, standard high-dose T-BMDC, low-dose T-BMDC, ethanol-treated BMDC (EtOH), and staurosporine-treated BMDC were tested for their capacity to produce MCP-1 (*p < 0.05, **p < 0.01, ***p < 0.001). KC, Keratinocyte chemoattractant.

FIGURE 3.

Inhibition of fatty-acid synthesis during DC development increases ER stress and alters DC production of inflammatory mediators. (A) Expression of markers of ER stress (GRP-78, eIF2α, p-eIF2α, XBP-1, and β-actin) was tested in day 8 BMDC and T-BMDC by Western blotting. (B) Human moDC generated in control or TOFA-enriched media were tested for expression of selected ER stress markers. Control or TOFA-treated murine BMDC were tested for expression of PPAR-γ by Western blotting (C) and PCR (D). (E) Human moDC generated in control media or TOFA-supplemented media were tested for expression of PPAR-γ by Western blotting. β-actin was used as a loading control. (FH) Twenty-four–hour cell-culture supernatants from day 8 BMDC and T-BMDC plated at equal densities were tested for the presence of numerous cytokines and chemokines. (I) Control BMDC, standard high-dose T-BMDC, low-dose T-BMDC, ethanol-treated BMDC (EtOH), and staurosporine-treated BMDC were tested for their capacity to produce MCP-1 (*p < 0.05, **p < 0.01, ***p < 0.001). KC, Keratinocyte chemoattractant.

Close modal

The respective roles of ER stress or endogenous fatty-acid synthesis on DC production of immune-modulatory cytokines and chemokines are uncertain. Because DC regulate immunity by production of soluble inflammatory mediators, we tested T-BMDC cytokine and chemokine production. BMDC production of an array of inflammatory mediators, including IL-1α, IL-1β, IL-6, IL-10, IL-12, IFN-γ, IP-10, keratinocyte chemoattractant, LIF, MCP-1, M-CSF, MIG, MIP-2, and G-CSF, were higher in T-BMDC compared with controls (Fig. 3F, 3G). However, the CC chemokines MIP-1α, MIP-1β, and RANTES were expressed at markedly lower levels in T-BMDC (Fig. 3H). Lower-dose TOFA (1 mg/dl) also increased BMDC cytokine production; however, 0.5% ethanol alone had no effect nor did staurosporine (Fig. 3I).

Ag uptake is a primary function of DC and a critical consideration in constructing DC vaccines for cancer immunotherapy (26, 27). To determine the role of fatty-acid synthesis in DC capacity to capture Ag, BMDC were grown alone or in media supplemented with TOFA, as above. Consistent with their relative immaturity, T-BMDC exhibited enhanced ability to capture Ag via generalized macropinocytosis (Fig. 4A) or using specialized mannose receptors (Fig. 4B, 4C). Similarly, serial treatment of mice with C75 resulted in markedly enhanced spleen DC capacity to capture Ag in vivo (Fig. 4D). Low-dose TOFA was similarly effective at enhancing DC capacity for Ag capture as high-dose TOFA (Supplemental Fig. 3A). Conversely, ethanol or staurosporine did not enhance DC ability to capture soluble Ag (Supplemental Fig. 3B). These data imply that blockade of fatty-acid synthesis enhances DC capacity for Ag capture in multiple contexts.

FIGURE 4.

Blockade of fatty-acid synthesis enhances DC capacity for Ag capture in vitro and in vivo. BMDC and T-BMDC were tested at various time points for uptake of fluorescent albumin (A), dextran (B), and mannosylated albumin (C). (D) Splenic CD11c+ cells from control or C75-treated mice were tested for uptake of FITC-albumin at 30 min after in vivo administration. Data are representative of experiments performed three times. *p < 0.05, **p < 0.01, ***p < 0.001. MFI, Median fluorescence intensity.

FIGURE 4.

Blockade of fatty-acid synthesis enhances DC capacity for Ag capture in vitro and in vivo. BMDC and T-BMDC were tested at various time points for uptake of fluorescent albumin (A), dextran (B), and mannosylated albumin (C). (D) Splenic CD11c+ cells from control or C75-treated mice were tested for uptake of FITC-albumin at 30 min after in vivo administration. Data are representative of experiments performed three times. *p < 0.05, **p < 0.01, ***p < 0.001. MFI, Median fluorescence intensity.

Close modal

Because blockade of fatty-acid synthesis augments BMDC ER stress and increases their production of inflammatory mediators, we postulated it would enhance their immune stimulatory function. In consort with our hypothesis, T-BMDC induced higher proliferation of allogeneic T cells in an MLR compared with controls (Fig. 5A). To determine the effect of inhibiting fatty-acid synthesis on DC capacity to stimulate Ag-restricted CD4+ T cell, control or T-BMDC were loaded with Ova323–339 and then cocultured in various concentrations with CD4+ OT-II T cells. Peptide-pulsed BMDC grown in TOFA induced more vigorous proliferation of Ag-restricted CD4+ T cells (Fig. 5B) and induced higher CD4+ T cell production of Th1 and Th17 cytokines (Fig. 5C) compared with peptide-pulsed control BMDC stimulators. Conversely, Th2 cytokines were uniformly expressed at low levels after stimulating OT-II cells using either BMDC or T-BMDC. There was similarly no significant difference between control and T-BMDC in their propensity to generate CD4+CD25+Foxp3+ regulatory T cells in coculture experiments (Fig. 5D). Low-dose TOFA was also effective at enhancing DC capacity for Ag presentation. Conversely, staurosporine diminished DC capacity for T cell stimulation (Supplemental Fig. 3C).

FIGURE 5.

T-BMDC induce enhanced allogeneic and Ag-restricted CD4+ T cell stimulation. (A)Various concentrations of BMDC and T-BMDC were tested for their ability to induce proliferation of allogeneic T cells in an MLR. BMDC and T-BMDC pulsed with Ova323–339 peptide were tested for their ability to induce Ag-restricted CD4+ T cell proliferation (B) and Th1, Th2, and Th17 cytokine production (C) in OT-II T cells. (D) CD4+ T cell coexpression of CD25 and Foxp3 was tested at 96 h after splenocytes were cocultured in a 1:1 ratio with BMDC or T-BMDC. *p < 0.05, **p < 0.01, ***p < 0.001. FSC, Forward light scatter.

FIGURE 5.

T-BMDC induce enhanced allogeneic and Ag-restricted CD4+ T cell stimulation. (A)Various concentrations of BMDC and T-BMDC were tested for their ability to induce proliferation of allogeneic T cells in an MLR. BMDC and T-BMDC pulsed with Ova323–339 peptide were tested for their ability to induce Ag-restricted CD4+ T cell proliferation (B) and Th1, Th2, and Th17 cytokine production (C) in OT-II T cells. (D) CD4+ T cell coexpression of CD25 and Foxp3 was tested at 96 h after splenocytes were cocultured in a 1:1 ratio with BMDC or T-BMDC. *p < 0.05, **p < 0.01, ***p < 0.001. FSC, Forward light scatter.

Close modal

To explore whether the increased immunogenicity of TOFA-treated BMDC extends to their interactions with CD8+ T cells, we loaded BMDC populations with Ova257–264 before coculture with CD8+ OT-I T cells. Peptide-pulsed T-BMDC induced ∼3-fold elevated CD8+ T cell proliferation (Fig. 6A). In addition, BMDC grown in TOFA induced higher CD44 expression in Ag-restricted CD8+ T cells (Fig. 6B) and markedly increased CD8+ T cell production of IFN-γ (Fig. 6C) and TNF-α (Fig. 6D) compared with peptide-pulsed control BMDC stimulators. Further, to determine the effect of blocking fatty-acid synthesis on DC capacity to cross-present Ag to CD8+ T cells, control or T-BMDC were loaded with OVA and cultured at various concentrations with CD8+ OT-I T cells. Again, T-BMDC induced enhanced cross presentation of OVA as evidenced by higher T cell proliferation (Fig. 6E) and production of IFN-γ (Fig. 6F).

FIGURE 6.

T-BMDC induce enhanced CD8+ T cell activation. (AD) BMDC and T-BMDC were loaded with Ova257–264 peptide and plated in various ratios with CD8+ OT-I T cells. (A) OT-I proliferation was measured by incorporation of [3H]thymidine. (B) OT-I T cell expression of CD44 was measured on flow cytometry (median fluorescence intensity is indicated). IFN-γ (C) and TNF-α (D) production by CD8+ OT-I T cells was measured in cell-culture supernatant. (E and F) To test DC capacity for cross-presentation, BMDC and T-BMDC were loaded with OVA and used in various ratios to stimulate CD8+ OT-I T cells. OT-I cellular proliferation (E) and production of IFN-γ (F) were measured. Restimulated CTL cultures from mice twice immunized by adoptive transfer of Ova257–264 peptide–pulsed BMDC or T-BMDC were tested for production of IFN-γ (G) and IL-10 (H). *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 6.

T-BMDC induce enhanced CD8+ T cell activation. (AD) BMDC and T-BMDC were loaded with Ova257–264 peptide and plated in various ratios with CD8+ OT-I T cells. (A) OT-I proliferation was measured by incorporation of [3H]thymidine. (B) OT-I T cell expression of CD44 was measured on flow cytometry (median fluorescence intensity is indicated). IFN-γ (C) and TNF-α (D) production by CD8+ OT-I T cells was measured in cell-culture supernatant. (E and F) To test DC capacity for cross-presentation, BMDC and T-BMDC were loaded with OVA and used in various ratios to stimulate CD8+ OT-I T cells. OT-I cellular proliferation (E) and production of IFN-γ (F) were measured. Restimulated CTL cultures from mice twice immunized by adoptive transfer of Ova257–264 peptide–pulsed BMDC or T-BMDC were tested for production of IFN-γ (G) and IL-10 (H). *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

DC are largely reliant on their capacity to induce CTL in their effort to target invading pathogens or cancer cells (28). To determine the requirement for fatty-acid synthesis during DC generation on their ability to generate CTL in vivo, mice were immunized twice at weekly intervals with Ova257–264 peptide–pulsed control BMDC or T-BMDC. Splenocytes were harvested from immunized mice 1 wk after the second immunization and were restimulated in vitro with Ova257–264 peptide. On day 5, CTL cultures were tested for production of IFN-γ and IL-10. Consistent with our previous findings, in vivo immunization using T-BMDC induced elevated production of IFN-γ in CTL supernatant (Fig. 6G). Moreover, T-BMDC immunization resulted in decreased production of IL-10, an inhibitory cytokine, in CTL cultures compared with immunization using peptide-pulsed control BMDC (Fig. 6H). Taken together, these data suggest that blockade of fatty-acid synthesis is an attractive strategy to enhance DC capacity for induction of immunogenic CTL responses.

Because DC immune-stimulatory capacity has been linked to the MAPK, NF-κB, and PI3K/Akt signaling pathways (29), we tested the effect of inhibition of fatty-acid synthesis on the cellular activation of these pathways. Consistent with their enhanced CD4+ and CD8+ T cell stimulatory capacity, we found that T-BMDC expressed elevated levels of pErk-1, an activated MAPK signaling intermediate (Supplemental Fig. 4A). We also found that T-BMDC expressed elevated levels of pAkt as well as p70 S6 kinase, which acts downstream of phosphatidylinositol 3,4,5-triphosphate, suggesting activation of the PI3K/Akt signaling pathway in BMDC in the context of fatty-acid synthesis blockade (Supplemental Fig. 4B). Phosphatase and tensin homolog, which negatively regulates PI3K/Akt signaling, was equally expressed in TOFA-treated and control BMDC (Supplemental Fig. 4B). However, activated NF-κB intermediates were expressed at lower levels after TOFA treatment, which is consistent with their elevated intracellular ER stress (30) (Supplemental Fig. 4C).

Elevated ER stress has been linked to enhanced Ag presentation by APC (31). Because the chaperone 4-phenylbutyrate inhibits adipogenesis by modulating the unfolded protein response and decreasing ER stress (32), we postulated this may mitigate the increased immunogenicity of TOFA-treated BMDC. Accordingly, we found that the preincubation with 4-phenylbutyrate significantly reduced the CD4+ and CD8+ T cell stimulatory capacity of T-BMDC (Fig. 7A, 7B). T cell activation by control BMDC was affected to a lesser extent by the chaperone. These data imply that the increased immunogenicity of T-BMDC may be related to their increased ER stress. Blockade of MAPK or PI3K/Akt signaling did not mitigate the augmented capacity of T-BMDC to induce T cell proliferation (Fig. 7C). However, MAPK inhibition lessened T cell activation (Fig. 7D). PI3K blockade had no effect (data not shown).

FIGURE 7.

Lowering ER stress reduces the enhanced T cell–stimulatory capacity of T-BMDC. BMDC or T-BMDC (1 × 104) were loaded with the appropriate Ova peptide and used to stimulate CD4+ OT-II T cells (A) or CD8+ OT-I T cells (B), respectively, for 72 h. In selected experiments, DC were preincubated with the chaperone 4-phenylbutyrate. (C) OTI-I T cell stimulation assays were repeated using soluble PI3K and MAPK inhibitors. T cell proliferation was measured by incorporation of [3H]thymidine during the last 24 h. (D) OT-I T cell activation after stimulation by peptide-pulsed BMDC in the context of MAPK inhibition or control was measured by production of IFN-γ. Representative data are shown from experiments repeated three times. *p < 0.05, **p < 0.01.

FIGURE 7.

Lowering ER stress reduces the enhanced T cell–stimulatory capacity of T-BMDC. BMDC or T-BMDC (1 × 104) were loaded with the appropriate Ova peptide and used to stimulate CD4+ OT-II T cells (A) or CD8+ OT-I T cells (B), respectively, for 72 h. In selected experiments, DC were preincubated with the chaperone 4-phenylbutyrate. (C) OTI-I T cell stimulation assays were repeated using soluble PI3K and MAPK inhibitors. T cell proliferation was measured by incorporation of [3H]thymidine during the last 24 h. (D) OT-I T cell activation after stimulation by peptide-pulsed BMDC in the context of MAPK inhibition or control was measured by production of IFN-γ. Representative data are shown from experiments repeated three times. *p < 0.05, **p < 0.01.

Close modal

We and others have demonstrated that BMDC are powerful activators of innate immune effector cells such as NK and NKT cells (33, 34). To examine the role of fatty-acid synthesis in BMDC capacity to activate NK cells, we cocultured T-BMDC and controls with equal numbers of NK cells before NK cell harvest and measurement of their phenotypic activation and production of IFN-γ. T-BMDC induced elevated NK cell expression of CD25 (Fig. 8A) and induced ∼4-fold higher production of IFN-γ compared with control BMDC (Fig. 8B). Because DC capacity to activate NK cells has recently been linked to their expression of Notch ligands (35), we tested whether blockade of acetyl CoA carboxylase secondarily increases Notch ligand expression in BMDC. As postulated, T-BMDC expressed higher Jagged-1 and Δ-4 compared with control BMDC on analysis by Western blotting (Fig. 8C) and flow cytometry (Fig. 8D).

FIGURE 8.

T-BMDC have enhanced capacity for NK cell activation. Spleen NK cells were harvested and coincubated with BMDC or T-BMDC. NK cell expression of CD25 (median fluorescence intensity is indicated) (A), production of IFN-γ (B), and expression of Notch ligands were measured by Western blotting (Jagged-1, Δ-4) (C) and flow cytometry (Jagged-1) (D). ***p < 0.001.

FIGURE 8.

T-BMDC have enhanced capacity for NK cell activation. Spleen NK cells were harvested and coincubated with BMDC or T-BMDC. NK cell expression of CD25 (median fluorescence intensity is indicated) (A), production of IFN-γ (B), and expression of Notch ligands were measured by Western blotting (Jagged-1, Δ-4) (C) and flow cytometry (Jagged-1) (D). ***p < 0.001.

Close modal

DC are a specialized population of APCs that link innate and adaptive immunity (1). DC can influence immune responses by both direct interaction with effector cells, such as T cells and NK cells, and via production of a wide array of inflammatory mediators. In this study, we found that blockade of fatty-acid synthesis markedly inhibits DC development from bone marrow or PBMC precursors in mice and humans, respectively, and induces apoptosis in DC precursors, which is associated with elevated cellular expression of cleaved caspase-3, BCL-xL, and downregulation of cyclin B1. For our in vivo experiments, we employed C75 in lieu of TOFA, as TOFA is highly toxic when administered systemically (7, 36). We found that in vivo blockade of fatty-acid synthesis hinders DC generation in peripheral tissues as well as primary and secondary lymphoid organs. Fatty-acid synthesis inhibition also has variable effects on DC surface phenotype including suppression of MHC II expression but increased CD40 expression. Further, T-BMDC express higher levels of selected MAPK and PI3K/Akt signaling intermediates and produce markedly elevated levels of numerous cytokines and chemokines (37, 38). Interestingly, the CC chemokines MIP-1α, MIP-1β, and RANTES, which are purported to play a role in granulocytic lineage proliferation or differentiation (39), are suppressed by fatty-acid synthesis inhibition. Understanding the mechanistic regulation of BMDC cytokine and chemokine production by fatty-acid synthesis requires more exact study; however, there are likely to be autocrine effects of specific cytokines on further BMDC production of additional inflammatory mediators. For example, Stober et al. (40) reported that IL-12 can influence IFN-γ production by BMDC. Taken together, our data suggest that the capacity for fatty-acid synthesis is important for DC generation and expression of their distinct immune phenotype.

The properties of DC generated in the context of fatty-acid synthesis blockade are relevant not only to understanding basic DC immunobiology but also for the development of vaccines for immunotherapy. In particular, TOFA-treated DC exhibited increased capacity to activate CD4+ and CD8+ T cells and NK cells, which can be exploited in the construction of DC cancer vaccines. T-BMDC induction of Ag-restricted CD8+ T cells led to increased production of IFN-γ and TNF-α, and decreased production of IL-10. DC immunotherapy regimens in cancer and benign diseases have largely been of limited clinical efficacy because of the modest adaptive immune responses and CTLs induced (28, 41). Varied cytokine cocktails and methods of exogenous DC stimulation have been employed to bolster the host’s Ag-restricted and innate immunogenic responses to DC vaccines (28); hence, our data suggest that inhibiting fatty-acid synthesis may be an attractive adjuvant in experimental immunotherapy.

The mechanism for the enhanced immune-stimulatory capacity of DC generated in the context of fatty-acid synthesis inhibition appears to be related in part to their elevated ER stress. ER stress is generated in response to an accumulation of unfolded or misfolded proteins in the lumen of the endoplasmic reticulum (42). ER stress attempts to restore normal cellular function by halting protein translation and activating signaling pathways, leading to production of molecular chaperones that facilitate protein folding. This process has been found to be conserved among all mammalian species and can result in cellular apoptosis if not resolved (43). There is an emerging role for ER stress in the function of APCs. Goodall et al. (44) reported that activation of ER stress, in combination with TLR ligation, markedly enhances DC expression of selected cytokines. Additionally, Oh et al. (45) recently reported that ER stress is a functional switch regulating M2 macrophage differentiation and phenotype including cellular cholesterol content. Our observations of elevated ER stress in TOFA-treated BMDC were made in both our murine and human models and were further adduced by higher DC expression of PPAR-γ after TOFA treatment. Further, our finding of increased expression of certain activated MAPK signaling intermediates in T-BMDC is consistent with a recent report showing involvement of Erk MAPK in ER stress in human neuroblastoma cells (46). Notably, Hayakawa et al. (30) recently found that ER stress depresses NF-κB activation, which is in consort with our finding of diminished levels activated NF-κB intermediates in T-BMDC. These findings are also consistent with the observation by Zeyda et al. (2), who reported that exogenous administration of PUFA lessened DC immune-stimulatory capacity independent of NF-κB signaling. However, our findings of enhanced Akt activation are surprising in this context because ER stress has been reported to negatively regulate the Akt/mTOR pathway (47). Further, PPAR-γ can also negatively regulate Akt phosphorylation (48). These data suggest that alternate mechanisms may be responsible for the elevated levels of pAkt in T-BMDC. It is also notable that MAPK inhibition but not PI3K/Akt signaling blockade mitigated the enhanced T cell immune-stimulatory capacity of T-BMDC. Our data also imply that, besides their elevated ER stress and increased inflammatory pathway signaling, the enhanced effector cell stimulatory capacity of T-BMDC may be a function of their augmented cytokine secretory profile. DC production of varied cytokine has well-established potent effects on their Ag-presenting and allostimulaotry capacity (49).

Our study on the effects of endogenous fatty-acid synthesis inhibition on DC function expands on previous reports showing that exogenous administration of PUFA results in diminished TNF-α production, CD40 expression, and T cell stimulatory capacity in developing human moDC (2). However, taken in the context of our recent work, there appears to be a dichotomy between the effects of fatty-acid synthesis inhibition on DC developing from cellular precursors versus blockade of fatty-acid synthesis on fully mature DC populations. In particular, we recently reported that fully mature liver DC can be divided into two distinct populations based on intracellular lipid content, including triglycerides and phospholipids (11). Further, we found that liver DC immunogenicity is determined by their lipid content, as lipid-rich liver DC were more immunogenic in comparison with lipid-poor liver DC. This was demonstrated by their higher secretion of cytokines and activation of Ag-restricted CD4+ and CD8+ T cells as well as NK cells and NKT cells. Moreover, blockade of fatty-acid synthesis in terminally differentiated lipid-rich liver DC using TOFA diminished their capacity for T cell and NK cell activation (11). Therefore, the effects of fatty acids, or blockade of their production, on DC properties in the current study appears to be limited to developmental effects rather than applicable to mature fully differentiated DC.

This work was supported in part by National Institutes of Health Awards DK085278 (to G.M.) and CA155649 (to G.M.).

The online version of this article contains supplemental material.

Abbreviations used in this article:

BMDC

bone marrow–derived dendritic cell

DC

dendritic cell

eIF2α

eukaryotic translation initiation factor 2α

ER

endoplasmic reticulum

MHC II

MHC class II

moDC

monocyte-derived dendritic cell

PB

PIPES buffer

PPAR-γ

peroxisome proliferator–activated receptor γ; PUFA, polyunsaturated fatty acid

T-BMDC

tall oil fatty acid–treated bone marrow–derived dendritic cell

TOFA

tall oil fatty acid.

1
Steinman
R. M.
,
Banchereau
J.
.
2007
.
Taking dendritic cells into medicine.
Nature
449
:
419
426
.
2
Zeyda
M.
,
Säemann
M. D.
,
Stuhlmeier
K. M.
,
Mascher
D. G.
,
Nowotny
P. N.
,
Zlabinger
G. J.
,
Waldhäusl
W.
,
Stulnig
T. M.
.
2005
.
Polyunsaturated fatty acids block dendritic cell activation and function independently of NF-kappaB activation.
J. Biol. Chem.
280
:
14293
14301
.
3
Herber
D. L.
,
Cao
W.
,
Nefedova
Y.
,
Novitskiy
S. V.
,
Nagaraj
S.
,
Tyurin
V. A.
,
Corzo
A.
,
Cho
H. I.
,
Celis
E.
,
Lennox
B.
, et al
.
2010
.
Lipid accumulation and dendritic cell dysfunction in cancer.
Nat. Med.
16
:
880
886
.
4
Miller
G.
,
Lahrs
S.
,
Pillarisetty
V. G.
,
Shah
A. B.
,
DeMatteo
R. P.
.
2002
.
Adenovirus infection enhances dendritic cell immunostimulatory properties and induces natural killer and T-cell-mediated tumor protection.
Cancer Res.
62
:
5260
5266
.
5
Kapadia
S. B.
,
Chisari
F. V.
.
2005
.
Hepatitis C virus RNA replication is regulated by host geranylgeranylation and fatty acids.
Proc. Natl. Acad. Sci. USA
102
:
2561
2566
.
6
Zhou
W.
,
Han
W. F.
,
Landree
L. E.
,
Thupari
J. N.
,
Pinn
M. L.
,
Bililign
T.
,
Kim
E. K.
,
Vadlamudi
A.
,
Medghalchi
S. M.
,
El Meskini
R.
, et al
.
2007
.
Fatty acid synthase inhibition activates AMP-activated protein kinase in SKOV3 human ovarian cancer cells.
Cancer Res.
67
:
2964
2971
.
7
Pizer
E. S.
,
Thupari
J.
,
Han
W. F.
,
Pinn
M. L.
,
Chrest
F. J.
,
Frehywot
G. L.
,
Townsend
C. A.
,
Kuhajda
F. P.
.
2000
.
Malonyl-coenzyme-A is a potential mediator of cytotoxicity induced by fatty-acid synthase inhibition in human breast cancer cells and xenografts.
Cancer Res.
60
:
213
218
.
8
Kashkar
H.
,
Krönke
M.
,
Jürgensmeier
J. M.
.
2002
.
Defective Bax activation in Hodgkin B-cell lines confers resistance to staurosporine-induced apoptosis.
Cell Death Differ.
9
:
750
757
.
9
Belmokhtar
C. A.
,
Hillion
J.
,
Ségal-Bendirdjian
E.
.
2001
.
Staurosporine induces apoptosis through both caspase-dependent and caspase-independent mechanisms.
Oncogene
20
:
3354
3362
.
10
Connolly
M. K.
,
Bedrosian
A. S.
,
Mallen-St Clair
J.
,
Mitchell
A. P.
,
Ibrahim
J.
,
Stroud
A.
,
Pachter
H. L.
,
Bar-Sagi
D.
,
Frey
A. B.
,
Miller
G.
.
2009
.
In liver fibrosis, dendritic cells govern hepatic inflammation in mice via TNF-alpha.
J. Clin. Invest.
119
:
3213
3225
.
11
Ibrahim
J.
,
Nguyen
A. H.
,
Rehman
A.
,
Ochi
A.
,
Jamal
M.
,
Graffeo
C. S.
,
Henning
J. R.
,
Zambirinis
C. P.
,
Fallon
N. C.
,
Barilla
R.
, et al
.
2012
.
Dendritic cell populations with different concentrations of lipid regulate tolerance and immunity in mouse and human liver.
Gastroenterology
143
:
1061
1072
.
12
Bedrosian, A. S., A. H. Nguyen, M. Hackman, M. K. Connolly, A. Malhotra, J. Ibrahim, N. E. Cieza-Rubio, J. R. Henning, R. Barilla, A. Rehman, et al. 2011. Dendritic cells promote pancreatic viability in mice with acute pancreatitis. Gastroenterology 141:1915–1926.e1911–1914
.
13
Landree
L. E.
,
Hanlon
A. L.
,
Strong
D. W.
,
Rumbaugh
G.
,
Miller
I. M.
,
Thupari
J. N.
,
Connolly
E. C.
,
Huganir
R. L.
,
Richardson
C.
,
Witters
L. A.
, et al
.
2004
.
C75, a fatty acid synthase inhibitor, modulates AMP-activated protein kinase to alter neuronal energy metabolism.
J. Biol. Chem.
279
:
3817
3827
.
14
Thupari
J. N.
,
Landree
L. E.
,
Ronnett
G. V.
,
Kuhajda
F. P.
.
2002
.
C75 increases peripheral energy utilization and fatty acid oxidation in diet-induced obesity.
Proc. Natl. Acad. Sci. USA
99
:
9498
9502
.
15
Guseva
N. V.
,
Rokhlin
O. W.
,
Glover
R. A.
,
Cohen
M. B.
.
2011
.
TOFA (5-tetradecyl-oxy-2-furoic acid) reduces fatty acid synthesis, inhibits expression of AR, neuropilin-1 and Mcl-1 and kills prostate cancer cells independent of p53 status.
Cancer Biol. Ther.
12
:
80
85
.
16
Wang
C.
,
Xu
C.
,
Sun
M.
,
Luo
D.
,
Liao
D. F.
,
Cao
D.
.
2009
.
Acetyl-CoA carboxylase-alpha inhibitor TOFA induces human cancer cell apoptosis.
Biochem. Biophys. Res. Commun.
385
:
302
306
.
17
Castilho
G.
,
Okuda
L. S.
,
Pinto
R. S.
,
Iborra
R. T.
,
Nakandakare
E. R.
,
Santos
C. X.
,
Laurindo
F. R.
,
Passarelli
M.
.
2012
.
ER stress is associated with reduced ABCA-1 protein levels in macrophages treated with advanced glycated albumin - reversal by a chemical chaperone.
Int. J. Biochem. Cell Biol.
44
:
1078
1086
.
18
Abbas
W.
,
Khan
K. A.
,
Tripathy
M. K.
,
Dichamp
I.
,
Keita
M.
,
Rohr
O.
,
Herbein
G.
.
2012
.
Inhibition of ER stress-mediated apoptosis in macrophages by nuclear-cytoplasmic relocalization of eEF1A by the HIV-1 Nef protein.
Cell Death Dis
3
:
e292
.
19
Seimon
T. A.
,
Kim
M. J.
,
Blumenthal
A.
,
Koo
J.
,
Ehrt
S.
,
Wainwright
H.
,
Bekker
L. G.
,
Kaplan
G.
,
Nathan
C.
,
Tabas
I.
,
Russell
D. G.
.
2010
.
Induction of ER stress in macrophages of tuberculosis granulomas.
PLoS ONE
5
:
e12772
.
20
Little
J. L.
,
Wheeler
F. B.
,
Fels
D. R.
,
Koumenis
C.
,
Kridel
S. J.
.
2007
.
Inhibition of fatty acid synthase induces endoplasmic reticulum stress in tumor cells.
Cancer Res.
67
:
1262
1269
.
21
Wu
Y.
,
Zhang
H.
,
Dong
Y.
,
Park
Y. M.
,
Ip
C.
.
2005
.
Endoplasmic reticulum stress signal mediators are targets of selenium action.
Cancer Res.
65
:
9073
9079
.
22
Han
K. L.
,
Choi
J. S.
,
Lee
J. Y.
,
Song
J.
,
Joe
M. K.
,
Jung
M. H.
,
Hwang
J. K.
.
2008
.
Therapeutic potential of peroxisome proliferators—activated receptor-alpha/gamma dual agonist with alleviation of endoplasmic reticulum stress for the treatment of diabetes.
Diabetes
57
:
737
745
.
23
Weber
S. M.
,
Chambers
K. T.
,
Bensch
K. G.
,
Scarim
A. L.
,
Corbett
J. A.
.
2004
.
PPARgamma ligands induce ER stress in pancreatic beta-cells: ER stress activation results in attenuation of cytokine signaling.
Am. J. Physiol. Endocrinol. Metab.
287
:
E1171
E1177
.
24
Zapata-Gonzalez
F.
,
Rueda
F.
,
Petriz
J.
,
Domingo
P.
,
Villarroya
F.
,
Diaz-Delfin
J.
,
de Madariaga
M. A.
,
Domingo
J. C.
.
2008
.
Human dendritic cell activities are modulated by the omega-3 fatty acid, docosahexaenoic acid, mainly through PPAR(gamma):RXR heterodimers: comparison with other polyunsaturated fatty acids.
J. Leukoc. Biol.
84
:
1172
1182
.
25
Appel
S.
,
Mirakaj
V.
,
Bringmann
A.
,
Weck
M. M.
,
Grünebach
F.
,
Brossart
P.
.
2005
.
PPAR-gamma agonists inhibit toll-like receptor-mediated activation of dendritic cells via the MAP kinase and NF-kappaB pathways.
Blood
106
:
3888
3894
.
26
Palucka
K.
,
Banchereau
J.
,
Mellman
I.
.
2010
.
Designing vaccines based on biology of human dendritic cell subsets.
Immunity
33
:
464
478
.
27
Pajtasz-Piasecka
E.
,
Indrová
M.
.
2010
.
Dendritic cell-based vaccines for the therapy of experimental tumors.
Immunotherapy
2
:
257
268
.
28
Frankenberger
B.
,
Schendel
D. J.
.
2012
.
Third generation dendritic cell vaccines for tumor immunotherapy.
Eur. J. Cell Biol.
91
:
53
58
.
29
Watts
C.
,
West
M. A.
,
Zaru
R.
.
2010
.
TLR signalling regulated antigen presentation in dendritic cells.
Curr. Opin. Immunol.
22
:
124
130
.
30
Hayakawa
K.
,
Nakajima
S.
,
Hiramatsu
N.
,
Okamura
M.
,
Huang
T.
,
Saito
Y.
,
Tagawa
Y.
,
Tamai
M.
,
Takahashi
S.
,
Yao
J.
,
Kitamura
M.
.
2010
.
ER stress depresses NF-kappaB activation in mesangial cells through preferential induction of C/EBP beta.
J. Am. Soc. Nephrol.
21
:
73
81
.
31
Granados
D. P.
,
Tanguay
P. L.
,
Hardy
M. P.
,
Caron
E.
,
de Verteuil
D.
,
Meloche
S.
,
Perreault
C.
.
2009
.
ER stress affects processing of MHC class I-associated peptides.
BMC Immunol.
10
:
10
.
32
Basseri
S.
,
Lhoták
S.
,
Sharma
A. M.
,
Austin
R. C.
.
2009
.
The chemical chaperone 4-phenylbutyrate inhibits adipogenesis by modulating the unfolded protein response.
J. Lipid Res.
50
:
2486
2501
.
33
Miller
G.
,
Lahrs
S.
,
Dematteo
R. P.
.
2003
.
Overexpression of interleukin-12 enables dendritic cells to activate NK cells and confer systemic antitumor immunity.
FASEB J.
17
:
728
730
.
34
Chan
C. W.
,
Crafton
E.
,
Fan
H. N.
,
Flook
J.
,
Yoshimura
K.
,
Skarica
M.
,
Brockstedt
D.
,
Dubensky
T. W.
,
Stins
M. F.
,
Lanier
L. L.
, et al
.
2006
.
Interferon-producing killer dendritic cells provide a link between innate and adaptive immunity.
Nat. Med.
12
:
207
213
.
35
Kijima
M.
,
Yamaguchi
T.
,
Ishifune
C.
,
Maekawa
Y.
,
Koyanagi
A.
,
Yagita
H.
,
Chiba
S.
,
Kishihara
K.
,
Shimada
M.
,
Yasutomo
K.
.
2008
.
Dendritic cell-mediated NK cell activation is controlled by Jagged2-Notch interaction.
Proc. Natl. Acad. Sci. USA
105
:
7010
7015
.
36
Zhou
W.
,
Simpson
P. J.
,
McFadden
J. M.
,
Townsend
C. A.
,
Medghalchi
S. M.
,
Vadlamudi
A.
,
Pinn
M. L.
,
Ronnett
G. V.
,
Kuhajda
F. P.
.
2003
.
Fatty acid synthase inhibition triggers apoptosis during S phase in human cancer cells.
Cancer Res.
63
:
7330
7337
.
37
Mäkelä
S. M.
,
Strengell
M.
,
Pietilä
T. E.
,
Osterlund
P.
,
Julkunen
I.
.
2009
.
Multiple signaling pathways contribute to synergistic TLR ligand-dependent cytokine gene expression in human monocyte-derived macrophages and dendritic cells.
J. Leukoc. Biol.
85
:
664
672
.
38
Caparrós
E.
,
Munoz
P.
,
Sierra-Filardi
E.
,
Serrano-Gómez
D.
,
Puig-Kröger
A.
,
Rodríguez-Fernández
J. L.
,
Mellado
M.
,
Sancho
J.
,
Zubiaur
M.
,
Corbí
A. L.
.
2006
.
DC-SIGN ligation on dendritic cells results in ERK and PI3K activation and modulates cytokine production.
Blood
107
:
3950
3958
.
39
Rice
C. M.
,
Scolding
N. J.
.
2010
.
Adult human mesenchymal cells proliferate and migrate in response to chemokines expressed in demyelination.
Cell Adhes. Migr.
4
:
235
240
.
40
Stober
D.
,
Schirmbeck
R.
,
Reimann
J.
.
2001
.
IL-12/IL-18-dependent IFN-gamma release by murine dendritic cells.
J. Immunol.
167
:
957
965
.
41
Ueno
H.
,
Klechevsky
E.
,
Schmitt
N.
,
Ni
L.
,
Flamar
A. L.
,
Zurawski
S.
,
Zurawski
G.
,
Palucka
K.
,
Banchereau
J.
,
Oh
S.
.
2011
.
Targeting human dendritic cell subsets for improved vaccines.
Semin. Immunol.
23
:
21
27
.
42
Hetz
C.
2012
.
The unfolded protein response: controlling cell fate decisions under ER stress and beyond.
Nat. Rev. Mol. Cell Biol.
13
:
89
102
.
43
Wu
J.
,
Kaufman
R. J.
.
2006
.
From acute ER stress to physiological roles of the Unfolded Protein Response.
Cell Death Differ.
13
:
374
384
.
44
Goodall
J. C.
,
Wu
C.
,
Zhang
Y.
,
McNeill
L.
,
Ellis
L.
,
Saudek
V.
,
Gaston
J. S.
.
2010
.
Endoplasmic reticulum stress-induced transcription factor, CHOP, is crucial for dendritic cell IL-23 expression.
Proc. Natl. Acad. Sci. USA
107
:
17698
17703
.
45
Oh
J.
,
Riek
A. E.
,
Weng
S.
,
Petty
M.
,
Kim
D.
,
Colonna
M.
,
Cella
M.
,
Bernal-Mizrachi
C.
.
2012
.
Endoplasmic reticulum stress controls M2 macrophage differentiation and foam cell formation.
J. Biol. Chem.
287
:
11629
11641
.
46
Arai
K.
,
Lee
S. R.
,
van Leyen
K.
,
Kurose
H.
,
Lo
E. H.
.
2004
.
Involvement of ERK MAP kinase in endoplasmic reticulum stress in SH-SY5Y human neuroblastoma cells.
J. Neurochem.
89
:
232
239
.
47
Qin
L.
,
Wang
Z.
,
Tao
L.
,
Wang
Y.
.
2010
.
ER stress negatively regulates AKT/TSC/mTOR pathway to enhance autophagy.
Autophagy
6
:
239
247
.
48
Goetze
S.
,
Eilers
F.
,
Bungenstock
A.
,
Kintscher
U.
,
Stawowy
P.
,
Blaschke
F.
,
Graf
K.
,
Law
R. E.
,
Fleck
E.
,
Gräfe
M.
.
2002
.
PPAR activators inhibit endothelial cell migration by targeting Akt.
Biochem. Biophys. Res. Commun.
293
:
1431
1437
.
49
Morelli
A. E.
,
Zahorchak
A. F.
,
Larregina
A. T.
,
Colvin
B. L.
,
Logar
A. J.
,
Takayama
T.
,
Falo
L. D.
,
Thomson
A. W.
.
2001
.
Cytokine production by mouse myeloid dendritic cells in relation to differentiation and terminal maturation induced by lipopolysaccharide or CD40 ligation.
Blood
98
:
1512
1523
.

The authors have no financial conflicts of interest.