Donor-specific induced pluripotent stem cells (iPSC) can be used to generate desired cell types, including naive immune effectors, for the treatment of different diseases. However, a greater understanding of the inherent immunogenicity of human iPSC and their cellular derivatives is needed for the development of safe and effective cell-replacement therapies, given that studies in mouse models claimed that the syngenic mouse iPSC lines can be immunogenic. We report the characterization of the innate and adaptive immune mechanisms in human iPSC lines derived from peripheral blood–derived dendritic cells using a nonintegrating RNA virus, Sendai virus. We show that these iPSC lines express mRNA of TLR molecules and the Ag-presentation pathway intermediates; however, these mRNA are not translated into functional proteins, and these iPSC lines do not induce TLR-mediated inflammatory cytokine responses or inflammasome activation. We also show that these iPSC lines do not activate T cells in an allogenic MLR; however, they express low levels of MHC class I molecules that can efficiently acquire antigenic peptides from their microenvironment and present them to Ag-specific T cells. In addition, we show that these iPSC lines can be efficiently differentiated into hematopoietic stem cell precursors, as well as APC, under appropriate culture conditions. Taken together, our data show that the dedifferentiation of human dendritic cells effectively shuts down their immunogenic pathways and implicates transcriptional and posttranscriptional mechanisms in this process.

Human pluripotent stem cells (hPSC) are defined by their ability to self-renew and to differentiate into different cell lineages under appropriate culture conditions. Derivation of human embryonic stem cell (hESC) lines from early-stage embryos opened up possibilities for the development of pluripotent stem cell–based cell-replacement therapies (CRT) (1), and successful reprogramming of somatic cells into induced pluripotent stem cells (iPSC) (2, 3) addressed the ethical issues associated with the therapeutic use of hESC. We are working on developing T cell–based cancer immunotherapy approaches utilizing the human melanoma–associated antigenic epitope MART-127–35 as a model human tumor Ag (48). Donor-derived immunogenic dendritic cells (DC) and engineered anti-tumor T cells produced encouraging clinical results in active specific immunization and T cell adoptive-transfer–based cancer immunotherapy settings (912); however, the efficacy of these approaches requires significant improvement. In this context, donor-derived iPSC can be a valuable resource for generating autologous naive immune effectors; however, efficient models are needed to systematically characterize the immunogenicity profiles of human iPSC lines and their differentiation potential.

The need for a systematic analysis of the inherent immunogenicity of human iPSC lines was highlighted by recent studies in mouse models that used rejection of iPSC line–induced teratoma in immune-competent mice as a model to examine the inherent immunogenicity of iPSC lines. These studies claimed that the syngenic mouse iPSC lines, especially those derived with genome-integrating viral vectors, are immunogenic (13) and that the inherent immunogenicity of these syngenic mouse iPSC lines correlates with their Ag profiles (14). Although other groups reported minimal or no immunogenicity of mouse syngenic iPSC lines and their cellular derivatives (1517), these reports highlighted the need for an in-depth characterization of the inherent immunogenicity of hPSC. hESC lines express low levels of MHC class I molecules (18), exhibit immune-privileged properties (19, 20), and are rejected in immune-competent mice through a T cell–mediated allograft-rejection process (19, 21, 22); however, the inherent immunogenicity of human iPSC lines has not been examined carefully. In this article, we show that the terminally differentiated gold standard for human immunogenic cells, DC, can be successfully dedifferentiated into the iPSC state using Sendai virus, a nonintegrating RNA virus. We used these iPSC lines to characterize the status of innate and adaptive immune mechanisms that are functional in human peripheral blood–derived DC and show that these iPSC lines are inherently nonimmunogenic. We also show that these iPSC lines express low levels of MHC class I molecules, and they can efficiently present antigenic peptides, acquired from their microenvironment, to corresponding Ag-specific T cells; as a result, they are also killed by the CTL. In addition, we show that these iPSC lines exhibit potent differentiation potential, as reflected by their ability to generate embryoid bodies (EB) that can produce hematopoietic stem cell (HSC) precursors and can be further differentiated into functional APC. Our findings provide a plausible explanation for the rejection of transplanted syngenic iPSC lines in animal models and demonstrate that DC-derived iPSC lines is an ideal system to characterize the immunogenicity profile of human iPSC lines. Furthermore, we believe that the dedifferentiation and redifferentiation model of human DC is also an ideal system to systematically characterize the molecular, cellular, and functional profile of human iPSC-derived donor-specific APC (iPSC-APC).

Human subject work was done in accordance with the University of Connecticut Health Center (UCHC) Institutional Review Board guidelines. HLA-A2+ individuals were enrolled in the study, with voluntary informed consent, to donate blood for the isolation of human PBL. CD4+ and CD8+ T cells and CD14+ monocytes were purified from human PBL by Ficoll-Hypaque gradient separation followed by magnetic-bead purification, as described previously (5, 23). hESC line H9 was obtained from the UCHC Stem Cell Core Facility, with an appropriate material transfer agreement from the WiCell Research Institute (Madison, WI). Mouse embryonic fibroblasts (MEF), CF-1, were procured from GlobalStem. Culture medium IMDM, DMEM, MEM-α, FBS, and KnockOut Serum Replacement were purchased from Life Technologies. Cytokines and growth supplements for hPSC culture and differentiation, such as GM-CSF, IL-4, IL-2, IL-3, IL-15, IL-7, Flt3, bone morphogenetic protein 4, stem cell factor (SCF), and vascular endothelial growth factor, were purchased from R&D Systems. Abs for FACS were purchased from BD Biosciences, BioLegend, and Affymetrix eBioscience.

A CytoTune-iPS 2.0 Sendai Reprogramming Kit (Life Technologies) was used to generate iPSC lines from human PBL, according to the manufacturer’s instructions, with some modifications. In brief, 50,000 whole PBL, purified CD4+ T cells, CD8+ T cells, and CD14+ monocyte-derived DC were transduced with recombinant Sendai virus, a nonintegrating RNA virus, expressing GFP to optimize the virus infection conditions. For the generation of iPSC lines, cells were infected with recombinant Sendai virus vectors expressing the reprogramming factor genes c-Myc, Klf-4, Oct-3/4, and Sox-2. Transduced cells were cultured in MEF-plated six-well culture plates, and the generation of iPSC clones was monitored. Potential iPSC clones generated 7–14 d later were transferred into fresh MEF-containing culture plates, propagated, and used to make frozen stocks. The karyotype and pluripotency status of the generated iPSC lines were established before characterizing their immunogenic profile and differentiation potential. To confirm that the iPSC lines generated were free of the Sendai virus vectors used for reprogramming, RT-PCR was done using primers specific for Sendai virus backbone that were provided with the kit, as described below. Uninfected human DC and the DC infected with GFP encoding Sendai virus were used as negative and positive controls, respectively. Primer sequences are listed in Supplemental Table I.

Irradiated MEF were plated in 0.1% gelatin–coated plates in DMEM containing 10% FBS a day before plating hPSC. The following day, MEF were washed three or four times, and hPSC clumps were plated in hPSC culture medium. Fresh hPSC medium was fed every other day. For passage, the hPSC culture plate was washed with PBS to remove the floating cells and fed with fresh medium, hPSC colonies were cut using a 1-ml syringe with a 16-gauge needle, and the cut colony clumps were plated in plates containing fresh CF-1 MEF.

To confirm the pluripotency status of hPSC by FACS, a single-cell suspension of hPSC was made by treating the hPSC cultures with 0.05% trypsin. Trypsin was neutralized by hPSC medium, cells were washed with PBS, and a single-cell suspension was made in PBS. Cells were stained with the indicated Ab, and surface phenotype was analyzed using a FACSCalibur (BD Biosciences). Ab against the pluripotency markers Oct-4, Sox-2, Nanog, Tra 1-61, and Tra 1-81 for immunofluorescence staining were purchased from Abcam. Intracellular staining was performed using Cytofix/Cytoperm (BD Biosciences).

RT-PCR was performed to confirm removal of Sendai virus vectors from the iPSC lines generated and pluripotency of the hPSC lines, as well as to characterize hPSC line–derived EB. In brief, hPSC cultures or hPSC-derived EB were washed with PBS and lysed using TRIzol Reagent (Life Technologies), and the RNA was made according to the manufacturer’s instructions, as described previously (4). RNA was quantified and cDNA was made with 500 ng of RNA per sample using the SuperScript III single-strand cDNA synthesis kit (Life Technologies). Primers used for RT-PCR analysis were procured from IDT Technologies and are listed in Supplemental Table I.

For immunofluorescence staining, hPSC were cultured on feeder-coated glass coverslips in 24-well plates. Colonies were fixed with 4% paraformaldehyde for 30 min at room temperature, washed three times with PBS, permeabilized with PBS containing 0.1% Triton X-100 for 30 min, and washed again with PBS three times. Fixed and permeabilized cells were kept in PBS at 4°C until used for staining. For staining, hPSC were incubated in blocking reagent (PBS with 10% BSA) for 30–45 min, followed by a 45–60-min incubation with primary Ab (1:50 dilution in PBS containing 1% BSA) in a humidified chamber in a 37°C incubator. Unbound primary Ab was removed by washing three times with PBS, followed by staining with the secondary Ab (1:100 dilution) in a humidified chamber for 45–60 min in a 37°C incubator, followed by three washes with PBS. Hoechst (1 μM final) was added at the second washing step. Slides were washed with water to remove traces of PBS before mounting with VECTASHIELD Mounting Medium (Vector Laboratories). Analysis was performed using a Zeiss LSM 780 confocal microscope at the UCHC Microscopy Core Facility.

The ability of hESC and iPSC to form three germ layers in vivo was confirmed by a teratoma assay. In brief, hPSC cultures were treated with 1 mg/ml dispase, collected in hPSC culture medium, and washed once with PBS, and 2 × 106 hPSC were injected i.m. into immunodeficient NOD-SCID mice (Jackson Laboratory, Bar Harbor, ME) for teratoma induction. Teratomas were excised 10–16 wk postinjection, fixed in 10% formalin, and embedded in frozen embedding medium (Shandon Cryomatrix; Thermo Scientific). Tissue sections were cut using a refrigerated microtome (Leica) and stained with H&E, and images were taken using an Olympus IX71 microscope fitted with a SPOT Flex camera (SPOT Imaging Solutions).

Immature DC (iDC) were derived from human PBL and cultured in the presence of LPS (100 ng/ml) and IFN-γ (1000 units/ml) for 12–16 h to generate mature DC (mDC), as described previously (4, 5, 24). In brief, PBL were put in six-well culture plates for 15–30 min to allow DC precursors to adhere to the wells. Nonadherent cells were removed by washing the wells three to five times with PBS. Alternatively, CD14+ monocytes were purified from blood-derived PBL using a CD14+ cell purification kit (STEMCELL Technologies) and were differentiated into iDC by culturing them in IMDM containing the DC-differentiating cytokines GM-CSF (1000 IU/ml) and IL-4 (1000 IU/ml) for 3–5 d.

The phagocytic ability of iPSC-APC and human blood monocyte-derived 3–5-d-old iDC was examined by phagocytosis of FITC-tagged dextran beads (40,000 m.w.; Molecular Probes), according to the manufacturer’s instructions. In brief, cells were coincubated with FITC-dextran beads for 2 h at 37°C and washed with PBS to remove free beads, and phagocytosis was quantified by FACS. To examine phagocytosis by immunofluorescence, following coincubation with the FITC-dextran beads, cells were mounted on glass slides using a Shandon cytospin centrifuge (700 rpm for 7 min), fixed using 4% paraformaldehyde, and imaged, following nucleus staining with 1 μM Hoechst, using a Zeiss 780 confocal microscope at the UCHC Microscopy Core Facility, as described previously.

Functional T cells specific for the human melanoma–associated MART-127–35 (M1) epitope and the influenza-associated MP58–66 (Flu) epitope were generated by coculturing peripheral blood–derived CD8+ T cells with mDC, as previously described (25, 26). In brief, human peripheral blood–derived CD8+ T cells were cocultured with autologous mDC in the presence of 100 U/ml IL-2. Once cells started to proliferate, cultures were maintained in the presence of 1000 U/ml IL-15. M1 and Flu epitope-specific CTL generated were quantified 7–10 d later by FACS staining with the respective tetramer reagents (BD Biosciences).

TCR-engineered (TCReng) CD4 and CD8 T cells specific for the M1 epitope were generated by engineering human CD4+ and CD8+ T cells, as described previously (7, 27). In brief, human PBL–derived T cells were activated with anti-CD3 and anti-CD28 Ab cross-linked on tissue culture wells (5 μg each per 24-well tissue culture plate well) and cultured in IMDM containing 100 IU/ml IL-2. Cells were transduced with retrovirus encoding the M1-specific TCR at 2–3 d postactivation. The TCReng T cells generated were quantified by FACS analysis with an M1 epitope–specific tetramer.

Functional characterization of Ag-specific T cells generated was done by coculturing them with T2-A2 cells, an HLA-A2 MHC class I molecule–expressing cell line described previously (6), human DC, hPSC (hESC and iPSC), or hPSC-derived APC (hPSC-APC) alone or following a 15–20-min pulse with a cognate peptide (M1 or Flu peptides for M1 Ag– or Flu Ag–specific T cells, respectively) or a control peptide (MAGE-3271–279 [M3]). Cytokines released in the supernatants were quantified 16 h postcoculture set-up by ELISA (R&D Systems), as described previously (6, 7, 23).

Expression of TLR mRNA in H9 hESC and human DC–derived iPSC lines was examined by RT-PCR using human blood–derived DC as control. Primers used are listed in Supplemental Table I. Expression of functional TLR proteins in H9 hESC and DC-derived iPSC lines was quantified by FACS staining. Expression of TLR3 and TLR9 was examined by intracellular FACS staining using Cytofix/Cytoperm (BD Biosciences). For functional characterization of TLRs in H9 hESC and human DC–derived iPSC lines, 1 × 104 cells were exposed to TLR ligands (InvivoGen) in IMDM, as recommended by the manufacturer, and cytokines released in the supernatants were quantified 16 h later by ELISA. Human blood–derived DC were used as control. TLR ligands used included 1 μg/ml Pam3CSK4 (TLR1/2), 108 cells per milliliter of heat-killed Listeria monocytogenes (TLR2), 0.1 μg/ml polyinosinic-polycytidylic acid (TLR3), 1 μg/ml LPS (TLR4), 1 μg/ml Salmonella typhimurium flagellin (TLR5), 1 μg/ml FSL-1, a synthetic lipoprotein representing the N-terminal part of the LP44 lipoprotein of Mycoplasma salivarium (TLR6/2), 1 μg/ml imiquimod (TLR7), 1 μg/ml 20 mer ssRNA coupled with cationic lipid (TLR8), and 5 μM CpG oligonucleotides (TLR9).

The primary MLR assay was set up using H9 hESC, DC-derived iPSC, or human peripheral blood–derived DC as stimulators; human PBL or human PBL purified CD4+CD25−ve T cells and CD8+ T cells were used as responders. In brief, 1 × 103 stimulator cells were cocultured with responder cells at different ratios (stimulatory responder ratios of 1:1, 1:10, 1:100, 1:200). Radiolabeled thymidine (1 μCi per well) was added 72 h postcoculture, and thymidine incorporated in the cells was quantified 24 h later using a liquid scintillation counter (Beckman Counter). For measurement of cytokines in MLR experiments, supernatants were taken out before adding radiolabeled thymidine and analyzed by ELISA. Activated CD4+CD25−ve and CD8+ T cells were used as control in the cytokine-release assays. Activated T cells were generated by coculturing naive T cells in 24-well culture plates with cross-linked anti-CD3 and anti-CD28 Ab (5 μg each), in the presence of 100 units/ml IL-2, for 5 d. For the cytokine-release assay, 1 × 103 activated T cells were re-exposed to anti-CD3/CD28 Ab (1 μg each) in a 96-well plate, and the cytokines released were quantified by ELISA.

H9 hESC or DC-derived iPSC were treated with 5-azacytidine (5-Aza; 10 ng/ml; Sigma-Aldrich) and/or trichostatin A (TSA; 10 ng/ml; InvivoGen) overnight (∼16 h), and the expression of MHC class I and MHC class II molecules, costimulatory molecules, and Ag-presentation pathway–associated molecules was quantified by FACS.

EB were generated by two methods. In the first method, single-cell suspensions of hPSC lines were made by treating hPSC cultures with collagenase (1 mg/ml). Next, cells were washed with PBS and resuspended in EB medium, followed by forced aggregation in AggreWell 400 plates (STEMCELL Technologies) by centrifugation in a swing bucket rotor at 2000 rpm for 12 min. The following day, hPSC clumps were gently transferred to low-adherence tissue culture plates in fresh EB medium. In the second method, hPSC colonies were gently scraped off, washed twice with PBS, and cultured in low-adherence tissue culture plates in the EB medium. For time-kinetics experiments, EB were used at the indicated time points, whereas the 20-d-old EB were used for the rest of the experiments. The quality of EB generated was validated by examining the expression of germ layer–specific markers by RT-PCR.

To confirm the generation of HSC precursors in hPSC-derived EB by FACS, single-cell suspensions of EB were generated by treating the hPSC-derived EB with 0.05% trypsin. Single-cell suspensions were washed with PBS and stained with the indicated Ab, and surface phenotype was examined by FACS staining using a FACSCalibur (BD Biosciences).

Generation of HSC in hPSC-derived EB was confirmed by a CFU assay using MethoCult medium (STEMCELL Technologies). In brief, a single-cell suspension of hPSC-derived EB was made as described above, and ∼2 × 104 to 5 × 104 cells were cultured in MethoCult medium in humidified six-well plates in a 37°C incubator. Generation of CFU was followed, and the colonies generated were counted using an Olympus IX71 microscope fitted with a SPOT Flex camera (SPOT Imaging Solutions).

For the generation of functional APC from hPSC, hPSC line–derived EB were cultured in tissue culture plates in IMDM containing 20% FBS with stage-specific differentiation-supporting cytokines. In stage 1, EB were cultured in the presence of vascular endothelial growth factor (50 ng/ml), bone morphogenetic protein 4 (50 ng/ml), and SCF (50 ng/ml) cytokines for 5 d to facilitate HSC precursor generation. At day 5, medium was supplemented with Flt3 (50 ng/ml), SCF (50 ng/ml), and GM-CSF (50 ng/ml) cytokines to facilitate the generation of APC. From day 10 onward, EB were cultured in GM-CSF (50 ng/ml) and IL-4 (20 ng/ml)–supplemented medium to generate mature APC. hPSC-derived APC were retrieved by straining cultures through 50-μm cell strainers (BD Biosciences) and photographed with an Olympus IX71 microscope equipped with a SPOT Flex camera; human blood–derived DC were used as controls. Functional characterization of the APC generated was done by examining their phagocytic ability, using FITC-tagged dextran beads (40,000 m.w.; Molecular Probes), and antigenic peptide epitope-presentation efficiency, as discussed before, using human blood–derived DC as positive controls.

Human PBL of HLA-A2+ donors were infected with recombinant Sendai virus vectors, a nongenome-integrating RNA virus, expressing the reprogramming factor genes c-Myc, Klf-4, Oct-3/4, and Sox-2 to generate iPSC lines. Fig. 1A shows our iPSC-generation schema (Fig. 1Ai) and the efficiency of Sendai virus infection in human PBL (Fig. 1Aii). To establish which cell population in human PBL is most likely to be reprogrammed by Sendai virus, we first examined the efficiency of GFP encoding Sendai virus infection in magnetic bead–purified CD4+ T cells, CD8+ T cells, and CD14+ monocyte-derived DC (Fig. 1B). In subsequent iPSC line–generation experiments, DC were infected with recombinant Sendai virus encoding GFP or the reprogramming factor genes. The first set of experiments was used to verify virus infection efficiency by FACS, and the second set of experiments was used to generate iPSC lines. Although we found that the Sendai virus could infect all three cell populations with comparable efficiencies (Fig. 1Ci), it was the DC fraction that consistently generated iPSC lines, in multiple experiments from different donors (Fig. 1Cii). Fig. 1Ciii shows the surface phenotype of human peripheral blood monocyte–derived DC used for iPSC line generation. Fig. 2Ai shows three iPSC lines derived from two individuals: a 37-y-old individual (NS02) and a 75-y-old individual (NS07). As shown in Fig. 2Aii, DC-derived iPSC lines became free of Sendai virus vector following ∼20 passages.

FIGURE 1.

(A) Schematic representation of derivation of iPSC lines from human PBL. (Ai) iPSC line derivation. (Aii) FACS analysis showing human PBL infected with recombinant Sendai virus expressing GFP. (Aiii) Bright-field images showing a potential iPSC clone derived from human PBL (left panel) and a well-established iPSC clone (right panel). Original magnification ×10. (B) Purity of human peripheral blood–derived T cells and monocytes used in the study. (Bi) FACS analysis showing the purity of human PBL-derived CD4+ and CD8+ T cells used for iPSC line generation, using a magnetic bead–purification method. (Bii) Purity of human PBL-derived CD14+ monocytes used for iPSC line generation, before and after purification with magnetic beads. (C) Efficiency of Sendai virus infection. (Ci) FACS analysis showing efficiency of GFP encoding Sendai virus infection in human PBL-derived CD4+ T cells, CD8+ T cells, and CD14+ monocyte-derived DC (multiplicity of infection = 5). (Cii) iPSC clone derivation data from PBL-derived T cells and DC from three donors. (Ciii) FACS analysis showing the phenotype of human blood monocyte-derived DC.

FIGURE 1.

(A) Schematic representation of derivation of iPSC lines from human PBL. (Ai) iPSC line derivation. (Aii) FACS analysis showing human PBL infected with recombinant Sendai virus expressing GFP. (Aiii) Bright-field images showing a potential iPSC clone derived from human PBL (left panel) and a well-established iPSC clone (right panel). Original magnification ×10. (B) Purity of human peripheral blood–derived T cells and monocytes used in the study. (Bi) FACS analysis showing the purity of human PBL-derived CD4+ and CD8+ T cells used for iPSC line generation, using a magnetic bead–purification method. (Bii) Purity of human PBL-derived CD14+ monocytes used for iPSC line generation, before and after purification with magnetic beads. (C) Efficiency of Sendai virus infection. (Ci) FACS analysis showing efficiency of GFP encoding Sendai virus infection in human PBL-derived CD4+ T cells, CD8+ T cells, and CD14+ monocyte-derived DC (multiplicity of infection = 5). (Cii) iPSC clone derivation data from PBL-derived T cells and DC from three donors. (Ciii) FACS analysis showing the phenotype of human blood monocyte-derived DC.

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FIGURE 2.

Generation and characterization of iPSC lines from human peripheral blood–derived DC of HLA-A2+ve donors. (Ai) Bright-field images of three iPSC lines derived from two donor-derived DC and the H9 hESC line. (ii) RT-PCR analysis for Sendai virus genome presence in DC-derived iPSC lines, ∼20 passages post-iPSC clone isolation. Human peripheral blood–derived DC that were infected with Sendai virus or were left uninfected were used as positive and negative controls, respectively. The iPSC lines generated were characterized for the expression of pluripotency markers by RT-PCR (B), FACS (C), and immunofluorescence microscopy (D), using the H9-hESC line as a positive control. In the RT-PCR analysis, coding and noncoding endogenous regions (depicted as Oct-E, Nanog-E, and Lin-28-E) were amplified. (E) Pluripotency characterization of iPSC lines by TaqMan quantitative RT-PCR–based hPSC pluripotency panels (Life Technologies) using H9 hESC as control. CT correlation analysis of the three DC-derived iPSC clones against the H9 hESC line. (F) Alkaline phosphatase staining of iPSC lines and H9 hESC line by VECTOR Red (Vector Laboratories). (G) Karyotype analysis of iPSC and H9 hESC lines. Data for hPSC line characterization are representative of at least three independent experiments each. Original magnification ×10 (A and F); original magnification ×20 (D).

FIGURE 2.

Generation and characterization of iPSC lines from human peripheral blood–derived DC of HLA-A2+ve donors. (Ai) Bright-field images of three iPSC lines derived from two donor-derived DC and the H9 hESC line. (ii) RT-PCR analysis for Sendai virus genome presence in DC-derived iPSC lines, ∼20 passages post-iPSC clone isolation. Human peripheral blood–derived DC that were infected with Sendai virus or were left uninfected were used as positive and negative controls, respectively. The iPSC lines generated were characterized for the expression of pluripotency markers by RT-PCR (B), FACS (C), and immunofluorescence microscopy (D), using the H9-hESC line as a positive control. In the RT-PCR analysis, coding and noncoding endogenous regions (depicted as Oct-E, Nanog-E, and Lin-28-E) were amplified. (E) Pluripotency characterization of iPSC lines by TaqMan quantitative RT-PCR–based hPSC pluripotency panels (Life Technologies) using H9 hESC as control. CT correlation analysis of the three DC-derived iPSC clones against the H9 hESC line. (F) Alkaline phosphatase staining of iPSC lines and H9 hESC line by VECTOR Red (Vector Laboratories). (G) Karyotype analysis of iPSC and H9 hESC lines. Data for hPSC line characterization are representative of at least three independent experiments each. Original magnification ×10 (A and F); original magnification ×20 (D).

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The pluripotency status of iPSC lines was characterized by examining the expression of pluripotency markers by RT-PCR, FACS, and immunofluorescence staining (Fig. 2B–D), using the H9 hESC line as a positive control. Expression of the coding and noncoding areas of the pluripotency genes was examined by RT-PCR to rule out the possibility of PCR amplification from open reading frames of pluripotency genes encoded by the reprogramming vectors (Fig. 2B). TaqMan quantitative RT-PCR–based hPSC pluripotency scorecard (Life Technologies) analysis was also performed on iPSC lines; it produced comparable CT correlation plots against the H9 hESC line (Fig. 2E). The iPSC lines were also confirmed for alkaline phosphatase activity (Fig. 2F). The karyotype of H9 hESC and human DC–derived iPSC lines was also examined; all hPSC lines exhibited normal karyotypes (Fig. 2G). Taken together, human DC–derived iPSC lines were thoroughly validated before proceeding to the characterization of their immunogenicity profiles and differentiation potential.

To characterize the immunogenicity profiles of human DC–derived iPSC lines, we first examined the expression and functional status of TLR molecules using human peripheral blood–derived DC as a positive control. As shown in Fig. 3A, human DC express TLR mRNA that are efficiently translated into corresponding functional proteins (Fig. 3E), and they can induce an inflammatory cytokine response upon exposure to corresponding TLR ligands (Fig. 3B). Interestingly, human DC–derived iPSC lines also express mRNA of TLR molecules (Fig. 3C); however, these mRNA are not translated into corresponding proteins (Fig. 3E), and exposure of these iPSC lines to different TLR ligands does not trigger an inflammatory cytokine response (Fig. 3D). Although Fig. 3D shows TLR-mediated cytokine-release data for just one iPSC line (NS07#1), other lines also did not produce any inflammatory cytokines upon TLR ligand exposure (data not shown).

FIGURE 3.

TLR-mediated innate immune mechanisms in DC-derived iPSC lines. (A) RT-PCR analysis of TLR expression in 5–7-d-old human peripheral blood–derived iDC from three donors (D1, D2, and D3). (B) Functional characterization of TLR in blood-derived iDC was done by quantification of the inflammatory cytokines IL-6 (Bi), TNF-α (Bii), and IL-10 (Biii) released in the supernatants following exposure to different TLR agonist ligands. Untreated DC were used as control. Minimum value of the y-axis is −100 (C). Expression of TLR mRNA in DC-derived iPSC lines was examined using blood-derived DC as control. (D) The functional status of TLR expressed in iPSC lines was examined by exposing them to different TLR agonist ligands and quantifying the cytokines IL-6 (Di), TNF-α (Dii), and IL-10 (Diii) released in the supernatant. Human blood–derived DC were used as positive control. (Control: untreated DC, H9 hESC, and iPSC). Minimum value of the y-axis is −100. (E) Expression of TLR proteins in DC-derived iPSC lines was examined by FACS staining, using human monocytes as control. Line graphs of respective protein expression (lines) overlaid on isotypes (filled graphs). (F) Activation of inflammasome in response to LPS exposure was examined in DC-derived iPSC lines by measuring IL-1β released upon LPS exposure. Human PBL were used as positive control. Data shown are representative of three independent experiments.

FIGURE 3.

TLR-mediated innate immune mechanisms in DC-derived iPSC lines. (A) RT-PCR analysis of TLR expression in 5–7-d-old human peripheral blood–derived iDC from three donors (D1, D2, and D3). (B) Functional characterization of TLR in blood-derived iDC was done by quantification of the inflammatory cytokines IL-6 (Bi), TNF-α (Bii), and IL-10 (Biii) released in the supernatants following exposure to different TLR agonist ligands. Untreated DC were used as control. Minimum value of the y-axis is −100 (C). Expression of TLR mRNA in DC-derived iPSC lines was examined using blood-derived DC as control. (D) The functional status of TLR expressed in iPSC lines was examined by exposing them to different TLR agonist ligands and quantifying the cytokines IL-6 (Di), TNF-α (Dii), and IL-10 (Diii) released in the supernatant. Human blood–derived DC were used as positive control. (Control: untreated DC, H9 hESC, and iPSC). Minimum value of the y-axis is −100. (E) Expression of TLR proteins in DC-derived iPSC lines was examined by FACS staining, using human monocytes as control. Line graphs of respective protein expression (lines) overlaid on isotypes (filled graphs). (F) Activation of inflammasome in response to LPS exposure was examined in DC-derived iPSC lines by measuring IL-1β released upon LPS exposure. Human PBL were used as positive control. Data shown are representative of three independent experiments.

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We next examined the functional status of the inflammasome pathway in these iPSC lines by examining the production of IL-1β in response to LPS signaling. As shown in Fig. 3F, human PBL produced considerable amounts of IL-1β when exposed to LPS, whereas DC-derived iPSC lines did not. Treatment with demethylating and/or deacetylating agents had no effect on TLR protein expression or inflammasome activation in these iPSC lines (data not shown).

Because T cells are the key immune mediators of adaptive immune responses, and DC are the best professional APC to orchestrate the generation of a protective Ag-specific T cell immune response, we next examined the ability of DC-derived iPSC lines to engage effector T cells and induce T cell responses. To this end, we first performed primary allogenic MLR by coculturing the human peripheral blood purified CD4+CD25 and CD8+ T cells with DC-derived iPSC lines, using human DC as a positive control. As shown in Fig. 4A, iPSC lines derived from human DC did not trigger proliferation of T cells, unlike human DC. As a secondary readout, we examined the production of effector cytokines by T cells in the MLR cultures, using activated T cells as a positive control. As shown in Fig. 4B, T cells activated with anti-CD3 and anti-CD28 Ab, upon restimulation with anti-CD3 Ab, produced significant amounts of cytokines; however, we did not detect any cytokines in the primary MLR cultures at 72 h post-MLR coculture set-up.

FIGURE 4.

Activation of human peripheral blood–derived naive T cells in an MLR. (A) Allogenic human PBL and purified CD4+25 and CD8+ T cells were cocultured with human DC–derived iPSC lines. Human peripheral blood–derived DC were used as positive control. Proliferation of cells was quantified by measuring incorporation of radioactively labeled thymidine. (B) Cytokines released in the MLR culture supernatants were quantified by ELISA at 72 h postcoculture set-up. Data shown are representative of two or more independent experiments. The minimum value of the y-axis is −100.

FIGURE 4.

Activation of human peripheral blood–derived naive T cells in an MLR. (A) Allogenic human PBL and purified CD4+25 and CD8+ T cells were cocultured with human DC–derived iPSC lines. Human peripheral blood–derived DC were used as positive control. Proliferation of cells was quantified by measuring incorporation of radioactively labeled thymidine. (B) Cytokines released in the MLR culture supernatants were quantified by ELISA at 72 h postcoculture set-up. Data shown are representative of two or more independent experiments. The minimum value of the y-axis is −100.

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DC express a range of costimulatory and coinhibitory molecules and Ag-presentation pathway intermediates that are necessary for optimum priming of Ag-specific T cell precursors (28, 29). Therefore, we next examined the expression of these molecules in DC-derived iPSC lines using human peripheral blood–derived DC as control (Fig. 5). As shown in Fig. 5A, blood-derived DC express mRNA of all of these molecules that are translated into corresponding proteins (Fig. 5D), and the expression of these molecules is upregulated in response to exposure to the DC maturation factors LPS and IFN-γ (Fig. 5B). Interestingly, we found that the DC-derived iPSC lines do not express mRNA of costimulatory molecules (Fig. 5C, left panels), but they do express mRNA of Ag-presentation pathway intermediaries (Fig. 5C, right panels). However, this could not be attributed to their DC origin, because the H9 hESC line also expressed mRNA of these Ag-presentation pathway molecules (Fig. 5C). More importantly, none of these mRNA, with the exception of LAMP-2, are translated into corresponding proteins in iPSC and H9 hESC lines (Fig. 5D).

FIGURE 5.

Analysis of costimulatory molecules and Ag-processing machinery in DC-derived iPSC lines. (A) RT-PCR analysis of the expression of costimulatory molecules and Ag-presentation pathway–associated molecules in the 5–7-d-old immature state (iDC). (B) FACS analysis for the expression of proteins corresponding to costimulatory molecules CD80 and CD86 and MHC class I molecules in human blood–derived DC and in iDC, as well as following maturation (mDC) with LPS and IFN-γ. As shown, iDC express these molecules, and their expression is upregulated upon maturation. (C) RT-PCR analysis of the expression of costimulatory molecules and Ag-presentation pathway intermediates in DC-derived iPSC lines. (D) FACS-mediated characterization of the expression of MHC class I and MHC class II molecules and costimulatory and coinhibitory molecules (Di) and Ag-presentation pathway intermediates (Dii) in human blood monocyte-derived DC and DC-derived iPSC lines. Line graphs of respective protein expression (lines) overlaid on isotypes (filled graphs). (E) Effect of IFN-γ, 5-Aza, and TSA treatment on DC-derived iPSC lines. FACS analysis of the effect of IFN-γ treatment (1000 IU/ml) on the expression of MHC class I (Ei) and MHC class II molecules (Eii). FACS analysis of the effect of 5-Aza (10 ng/ml) and/or TSA (10 ng/ml) treatment on the expression of MHC class I molecules in DC-derived iPSC lines and the H9 hESC line (Eiii) and on the expression of Ag-presentation pathway intermediates TAP-2 and LAMP-2 in the NS07#1 iPSC line (Eiv). In (Eiii) and (Eiv), isotypes are shown as filled graphs; left panel shows the expression of MHC class I (Eiii) and TAP-2 and LAMP-2 (Eiv) in control hESC or iPSC lines. Effect of 5-Aza and/or TSA treatment on the expression of these proteins under different treatment condition (lines) and untreated control cells (filled graphs). Data shown are representative of at least three independent experiments.

FIGURE 5.

Analysis of costimulatory molecules and Ag-processing machinery in DC-derived iPSC lines. (A) RT-PCR analysis of the expression of costimulatory molecules and Ag-presentation pathway–associated molecules in the 5–7-d-old immature state (iDC). (B) FACS analysis for the expression of proteins corresponding to costimulatory molecules CD80 and CD86 and MHC class I molecules in human blood–derived DC and in iDC, as well as following maturation (mDC) with LPS and IFN-γ. As shown, iDC express these molecules, and their expression is upregulated upon maturation. (C) RT-PCR analysis of the expression of costimulatory molecules and Ag-presentation pathway intermediates in DC-derived iPSC lines. (D) FACS-mediated characterization of the expression of MHC class I and MHC class II molecules and costimulatory and coinhibitory molecules (Di) and Ag-presentation pathway intermediates (Dii) in human blood monocyte-derived DC and DC-derived iPSC lines. Line graphs of respective protein expression (lines) overlaid on isotypes (filled graphs). (E) Effect of IFN-γ, 5-Aza, and TSA treatment on DC-derived iPSC lines. FACS analysis of the effect of IFN-γ treatment (1000 IU/ml) on the expression of MHC class I (Ei) and MHC class II molecules (Eii). FACS analysis of the effect of 5-Aza (10 ng/ml) and/or TSA (10 ng/ml) treatment on the expression of MHC class I molecules in DC-derived iPSC lines and the H9 hESC line (Eiii) and on the expression of Ag-presentation pathway intermediates TAP-2 and LAMP-2 in the NS07#1 iPSC line (Eiv). In (Eiii) and (Eiv), isotypes are shown as filled graphs; left panel shows the expression of MHC class I (Eiii) and TAP-2 and LAMP-2 (Eiv) in control hESC or iPSC lines. Effect of 5-Aza and/or TSA treatment on the expression of these proteins under different treatment condition (lines) and untreated control cells (filled graphs). Data shown are representative of at least three independent experiments.

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We also found that these iPSC lines express low levels of MHC class I molecules and significant levels of β-2 microglobulin, but they do not express MHC class II molecules, similar to the H9 hESC line (Fig. 5Di), in agreement with previous reports (19, 30). However, it should be noted that the expression of MHC class I and β-2 microglobulin molecules in hPSC lines was significantly lower than in the human blood–derived monocytes. Although the human DC-derived iPSC lines and the H9 hESC line do not express the MHC class I pathway–associated Ag-presentation intermediary, TAP-2, they express the MHC class II pathway–associated molecule LAMP-2, although at much lower levels than do human blood–derived monocytes (Fig. 5Dii). Expression of MHC class I molecules was inducible by IFN-γ treatment (Fig. 5Ei), as previously reported in hESC (30), but expression of MHC class II molecules was not (Fig. 5Eii). Of further interest, pretreatment with the demethylating agent 5-Aza and/or the deacetylating agent TSA had no effect on the expression of costimulatory molecules (Supplemental Fig. 1) or MHC class I molecules (Fig. 5Eiii), nor did it affect expression of the Ag-presentation pathway intermediates TAP-2 and LAMP-2 in iPSC lines or the H9 hESC line (Fig. 5Eiv). Fig. 5Eiv shows the effect of 5-Aza and/or TSA treatment on TAP-2 and LAMP-2 expression in the NS07#1 iPSC line; a similar effect was observed in other iPSC lines and the H9 hESC line (data not shown).

We used HLA-A2+ donor-derived DC to generate iPSC lines to examine the ability of these iPSC lines to present HLA-A2–restricted MHC class I peptide epitopes to corresponding Ag-specific T cells. For this purpose, we used M1 epitope– and Flu epitope–specific natural CTL generated from human blood–derived CD8+ T cells (Fig. 6Ai) in an in vitro coculture system, as described previously (4, 5, 25). Because human blood–derived CTL populations are heterogenous in their TCR repertoire, we also used the M1 epitope–specific transgenic TCReng CD8+ CTL (Fig. 6Bi), which were generated according to our previously published methods (6, 7). As shown in Fig. 6, despite expressing significantly low levels of HLA-A2–restricted MHC class I molecules (Fig. 6Aii), these iPSC and H9 hESC lines efficiently presented the M1 peptide epitope to M1 epitope–specific natural CTL (Fig. 6Aiii), as well as to M1 epitope–specific TCReng monoclonal CD8+ antitumor T cells (Fig. 6Bii) and efficiently presented the Flu peptide epitope to corresponding Flu Ag-specific natural CTL (Fig. 6Aiv). M3 was used as a control peptide in functional assays in Fig. 6, and all of the CTL used exhibited Ag-specific effector cytokine responses. Peptide dose kinetics for the presentation of M1 peptide epitope to M1 epitope–specific TCReng CD8 T cells showed that the iPSC lines exhibit peptide dose kinetics that are similar to those of T2-A2 cells (Fig. 6C), a human HLA-A2 MHC class I molecule–expressing cell line that is routinely used in functional assays (4, 7, 27). However, in agreement with our data showing a lack of costimulatory molecule expression in human DC–derived iPSC lines (Fig. 5Di), their inability to trigger T cell proliferation in MLR settings (Fig. 4), and the published reports showing immune-privileged properties in hESC lines (19), peptide-pulsed iPSC lines could not generate M1 epitope–specific functional CTL from autologous human peripheral blood–derived naive CD8+ T cells in an in vitro culture assay (data not shown). As expected, these peptide-pulsed iPSC lines were also killed by the CTL in an epitope-specific manner (data not shown).

FIGURE 6.

Presentation of HLA-A2–restricted peptide epitopes to Ag-specific T cells by human DC–derived iPSC lines. (Ai) M1 epitope– and Flu epitope–specific natural CTL were generated from HLA-A2+ve donor blood-derived CD8+ naive T cells and quantified by FACS using the respective epitope-specific tetramers. Isotype is overlaid on the upper left panel (upper left quadrant). (Aii) Expression of HLA-A2–restricted MHC class I molecules on DC-derived iPSC lines and the H9 hESC line was quantified by FACS using HLA-A2+ve and HLA-A2−ve donor-derived PBL as positive and negative controls, respectively. HLA-A2 protein expression (lines) overlaid on isotypes (filled graphs). (Aiii) M1 epitope–specific CTL were cocultured with H9 hESC or DC-derived iPSC pulsed with control or cognate peptides; IFN-γ and TNF-α cytokines released in the supernatants were quantified by ELISA. (Control cells, cells pulsed with control peptide, M3, or the M1 cognate peptide.) The minimum value of the y-axis in the graphs is −100. (Aiv) Flu epitope–specific CTL were cocultured with peptide-pulsed H9 hESC or DC-derived iPSC as in (Aiii) using Flu as the cognate peptide, and IFN-γ cytokine released in the supernatants was quantified by ELISA. Peptide-pulsed HLA-A2+ve donor-derived DC and/or HLA-A2+ surrogate target T2 cells were used as positive control in (Aiii) and (Aiv). The minimum value of the y-axis in the graphs is −100. (Bi) M1 epitope–specific TCReng CD8+ CTL were generated according to our published methods (7, 27) and quantified by M1 epitope–specific tetramer staining. (Bii) TCReng CD8+ CTL were used in functional assays against the peptide-pulsed iPSC and H9 hESC lines, as in (A), for the natural CTL. TCReng CD8 T cells also effectively recognized peptide epitope presented on human DC-derived iPSC lines. (C) Different dosages of M1 peptide–pulsed H9 hESC, DC-derived iPSC, and T2-A2 cells were cocultured with M1 epitope–specific TCReng CD8 T cells; IFN-γ and TNF-α released in the supernatants were quantified. The minimum values of the y-axis in the graphs in (Ci) and (Cii) are −20 and −100, respectively. Data shown are representative of at least two independent experiments each.

FIGURE 6.

Presentation of HLA-A2–restricted peptide epitopes to Ag-specific T cells by human DC–derived iPSC lines. (Ai) M1 epitope– and Flu epitope–specific natural CTL were generated from HLA-A2+ve donor blood-derived CD8+ naive T cells and quantified by FACS using the respective epitope-specific tetramers. Isotype is overlaid on the upper left panel (upper left quadrant). (Aii) Expression of HLA-A2–restricted MHC class I molecules on DC-derived iPSC lines and the H9 hESC line was quantified by FACS using HLA-A2+ve and HLA-A2−ve donor-derived PBL as positive and negative controls, respectively. HLA-A2 protein expression (lines) overlaid on isotypes (filled graphs). (Aiii) M1 epitope–specific CTL were cocultured with H9 hESC or DC-derived iPSC pulsed with control or cognate peptides; IFN-γ and TNF-α cytokines released in the supernatants were quantified by ELISA. (Control cells, cells pulsed with control peptide, M3, or the M1 cognate peptide.) The minimum value of the y-axis in the graphs is −100. (Aiv) Flu epitope–specific CTL were cocultured with peptide-pulsed H9 hESC or DC-derived iPSC as in (Aiii) using Flu as the cognate peptide, and IFN-γ cytokine released in the supernatants was quantified by ELISA. Peptide-pulsed HLA-A2+ve donor-derived DC and/or HLA-A2+ surrogate target T2 cells were used as positive control in (Aiii) and (Aiv). The minimum value of the y-axis in the graphs is −100. (Bi) M1 epitope–specific TCReng CD8+ CTL were generated according to our published methods (7, 27) and quantified by M1 epitope–specific tetramer staining. (Bii) TCReng CD8+ CTL were used in functional assays against the peptide-pulsed iPSC and H9 hESC lines, as in (A), for the natural CTL. TCReng CD8 T cells also effectively recognized peptide epitope presented on human DC-derived iPSC lines. (C) Different dosages of M1 peptide–pulsed H9 hESC, DC-derived iPSC, and T2-A2 cells were cocultured with M1 epitope–specific TCReng CD8 T cells; IFN-γ and TNF-α released in the supernatants were quantified. The minimum values of the y-axis in the graphs in (Ci) and (Cii) are −20 and −100, respectively. Data shown are representative of at least two independent experiments each.

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Generation of EB and formation of teratoma in mice are used to characterize the differentiation potential of hPSC lines by characterizing the expression of ectoderm, mesoderm, and endoderm markers in EB and the corresponding cellular structures in hPSC-induced teratomas. As shown in Fig. 7A, all of the iPSC lines generated EB that expressed characteristic ectoderm, mesoderm, and endoderm markers (Fig. 7B). Fig. 7C shows the histological analysis of teratomas generated from an iPSC line, showing all three germ layer–associated structures. EB are known to produce HSC precursors, and the CFU assay has been used to test the hematopoietic differentiation potential of EB-derived HSC. We found that EB derived from these DC-derived iPSC lines produced HSC precursors (Fig. 7D) that generated different types of colonies in CFU assay (Fig. 7E). We next examined the ability of these iPSC lines to generate human APC precursors by coculturing the iPSC-derived EB in appropriate cytokines (Fig. 7F–H) in a stage-specific differentiation system (Supplemental Fig. 3). As shown in Fig. 7Fi, we generated cells with DC morphology that are CD14+, CD11c+, and CD1c+ (Fig. 7Fii). To examine the phagocytic potential of hPSC-APC, cells were incubated with FITC-tagged dextran beads at 37°C for 2 h, and phagocytosis of beads was examined by FACS and immunofluorescence microscopy (Fig. 7G). hPSC-APC exhibit phagocytic potential (Fig. 7Gib), as do human blood–derived iDC (Fig. 7Gii). It should be noted that in a forward and side scatter plot analysis of human PBL (Fig. 7Gia, lower panel), monocytes (R2 gate) are located in the upper left portion of the plot since they are larger in size than the lymphocytes (R1 gate). Similarly, in the FITC-dextran bead phagocytosis assay shown in Fig. 7Gib, most phagocytic iPSC-APC also were the larger-sized cells (Fig. 7Gia, upper panel, R2 gate). To examine the ability of iPSC-APC to present peptide epitopes to Ag-specific T cells, we used human tumor Ag-specific TCReng CD4+ and CD8+ T cells (Supplemental Fig. 2). iPSC-APC efficiently presented the peptide epitopes to Ag-specific CD4 (Fig. 7Hi) and CD8 (Fig. 7Hii) T cells. Human PBL–derived monocytes were used as controls in the bright-field imaging (Fig. 7Fi), Giemsa staining (Fig. 7E), and phagocytosis assay (Fig. 7Gii) experiments. These sets of data confirm the differentiation potential of human DC–derived iPSC lines. Taken together, we have found that the dedifferentiation of human DC into iPSC lines shuts down their innate and adaptive immune mechanisms (Fig. 8), and that these DC-derived iPSC lines are pluripotent and exhibit potent differentiation potential.

FIGURE 7.

Characterization of differentiation potential of DC-derived iPSC lines. (A) EB derived from iPSC lines and the H9 hESC line. (B) RT-PCR analysis of iPSC and H9 hESC line–derived EB confirmed the expression of ectoderm, mesoderm, and endoderm markers. (C) Analysis of teratoma generated by the NS07#1 iPSC line with germ layer–associated structures. (D) FACS analysis of single-cell suspension of EB shows generation of CD34+ve HSC precursors. (E) Differentiation potential of EB-derived HSC precursors in a CFU assay. The CFU colonies generated (upper panels) were scored as CFU-GM, BFU-E, and CFU-GEMM (ref., Atlas on hematopoietic colonies, STEMCELL Technologies). Cells from these CFU colonies were stained with Wright–Giemsa stain (lower panels); insets show data from different respective planes. (FH). Generation of phagocytic APC from DC-derived iPSC lines. (Fi) iPSC-derived EB were cultured in a three-step method (Supplemental Fig. 3) to generate APC (iPSC-APC). Bright-field images (upper panels) and Giemsa staining (lower panels) of iPSC-APC and human blood–derived DC. Insets show data from different areas of the stained slides. Cells with morphological features of human DC were generated. (Fii) FACS analysis of iPSC-APC cultures confirmed the presence of CD14+, CD1c+, and CD11c+ precursors. (Gi) Phagocytic ability of iPSC-APC was examined by phagocytosis of FITC-tagged dextran beads (40,000 m.w.; Molecular Probes). (Gia) Gating strategy (upper panel). (Gib) As expected, larger-sized cells (R2 gate) contained more CD11c+ve cells with more phagocytic ability than did the smaller-sized cells. (Gia) Scatter plot of human PBL, with the R1 gate corresponding to the smaller-sized lymphocyte fraction and the R2 gate corresponding to the larger-sized monocyte fraction (lower panel). Human peripheral blood–derived CD11c+ iDC were used as positive control. Phagocytosis was examined by FACS (Giia) and by immunofluorescence microscopy (Giib). M1 epitope–specific TCReng antitumor CD4+ (Hi) and CD8+ (Hii) T cells were generated (Supplemental Fig. 2) and used to recognize the M1 epitope on peptide-pulsed iPSC-APC (iAPC). iPSC-APC efficiently presented M1 peptide to TCReng CD4 and CD8 T cells, as did T2-A2 cells (Supplemental Fig. 2). The minimum value of the y-axis in the graphs is −100. Data shown are representative of at least two independent experiments each. Original magnification ×4 (A, left, and E, top left); original magnification ×10 (A, right, and E, top middle and top right); original magnification ×40 (E, bottom, and Fi).

FIGURE 7.

Characterization of differentiation potential of DC-derived iPSC lines. (A) EB derived from iPSC lines and the H9 hESC line. (B) RT-PCR analysis of iPSC and H9 hESC line–derived EB confirmed the expression of ectoderm, mesoderm, and endoderm markers. (C) Analysis of teratoma generated by the NS07#1 iPSC line with germ layer–associated structures. (D) FACS analysis of single-cell suspension of EB shows generation of CD34+ve HSC precursors. (E) Differentiation potential of EB-derived HSC precursors in a CFU assay. The CFU colonies generated (upper panels) were scored as CFU-GM, BFU-E, and CFU-GEMM (ref., Atlas on hematopoietic colonies, STEMCELL Technologies). Cells from these CFU colonies were stained with Wright–Giemsa stain (lower panels); insets show data from different respective planes. (FH). Generation of phagocytic APC from DC-derived iPSC lines. (Fi) iPSC-derived EB were cultured in a three-step method (Supplemental Fig. 3) to generate APC (iPSC-APC). Bright-field images (upper panels) and Giemsa staining (lower panels) of iPSC-APC and human blood–derived DC. Insets show data from different areas of the stained slides. Cells with morphological features of human DC were generated. (Fii) FACS analysis of iPSC-APC cultures confirmed the presence of CD14+, CD1c+, and CD11c+ precursors. (Gi) Phagocytic ability of iPSC-APC was examined by phagocytosis of FITC-tagged dextran beads (40,000 m.w.; Molecular Probes). (Gia) Gating strategy (upper panel). (Gib) As expected, larger-sized cells (R2 gate) contained more CD11c+ve cells with more phagocytic ability than did the smaller-sized cells. (Gia) Scatter plot of human PBL, with the R1 gate corresponding to the smaller-sized lymphocyte fraction and the R2 gate corresponding to the larger-sized monocyte fraction (lower panel). Human peripheral blood–derived CD11c+ iDC were used as positive control. Phagocytosis was examined by FACS (Giia) and by immunofluorescence microscopy (Giib). M1 epitope–specific TCReng antitumor CD4+ (Hi) and CD8+ (Hii) T cells were generated (Supplemental Fig. 2) and used to recognize the M1 epitope on peptide-pulsed iPSC-APC (iAPC). iPSC-APC efficiently presented M1 peptide to TCReng CD4 and CD8 T cells, as did T2-A2 cells (Supplemental Fig. 2). The minimum value of the y-axis in the graphs is −100. Data shown are representative of at least two independent experiments each. Original magnification ×4 (A, left, and E, top left); original magnification ×10 (A, right, and E, top middle and top right); original magnification ×40 (E, bottom, and Fi).

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FIGURE 8.

Diagram showing the immunogenicity profile of human DC and the iPSC lines derived from them. Human DC harbor well-developed innate and adaptive immune mechanisms. The innate immune receptors (e.g., TLR) trigger an immediate response upon encountering pathogen-associated molecular patterns/danger-associated molecular patterns. Adaptive immune mechanisms process the internally synthesized and exogenously acquired Ag through MHC class I and MHC class II pathways and present the processed antigenic epitopes to CD8+ and CD4+ T cells, respectively, resulting in generation of CD8+ cytolytic responses and CD4+ helper responses. These immune mechanisms are shut down as a result of dedifferentiation of human DC; however, these iPSC express MHC class I molecules that can acquire antigenic peptide epitopes from the microenvironment and present them to Ag-specific T cells. The IFN-γ produced by T cells further induces the expression of MHC class I on these iPSC lines in a positive-feedback loop, and the peptide-bound iPSC lines are also killed by cytolytic T cells in an Ag-specific manner.

FIGURE 8.

Diagram showing the immunogenicity profile of human DC and the iPSC lines derived from them. Human DC harbor well-developed innate and adaptive immune mechanisms. The innate immune receptors (e.g., TLR) trigger an immediate response upon encountering pathogen-associated molecular patterns/danger-associated molecular patterns. Adaptive immune mechanisms process the internally synthesized and exogenously acquired Ag through MHC class I and MHC class II pathways and present the processed antigenic epitopes to CD8+ and CD4+ T cells, respectively, resulting in generation of CD8+ cytolytic responses and CD4+ helper responses. These immune mechanisms are shut down as a result of dedifferentiation of human DC; however, these iPSC express MHC class I molecules that can acquire antigenic peptide epitopes from the microenvironment and present them to Ag-specific T cells. The IFN-γ produced by T cells further induces the expression of MHC class I on these iPSC lines in a positive-feedback loop, and the peptide-bound iPSC lines are also killed by cytolytic T cells in an Ag-specific manner.

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Human DC express pattern recognition receptors on their cell surface, as well as in intracellular compartments, to sense the presence of foreign pathogens via recognition of pathogen-associated molecular patterns. In doing so, DC contribute to the development of an immediate inflammatory innate immune response, as well as undergo maturation to facilitate the subsequent generation of adaptive immune responses. mDC present endogenously synthesized antigenic peptide epitopes via the MHC class I pathway and exogenously acquired Ags via the MHC class II pathway to CD8+ and CD4+ effector T cells, respectively, to generate protective cellular immunity. In addition to Ag-presentation machinery, DC express several other accessory molecules, the costimulatory and coinhibitory molecules, which play essential roles in the generation of a protective or an inhibitory immune response. These unique properties make DC a central figure in immune-intervention strategies, including cancer immunotherapy (10, 29, 31).

As discussed previously, studies in mouse models describing rejection of iPSC-based teratomas claimed that the syngenic mouse iPSC lines can be inherently immunogenic (13, 14). Other groups showed minimal or no immunogenicity of syngenic mouse iPSC lines or their cellular derivatives (1517). These reports highlighted the need for a detailed characterization of the inherent immunogenicity of human iPSC lines for the development of safe and effective CRT. Because of the well-characterized immunologic properties of human DC, we generated iPSC lines from human peripheral blood–derived DC (Fig. 2) and examined the status of innate and adaptive immune mechanisms in these iPSC lines, using human DC as controls, to address the issue of immunogenicity of human iPSC lines. We show that the dedifferentiation of human DC effectively shuts down their innate immune response mechanisms, because no functional TLR proteins were detected in these iPSC lines, and the exposure to different TLR ligands does not trigger inflammatory cytokine release or inflammasome activation in these iPSC lines, unlike in the peripheral blood–derived DC (Fig. 3). Although we found that these iPSC lines express mRNA for the TLR molecules, this could not be attributed to their DC origin, because the H9 hESC line also expressed TLR mRNA. Treatment with demethylating and/or deacetylating agents had no effect on the expression of functional TLR proteins or on the activation of inflammasomes in human DC–derived iPSC lines (data not shown). The presence of TLR mRNA but the absence of functional TLR proteins also suggest the involvement of posttranscriptional mechanisms for an effective translation of these mRNA, such as requirement of lineage-specific molecular mediators/transcription factors. Taken together, our data show that the human DC–derived iPSC lines do not possess functional innate immune mechanisms.

DC harbor MHC class I and MHC class II Ag-presentation pathways that are tasked with processing of internally synthesized and exogenously acquired Ags, respectively, with avenues for cross-presentation (28). DC are well known as the best professional APC to optimally prime Ag-specific T cell precursors for the generation of protective T cell immunity, as a result of their ability to provide signal 1 (engagement of TCR with MHC class I and MHC class II molecules bound antigenic peptide epitopes), signal 2 (costimulatory signals), and signal 3 (production of immune-stimulatory cytokines, such as IL-12) (32). Therefore, we next examined the status of coreceptor molecules and Ag- processing and -presentation intermediates in human DC–derived iPSC lines. We found that the dedifferentiation of human DC results in dramatically lower expression of MHC class I molecules in DC-derived iPSC lines and no expression of MHC class II molecules (Fig. 5Di). Although the expression of MHC class I molecules is induced by IFN-γ treatment, it had no effect on the expression of MHC class II molecules (Fig. 5Ei, ii). In addition, treatment with demethylating and/or deacetylating agents had no effect on the expression of functional MHC class I and MHC class II proteins (Fig. 5Eiii, data not shown). These DC-derived iPSC do not express the mRNA or the corresponding proteins of costimulatory molecules (Fig. 5C, 5Di), and treatment with demethylating and/or deacetylating agents also has no effect on the expression of these molecules (Supplemental Fig. 1). Interestingly, these iPSC lines express mRNA for the MHC class I peptide-loading intermediate TAP-2; however, this could not be attributed to their DC origin because the H9 hESC line exhibited a similar profile (Fig. 5C). Of further interest, TAP-2 mRNA were not translated into corresponding functional TAP-2 protein in these lines (Fig. 5Dii), and treatment with demethylating and/or deacetylating agents had no effect on TAP-2 protein expression (Fig. 5Eiv). Although we found mRNA and protein expression for the lysosomal pathway–associated intermediate LAMP-2 in these iPSC lines (Fig. 5C, 5Dii), this also could not be attributed to their DC origin because the H9 hESC line also expressed it. As discussed previously, engagement of TLR molecules on DC modulates the expression of Ag-processing and -presentation pathway intermediates and results in their functional maturation (33); however, engagement of TLR ligands had no effect on the immunogenic profile of human DC–derived iPSC lines, because these cells do not express functional TLR proteins. Taken together, our data show that the coreceptor engagement ability, as well as the Ag-processing and -presentation machinery, of human DC is effectively shut down during the course of their dedifferentiation into the iPSC state.

Our data showing that the DC-derived iPSC lines, unlike human DC, do not trigger T cell proliferation in an allogenic MLR, even at an E:T ratio of 200:1 (Fig. 4A), is in agreement with previous findings in hESC (19). We used an HLA-A2+ donor population for iPSC derivation, which allowed us to examine their ability to present HLA-A2 MHC class I molecule–restricted peptide epitopes to Ag-specific T cells (Fig. 6). We found that, despite expressing low levels of MHC class I molecules in comparison with DC (Fig. 6Aii), these iPSC lines can efficiently present the peptide epitopes to Ag-specific T cells (Fig. 6Aiii, iv); in doing so, peptide-pulsed iPSC lines are also killed by cytolytic T cells (data not shown). Taken together, our findings (summarized schematically in Fig. 8) demonstrate that the dedifferentiation of human terminally differentiated immunogenic DC effectively shuts down their innate and adaptive immune mechanisms. Our data also suggest that the presentation of antigenic peptides acquired from the microenvironment to cytolytic T cells might result in the targeted killing of the transplanted iPSC and rejection of iPSC-induced teratomas in immune-competent animals.

To characterize the differentiation potential of the iPSC lines generated, we examined their ability to generate HSC precursors and functional APC. Human DC were generated from the HSC precursors derived from different anatomical locations, such as peripheral blood, bone marrow, and umbilical cord blood, and they were shown to be effective immune stimulators (3436). However, donor-specific iPSC lines can be used as an off-the-shelf resource for deriving autologous immunologically matched HSC precursors to generate donor-specific cells of choice, including APC. Given that APC can be immunogenic or tolerogenic (28, 37), incorporation of appropriate growth supplements in differentiating medium could also be used to generate naive APC with defined immunological properties. It should be pointed out that APC were generated from hESC lines (3840); however, efficient models are needed for in-depth characterization of the molecular, cellular, and functional profiles of human iPSC-derived APC. This is important given that the molecular, cellular, and functional profiles of cellular derivatives derived from iPSC lines, from diverse somatic cell sources, and/or by various methods can be different (41). In this context, our data showing that young and elderly donor DC-derived iPSC lines exhibit comparable differentiation potential are noteworthy, given that they yield EB that can produce HSC precursors that can be further differentiated into cells that exhibit DC morphology, are phagocytic, and can present antigenic peptide epitopes to Ag-specific T cells (Fig. 7). Our findings suggest that the dedifferentiation and redifferentiation of human DC can be used as an efficient model to systematically characterize the cellular, functional, and molecular profiles of human iPSC-derived APC, using autologous blood–derived DC as control. We believe that our findings have implications for understanding the development of human DC lineages and for DC-based CRT.

We thank UCHC General Clinical Core staff members Susan Walters, Kathleen Curley, Harriet Zawistowski, and Tom Kiley for human subject blood samples. We also thank Leann Crandell (University of Connecticut [UConn] Health Stem Cell Core Facility) for providing hESC lines and technical insights, John Glynn (UConn Health Molecular Core Facility) for help with real-time PCR for stem cell pluripotency analysis, Susan Krueger (UConn Health Microscopy Core Facility) for help with confocal microscopy, Dr. Judy Brown (UConn Chromosome Core Facility) for karyotype analysis of hESC and iPSC lines, and Dr. Evan Jellison (UConn Health FACS Core Facility) for help with FACS analysis. We also thank the laboratories of Dr. Alexander Lichtler and Dr. Ernst Reichenberger for help during the course of the study.

This work was supported by Stem Cell Seed Grant 10-SCA-23 and Stem Cell Established Investigator Grant 13-SCB-05 (both to A.C.) from the State of Connecticut, as well as by Grant 1RR06192 from the National Institutes of Health to the Clinical Research Center, University of Connecticut Health Center.

The online version of this article contains supplemental material.

Abbreviations used in this article:

5-Aza

5-azacytidine

CRT

cell-replacement therapy

DC

dendritic cell

EB

embryoid body

Flu

influenza-associated MP58–66

hESC

human embryonic stem cell

hPSC

human pluripotent stem cell

hPSC-APC

hPSC-derived APC

HSC

hematopoietic stem cell

iDC

immature DC

iPSC

induced pluripotent stem cell

iPSC-APC

iPSC-derived donor-specific APC

M1

human melanoma–associated MART-127–35

M3

MAGE-3271–279

mDC

mature DC

MEF

mouse embryonic fibroblast

SCF

stem cell factor

TCReng

TCR-engineered

TSA

trichostatin A

UCHC

University of Connecticut Health Center

UConn

University of Connecticut.

1
Thomson
J. A.
,
Itskovitz-Eldor
J.
,
Shapiro
S. S.
,
Waknitz
M. A.
,
Swiergiel
J. J.
,
Marshall
V. S.
,
Jones
J. M.
.
1998
.
Embryonic stem cell lines derived from human blastocysts.
Science
282
:
1145
1147
.
2
Takahashi
K.
,
Tanabe
K.
,
Ohnuki
M.
,
Narita
M.
,
Ichisaka
T.
,
Tomoda
K.
,
Yamanaka
S.
.
2007
.
Induction of pluripotent stem cells from adult human fibroblasts by defined factors.
Cell
131
:
861
872
.
3
Yu
J.
,
Vodyanik
M. A.
,
Smuga-Otto
K.
,
Antosiewicz-Bourget
J.
,
Frane
J. L.
,
Tian
S.
,
Nie
J.
,
Jonsdottir
G. A.
,
Ruotti
V.
,
Stewart
R.
, et al
.
2007
.
Induced pluripotent stem cell lines derived from human somatic cells.
Science
318
:
1917
1920
.
4
Chhabra
A.
,
Chakraborty
N. G.
,
Mukherji
B.
.
2008
.
Silencing of endogenous IL-10 in human dendritic cells leads to the generation of an improved CTL response against human melanoma associated antigenic epitope, MART-1 27-35.
Clin. Immunol.
126
:
251
259
.
5
Chhabra
A.
,
Mehrotra
S.
,
Chakraborty
N. G.
,
Mukherji
B.
,
Dorsky
D. I.
.
2004
.
Cross-presentation of a human tumor antigen delivered to dendritic cells by HSV VP22-mediated protein translocation.
Eur. J. Immunol.
34
:
2824
2833
.
6
Chhabra
A.
,
Yang
L.
,
Wang
P.
,
Comin-Anduix
B.
,
Das
R.
,
Chakraborty
N. G.
,
Ray
S.
,
Mehrotra
S.
,
Yang
H.
,
Hardee
C. L.
, et al
.
2008
.
CD4+CD25− T cells transduced to express MHC class I-restricted epitope-specific TCR synthesize Th1 cytokines and exhibit MHC class I-restricted cytolytic effector function in a human melanoma model.
J. Immunol.
181
:
1063
1070
.
7
Ray
S.
,
Chhabra
A.
,
Chakraborty
N. G.
,
Hegde
U.
,
Dorsky
D. I.
,
Chodon
T.
,
von Euw
E.
,
Comin-Anduix
B.
,
Koya
R. C.
,
Ribas
A.
, et al
UCLA-CALTECH-CHLA-USC-UCONN Consortium on Translational Program in Engineered Immunity
.
2010
.
MHC-I–restricted melanoma antigen specific TCR-engineered human CD4+ T cells exhibit multifunctional effector and helper responses, in vitro.
Clin. Immunol.
136
:
338
347
.
8
Chhabra
A.
2009
.
MHC class I TCR engineered anti-tumor CD4 T cells: implications for cancer immunotherapy.
Endocr. Metab. Immune Disord. Drug Targets
9
:
344
352
.
9
Mukherji
B.
,
Chakraborty
N. G.
,
Yamasaki
S.
,
Okino
T.
,
Yamase
H.
,
Sporn
J. R.
,
Kurtzman
S. K.
,
Ergin
M. T.
,
Ozols
J.
,
Meehan
J.
, et al
.
1995
.
Induction of antigen-specific cytolytic T cells in situ in human melanoma by immunization with synthetic peptide-pulsed autologous antigen presenting cells.
Proc. Natl. Acad. Sci. USA
92
:
8078
8082
.
10
Gilboa
E.
2007
.
DC-based cancer vaccines.
J. Clin. Invest.
117
:
1195
1203
.
11
Morgan
R. A.
,
Dudley
M. E.
,
Wunderlich
J. R.
,
Hughes
M. S.
,
Yang
J. C.
,
Sherry
R. M.
,
Royal
R. E.
,
Topalian
S. L.
,
Kammula
U. S.
,
Restifo
N. P.
, et al
.
2006
.
Cancer regression in patients after transfer of genetically engineered lymphocytes.
Science
314
:
126
129
.
12
Porter
D. L.
,
Kalos
M.
,
Zheng
Z.
,
Levine
B.
,
June
C.
.
2011
.
Chimeric antigen receptor therapy for B-cell malignancies.
J. Cancer
2
:
331
332
.
13
Zhao
T.
,
Zhang
Z. N.
,
Rong
Z.
,
Xu
Y.
.
2011
.
Immunogenicity of induced pluripotent stem cells.
Nature
474
:
212
215
.
14
Zhao
T.
,
Zhang
Z. N.
,
Westenskow
P. D.
,
Todorova
D.
,
Hu
Z.
,
Lin
T.
,
Rong
Z.
,
Kim
J.
,
He
J.
,
Wang
M.
, et al
.
2015
.
Humanized mice reveal differential immunogenicity of cells derived from autologous induced pluripotent stem cells.
Cell Stem Cell
17
:
353
359
.
15
Araki
R.
,
Uda
M.
,
Hoki
Y.
,
Sunayama
M.
,
Nakamura
M.
,
Ando
S.
,
Sugiura
M.
,
Ideno
H.
,
Shimada
A.
,
Nifuji
A.
,
Abe
M.
.
2013
.
Negligible immunogenicity of terminally differentiated cells derived from induced pluripotent or embryonic stem cells.
Nature
494
:
100
104
.
16
Guha
P.
,
Morgan
J. W.
,
Mostoslavsky
G.
,
Rodrigues
N. P.
,
Boyd
A. S.
.
2013
.
Lack of immune response to differentiated cells derived from syngeneic induced pluripotent stem cells.
Cell Stem Cell
12
:
407
412
.
17
Morizane
A.
,
Doi
D.
,
Kikuchi
T.
,
Okita
K.
,
Hotta
A.
,
Kawasaki
T.
,
Hayashi
T.
,
Onoe
H.
,
Shiina
T.
,
Yamanaka
S.
,
Takahashi
J.
.
2013
.
Direct comparison of autologous and allogeneic transplantation of iPSC-derived neural cells in the brain of a non-human primate.
Stem Cell Rep.
1
:
283
292
.
18
Drukker
M.
2004
.
Immunogenicity of human embryonic stem cells: can we achieve tolerance?
Springer Semin. Immunopathol.
26
:
201
213
.
19
Li
L.
,
Baroja
M. L.
,
Majumdar
A.
,
Chadwick
K.
,
Rouleau
A.
,
Gallacher
L.
,
Ferber
I.
,
Lebkowski
J.
,
Martin
T.
,
Madrenas
J.
,
Bhatia
M.
.
2004
.
Human embryonic stem cells possess immune-privileged properties.
Stem Cells
22
:
448
456
.
20
Mohib
K.
,
Allan
D.
,
Wang
L.
.
2010
.
Human embryonic stem cell-extracts inhibit the differentiation and function of monocyte-derived dendritic cells.
Stem Cell Rev.
6
:
611
621
.
21
Drukker
M.
,
Katchman
H.
,
Katz
G.
,
Even-Tov Friedman
S.
,
Shezen
E.
,
Hornstein
E.
,
Mandelboim
O.
,
Reisner
Y.
,
Benvenisty
N.
.
2006
.
Human embryonic stem cells and their differentiated derivatives are less susceptible to immune rejection than adult cells.
Stem Cells
24
:
221
229
.
22
Swijnenburg
R. J.
,
Schrepfer
S.
,
Govaert
J. A.
,
Cao
F.
,
Ransohoff
K.
,
Sheikh
A. Y.
,
Haddad
M.
,
Connolly
A. J.
,
Davis
M. M.
,
Robbins
R. C.
,
Wu
J. C.
.
2008
.
Immunosuppressive therapy mitigates immunological rejection of human embryonic stem cell xenografts.
Proc. Natl. Acad. Sci. USA
105
:
12991
12996
.
23
Mehrotra
S.
,
Chhabra
A.
,
Chattopadhyay
S.
,
Dorsky
D. I.
,
Chakraborty
N. G.
,
Mukherji
B.
.
2004
.
Rescuing melanoma epitope-specific cytolytic T lymphocytes from activation-induced cell death, by SP600125, an inhibitor of JNK: implications in cancer immunotherapy.
J. Immunol.
173
:
6017
6024
.
24
Mehrotra
S.
,
Chhabra
A.
,
Chakraborty
A.
,
Chattopadhyay
S.
,
Slowik
M.
,
Stevens
R.
,
Zengou
R.
,
Mathias
C.
,
Butterfield
L. H.
,
Dorsky
D. I.
, et al
.
2004
.
Antigen presentation by MART-1 adenovirus-transduced interleukin-10-polarized human monocyte-derived dendritic cells.
Immunology
113
:
472
481
.
25
Chhabra
A.
,
Mehrotra
S.
,
Chakraborty
N. G.
,
Dorsky
D. I.
,
Mukherji
B.
.
2006
.
Activation-induced cell death of human melanoma specific cytotoxic T lymphocytes is mediated by apoptosis-inducing factor.
Eur. J. Immunol.
36
:
3167
3174
.
26
Mehrotra
S.
,
Chhabra
A.
,
Hegde
U.
,
Chakraborty
N. G.
,
Mukherji
B.
.
2007
.
Inhibition of c-Jun N-terminal kinase rescues influenza epitope-specific human cytolytic T lymphocytes from activation-induced cell death.
J. Leukoc. Biol.
81
:
539
547
.
27
Chhabra
A.
,
Mukherji
B.
.
2013
.
Death receptor-independent activation-induced cell death in human melanoma antigen-specific MHC class I–restricted TCR-engineered CD4 T cells.
J. Immunol.
191
:
3471
3477
.
28
Schuurhuis
D. H.
,
Fu
N.
,
Ossendorp
F.
,
Melief
C. J.
.
2006
.
Ins and outs of dendritic cells.
Int. Arch. Allergy Immunol.
140
:
53
72
.
29
Steinman
R. M.
2012
.
Decisions about dendritic cells: past, present, and future.
Annu. Rev. Immunol.
30
:
1
22
.
30
Drukker
M.
,
Katz
G.
,
Urbach
A.
,
Schuldiner
M.
,
Markel
G.
,
Itskovitz-Eldor
J.
,
Reubinoff
B.
,
Mandelboim
O.
,
Benvenisty
N.
.
2002
.
Characterization of the expression of MHC proteins in human embryonic stem cells.
Proc. Natl. Acad. Sci. USA
99
:
9864
9869
.
31
Banchereau
J.
,
Palucka
A. K.
.
2005
.
Dendritic cells as therapeutic vaccines against cancer.
Nat. Rev. Immunol.
5
:
296
306
.
32
Reis e Sousa
C.
2006
.
Dendritic cells in a mature age.
Nat. Rev. Immunol.
6
:
476
483
.
33
Vyas
J. M.
,
Van der Veen
A. G.
,
Ploegh
H. L.
.
2008
.
The known unknowns of antigen processing and presentation.
Nat. Rev. Immunol.
8
:
607
618
.
34
Caux
C.
,
Dezutter-Dambuyant
C.
,
Schmitt
D.
,
Banchereau
J.
.
1992
.
GM-CSF and TNF-alpha cooperate in the generation of dendritic Langerhans cells.
Nature
360
:
258
261
.
35
Szabolcs
P.
,
Moore
M. A.
,
Young
J. W.
.
1995
.
Expansion of immunostimulatory dendritic cells among the myeloid progeny of human CD34+ bone marrow precursors cultured with c-kit ligand, granulocyte-macrophage colony-stimulating factor, and TNF-alpha.
J. Immunol.
154
:
5851
5861
.
36
Strobl
H.
,
Riedl
E.
,
Scheinecker
C.
,
Bello-Fernandez
C.
,
Pickl
W. F.
,
Rappersberger
K.
,
Majdic
O.
,
Knapp
W.
.
1996
.
TGF-beta 1 promotes in vitro development of dendritic cells from CD34+ hemopoietic progenitors.
J. Immunol.
157
:
1499
1507
.
37
Steinman
R. M.
,
Hawiger
D.
,
Nussenzweig
M. C.
.
2003
.
Tolerogenic dendritic cells.
Annu. Rev. Immunol.
21
:
685
711
.
38
Zhan
X.
,
Dravid
G.
,
Ye
Z.
,
Hammond
H.
,
Shamblott
M.
,
Gearhart
J.
,
Cheng
L.
.
2004
.
Functional antigen-presenting leucocytes derived from human embryonic stem cells in vitro.
Lancet
364
:
163
171
.
39
Slukvin
I. I.
,
Vodyanik
M. A.
,
Thomson
J. A.
,
Gumenyuk
M. E.
,
Choi
K. D.
.
2006
.
Directed differentiation of human embryonic stem cells into functional dendritic cells through the myeloid pathway.
J. Immunol.
176
:
2924
2932
.
40
Tseng
S. Y.
,
Nishimoto
K. P.
,
Silk
K. M.
,
Majumdar
A. S.
,
Dawes
G. N.
,
Waldmann
H.
,
Fairchild
P. J.
,
Lebkowski
J. S.
,
Reddy
A.
.
2009
.
Generation of immunogenic dendritic cells from human embryonic stem cells without serum and feeder cells.
Regen. Med.
4
:
513
526
.
41
Polo
J. M.
,
Liu
S.
,
Figueroa
M. E.
,
Kulalert
W.
,
Eminli
S.
,
Tan
K. Y.
,
Apostolou
E.
,
Stadtfeld
M.
,
Li
Y.
,
Shioda
T.
, et al
.
2010
.
Cell type of origin influences the molecular and functional properties of mouse induced pluripotent stem cells.
Nat. Biotechnol.
28
:
848
855
.

The authors have no financial conflicts of interest.

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