Generation of Mouse Pluripotent Stem Cell–Derived Proliferating Myeloid Cells as an Unlimited Source of Functional Antigen-Presenting Cells

Dendritic cells (DC) are commonly used as biologic therapy for cancer because of their physiologic roles in initiating and modulating the host immune response (1, 2). In many cases, monocytes obtained from patients by apheresis have been used as a source of DCs (3, 4). However, the number of monocytes obtained from peripheral blood, their ability to differentiate into DCs, and the quality of the resulting DCs vary among patients. Therefore, it can be difficult to stably generate a sufficient number of high-quality DCs for use in cancer therapy. In addition, the requirement to prepare DCs separately for each patient prevents the broader application of this strategy.

Pluripotent stem cells (PSC), such as embryonic stem cells (ESC) and the recently developed induced PSCs (iPSC), have the potential to propagate indefinitely and can differentiate into various somatic cell types (5–7). Previous studies have established methods of generating DCs from PSCs, and demonstrated the utility of PSC-derived DCs in cancer immunotherapy (8–25). However, generating a large number of DCs from ESCs or iPSCs has required a scaling-up of the initial volume of undifferentiated PSCs. In addition, these methods are too laborious for practical application in the clinical setting.

We recently developed a simple and efficient method for obtaining a large number of functional antigen-presenting cells (APC) from human iPSCs (26, 27). By introducing the c-Myc gene, along with antisenescence factors, such as Bmi-1 or Ezh2, the myeloid cells generated from iPSCs could proliferate almost indefinitely while retaining the capacity to differentiate into potent APCs. In addition, cells derived from transporter associated with antigen processing 2 (TAP2)–deficient iPSCs downregulated the expression of endogenous human leukocyte antigen (HLA)-I and avoided recognition by alloreactive CD8⁺ T cells, and cells expressing exogenously introduced HLA-I genes stimulated tumor-associated antigen-specific CD8⁺ T cells in vitro (26). Genetic modification of macrophages derived from the proliferating myeloid cells (pMC) to express an anti-HER2 antibody or interferon (IFN) could also exert therapeutic effects against peritoneally disseminated gastric and pancreatic cancer in xenograft models (27). However, their in vivo therapeutic potential against cancer and in vivo safety have not been examined in syngeneic mouse models. In addition, whether pMCs can be generated from mouse PSCs remains unclear.

In this study, we generated pMCs from mouse ESCs and iPSCs, and showed their potential to differentiate into functional APCs. This new method enabled us to obtain a large number of APCs from a small number of undifferentiated PSCs. Moreover, once the pMCs were established, it was possible to supply APCs in a few days using the pMCs as a cell source. Using a model antigen, we demonstrated that vaccination with APCs was safe and feasible for cancer immunotherapy, without any induction of autoimmunity or leukemia in vivo.

C57BL/6 and BALB/c mice were purchased from Japan SLC, Inc. or CLEA Japan, Inc., and were maintained under specific pathogen-free conditions. All animal experiments were performed with approval from the Animal Experiment Committee of the Aichi Cancer Center.

The mouse ESC line, B6-ES, derived from C57BL/6 blastocysts (28), and the mouse iPSC lines, 2A-4F-100 and 2A-4F-136 (29), derived from C57BL/6 embryonic fibroblasts, were maintained as described previously (28, 29). The M-CSF–defective bone marrow–derived stromal cell line, OP9 (30), was maintained in α-minimum essential medium (α-MEM; Life Technologies) supplemented with 20% fetal bovine serum (FBS), and the cells were seeded onto gelatin-coated dishes before being used as feeder cells. MO4 (31), a C57BL/6-derived B16 melanoma cell line expressing OVA, luciferase-transduced MO4 (MO4-Luc), and the EL-4 thymoma cell line, were maintained in RPMI-1640 medium (Sigma-Aldrich) supplemented with 10% FBS.

A lentivirus vector, CSII-EF, and the plasmids used for lentiviral vector packaging, pCMV-VSV-G-RSV-Rev and pCAG-HIVgp, were kindly provided by Dr. H. Miyoshi (RIKEN BioResource Center; Tsukuba, Japan). CSII-EF containing a human c-Myc cDNA insert was used as described previously (32). Plasmid constructs were introduced into 293T cells using lipofection (Lipofectamine 2000; Life Technologies). Three days later, recombinant lentiviruses were recovered from the culture supernatant using a Lenti-X Concentrator (Clontech).

ESCs or iPSCs were induced to differentiate into myeloid cells according to an established procedure (Fig. 1; refs. 9, 11–14, 18, 19). Briefly, the ESCs or iPSCs were cultured on feeder layers of OP9 cells for 6 to 7 days in α-MEM supplemented with 20% FBS. The mesodermally differentiated cells were then harvested, reseeded onto fresh OP9 cell layers, and cultured in α-MEM supplemented with 20% FBS, 20 ng/mL GM-CSF, and 50 μmol/L 2-ME. On day 13 to 14, floating cells were recovered by pipetting. These cells were considered to be ESC- or iPSC-derived myeloid cells (ES-MCs or iPS-MCs, respectively). The cells were infected with lentivirus vectors expressing the c-Myc gene in the presence of 8 ng/mL polybrene (Sigma-Aldrich), and were cultured in α-MEM supplemented with 20% FBS, 30 ng/mL GM-CSF, and 30 ng/mL M-CSF. After 5 to 6 days, proliferating cells appeared and were considered to be ESC- or iPSC-derived pMCs (ES-pMC or iPS-pMC, respectively). To induce the differentiation of these cells into DC-like cells (pMC-DC), they were cultured in RPMI-1640 supplemented with 20% FBS in the presence of 20 ng/mL IL4 plus 30 ng/mL GM-CSF for 3 days.

Cell samples were treated with a Fc-receptor blocking reagent (Miltenyi Biotec) for 10 minutes, stained with the fluorochrome-conjugated monoclonal antibody (mAb; Supplementary Materials and Methods) for 20 minutes, and washed three times with phosphate-buffered saline/2% FBS. The stained cell samples were analyzed on a FACSCalibur flow cytometer, and the data were analyzed using the BD CellQuest Pro software program (BD Biosciences) or FlowJo software program (TreeStar Inc.). For the morphologic analysis, cytospin specimens were stained with May-Grünwald/Giemsa (Merck).

Cell proliferation was evaluated by the direct enumeration of cells, the standard 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, and the [³H]-thymidine incorporation assay (Supplementary Materials and Methods).

Cytokine levels in the culture supernatants were evaluated using enzyme-linked immunosorbent assays (ELISA; mouse TNFα and IL12p70: Affymetrix eBioscience).

ES-pMC- or iPS-pMC–derived DC-like cells (pMC-DC) were loaded with vehicle or 10 μmol/L OVA_257-264 peptide for 3 hours, and were injected i.p. into C57BL/6 mice (1.0 × 10⁵ cells/mouse) twice at 7-day intervals. In one experiment, the pMC-DCs were heat-killed by incubation at 70°C for 20 minutes or were irradiated (45 Gy) before injection. Seven days after the second injection, spleen cells were isolated from the mice and pooled for each group of 3 mice. After hemolysis, the spleen cells were cultured in 24-well culture plates (2.5 × 10⁶/well) in RPMI-1640 supplemented with 10% horse serum, 2-ME (50 μmol/L), IL2 (10 U/mL), and OVA_257–264 peptide (0.1 μmol/L). After 5 days, the cells were recovered, and the OVA-specific cytotoxic T lymphocyte (CTL) activity was analyzed using the ⁵¹Cr-release assay using EL-4 thymoma cells or MO4 melanoma cells as targets. The percentage of specific lysis was calculated as: 100 × (experimental release − spontaneous release)/(maximal release − spontaneous release). The spontaneous release and maximal release were determined in the presence of medium alone and with 1% Triton X-100, respectively.

The OVA_257–264 peptide (10 μmol/L)- or vehicle-loaded pMC-DCs were injected i.p. into C57BL/6 mice (1.0 × 10⁵ cells/mouse) twice at 7-day intervals. Seven days after the second transfer, 2.0 × 10⁵ luciferase-transduced MO4 cells were inoculated s.c. into the shaved flanks. The mice were monitored for tumor growth and survival; tumor size was measured at 5-day intervals until the mice either died or were euthanized because of tumor progression.

Tumor-bearing mice were injected with 200-μL d-luciferin (15 mg/mL, VivoGlo Luciferin; Promega) under 2% inhaled isoflurane anesthesia, and bioluminescence images were obtained using the IVIS Lumina II instrument with the Living Image software version 3.2 (Xenogen).

The Prism version 6.0e software program (GraphPad Software) was used for all of the statistical analyses. To compare multiple experimental groups, a one-way ANOVA with the Bonferroni post hoc test was used to assess statistical significance. For statistical analysis of the Kaplan–Meier survival curves, a log-rank (Mantel–Cox) test was used to calculate P values. P values < 0.05 were considered to denote statistical significance, and are indicated in figures by asterisks (*, P < 0.05; **, P < 0.01).

We previously established a method for generating human myelomonocytic cells with the capacity for proliferation using the simultaneous transduction of c-Myc together with Bmi-1 or Ezh2 into human iPSC-derived myeloid cells (26, 27). In that preliminary study, mouse ES-MCs were transduced with several genes to identify those that induced proliferation. On the basis of these results, we found that mouse ES-MCs could proliferate after lentivirus-mediated transduction of c-Myc alone (data not shown). We then attempted to induce the proliferation of mouse ES-MCs and iPS-MCs through the transduction of c-Myc alone, and successfully stimulated their differentiation into DC-like APCs. The procedure for the generation of proliferating ES- or iPS-MCs (ES- or iPS-pMCs) and the subsequent differentiation of these cells into DC-like cells (pMC-DC) are shown in Fig. 1A. ESCs or iPSCs (day 0) were seeded on the OP9 feeder cell layers. After 6 to 7 days, the mesodermally differentiated cells (day 6–7) were transferred onto newly prepared OP9 cell layers in the presence of GM-CSF. On days 13 to 14, the differentiated cells (myeloid cells) were transduced with c-Myc and cultured in the presence of M-CSF and GM-CSF. After about 5 to 6 days, pmCs with protrusions appeared and propagated continuously (Supplementary Fig. S1A). The pMCs were treated with GM-CSF and IL4 (days 23–24). After 3 days of culture, the pMCs differentiated into DC-like cells (pMC-DC; days 26–27). The fold-increase in the number of differentiated cells from PSCs at each stage and the phase contrast micrographs of the cells at each differentiation stage are shown in Fig. 1B and C, respectively. The number of pMC-DCs generated via the c-Myc transduction of myeloid cells ranged from 8,000 to 40,000 times that obtained from undifferentiated stem cells. This total was estimated to be at least 160 times more than that of the MC-DCs (without c-Myc transduction).

The myeloid cells (days 13–14) were a heterogeneous population, almost 90% of which was characterized to be CD11b⁺ (Fig. 2A). In this stage, CD11b⁺Gr-1^high, CD11c⁺Gr-1^low, and DEC205^lowGr-1^+/− cells were present as minor populations. The pMCs (days 23–24) obtained by transduction of the c-Myc gene showed the upregulation of both CD11c and DEC205, and decreases in the Gr-1^high population (Fig. 2B). The pMC-DCs (days 26–27) obtained after 3 days of culture in the presence of GM-CSF and IL4 showed an enhanced expression of CD11c and a downregulation of Gr-1 (Fig. 2B). In contrast to the BM-DCs, which were composed of a heterogeneous population (Fig. 2C), the pMC-DCs (GM-CSF/IL4-treated pMCs) were largely composed of a relatively homogeneous population, thus indicating that they were CD11b⁺/CD11c⁺/DEC205⁺/Gr-1^low DC-like cells (Supplementary Fig. S1B).

In the maintenance culture medium containing M-CSF and GM-CSF, mouse ES- and iPS-pMCs proliferated for more than 3 months. The doubling times of the ES-pMCs and iPS-pMCs after 15 days of c-Myc transduction were 16.73 and 17.60 hours, respectively (Fig. 3A). The doubling times of the iPS-pMCs cultured for 1, 2, and 3 months were 18.74, 17.71, and 20.06 hours, respectively (Fig. 3B). In addition, there were no significant changes in the phenotypes of these cultured pMCs (Supplementary Fig. S1C).

To increase our understanding of the proliferation of these cells, we examined their proliferation in the presence or absence of each cytokine or with combinations of cytokines (Fig. 3C). The results showed that proliferation was dependent on GM-CSF, but not M-CSF. The additional presence of M-CSF had no effect on the proliferation of these cells. The cells proliferated in a GM-CSF concentration–dependent manner, and the maximal proliferation was induced by 10 ng/mL GM-CSF (Fig. 3D).

In the maintenance culture medium containing M-CSF and GM-CSF, mouse ES- and iPS-pMCs proliferated and expressed major histocompatibility complex (MHC)-I, MHC-II, CD80, and CD86 (Fig. 4A). However, CD40 was expressed at a low level. Three days after treatment with IL4 plus GM-CSF, the proliferation of pMCs was impaired (Supplementary Fig. S2). In the additional presence of lipopolysaccharide (LPS), which is known to be an inducer of DC maturation, the pMCs showed enhanced expression of MHC-II, CD40, and CD86, and their proliferation was further impaired (Fig. 4B and Supplementary Fig. S2). The iPSC (2A-4F-100)–derived pMCs showed a tendency to have a substantially lower expression of MHC-II. This tendency was also observed when the cells were stimulated with LPS.

To determine whether the lower expression of MHC-II is a property common to all iPSC-derived pMC lines, we examined the expression of surface antigens in other pMCs derived from a different iPSC clone (2A-4F-136). The levels of MHC-II expression of 2A-4F-136–derived iPS-pMCs were generally low, but were higher than those from 2A-4F-100–derived iPS-pMCs (Fig. 4A). Therefore, low expression of MHC-II may be a common property, but levels of expression may differ depending on the parental iPSC clone. LPS or OK432 (penicillin-killed Streptococcus pyogenes) stimulation induced the production of TNFα and IL12p70 in the IL4/GM-CSF–treated pMCs, similar to that observed in BM-DCs (Fig. 4C).

To determine the capacity of the cells to activate naïve T cells, primary MLR assays were performed. LPS-treated pMC-DCs stimulated proliferative responses in allogeneic T cells, but the responses were lower than those induced by LPS-treated BM-DCs (Fig. 4D). These findings collectively indicate that pMCs treated with IL4 plus GM-CSF exhibited some features of DCs.

To determine whether pMC-DCs could prime antigen-specific T-cell responses in vivo, OVA_257–264 peptide- or vehicle-loaded pMC-DCs were i.p. transferred into syngeneic C57BL/6 mice twice at 7-day intervals. Spleen cells were isolated 7 days after the second transfer and were cultured in vitro in the presence of the OVA_257–264 peptide. After 5 days, the cells were recovered and assayed for their capacity to kill OVA_257–264 peptide-loaded EL-4 thymoma cells or OVA-expressing melanoma cells (MO4; Fig. 5). The CTLs primed in vivo by OVA_257–264 peptide-loaded pMC-DCs or OVA_257–264 peptide-loaded BM-DCs killed OVA peptide-loaded EL-4 cells, but not vehicle-loaded EL-4 cells (Fig. 5A–C). In contrast, the CTLs primed by vehicle-loaded pMC-DCs killed neither the OVA peptide-loaded nor the vehicle-loaded EL-4 cells. No differences in the capacity for antigen-specific CTL priming were observed among the DCs derived from pMCs cultured for 1, 2, or 3 months (Fig. 5D). In addition, the CTLs primed in vivo by OVA_257–264 peptide-loaded pMC-DCs killed OVA-expressing melanoma, but the CTLs primed by vehicle-loaded pMC-DCs did not (Fig. 5E). Moreover, the CTLs primed in vivo by OVA_257–264 peptide-loaded, irradiated pMC-DCs killed OVA-expressing melanoma. In contrast, OVA_257–264 peptide-loaded, heat-killed pMC-DCs did not result in the priming of OVA-specific CTLs (Fig. 5E). It is conceivable that heat-killed pMC-DCs may fail to induce immunity due to their loss of an adjuvant effect, whereas irradiated pMC-DCs survived for a certain period in vivo and were able to prime antigen-specific CTLs. These results suggest that peptide antigen-loaded pMC-DCs could prime the antigen-specific CTL response in vivo, and their function was maintained for at least 3 months.

To determine whether CTLs primed by pMC-DCs adoptively transferred into syngeneic C57BL/6 mice could protect against a subsequent challenge with OVA-expressing tumor cells, OVA_257–264 peptide- or vehicle-loaded pMC-DCs were i.p. transferred into mice twice at 7-day intervals. Seven days after the second transfer, the mice were inoculated s.c. with MO4 (Fig. 6 and Supplementary Fig. S3). Treatment with OVA_257–264 peptide-loaded ES- or iPS-pMC-DCs inhibited the growth of the inoculated MO4 for 1 month (Fig. 6A–C and Supplementary Fig. S3A–S3C) and significantly prolonged the survival of the mice compared with the vehicle-loaded pMC-DC treatment (Fig. 6D and Supplementary Fig. S3D). These observations indicate that peptide antigen–loaded pMC-DCs induced antigen-specific protective immunity against melanoma in vivo.

We next explored whether the mice were adversely affected by pMC-DC i.p. treatment. Two groups of mice treated with 2.0 × 10⁵ or 1.0 × 10⁷ iPS-pMC-DCs were monitored. No tumor formation or immune-related toxicities were observed for 3 months, even at a dose that was 100-fold higher than the doses used in the immunization protocols (Supplementary Fig. S4). Collectively, these observations indicate that peptide antigen-loaded pMC-DCs exhibited antigen-specific antitumor reactivity and were safe in vivo.

Mouse and human ESCs or iPSCs are capable of differentiating into functional DCs, thus providing an unlimited novel source of DCs for cancer immunotherapy (9, 17, 19). However, the process of inducing differentiation was relatively complicated, and generating practical volumes of DCs from ESCs or iPSCs has required a large number of PSCs at the start of the differentiation culture. Therefore, this process has not been suitable for the rapid mass production of cell preparations. To address this issue, we recently established a method for generating human pMCs (the precursors of DC-like cells) by transduction of c-Myc, together with antisenescence factors, such as Bmi-1 and Ezh2, into human iPSC-derived myeloid cells (26). To evaluate the possible application of these cells and their safety, we required an in vivo model not requiring xenotransplantation, which we developed for this study. Using this model, we demonstrated that pMCs could be generated from mouse ESCs or iPSCs, and that the cells could differentiate into functional APCs to induce antitumor immune responses in vivo without inducing autoimmunity or leukemia.

Mouse ESC-derived myeloid cells could acquire the capacity to proliferate in a cytokine-dependent manner after lentivirus-mediated transduction of c-Myc alone. Cell proliferation could also be induced in mouse iPSC-derived myeloid cells using the same procedure (Figs. 1 and 3). Because both the ESC- and iPSC-derived pMCs were of C57BL/6 origin, it is possible that these observations were unique to this strain. However, we made similar observations for the 129/Sv strain (data not shown). On the basis of these results, it appears that mouse PSC-derived myeloid cells acquire the capacity to proliferate continuously through the increased expression of c-Myc alone, irrespective of the mouse strain.

The c-Myc protein is a transcription factor that induces cell proliferation and cell senescence (33). Other than c-Myc, antisenescence genes, such as Bmi-1, Mdm2, and Ezh2, may be indispensable for the continuous proliferation of human ESC- or iPSC-derived myeloid cells (34–36). However, antisenescence factors were not required for acquisition of the capacity for long-term proliferation in the mouse PSC-derived myeloid cells. In addition, the increased expression of Bmi-1 had no effect on the cell proliferation, morphology, or surface markers in mouse ES- or iPS-pMCs (data not shown). This may be explained by differences between species, but the precise mechanism remains unclear.

When the medium was supplemented with GM-CSF with or without M-CSF, the mouse ES- and iPS-pMCs proliferated indefinitely. In the additional presence of IL4, the proliferation was impaired, and the cells functioned as APCs. LPS exposure induced the production of TNFα and IL12p70, and further enhanced the expression of antigen-presenting molecules and costimulatory molecules (Fig. 4). These findings were similar to the results of DC maturation. However, the T-cell stimulatory activity of ES- or iPS-pMC-DCs was slightly inferior to that of BM-DCs, based on their lower expression of T cell–stimulating molecules and the decreased production of cytokines (Fig. 4). In particular, iPSC-derived pMCs showed a tendency to have substantially lower expression of MHC-II than BM-DCs or ESC-derived pMCs (Fig. 4A). At present, the mechanisms by which iPSC-derived pMCs have downregulated MHC-II expression are not clear. The expression levels of MHC-II in iPS-pMC-DCs varied depending on the iPSC lines used as the source, and the epigenetic memory of iPSCs might affect the expression of MHC-II (37). In any case, despite the relatively lower levels of MHC-II expression, the iPS-pMC-DCs could prime the antigen-specific T cells in vivo (Fig. 5).

We successfully stimulated OVA_257–264 peptide-specific CTLs in vivo by administration of OVA_257–264 peptide-prepulsed ES- or iPS-pMC-DCs, leading to protection against OVA-expressing melanoma (Fig. 6 and Supplementary Fig. S3). DCs transduced with the entire tumor antigen gene expressed diverse T-cell epitopes via different MHC molecules and efficiently stimulated diverse antigen-specific T cells (38). Therefore, vaccination with pMC-DCs expressing the entire tumor antigen may be more effective for inducing antigen-specific CTLs, which could elicit potent antitumor responses. Moreover, pMC-DCs expressing multiple tumor antigens may exert a higher therapeutic effect (39–41). In addition to manipulation of the tumor antigens, modification of pMC-DCs through the transduction of immunostimulatory molecules (such as IL21) may improve the cellular immune responses (25, 42). To this end, genetic modification of pMC-DCs can be performed by introducing exogenous genes into pMCs via lentivirus transduction, followed by subsequent induction of pMC-DCs.

The human iPS-pMCs proliferated in an M-CSF–dependent manner (26), whereas the pMCs established from mouse ESCs or iPSCs in this study proliferated in a GM-CSF–dependent manner (Fig. 3C and D). There is a good concordance between the human and mouse pMCs in terms of their cytokine-dependent proliferation. In either case, the long-term proliferating capacity of pMCs may lead to the development of leukemia after administration to a patient. However, proliferation was induced by cytokine concentrations higher (>10 ng/mL) than the physiologic levels. The observation that the administration of a large amount of pMC-DCs (1.0 × 10⁷) did not induce the onset of leukemia in this study is suggestive of their in vivo safety (Supplementary Fig. S4). Although pMCs may not proliferate in the absence of GM-CSF, we can further enhance their safety by irradiating such pMCs before they are administered as therapy.

Despite the observed in vivo safety of the treatment, there are several challenges to the possible clinical application of this technique in humans. The lentivirus-mediated delivery of the c-Myc gene permanently integrates the transgene into the genome, potentially altering the genomic features. In addition, the oncogenic c-Myc gene can lead to the development of certain cancers. In this regard, a genome-integrating but excisable system (43), and transient expression of c-Myc with Sendai-virus (44), adenoviral vectors (45), or nonviral expression [such as transduction of the c-Myc protein (46) and the use of modified mRNA (47)] may provide safer pMCs in the future. In addition, using other genes that promote the proliferation of pMCs may secure their safety.

Construction of an extensive bank of iPSCs, including the most common HLA haplotypes, is currently in progress. With this resource it will be possible to generate pMCs from the cells in the iPSC bank. When the iPS-pMC bank is established, functional APCs can be supplied within a short period, which will permit broader clinical applications of these cells. Considering the use of pMC-DCs in an allogeneic or semi-allogeneic setting as a cancer vaccine, one potential hurdle is that the alloreactive CTLs may eliminate pMC-DCs, thereby compromising their efficacy. To avoid such issues, the genes associated with the cell-surface expression of HLA-I molecules, such as TAP or β2-microglobulin, can be disrupted. Alternatively, recipient-matched HLA-I heavy chain or the β2-microglobulin–linked form of the HLA-I heavy chain can be transduced into the TAP- or β2-microglobulin gene–deficient iPS-pMCs to induce suitable CTL responses (18).

iPSC-derived DCs may resolve the problems associated with the number and quality of DCs derived from patients with cancer. However, considering the cost, labor, and preparation time, iPSC-derived DC vaccine therapy as personalized medicine is not yet suitable as a widely applicable form of cancer immunotherapy. In contrast, production of iPS-pMCs is easily scalable to rapidly provide an unlimited number of functional APCs, which would benefit patients requiring multiple vaccine doses. The iPSC-derived proliferating APCs with the capacity to stimulate tumor antigen–specific T-cell responses and with good in vivo safety profiles may provide a novel vaccine strategy for clinical cancer therapy, although further investigation is required to improve their effectiveness and to further confirm their safety.

No potential conflicts of interest were disclosed.

Conception and design: N. Hirosawa, Y. Sakamoto, Y. Uemura

Development of methodology: R. Zhang, T.-Y. Liu, S. Senju, M. Haruta, M. Suzuki

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): R. Zhang, T.-Y. Liu, M. Tatsumi, N. Ueda, H. Maki, H. Nakanishi, Y. Akatsuka

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): R. Zhang, T.-Y. Liu, N. Hirosawa, Y. Sakamoto, K. Kuzushima, Y. Uemura

Writing, review, and/or revision of the manuscript: R. Zhang, T.-Y. Liu, S. Senju, N. Hirosawa, Y. Akatsuka, Y. Sakamoto, Y. Uemura

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): R. Nakatsuka, Y. Matsuoka, Y. Sasaki, S. Tsuzuki, R. Araki, M. Abe, Y. Sonoda, K. Kuzushima

Study supervision: N. Hirosawa, Y. Sakamoto, Y. Nishimura, Y. Uemura

pCSII-EF, pCMV-VSV-G-RSV-Rev, and pCAG-HIVgp constructs were kindly provided by Dr. H. Miyoshi (RIKEN BioResource Center).

This study was supported by grants from the Japan Science and Technology Agency (JST), a Kansai Medical University Internal grant C (9), the Osaka Cancer Research Foundation (9), the Takamatsu Cancer Research Foundation (9), the Aichi Cancer Research Foundation (12, 13), the Nagono Medical Foundation (12, 13), the Daiwa Securities Health Foundation (13), the Pancreas Research Foundation of Japan (13), and the Foundation for Promotion of Cancer Research in Japan (13). R. Zhang, M. Suzuki, and Y. Uemura were supported, in part, by Grants-in-Aid 25861253, 23791850, or 23592022, respectively, from the Ministry of Education, Culture, Sports, Science and Technology (MEXT) of Japan. T.-Y. Liu was supported by the National Natural Science Foundation of China (81101882). K. Kuzushima was supported by the Takeda Science Foundation (12–14).

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