HTLV-1 Tax-Specific CTL Epitope–Pulsed Dendritic Cell Therapy Reduces Proviral Load in Infected Rats with Immune Tolerance against Tax

Satomi Ando, Atsuhiko Hasegawa, Yuji Murakami, Na Zeng, Natsuko Takatsuka, Yasuhiro Maeda, Takao Masuda, Youko Suehiro and Mari Kannagi
J Immunol February 1, 2017, 198 (3) 1210-1219; DOI: https://doi.org/10.4049/jimmunol.1601557

Abstract

Adult T cell leukemia/lymphoma (ATL), a CD4+ T cell malignancy with a poor prognosis, is caused by human T cell leukemia virus type 1 (HTLV-1) infection. High proviral load (PVL) is a risk factor for the progression to ATL. We previously reported that some asymptomatic carriers had severely reduced functions of CTLs against HTLV-1 Tax, the major target Ag. Furthermore, the CTL responses tended to be inversely correlated with PVL, suggesting that weak HTLV-1–specific CTL responses may be involved in the elevation of PVL. Our previous animal studies indicated that oral HTLV-1 infection, the major route of infection, caused persistent infection with higher PVL in rats compared with other routes. In this study, we found that Tax-specific CD8+ T cells were present, but not functional, in orally infected rats as observed in some human asymptomatic carriers. Even in the infected rats with immune unresponsiveness against Tax, Tax-specific CTL epitope–pulsed dendritic cell (DC) therapy reduced the PVL and induced Tax-specific CD8+ T cells capable of proliferating and producing IFN-γ. Furthermore, we found that monocyte-derived DCs from most infected individuals still had the capacity to stimulate CMV-specific autologous CTLs in vitro, indicating that DC therapy may be applicable to most infected individuals. These data suggest that peptide-pulsed DC immunotherapy will be useful to induce functional HTLV-1–specific CTLs and decrease PVL in infected individuals with high PVL and impaired HTLV-1–specific CTL responses, thereby reducing the risk of the development of ATL.

Introduction

Adult T cell leukemia/lymphoma (ATL) is a highly aggressive CD4+ T cell malignancy with an extremely poor prognosis, which is caused by human T cell leukemia virus type 1 (HTLV-1) (1–3). Approximately 10 million individuals have been infected with HTLV-1 worldwide, especially in southern Japan, the Caribbean basin, South America, Melanesia, and equatorial Africa. Approximately 5% of HTLV-1 seropositive individuals develop ATL, although most HTLV-1–infected individuals remain asymptomatic throughout their lives. High proviral load (PVL), advanced age, and family history of ATL have been suggested to be risk factors for ATL development (4).

HTLV-1 is mainly transmitted via breast milk from an infected mother to her child (5, 6). Indeed, almost all ATL patients appear to have acquired HTLV-1 infection early in life (7). Our previous studies using an animal model demonstrated that rats orally infected with HTLV-1 developed persistent infection with high PVL, a risk factor for ATL development (8, 9). This suggests that oral/mucosal HTLV-1 infection may be involved in the development of ATL.

ATL, especially the acute or lymphoma types of ATL, is characterized by rapid progression of disease. Intensive multidrug chemotherapy, zidovudine/IFN-α, or mogamulizumab (Poteligeo; Kyowa Hakko Kirin, Tokyo, Japan) is now available as the first line of treatment for ATL (10–12). Additionally, ATL frequently relapses, which is probably one reason for the poor prognosis of this disease. Allogeneic hematopoietic stem cell transplantation is widely used as a second line treatment for ATL and achieves long-term remission in 30–40% of ATL patients (13, 14). However, this treatment has been reported to induce treatment-related mortality in a similar percentage of ATL patients (13, 15–17), suggesting the requirement for novel second line treatments for ATL. Additionally, given the complexity of the ATL treatment, it is also important to prevent asymptomatic carriers (ACs) from progressing to ATL. However, no prophylactic vaccines have been established.

HTLV-1–specific CTLs play an important role in controlling the expansion of HTLV-1–infected cells (18–20). Tax is the major target Ag for HTLV-1–specific CTLs (18). However, we and other groups previously reported severely impaired Tax-specific CTL responses in some ACs and most ATL patients (21–24), although the precise mechanism has not yet been elucidated. We also observed that the impaired functions of Tax-specific CD8+ T cells were likely to be correlated with elevated PVL in ACs (21). Additionally, our animal study showed that blockade of T cell responses with anti-CD80/86 Abs led to vigorous growth of HTLV-1–infected tumor cells in immunocompetent rats (25). These observations suggest that a weak HTLV-1–specific CTL response allows infected cells to escape from immune surveillance, survive, and expand, which leads to elevation of the PVL.

Dendritic cells (DCs) are professional APCs that induce primary immune responses and potentiate the effector functions of previously primed memory T cells (26). However, decreased numbers of both myeloid and plasmacytoid DCs, as well as reduced function of plasmacytoid DCs, have been observed in ATL patients compared with uninfected individuals (27), which may be one of the reasons why HTLV-1–specific CTL responses are severely reduced in most ATL patients.

DCs have been used in clinical trials as cellular mediators for therapeutic vaccines in cancer patients. A number of clinical trials have demonstrated that DC-based cancer vaccines are safe with little or no toxicity (28). Several clinical trials have shown that vaccines using monocyte-derived DCs (MoDCs) are effective for hepatitis B virus–infected individuals and patients with cancer, including melanoma, breast cancer, kidney cancer, and glioblastoma (29–33). This evidence strongly encourages us to expect that a DC-based vaccine has the potential to induce functional HTLV-1–specific CTLs capable of reducing ATL/infected cells in HTLV-1–infected individuals with decreased HTLV-1–specific CTL responses. However, it has been reported that monocytes from HTLV-1–infected individuals cannot fully mature into DCs (34, 35). Although we recently reported that all three ATL patients who received a Tax-specific CTL epitope–pulsed DC vaccine showed favorable clinical outcomes without severe adverse events (36), it remains undetermined whether MoDCs generated from HTLV-1–infected individuals have the capacity to stimulate Ag-specific CD8+ T cells. The direct effects of this vaccine on controlling the expansion of ATL and/or infected cells are also unclear, as these patients had received multiple other treatments before the vaccine inoculation.

In the present study, we show severely impaired functions of dominant Tax-specific CD8+ T cells in orally HTLV-1–infected rats, as seen in some human ACs, and demonstrate that s.c. immunization of CTL epitope–pulsed bone marrow–derived DCs (BMDCs) reduced PVL via induction of functional Tax-specific CD8+ T cells in these infected rats. We also found that MoDCs from most HTLV-1–infected individuals, including ATL patients and ACs, still had the capacity to stimulate Ag-specific CD8+ T cells. Our results suggest that a peptide-pulsed DC vaccine would be an effective strategy to strengthen Tax-specific CTL responses and thus reduce PVL in HTLV-1–infected individuals.

Materials and Methods

Animals

Three- to six-week-old female rats (F344/N Jcl-rnu/+) were purchased from Japan Clea (Tokyo, Japan) and maintained at the Center for Experimental Animals in the Tokyo Medical and Dental University. All animal experiments were approved by the Institutional Animal Care and Use Committee of the Tokyo Medical and Dental University.

Subjects

Blood samples from 4 healthy donors and 10 HTLV-1–infected individuals were used in this study (Table I). All blood samples were collected after obtaining written informed consent, and this study was reviewed and approved by the Institutional Review Board of the Tokyo Medical and Dental University.

Cell lines

An HTLV-1–infected human T cell line, MT-2, was maintained in RPMI 1640 (Life Technologies, Carlsbad, CA) supplemented with 10% heat-inactivated FCS (Sigma-Aldrich, St. Louis, MO), 100 IU/ml penicillin (Life Technologies), and 100 μg/ml streptomycin (Life Technologies). Aliquots of the cells were cryopreserved as a master stock immediately after characterization and authentication by phenotypic analysis including HLA typing. The cells were cultured for 2–3 wk before using for HTLV-1 infection of rats. FPM1-V1AX is an HTLV-1–immortalized rat T cell line, which was previously established in our laboratory (37). The cells were maintained in RPMI 1640 containing 10% FCS and 2-ME (10−5 M; Life Technologies). G14-Tax is an HTLV-1 Tax-expressing transfectant of an HTLV-1–negative rat T cell line, G14. Both G14 and G14-Tax were established as described in our previous study (38). G14-Tax was maintained in RPMI 1640 with 10% FCS, 10−5 M 2-ME, and 10 U/ml recombinant human (rh)IL-2 (Shionogi Pharmaceutical, Osaka, Japan). FPM1-V1AX and G14-Tax were used to stimulate Tax-specific rat CD8+ T cells.

Peptides

Peptides used in this study were the RT1.Al-restricted CTL epitope (39) Tax180–188 (LLFGYPVYV; Hokudo, Sapporo, Japan), the HLA-A2–restricted CTL epitope (40) CMV495–503 (NLVPMVATV; Sigma-Aldrich), and the HLA-A24–restricted CTL epitope (41) CMV341–349 (QYDPVAALF; Sigma-Aldrich).

HTLV-1 infection

Rats were infected with HTLV-1 by inoculation of HTLV-1–producing MT-2 cells. For oral HTLV-1 infection, MT-2 cells (2 × 107 cells) were directly administered into the esophagus of 6- or 7-wk-old rats through a feeder tube. For i.p. infection, MT-2 cells (2 × 107 cells) were percutaneously injected into the abdominal cavity. To generate BMDCs, some infected rats were euthanized 4 wk postinoculation with MT-2 cells. For comparison of Tax-specific CD8+ T cell responses between orally and i.p. HTLV-1–infected rats (i.p. rats), the rats were euthanized 4 wk postinfection. In vaccine studies, the infected rats were used 4 wk after HTLV-1 infection.

Preparation of rat BMDCs

Four-week-old rats were orally infected with or without HTLV-1 and sacrificed 4 wk later to obtain bone marrow cells. BMDCs were generated from bone marrow cells of the rats as previously described with slight modifications (42). In brief, bone marrow cells (1–2 × 107 cells) obtained from femurs and tibias were cultured in the presence of recombinant rat (rr)GM-CSF (10 ng/ml; PeproTech, Rocky Hill, NJ) and rrIL-4 (5 ng/ml; PeproTech) in a 100-mm cell culture dish. On day 6, nonadherent cells were gently removed and adherent cells were cultured for an additional 5 d in the presence of 10 ng/ml rrGM-CSF and 5 ng/ml rrIL-4. For maturation of BMDCs, BMDCs were stimulated with LPS (1 μg/ml; Sigma-Aldrich) for the last 24 h.

Immunization with Tax180–188 peptide–pulsed BMDCs

The mature BMDCs were pulsed with 10 μM RT1.A-restricted CTL epitope, Tax180–188, for 1 h at room temperature, washed, and resuspended in PBS. The cells (1 × 106 cells) were s.c. inoculated once a week for 3 wk into rats that had been orally infected 4 wk before with or without HTLV-1. At 3 wk after the last immunization, the rats were sacrificed to analyze Tax-specific CD8+ T cell responses and PVL.

Flow cytometry

For phenotypic analyses of rat BMDCs and human MoDCs, the following fluorochrome-conjugated anti-rat and anti-human mAbs were used: rat CD80-PE (3H5; BioLegend, San Diego, CA), rat CD86-PE (24F; BioLegend), rat RT1.A-PE (OX-18; BioLegend), rat RT1.B-PE (OX-6; BioLegend), human CD40-FITC (5C3; BD Biosciences, San Jose, CA), human CD80-FITC (2D10; BD Biosciences), human CD83-PE (HB15a; BD Biosciences), human CD86-FITC (FUN-1; BD Biosciences), human HLA-ABC–PE (G46-2.6; BD Biosciences), and human HLA-DR–FITC (L243, BD Biosciences). To detect Tax-specific rat CD8+ T cells, PE-conjugated RT1.A/Tax180–188 tetramer was generated through the custom service of Medical and Biological Laboratories (Nagoya, Japan). Splenic T cells (2 × 105 cells) were stained with RT1.A/Tax180–188 tetramer–PE, rat CD3-FITC (1F4; BD Biosciences), and rat CD8-PerCP (OX-8; BioLegend). CMVpp65-specific human CD8+ T cells were detected with PE-conjugated HLA-A*02:01/CMVpp65:395–403 or HLA-A*24:02/CMVpp65:341–349 tetramers (Medical and Biological Laboratories). Peripheral blood or cultured cells were stained with HLA-I/CMVpp65 tetramer–PE in conjunction with human CD3-FITC (UCHT1; BioLegend) and human CD8-PerCP (RPA-T8; BioLegend). For whole-blood samples, RBCs were lysed and leukocytes were fixed in BD FACS lysing solution (BD Biosciences) before washing. Samples were analyzed on a FACSCalibur (BD Biosciences) and data analyses were performed using FlowJo software (Tree Star, Ashland, OR).

Proliferation of Tax-specific rat CD8+ T cells

Rat splenic T cells were enriched with a nylon-wool column and used as responder cells. To prepare stimulator cells, FPM1-V1AX cells (5 × 105 cells/ml) were treated with mitomycin C (MMC; 50 μg/ml; Kyowa Hakko Kirin) at 37°C for 30 min. The splenic T cells (1 × 106 cells/ml) were cocultured with MMC-treated FPM1-V1AX (5 × 105 cells/ml) for 7 d in the presence of 10 U/ml rhIL-2. The cells were then stained with RT1.A/Tax180–188 tetramer–PE, rat CD3-FITC, and rat CD8-PerCP.

IFN-γ production in Tax-specific rat CD8+ T cells

The splenic T cells were cocultured with MMC-treated FPM1-V1AX or G14-Tax for 7–10 d in the presence of 10 U/ml rhIL-2, harvested, and stimulated for 6 h with PMA (50 ng/ml; Sigma-Aldrich) and ionomycin (1 μg/ml; Sigma-Aldrich) in the presence of brefeldin A (1 μg/ml; Sigma-Aldrich). The cells were then stained with rat IFN-γ–FITC (DB-1; BioLegend), RT1.A/Tax180–188 tetramer–PE, and rat CD8-PerCP using the BD Cytofix/Cytoperm kit (BD Biosciences).

Rat IL-12p40 and IL-10 ELISA

Immature BMDCs (2.5 × 105 cells/ml) were cultured in the presence or absence of 1 μg/ml LPS for 2 d and the supernatant was then collected. IL-12 and IL-10 concentrations in the supernatant were measured with rat IL-12 and rat IL-10 ELISA kits (Life Technologies), respectively.

Generation of human MoDCs

PBMCs were isolated from heparinized peripheral blood of healthy volunteers and HTLV-1–infected individuals by the density gradient centrifugation with Ficoll-Paque Plus (GE Healthcare UK, Buckinghamshire, U.K.). Monocytes were then enriched from PBMCs by the plastic adherence method and cultured for 5 d in RPMI 1640 supplemented with 10% FCS in the presence of 1000 U/ml rhGM-CSF (R&D Systems, Minneapolis, MN), 500 U/ml rhIL-4 (R&D Systems), and 1 μM zidovudine (Retrovir; GlaxoSmithKline, Middlesex, U.K.). For maturation of MoDCs, the cells were stimulated for 2 d with 10 ng/ml rhTNF-α (PeproTech), 0.05 clinical units/ml OK432 (Picibanil; Chugai Pharmaceutical, Tokyo, Japan), and 25 μg/ml keyhole limpet hemocyanin (KLH; Calbiochem, San Diego, CA).

Human IL-12p70 ELISA

Immature MoDCs (2.5 × 105 cells/ml) were stimulated for 48 h in the presence or absence of 10 ng/ml TNF-α, 0.05 clinical units/ml OK432, and 25 μg/ml KLH. IL-12p70 in the supernatant was then measured with the BD OptEIA human IL-12p70 ELISA set (BD Biosciences).

Allogeneic MLR

Allogeneic CD4+ T cells were isolated from HTLV-1 seronegative healthy donors, labeled with CFSE (Sigma-Aldrich), and cocultured with mature MoDCs for 4 d at responder/stimulator ratios of 100 and 1000. The cells were then stained with human CD4-PE/Cy5 and analyzed by flow cytometry.

Autologous MLR

Mature MoDCs were pulsed with or without 1 μM CMV-specific CTL epitope for 1 h at room temperature and cocultured with autologous PBLs in the presence of 10 U/ml rhIL-2 for 13 d at a responder/stimulator ratio of 10. The cells were stained with HLA-I/CMVpp65 tetramer–PE, human CD3-FITC, and human CD8-PerCP.

Proliferation assay

PBMCs from HTLV-1–infected individuals were stimulated with 100 nM CMV-specific CTL epitope for 13 d in the presence of 10 U/ml rhIL2. The cells were stained with HLA-I/CMVpp65 tetramer–PE, human CD3-FITC, and human CD8-PerCP.

Quantification of HTLV-1 PVL by LightCycler-based real-time PCR

The HTLV-1 PVL in splenic T cells of HTLV-1–infected rats was quantified using the QuantiTect SYBR Green PCR kit (Qagen, Courtaboeuf, France) with a LightCycler (Roche Diagnostics, Mannheim, Germany) as previously described (8). The data were normalized to the copy number of β-actin in each sample. The primers used for quantitative PCR were as follows: HTLV-1 pX2, 5′-CGG ATA CCC AGT CTA CGT GTT TGG AGA CTG T-3′ (sense); HTLV-1 pX3, 5′-GAG CCG ATA ACG CGT CCA TCG ATG GGG TCC-3′ (antisense); β-actin forward primer, 5′-CCT GTA TGC CTC TGG TCG TA-3′; and β-actin reverse primer, 5′-CCA TCT CTT GCT CGA AGT CT-3′.

Statistical analysis

The Mann–Whitney U test and the unpaired t test were performed to determine whether differences between groups were statistically significant using GraphPad Prism software (GraphPad Software, La Jolla, CA). In all cases, two-tailed p values of <0.05 were considered statistically significant.

Results

Orally HTLV-1–infected rats showed limited Tax-specific CD8+ T cell responses

We previously reported that orally (per os) HTLV-1–infected rats (p.o. rats) carried higher PVL than did i.p. rats (8). We also identified an MHC class I (RT1.Al)–restricted epitope, Tax180–188, which was predominantly recognized by HTLV-1–specific CD8+ CTLs in rats immunized with HTLV-1–infected syngeneic cells (39). First, we newly generated the RT1.A/Tax180–188 tetramer and examined the proliferative capacity of Tax-specific splenic CD8+ T cells in p.o. and i.p. rats. In representative rats, tetramer+ cells were detected at day 0 but were only present at low frequencies (0.0301 and 0.0148% in a p.o. and an i.p. rat, respectively, Fig. 1A). Interestingly, at day 7 after stimulation of splenic T cells with HTLV-1–infected syngeneic FPM1-V1AX cells, Tax-specific CD8+ T cells in the p.o. rat only slightly proliferated to 0.0922% of total CD8+ T cells whereas tetramer+ cells in the i.p. rat vigorously expanded to 7.84% (Fig. 1A). As summarized in Fig. 1B, Tax-specific CD8+ T cells in p.o. rats had a limited proliferative capacity when compared with those in i.p. rats. The average frequency of tetramer+ cells at day 7 was significantly lower in p.o. rats than that in i.p. rats (p = 0.0003, Fig. 1B, right panel). Furthermore, in p.o. rats, tetramer+CD8+ T cells that had been detected at day 7 poststimulation with a Tax Ag became undetectable after restimulation with PMA and ionomycin, whereas a high percentage of Tax-specific CD8+ T cells produced IFN-γ in i.p. rats (Fig. 1C). These data suggest severely reduced functions of Tax-specific CD8+ T cells in p.o. rats, as observed in some ACs and most ATL patients (21). This impaired Tax-specific CD8+ T cell response could be one of the reasons why p.o. rats carry a significantly higher PVL than do i.p. rats. As a higher PVL is associated with an elevated risk for the development of ATL, hereafter we used p.o. rats for the vaccine study.

FIGURE 1.
FIGURE 1.

Reduced function of Tax180–188-specific CD8+ T cells in p.o. rats. Six- to seven-week-old rats were orally (p.o., top panel) or i.p. (bottom panel) infected with HTLV-1. Four weeks later, splenic T cells were isolated from both infected rats and stimulated with an MMC-treated HTLV-1–infected syngeneic T cell line, FPM1-V1AX, for 7 d. (A) Representative dot plots of RT1.A/Tax180–188 tetramer+ cells before (day 0, left) and after (day 7, right) stimulation. The number represents the percentage of tetramer+ cells of total CD8+ T cells. (B) Summary of data on frequency of tetramer+ cells in p.o. (n = 8, left panel) and i.p. (n = 7, middle panel) rats before and after stimulation. Each dot indicates the percentage of tetramer+ cells of total CD8+ T cells. In the right panel, the tetramer+ cell frequency shown in the left panel (day 7, p.o. rats) is compared with that shown in the middle panel (day 7, i.p. rats). The bars indicate the average percentage of tetramer+ cells. The statistical significance of differences was determined with the Mann–Whitney U test. (C) Splenic T cells from p.o. (top panel) or i.p. (bottom panel) rats were cocultured with MMC-treated G14-Tax, a Tax-expressing rat T cell line for 7 d, and stained with the RT1.A/Tax180–188 tetramer (left panel). The number indicates the percentage of tetramer+ cells of total CD8+ T cells. The cells were then stimulated with PMA and ionomycin (P/I) for 6 h in the presence of brefeldin A. The number indicates the percentage of IFN-γ–producing cells of tetramer+CD8+ cells (right panel). The data shown are representative of two animals per group. In the other six p.o. rats, IFN-γ production was not evaluated because the frequency of Tax-specific CD8+ T cells was very low.

BMDCs generated from orally infected rats have the capacity to stimulate Ag-specific CD8+ T cells in vivo

In human clinical studies, Ag-pulsed MoDCs are one of the vaccine candidates to elicit Ag-specific T cell immunity (28). However, in humans, it has been reported that MoDCs were functionally impaired in HTLV-1–infected individuals compared with healthy donors (34, 35). Therefore, we performed phenotypic and functional analyses of BMDCs generated from both uninfected and p.o. rats. As shown in Fig. 2A, there was no significant difference in the mean expression of costimulatory molecules (CD80 and CD86) and RT1.A between infected and uninfected rat–derived BMDCs after LPS stimulation. Although the histogram plot revealed a reduction of RT1.B (MHC class II) expression on the infected rat–derived BMDCs (Fig. 2A, top), the mean level of RT1.B expression on infected rat–derived BMDCs was comparable to that on uninfected rat–derived BMDCs (Fig. 2A, bottom)

FIGURE 2.
FIGURE 2.

Phenotypic analysis and cytokine production of BMDCs from both HTLV-1–uninfected and –infected rats. (A) Representative histogram plots (top) of the expression of indicated surface molecules on LPS-stimulated BMDCs from uninfected (dotted line) and p.o. (solid line) rats. Isotype control Abs were used as a negative control (filled histogram). Mean florescence intensity (MFI; bottom) of the indicated surface molecules on LPS-stimulated BMDCs derived from uninfected (Uninf; n = 4 for CD80 and CD86, n = 3 for RT1.A and RT1.B) and p.o. (Inf; n = 7 for CD80, CD86, and RT1.B, n = 4 for RT1.A) rats. The graphs shown represent the MFI of each molecule divided by the MFI of corresponding isotype control. Error bars indicate the SD. The statistical significance of differences was determined with the unpaired t test. (B) IL-12p40 (left) and IL-10 production (right) of BMDCs from uninfected (UN) and infected (PO) rats. BMDCs were stimulated with (ST) or without LPS for 48 h, and the culture supernatant was collected. IL-12p40 and IL-10 in the supernatant were then measured by ELISA. The data shown are representative of two animals per group. The statistical significance of differences was determined with the unpaired t test.

We also examined IL-12 and IL-10 production by BMDCs from p.o. rats. The cells produced a large amount of IL-12 after LPS stimulation, the level of which was comparable to that of uninfected rat–derived BMDCs (p = 0.4358, Fig. 2B, left). In contrast to IL-12 production, a higher IL-10 production was observed in infected rat–derived BMDCs than in uninfected rat–derived BMDCs, regardless of LPS stimulation (Fig. 2B, right).

We further examined the ability of BMDCs to stimulate Ag-specific naive CD8+ T cells in vivo. BMDCs from p.o. rats were pulsed with Tax180–188 peptide and s.c. injected into uninfected rats once a week for 3 wk. As expected, Tax-specific CD8+ T cells were induced by immunization with infected rat–derived BMDCs as well as with uninfected rat–derived cells, although their frequencies were quite low (day 0, Fig. 3A, 3B). The expansion of CD8+ T cells in immunized rats varied at 7 d poststimulation with FPM1-V1AX (Fig. 3A, 3B). There was no statistically significant difference in the average frequency of Tax-specific CD8+ T cells between rats immunized with infected and uninfected rat–derived BMDCs (p = 0.4857, Fig. 3B). Furthermore, cytokine flow cytometry revealed that the CD8+ T cells in rats produced similar levels of IFN-γ (p = 0.3429, Fig. 3C). Notably, immunization with infected rat–derived BMDCs that were not pulsed with the Tax peptide did not induce any Tax-specific CD8+ T cell responses in uninfected rats (Supplemental Fig. 1), suggesting that few, if any, BMDCs generated from infected rats are infected with HTLV-1. These data indicate that BMDCs derived from both infected and uninfected rats have a comparable capacity to induce functional Tax-specific CD8+ T cells capable of proliferating and producing IFN-γ in vivo.

FIGURE 3.
FIGURE 3.

Induction of Tax180–188-specific CD8+ T cells in uninfected rats immunized with Tax180–188-pulsed BMDCs. (A and B) Tax180–188 peptide (pTax)–pulsed BMDCs (1 × 106 cells) were generated from rats that had been orally infected 4 wk before with or without HTLV-1 and s.c. injected into other uninfected rats once a week for 3 wk. Splenic T cells were isolated from the immunized rats 3 wk after the last injection. Proliferative capacity of tetramer+ cells in the immunized rats was then assessed as in Fig. 1. (A) Representative dot plots of tetramer+ cells in splenic T cells of rats immunized without (No DC; bottom panel) or with pTax-pulsed BMDCs that were generated from uninfected (pTax/uninf DC; top panel) or infected (pTax/inf DC; middle panel) rats. The number represents the percentage of tetramer+ cells of total CD8+ T cells (B) Summary of data on frequency of tetramer+ cells in rats immunized with pTax/uninf DC (n = 4, left) or pTax/inf DC (n = 4, right). The statistical significance of differences was determined with the Mann–Whitney U test. (C) Representative dot plots (top panel) of IFN-γ production of Tax-specific CD8+ T cells from rats immunized with pTax/uninf DC (left panel) or pTax/inf DC (right panel). The number indicates the percentage of IFN-γ–producing cells of total tetramer+ cells. Summary of data on IFN-γ production of Tax-specific CD8+ T cells from both rats (n = 4 rats per group, bottom panel) are shown. The bars indicate the average percentage of tetramer+ cells. The statistical significance of differences was determined with the Mann–Whitney U test.

Orally infected rats vaccinated with Tax epitope–pulsed BMDCs showed functional Tax-specific CD8+ T cell responses and had a low PVL

In humans, high PVL was reported to be the risk factor for the progression to ATL (4). As shown in Fig. 1, the function of Tax-specific CD8+ T cells was severely impaired in p.o. rats. Therefore, to strengthen Tax-specific CD8+ T cell responses in p.o. rats, we s.c. vaccinated three times with Tax180–188 peptide–pulsed, infected rat–derived BMDCs. On day 0, the frequency of Tax-specific CD8+ T cells in fresh splenic CD8+ T cells was very low in vaccinated rats (Fig. 4A, 4B), which was comparable to that in unvaccinated rats (Fig. 1A, 1B). As the level of Tax-specific CD8+ T cell expansion and PVL are found to be stable in p.o. rats between 4 and 8 wk postinfection, we compared the level of Tax-specific CD8+ T cell expansion in vaccinated rats with that in p.o. rats shown in Fig. 1B. At day 7 after Ag stimulation, the average frequency of Tax-specific CD8+ T cells in vaccinated rats was significantly higher than that in unvaccinated rats (p = 0.0173, Fig. 4C). Notably, among the six vaccinated rats, half showed a strong expansion and vigorous IFN-γ production of Tax-specific T cells (Fig. 4C and 4D, respectively). The other half showed a weak Tax-specific CD8+ T cell expansion, the level of which was only slightly higher than that of the unvaccinated rats (Fig. 4C). However, in contrast to Tax-specific CD8+ T cells in p.o. rats (Fig. 1C), ∼10% of Tax-specific CD8+ T cells produced IFN-γ in these rats (Fig. 4D), suggesting that functional Tax-specific CD8+ T cells are induced in these vaccinated rats even though their frequency is low. As shown in Fig. 4E, the PVL in all vaccinated rats was significantly lower than that in the unvaccinated rats (p = 0.0173). Intriguingly, the vaccinated rats with a strong Tax-specific CD8+ T cell expansion had an undetectable PVL. The other vaccinated rats showing a weak Tax-specific CD8+ T cell response carried a detectable PVL, although it tended to be lower than that in unvaccinated rats (Fig. 4E). These results suggest that the PVL can be reduced because of functional Tax-specific CD8+ T cell responses restored by the Tax peptide–pulsed BMDC vaccine.

FIGURE 4.
FIGURE 4.

Orally infected rats vaccinated with Tax epitope–pulsed BMDCs showed functional Tax-specific CD8+ T cell responses and carried low PVL. Rats that had been infected 4 wk before with HTLV-1 were immunized with Tax180–188 peptide–pulsed BMDCs (1 × 106 cells) generated from other orally infected rats once a week for 3 wk. Three weeks after immunization, splenic T cells were isolated from the vaccinated rats. (A and B) The proliferative capacity of tetramer+ cells from the vaccinated rats was assessed as in Fig. 1. (A) Experimental scheme (top) and representative dot plots of tetramer+ cells in vaccinated rats at day 0 and day 7 after Ag stimulation (bottom). The number indicates the percentage of tetramer+ cells of total CD8+ T cells. (B) Summary of data on frequency of tetramer+ cells in the vaccinated rats (n = 6) before and after Ag stimulation. The number indicates the percentage of tetramer+ cells at day 7 after Ag stimulation. (C) The frequency of tetramer+ cells in unvaccinated (●; n = 5) and vaccinated (▴ and ▪) rats at day 7 after Ag stimulation. Half the vaccinated rats (▪) had a strong expansion of tetramer+ cells. The other half (▴) showed only slightly higher tetramer+ cell responses than did unvaccinated rats (●). Each dot indicates the percentage of tetramer+ cells of total CD8+ T cells. The bars indicate the average percentage of tetramer+ cells. (D) Representative dot plot (top panel) of IFN-γ production of tetramer+ cells in the vaccinated rats was examined as in Fig. 3C. The number indicates IFN-γ+ cells of tetramer+ cells. A summary of data on IFN-γ production of Tax-specific CD8+ T cells in the vaccinated rats with a strong (n = 3) or weak (n = 2) Tax-specific CD8+ T cell expansion (bottom panel) is shown. The bars indicate the average percentage of tetramer+ cells. In one of the vaccinated rats with a weak Tax-specific T cell expansion, IFN-γ production was not evaluated because of the limited availability of the sample. (E) PVL in p.o. rats with (▴ and ▪) or without (●) the Tax180–188 peptide–infected DC vaccination (top). PVL was further compared between vaccinated rats with a strong (▪) and a weak (▴) tetramer+ cell expansion (bottom). Each dot indicates the copy number of HTLV-1 proviruses per 105 copies of β-actin. The bars indicate the average of PVL. The statistical significance of differences between groups was determined with the Mann–Whitney U test. (C and E) Among the data shown in Fig. 1B, the results on Tax-specific CD8+ T cell expansion in five p.o. rats are used as an unvaccinated control (C). The PVL in the same p.o. rats are shown as an unvaccinated control (E).

MoDCs from HTLV-1–infected individuals had the capacity to stimulate CMV-specific autologous CD8+ T cell expansion

In the p.o. rat model, peptide-pulsed BMDCs induced functional Tax-specific CD8+ T cells, which led to a reduction of PVL. This implies that an elevated PVL in human HTLV-1 carriers, a risk factor for disease progression, might be reduced by a similar vaccination strategy. However, reduced function of MoDCs generated from HTLV-1–infected individuals has been reported (34, 35). We therefore generated MoDCs from HTLV-1–infected individuals (Table I) to examine whether the MoDCs had the ability to stimulate Ag-specific autologous CD8+ T cells. As HTLV-1–infected cells produce a detectable amount of viral protein, including Tax, during in vitro culture of PBMCs and function as APCs, it is difficult to precisely evaluate the stimulatory capacity of MoDCs to HTLV-1 Ag-specific memory CD8+ T cells. Additionally, in all HTLV-1–infected individuals tested, CMV-specific CD8+ T cells were detected in fresh PBMCs (Fig. 5) but did not expand when stimulated without or with autologous MoDCs not pulsed with a CMV-specific CTL epitope (Supplemental Fig. 2), indicating that all cases tested in this study were infected with CMV at the time of blood collection. We therefore used CMV-specific CTL epitope as an Ag in this experiment. In all HTLV-1–infected individuals tested, stimulation of MoDCs with TNF-α, KLH, and OK432 revealed that MoDCs generated from infected individuals as well as healthy donors expressed several surface molecules (CD40, CD80, CD86, HLA classes I and II, and CD83), produced IL-12, and stimulated allogeneic CD4+ T cell expansion (Supplemental Figs. 3, 4). In an AC (AC no. 1), 2.31% of CD8+ T cells in fresh blood were CMV-specific CD8+ T cells (Fig. 5A). When PBMCs from AC no. 1 were stimulated for 13 d with a CMV-specific CTL epitope peptide alone, CMV-specific CD8+ T cells proliferated to 7.52% of total CD8+ T cells. In contrast to AC no. 1, CMV-specific CD8+ T cells in two ATL patients (acute type of ATL [aATL] no. 1 and no. 2) could not expand at 13 d poststimulation with CMV epitope peptide alone (10.9 and 0.284% in aATL no. 1 and no. 2, respectively), although 11.5 and 5.09% of the cells were present in fresh blood of aATL no. 1 and no. 2, respectively. This is consistent with our previous finding that, in most ATL patients, not only HTLV-1–specific but also CMV-specific CD8+ T cells did not proliferate when only stimulated with CTL epitope peptide (21). Surprisingly, stimulation of PBLs with CMV epitope–pulsed MoDCs revealed vigorous expansion of CMV-specific CD8+ T cells in these three HTLV-1–infected individuals (50.9, 45.9, and 28.5% in AC no. 1, aATL no. 1, and aATL no. 2, respectively; Fig. 5A). We also observed that CMV-specific CD8+ T cells proliferated vigorously upon stimulation with CMV peptide–pulsed MoDCs in two other infected individuals (AC no. 2 and aATL no. 4, Fig. 5B). However, CMV-specific CD8+ T cells in aATL patient no. 3 did not proliferate even when stimulated with CMV peptide–pulsed MoDCs (Fig. 5B). These results suggest that MoDCs generated from most HTLV-1–infected individuals have the capacity to stimulate Ag-specific CD8+ T cells.

View this table:
Table I. Disease status and HLA-A alleles of blood samples tested in this study
FIGURE 5.
FIGURE 5.

MoDCs from HTLV-1–infected individuals have the capacity to stimulate CMV-specific autologous CD8+ T cell expansion. (A) CMV-specific CD8+ T cells in an asymptomatic HTLV-1 carrier (AC no. 1) and two aATL patients (aATL no. 1 and no. 2) carrying an HLA-A2 allele were stimulated in two different ways. 1) PBMCs were stimulated with HLA-A2–restricted CMV epitope (A2 CMV epitope) for 13 d (middle panel). 2) PBLs were cocultured for 13 d with A2 CMV epitope–pulsed autologous MoDCs (right panel). Frequency of CMV-specific CD8+ T cells was assessed at day 0 (left panel) and day 13 (middle and right panels). (B) PBLs in a healthy donor (Healthy no. 4, HLA-A24), an asymptomatic HTLV-1 carrier (AC no. 2, HLA-A2), and two aATL patients (aATL no. 3 and no. 4, both HLA-A2) were stimulated for 13 d with CMV epitope–pulsed autologous MoDCs. The frequency of CMV-specific CD8+ T cells was assessed before (day 0, left) and after (day 13, right) MoDC stimulation. The number indicates the percentage of tetramer+ cells of total CD8+ T cells.

Discussion

We previously demonstrated that, in some ACs, the function of Tax-specific CD8+ T cells was severely impaired, which appeared to be correlated with increased PVL, a risk factor for the progression to ATL (21). Vaccines that elicit HTLV-1–specific CD8+ T cells may have not only therapeutic but also prophylactic potentials to prevent ATL development by reducing PVL in infected individuals with low specific CD8+ T cell responses. However, such vaccines have not yet been established. DCs are the most effective APCs to induce and regulate immune responses (26). Additionally, as large-scale DC generation is feasible, ex vivo–educated DCs can be applied to bypass endogenous DC dysfunction in patients with tumors and to induce therapeutic antitumor immunity (28). In the present study, we demonstrated that s.c. immunization of BMDCs pulsed with a dominant CTL epitope, Tax180–188, induced epitope-specific CD8+ T cell responses in p.o. rats showing immune suppression against Tax. Additionally, the PVL in immunized rats was significantly lower than those in unimmunized rats. Furthermore, we also showed that MoDCs generated from most infected individuals had the potential to stimulate Ag-specific memory CD8+ T cells in vitro, although functional impairment of MoDCs from infected individuals has been reported (34, 35). These results suggest that a peptide-pulsed DC vaccine would be an effective strategy to reduce ATL and/or infected cells expressing HTLV-1 viral Ags via induction of specific CD8+ T cell responses in infected individuals.

In humans, it is thought that individuals infected with HTLV-1 during childhood through breastfeeding (oral/mucosal infection) may have a higher risk of developing ATL (7, 43). Our previous animal studies demonstrated that p.o. rats showed low HTLV-1–specific T cell responses and high PVL (8, 9). In this study, we directly analyzed Tax-specific CD8+ T cells using newly generated tetramer and found that Tax-specific CD8+ T cells were present in p.o. rats but they neither proliferated well nor produced IFN-γ (Fig. 1). This is also likely to be a feature of severely reduced Tax-specific CTL responses observed in most ATL patients and some ACs (21). Furthermore, the p.o. rats after the Tax peptide–pulsed BMDC immunization showed a lower proliferative capacity of the Tax-specific CTLs than did uninfected rats that had received the same BMDC immunization (Figs. 3B, 4B), indicating that some of the suppression mechanisms may underlie the impairment of virus-specific CTL responses in p.o. rats.

DCs have the capacity to prime naive and boost memory Ag-specific T cells (26). In this study, immunization with Tax peptide–pulsed BMDCs derived from both uninfected and p.o. rats into uninfected rats comparably mounted Tax-specific CD8+ T cell responses (Fig. 3), implying that BMDCs generated from both rats have a similar capacity to stimulate Ag-specific naive CD8+ T cells. Moreover, the Tax peptide–pulsed BMDCs stimulated the in vitro expansion of a Tax-specific CTL line that was previously established from a Tax-coding DNA-vaccinated rat (data not shown) (44), indicating that BMDCs can stimulate Ag-specific memory CD8+ T cell expansion as well. However, Tax-specific memory CD8+ T cells from p.o. rats failed to proliferate in vitro after stimulation with the Tax peptide–pulsed BMDCs (data not shown), suggesting that Tax-specific memory CD8+ T cells that existed in p.o. rats may be functionally impaired. In humans, there is accumulating evidence of HTLV-1–specific CTL suppression in ATL patients. First, HTLV-1–specific CTLs are dysfunctional. The CTLs express inhibitory molecules, such as programmed death-1 on the surface (45), and HTLV-1 p8 leads to downregulation of TCRs (46). Second, more HTLV-1–uninfected regulatory T cells are observed in HTLV-1–infected individuals (47). Third, ATL and/or HTLV-1–infected cells have a regulatory T cell–like suppressive function (48). Lastly, endogenous DCs, such as plasmacytoid and myeloid DCs, are found in decreased number and with reduced functions (27). In our rat model, it is also possible that oral HTLV-1 infection induces oral tolerance or T cell anergy (49). However, in this study, we did not investigate the mechanisms underlying the severely impaired HTLV-1–specific CD8+ T cell responses observed in p.o. rats. Thus, further studies are required to clarify the mechanisms.

CD4+ Th cells are important to induce and maintain Ag-specific CD8+ T cell responses (50). Ag presentation of only CTL epitope peptides is thought to induce suboptimal CTL responses because it does not recruit specific CD4+ T cell help, and because the half-life of binding between the CTL epitope peptide and MHC class I is relatively short. Indeed, in this study, half the rats immunized with the Tax peptide–pulsed BMDCs showed a lower Tax-specific CD8+ T cell proliferation than did the other half (Fig. 4C), indicating that further improvement of BMDC immunization strategy is required to induce optimal Tax-specific CD8+ T cell responses in p.o. rats.

Several studies have reported that MoDCs generated from HTLV-1–infected individuals are functionally reduced (34, 35). In this study, to evaluate the ability of MoDCs to stimulate autologous CD8+ T cells, we did not use Tax-specific CTL epitope peptide as an Ag for the following reasons. First, HTLV-1–infected cells in PBMCs may function as APCs because of their production of Tax protein during the in vitro culture of PBMCs. Second, CMV-infected cells in PBMCs from all donors tested did not provoke CMV-specific memory T cell expansion in the absence of exogenous CMV Ags (Supplemental Fig. 2). Third, Tax-specific memory CD8+ T cells hardly expand in vitro, due in part to the impairment of Tax-specific memory CD8+ T cell responses (21) and other mechanisms underlying the Tax-specific T cell suppression. Lastly, generally, Ag-specific naive T cells do not expand to a detectable level by in vitro single stimulation of PBMCs with Ag-loaded DCs. Although we did not directly compare the function of MoDCs from infected and uninfected donors, we demonstrated that MoDCs from most HTLV-1–infected individuals still had the potential to stimulate Ag-specific CD8+ T cell expansion using CMV-specific CTL epitopes (Fig. 5). This result suggests that peptide-pulsed DC vaccination is applicable to most infected individuals showing weak Tax-specific CD8+ T cell responses.

High PVL is a risk factor for the onset of ATL (4). Weak HTLV-1–specific CD8+ T cell responses could be one of the reasons for elevated PVL. In this case, induction of optimal HTLV-1–specific CD8+ T cell responses by a peptide-pulsed DC vaccine could prevent infected individuals from developing ATL and treat ATL patients by reducing ATL and/or HTLV-1–infected cells. Our recent pilot clinical study demonstrated that all three ATL patients that received our Tax-specific CTL epitope–pulsed DC vaccine showed favorable clinical outcomes without severe adverse events (36). Although there is no direct evidence that Tax-specific CTLs were reinduced by the vaccine, the expansion of ATL and/or HTLV-1–infected cells was subsequently controlled. In this study, we demonstrated that Tax-specific CTL epitope–pulsed DC immunization reduced PVL, probably through induction of Tax-specific CTL responses. Our findings will assist the development of novel immunotherapies against HTLV-1 infection.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Julia Zimmermann (Imperial College London, London, U.K.) for helpful comments and suggestions.

Footnotes

  • This work was supported by Ministry of Education, Culture, Sports, Science and Technology in Japan Grants 23591414 (to A.H.) and 221S0001 (to M.K.), Japan Agency for Medical Research and Development Grants 4A135 and Project for Cancer Research and Therapeutic Evolution (to M.K.), and by funding from the Takeda Science Foundation (to A.H.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    aATL
    acute type of ATL
    AC
    asymptomatic carrier
    ATL
    adult T cell leukemia/lymphoma
    BMDC
    bone marrow–derived DC
    DC
    dendritic cell
    HTLV-1
    human T cell leukemia virus type 1
    i.p. rat
    i.p. infected human T cell leukemia virus type 1–infected rat
    KLH
    keyhole limpet hemocyanin
    MMC
    mitomycin C
    MoDC
    monocyte-derived DC
    p.o. rat
    orally (per os) human T cell leukemia virus type 1–infected rat
    PVL
    proviral load
    rh
    recombinant human
    rr
    recombinant rat.

  • Received September 6, 2016.
  • Accepted November 28, 2016.

References

  1. ↵
    1. Uchiyama T.,
    2. J. Yodoi,
    3. K. Sagawa,
    4. K. Takatsuki,
    5. H. Uchino
    . 1977. Adult T-cell leukemia: clinical and hematologic features of 16 cases. Blood 50: 481–492.
    OpenUrlFREE Full Text
    1. Hinuma Y.,
    2. K. Nagata,
    3. M. Hanaoka,
    4. M. Nakai,
    5. T. Matsumoto,
    6. K. I. Kinoshita,
    7. S. Shirakawa,
    8. I. Miyoshi
    . 1981. Adult T-cell leukemia: antigen in an ATL cell line and detection of antibodies to the antigen in human sera. Proc. Natl. Acad. Sci. USA 78: 6476–6480.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Poiesz B. J.,
    2. F. W. Ruscetti,
    3. A. F. Gazdar,
    4. P. A. Bunn,
    5. J. D. Minna,
    6. R. C. Gallo
    . 1980. Detection and isolation of type C retrovirus particles from fresh and cultured lymphocytes of a patient with cutaneous T-cell lymphoma. Proc. Natl. Acad. Sci. USA 77: 7415–7419.
    OpenUrlAbstract/FREE Full Text
  4. ↵
    1. Iwanaga M.,
    2. T. Watanabe,
    3. A. Utsunomiya,
    4. A. Okayama,
    5. K. Uchimaru,
    6. K. R. Koh,
    7. M. Ogata,
    8. H. Kikuchi,
    9. Y. Sagara,
    10. K. Uozumi,
    11. et al,
    12. Joint Study on Predisposing Factors of ATL Development investigators
    . 2010. Human T-cell leukemia virus type I (HTLV-1) proviral load and disease progression in asymptomatic HTLV-1 carriers: a nationwide prospective study in Japan. Blood 116: 1211–1219.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Kinoshita K.,
    2. T. Amagasaki,
    3. S. Hino,
    4. H. Doi,
    5. K. Yamanouchi,
    6. N. Ban,
    7. S. Momita,
    8. S. Ikeda,
    9. S. Kamihira,
    10. M. Ichimaru,
    11. et al
    . 1987. Milk-borne transmission of HTLV-I from carrier mothers to their children. Jpn. J. Cancer Res. 78: 674–680.
    OpenUrl
  6. ↵
    1. Yamanouchi K.,
    2. K. Kinoshita,
    3. R. Moriuchi,
    4. S. Katamine,
    5. T. Amagasaki,
    6. S. Ikeda,
    7. M. Ichimaru,
    8. T. Miyamoto,
    9. S. Hino
    . 1985. Oral transmission of human T-cell leukemia virus type-I into a common marmoset (Callithrix jacchus) as an experimental model for milk-borne transmission. Jpn. J. Cancer Res. 76: 481–487.
    OpenUrl
  7. ↵
    1. Murphy E. L.,
    2. B. Hanchard,
    3. J. P. Figueroa,
    4. W. N. Gibbs,
    5. W. S. Lofters,
    6. M. Campbell,
    7. J. J. Goedert,
    8. W. A. Blattner
    . 1989. Modelling the risk of adult T-cell leukemia/lymphoma in persons infected with human T-lymphotropic virus type I. Int. J. Cancer 43: 250–253.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Hasegawa A.,
    2. T. Ohashi,
    3. S. Hanabuchi,
    4. H. Kato,
    5. F. Takemura,
    6. T. Masuda,
    7. M. Kannagi
    . 2003. Expansion of human T-cell leukemia virus type 1 (HTLV-1) reservoir in orally infected rats: inverse correlation with HTLV-1-specific cellular immune response. J. Virol. 77: 2956–2963.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Kato H.,
    2. Y. Koya,
    3. T. Ohashi,
    4. S. Hanabuchi,
    5. F. Takemura,
    6. M. Fujii,
    7. H. Tsujimoto,
    8. A. Hasegawa,
    9. M. Kannagi
    . 1998. Oral administration of human T-cell leukemia virus type 1 induces immune unresponsiveness with persistent infection in adult rats. J. Virol. 72: 7289–7293.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    1. Gill P. S.,
    2. W. Harrington Jr..,
    3. M. H. Kaplan,
    4. R. C. Ribeiro,
    5. J. M. Bennett,
    6. H. A. Liebman,
    7. M. Bernstein-Singer,
    8. B. M. Espina,
    9. L. Cabral,
    10. S. Allen,
    11. et al
    . 1995. Treatment of adult T-cell leukemia–lymphoma with a combination of interferon alfa and zidovudine. N. Engl. J. Med. 332: 1744–1748.
    OpenUrlCrossRefPubMed
    1. Hermine O.,
    2. D. Bouscary,
    3. A. Gessain,
    4. P. Turlure,
    5. V. Leblond,
    6. N. Franck,
    7. A. Buzyn-Veil,
    8. B. Rio,
    9. E. Macintyre,
    10. F. Dreyfus,
    11. et al
    . 1995. Brief report: treatment of adult T-cell leukemia–lymphoma with zidovudine and interferon alfa. N. Engl. J. Med. 332: 1749–1751.
    OpenUrlCrossRefPubMed
  12. ↵
    1. Ishida T.,
    2. T. Joh,
    3. N. Uike,
    4. K. Yamamoto,
    5. A. Utsunomiya,
    6. S. Yoshida,
    7. Y. Saburi,
    8. T. Miyamoto,
    9. S. Takemoto,
    10. H. Suzushima,
    11. et al
    . 2012. Defucosylated anti-CCR4 monoclonal antibody (KW-0761) for relapsed adult T-cell leukemia-lymphoma: a multicenter phase II study. J. Clin. Oncol. 30: 837–842.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Utsunomiya A.,
    2. Y. Miyazaki,
    3. Y. Takatsuka,
    4. S. Hanada,
    5. K. Uozumi,
    6. S. Yashiki,
    7. M. Tara,
    8. F. Kawano,
    9. Y. Saburi,
    10. H. Kikuchi,
    11. et al
    . 2001. Improved outcome of adult T cell leukemia/lymphoma with allogeneic hematopoietic stem cell transplantation. Bone Marrow Transplant. 27: 15–20.
    OpenUrlCrossRefPubMed
  14. ↵
    1. Tanosaki R.,
    2. N. Uike,
    3. A. Utsunomiya,
    4. Y. Saburi,
    5. M. Masuda,
    6. M. Tomonaga,
    7. T. Eto,
    8. M. Hidaka,
    9. M. Harada,
    10. I. Choi,
    11. et al
    . 2008. Allogeneic hematopoietic stem cell transplantation using reduced-intensity conditioning for adult T cell leukemia/lymphoma: impact of antithymocyte globulin on clinical outcome. Biol. Blood Marrow Transplant. 14: 702–708.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Hishizawa M.,
    2. J. Kanda,
    3. A. Utsunomiya,
    4. S. Taniguchi,
    5. T. Eto,
    6. Y. Moriuchi,
    7. R. Tanosaki,
    8. F. Kawano,
    9. Y. Miyazaki,
    10. M. Masuda,
    11. et al
    . 2010. Transplantation of allogeneic hematopoietic stem cells for adult T-cell leukemia: a nationwide retrospective study. Blood 116: 1369–1376.
    OpenUrlAbstract/FREE Full Text
    1. Ishida T.,
    2. M. Hishizawa,
    3. K. Kato,
    4. R. Tanosaki,
    5. T. Fukuda,
    6. Y. Takatsuka,
    7. T. Eto,
    8. Y. Miyazaki,
    9. M. Hidaka,
    10. N. Uike,
    11. et al
    . 2013. Impact of graft-versus-host disease on allogeneic hematopoietic cell transplantation for adult T cell leukemia-lymphoma focusing on preconditioning regimens: nationwide retrospective study. Biol. Blood Marrow Transplant. 19: 1731–1739.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Okamura J.,
    2. A. Utsunomiya,
    3. R. Tanosaki,
    4. N. Uike,
    5. S. Sonoda,
    6. M. Kannagi,
    7. M. Tomonaga,
    8. M. Harada,
    9. N. Kimura,
    10. M. Masuda,
    11. et al
    . 2005. Allogeneic stem-cell transplantation with reduced conditioning intensity as a novel immunotherapy and antiviral therapy for adult T-cell leukemia/lymphoma. Blood 105: 4143–4145.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Jacobson S.,
    2. H. Shida,
    3. D. E. McFarlin,
    4. A. S. Fauci,
    5. S. Koenig
    . 1990. Circulating CD8+ cytotoxic T lymphocytes specific for HTLV-I pX in patients with HTLV-I associated neurological disease. Nature 348: 245–248.
    OpenUrlCrossRefPubMed
    1. Kannagi M.,
    2. N. Harashima,
    3. K. Kurihara,
    4. T. Ohashi,
    5. A. Utsunomiya,
    6. R. Tanosaki,
    7. M. Masuda,
    8. M. Tomonaga,
    9. J. Okamura
    . 2005. Tumor immunity against adult T-cell leukemia. Cancer Sci. 96: 249–255.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Bangham C. R.
    2008. HTLV-1 infection: role of CTL efficiency. Blood 112: 2176–2177.
    OpenUrlFREE Full Text
  21. ↵
    1. Takamori A.,
    2. A. Hasegawa,
    3. A. Utsunomiya,
    4. Y. Maeda,
    5. Y. Yamano,
    6. M. Masuda,
    7. Y. Shimizu,
    8. Y. Tamai,
    9. A. Sasada,
    10. N. Zeng,
    11. et al
    . 2011. Functional impairment of Tax-specific but not cytomegalovirus-specific CD8+ T lymphocytes in a minor population of asymptomatic human T-cell leukemia virus type 1-carriers. Retrovirology 8: 100.
    OpenUrlCrossRefPubMed
    1. Arnulf B.,
    2. M. Thorel,
    3. Y. Poirot,
    4. R. Tamouza,
    5. E. Boulanger,
    6. A. Jaccard,
    7. E. Oksenhendler,
    8. O. Hermine,
    9. C. Pique
    . 2004. Loss of the ex vivo but not the reinducible CD8+ T-cell response to Tax in human T-cell leukemia virus type 1-infected patients with adult T-cell leukemia/lymphoma. Leukemia 18: 126–132.
    OpenUrlCrossRefPubMed
    1. Manuel S. L.,
    2. M. Sehgal,
    3. J. Connolly,
    4. G. Makedonas,
    5. Z. K. Khan,
    6. J. Gardner,
    7. M. R. Betts,
    8. P. Jain
    . 2013. Lack of recall response to Tax in ATL and HAM/TSP patients but not in asymptomatic carriers of human T-cell leukemia virus type 1. J. Clin. Immunol. 33: 1223–1239.
    OpenUrl
  24. ↵
    1. Shimizu Y.,
    2. A. Takamori,
    3. A. Utsunomiya,
    4. M. Kurimura,
    5. Y. Yamano,
    6. M. Hishizawa,
    7. A. Hasegawa,
    8. F. Kondo,
    9. K. Kurihara,
    10. N. Harashima,
    11. et al
    . 2009. Impaired Tax-specific T-cell responses with insufficient control of HTLV-1 in a subgroup of individuals at asymptomatic and smoldering stages. Cancer Sci. 100: 481–489.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Hanabuchi S.,
    2. T. Ohashi,
    3. Y. Koya,
    4. H. Kato,
    5. F. Takemura,
    6. K. Hirokawa,
    7. T. Yoshiki,
    8. H. Yagita,
    9. K. Okumura,
    10. M. Kannagi
    . 2000. Development of human T-cell leukemia virus type 1-transformed tumors in rats following suppression of T-cell immunity by CD80 and CD86 blockade. J. Virol. 74: 428–435.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Banchereau J.,
    2. R. M. Steinman
    . 1998. Dendritic cells and the control of immunity. Nature 392: 245–252.
    OpenUrlCrossRefPubMed
  27. ↵
    1. Hishizawa M.,
    2. K. Imada,
    3. T. Kitawaki,
    4. M. Ueda,
    5. N. Kadowaki,
    6. T. Uchiyama
    . 2004. Depletion and impaired interferon-α-producing capacity of blood plasmacytoid dendritic cells in human T-cell leukaemia virus type I-infected individuals. Br. J. Haematol. 125: 568–575.
    OpenUrlCrossRefPubMed
  28. ↵
    1. Palucka K.,
    2. J. Banchereau
    . 2013. Dendritic-cell-based therapeutic cancer vaccines. Immunity 39: 38–48.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Luo J.,
    2. J. Li,
    3. R. L. Chen,
    4. L. Nie,
    5. J. Huang,
    6. Z. W. Liu,
    7. L. Luo,
    8. X. J. Yan
    . 2010. Autologus dendritic cell vaccine for chronic hepatitis B carriers: a pilot, open label, clinical trial in human volunteers. Vaccine 28: 2497–2504.
    OpenUrl
    1. Liau L. M.,
    2. R. M. Prins,
    3. S. M. Kiertscher,
    4. S. K. Odesa,
    5. T. J. Kremen,
    6. A. J. Giovannone,
    7. J. W. Lin,
    8. D. J. Chute,
    9. P. S. Mischel,
    10. T. F. Cloughesy,
    11. M. D. Roth
    . 2005. Dendritic cell vaccination in glioblastoma patients induces systemic and intracranial T-cell responses modulated by the local central nervous system tumor microenvironment. Clin. Cancer Res. 11: 5515–5525.
    OpenUrlAbstract/FREE Full Text
    1. Tuettenberg A.,
    2. C. Becker,
    3. E. Huter,
    4. J. Knop,
    5. A. H. Enk,
    6. H. Jonuleit
    . 2006. Induction of strong and persistent MelanA/MART-1-specific immune responses by adjuvant dendritic cell-based vaccination of stage II melanoma patients. Int. J. Cancer 118: 2617–2627.
    OpenUrlCrossRefPubMed
    1. Czerniecki B. J.,
    2. G. K. Koski,
    3. U. Koldovsky,
    4. S. Xu,
    5. P. A. Cohen,
    6. R. Mick,
    7. H. Nisenbaum,
    8. T. Pasha,
    9. M. Xu,
    10. K. R. Fox,
    11. et al
    . 2007. Targeting HER-2/neu in early breast cancer development using dendritic cells with staged interleukin-12 burst secretion. Cancer Res. 67: 1842–1852.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Wierecky J.,
    2. M. R. Müller,
    3. S. Wirths,
    4. E. Halder-Oehler,
    5. D. Dörfel,
    6. S. M. Schmidt,
    7. M. Häntschel,
    8. W. Brugger,
    9. S. Schröder,
    10. M. S. Horger,
    11. et al
    . 2006. Immunologic and clinical responses after vaccinations with peptide-pulsed dendritic cells in metastatic renal cancer patients. Cancer Res. 66: 5910–5918.
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. Nascimento C. R.,
    2. M. A. Lima,
    3. M. J. de Andrada Serpa,
    4. O. Espindola,
    5. A. C. Leite,
    6. J. Echevarria-Lima
    . 2011. Monocytes from HTLV-1-infected patients are unable to fully mature into dendritic cells. Blood 117: 489–499.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Makino M.,
    2. S. Wakamatsu,
    3. S. Shimokubo,
    4. N. Arima,
    5. M. Baba
    . 2000. Production of functionally deficient dendritic cells from HTLV-I-infected monocytes: implications for the dendritic cell defect in adult T cell leukemia. Virology 274: 140–148.
    OpenUrlCrossRefPubMed
  36. ↵
    1. Suehiro Y.,
    2. A. Hasegawa,
    3. T. Iino,
    4. A. Sasada,
    5. N. Watanabe,
    6. M. Matsuoka,
    7. A. Takamori,
    8. R. Tanosaki,
    9. A. Utsunomiya,
    10. I. Choi,
    11. et al
    . 2015. Clinical outcomes of a novel therapeutic vaccine with Tax peptide-pulsed dendritic cells for adult T cell leukaemia/lymphoma in a pilot study. Br. J. Haematol. 169: 356–367.
    OpenUrlCrossRefPubMed
  37. ↵
    1. Ohashi T.,
    2. S. Hanabuchi,
    3. H. Kato,
    4. Y. Koya,
    5. F. Takemura,
    6. K. Hirokawa,
    7. T. Yoshiki,
    8. Y. Tanaka,
    9. M. Fujii,
    10. M. Kannagi
    . 1999. Induction of adult T-cell leukemia-like lymphoproliferative disease and its inhibition by adoptive immunotherapy in T-cell-deficient nude rats inoculated with syngeneic human T-cell leukemia virus type 1-immortalized cells. J. Virol. 73: 6031–6040.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Ohashi T.,
    2. S. Hanabuchi,
    3. H. Kato,
    4. H. Tateno,
    5. F. Takemura,
    6. T. Tsukahara,
    7. Y. Koya,
    8. A. Hasegawa,
    9. T. Masuda,
    10. M. Kannagi
    . 2000. Prevention of adult T-cell leukemia-like lymphoproliferative disease in rats by adoptively transferred T cells from a donor immunized with human T-cell leukemia virus type 1 Tax-coding DNA vaccine. J. Virol. 74: 9610–9616.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Hanabuchi S.,
    2. T. Ohashi,
    3. Y. Koya,
    4. H. Kato,
    5. A. Hasegawa,
    6. F. Takemura,
    7. T. Masuda,
    8. M. Kannagi
    . 2001. Regression of human T-cell leukemia virus type I (HTLV-I)-associated lymphomas in a rat model: peptide-induced T-cell immunity. J. Natl. Cancer Inst. 93: 1775–1783.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Wills M. R.,
    2. A. J. Carmichael,
    3. K. Mynard,
    4. X. Jin,
    5. M. P. Weekes,
    6. B. Plachter,
    7. J. G. Sissons
    . 1996. The human cytotoxic T-lymphocyte (CTL) response to cytomegalovirus is dominated by structural protein pp65: frequency, specificity, and T-cell receptor usage of pp65-specific CTL. J. Virol. 70: 7569–7579.
    OpenUrlAbstract/FREE Full Text
  41. ↵
    1. Kuzushima K.,
    2. N. Hayashi,
    3. H. Kimura,
    4. T. Tsurumi
    . 2001. Efficient identification of HLA-A*2402-restricted cytomegalovirus-specific CD8+ T-cell epitopes by a computer algorithm and an enzyme-linked immunospot assay. Blood 98: 1872–1881.
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Grauer O.,
    2. G. Wohlleben,
    3. S. Seubert,
    4. A. Weishaupt,
    5. E. Kämpgen,
    6. R. Gold
    . 2002. Analysis of maturation states of rat bone marrow-derived dendritic cells using an improved culture technique. Histochem. Cell Biol. 117: 351–362.
    OpenUrlCrossRefPubMed
  43. ↵
    1. Iwanaga M.,
    2. T. Watanabe,
    3. K. Yamaguchi
    . 2012. Adult T-cell leukemia: a review of epidemiological evidence. Front. Microbiol. 3: 322.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Ohashi T.,
    2. S. Hanabuchi,
    3. R. Suzuki,
    4. H. Kato,
    5. T. Masuda,
    6. M. Kannagi
    . 2002. Correlation of major histocompatibility complex class I downregulation with resistance of human T-cell leukemia virus type 1-infected T cells to cytotoxic T-lymphocyte killing in a rat model. J. Virol. 76: 7010–7019.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Kozako T.,
    2. M. Yoshimitsu,
    3. H. Fujiwara,
    4. I. Masamoto,
    5. S. Horai,
    6. Y. White,
    7. M. Akimoto,
    8. S. Suzuki,
    9. K. Matsushita,
    10. K. Uozumi,
    11. et al
    . 2009. PD-1/PD-L1 expression in human T-cell leukemia virus type 1 carriers and adult T-cell leukemia/lymphoma patients. Leukemia 23: 375–382.
    OpenUrlCrossRefPubMed
  46. ↵
    1. Van Prooyen N.,
    2. V. Andresen,
    3. H. Gold,
    4. I. Bialuk,
    5. C. Pise-Masison,
    6. G. Franchini
    . 2010. Hijacking the T-cell communication network by the human T-cell leukemia/lymphoma virus type 1 (HTLV-1) p12 and p8 proteins. Mol. Aspects Med. 31: 333–343.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Toulza F.,
    2. A. Heaps,
    3. Y. Tanaka,
    4. G. P. Taylor,
    5. C. R. Bangham
    . 2008. High frequency of CD4+FoxP3+ cells in HTLV-1 infection: inverse correlation with HTLV-1-specific CTL response. Blood 111: 5047–5053.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. Chen S.,
    2. N. Ishii,
    3. S. Ine,
    4. S. Ikeda,
    5. T. Fujimura,
    6. L. C. Ndhlovu,
    7. P. Soroosh,
    8. K. Tada,
    9. H. Harigae,
    10. J. Kameoka,
    11. et al
    . 2006. Regulatory T cell-like activity of Foxp3+ adult T cell leukemia cells. Int. Immunol. 18: 269–277.
    OpenUrlAbstract/FREE Full Text
  49. ↵
    1. Mayer L.,
    2. L. Shao
    . 2004. Therapeutic potential of oral tolerance. Nat. Rev. Immunol. 4: 407–419.
    OpenUrlCrossRefPubMed
  50. ↵
    1. Matloubian M.,
    2. R. J. Concepcion,
    3. R. Ahmed
    . 1994. CD4+ T cells are required to sustain CD8+ cytotoxic T-cell responses during chronic viral infection. J. Virol. 68: 8056–8063.
    OpenUrlAbstract/FREE Full Text
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