Pilot study of Wilms tumor 1-targeted human platelet lysate-induced antigen-presenting cells in advanced solid cancer patients

Abstract

Introduction

For the wide implementation of dendritic cell-based cancer immunotherapy, a clinical phase I study was conducted to explore its safety and tolerability using Wilms’ tumor 1 (WT1) antigen-targeted human platelet lysate-induced antigen-presenting cells (WT1-HPL-APCs), developed by our team, in patients with advanced solid cancers.

Methods

WT1-HPL-APCs were manufactured from apheresed peripheral blood mononuclear cells administered at 4 × 10⁷ cells biweekly for three sessions (one course). Adverse events and the induction of WT1-specific T cells (WT1-Cytotoxic T lymphocytes [CTLs]) were evaluated. Furthermore, the potential immunoinducibility of WT1-HPL-APC was examined in vitro and regulatory T cells and monocytic myeloid-derived suppressor cells (M-MDSCs) in the tumor microenvironment (TME) and peripheral blood were assessed to monitor immune cells.

Results

The adverse events associated with WT1-HPL-APC vaccination included fever, erythema at the injection site, arthralgia, and chills, all of which were lower than grade 2 in two patients who completed three sessions. The WT1 expression levels and infiltration of CD4⁺ T cells, CD8⁺ T cells, and macrophages in the TME differed among the patients. In only one of the two patients with HLA-A*24:02, WT1-HPL-APC induced WT1-CTLs in vitro. Although a WT1-CTL response could not be detected following three sessions of WT1-HPL-APC administration, the percentage of M-MDSCs in the PB tended to decrease.

Conclusions

As this pilot study included only three patients with different cancer types, evidence supporting the effectiveness or functionality of WT1-HPL-APCs remains limited. WT1-HPL-APC vaccination was tolerable among patients with advanced solid cancers. Our findings suggest that the immunoinducibility of WT1-HPL-APC vary across individuals and that the preexisting immunosuppressive environment could limit WT1-CTL induction. Future studies are warranted to determine whether WT1-HPL-APC vaccination can improve the patient's immunosuppressive environment and induce WT1-CTLs in combination with optimized chemotherapy or adjuvant therapy in large-scale clinical trials involving multiple doses.

Clinical trial registration number

jRCTc040200005.

Abbreviations

Abs

Antibodies

APC

Antigen-presenting cells

CA

Cancer antigen

CTLs

Cytotoxic T lymphocytes

DC

Dendritic cell

DTH

Delayed-type hypersensitivity

ELISpot

Enzyme-linked immunospot

EMA

Epithelial membrane antigen

FITC

Fluorescein isothiocyanate

GEM

Gemcitabine

GM-CSF

Granulocyte–macrophage colony-stimulating factor

HLA

Human leukocyte antigen

HPL

Human platelet lysate

IFN

Interferon

IFN-DCs

type I IFN-induced DCs

IHC

Immunohistochemistry

M-MDSCs

Monocytic myeloid-derived suppressor cells

Nab-paclitaxel

nab-PTX

OS

Overall survival

PBMCs

Peripheral blood mononuclear cells

PCA

Personalized cancer antigen

PE

Phycoerythrin

TME

Tumor microenvironment

Tregs

Regulatory T cells

WT1

Wilms tumor 1

WT1-CTLs

WT1-specific CTLs

WT1-HP

WT1 helper peptide

WT1-HPL-APCs

WT1-loaded human platelet lysate-induced antigen-presenting cells

WT1-KP

WT1 killer peptide

7-AAD

7-aminoactinomycin D

1. Introduction

Antigen-presenting cells (APCs), such as dendritic cells (DCs), play a pivotal role in acquired immunity by activating T cells through major histocompatibility complex-bound peptides on compatible T-cell receptors, which also interact with CD28 on T cells, binding to CD80/CD86 on APCs [[1], [2], [3]]. Cytotoxic T lymphocytes (CTLs), a subtype of antigen-specific T cells, exhibit strong effector activity and can directly target and kill tumor cells expressing specific antigens [4,5]. DC-based cancer vaccines have been conventionally developed by differentiating apheresed peripheral blood mononuclear cells (PBMCs) into monocyte-derived DCs. These monocyte-derived DC vaccines have been used as active immunotherapies for solid cancers [[6], [7], [8], [9]]. Interleukin-4 (IL-4) and granulocyte–macrophage colony-stimulating factor (GM-CSF) have long been used to generate monocyte-derived DCs [10]. Type 1 interferon (IFN)-induced DCs (IFN-DCs) can also be used as alternatives [11]. IFN-DCs not only induce CD8⁺ T cells [12,13] but also direct cytotoxic activity against tumor cells [14]. IFN-DC vaccines have also been used in clinical trials for cancer therapies [[15], [16], [17]].

Wilms’ tumor 1 (WT1) antigen is expressed in various tumor cells [18]; thus, it has been used as a common targeted antigen for DC-based cancer therapies [[19], [20], [21]]. WT1 is particularly useful as a priority target owing to its expression in not only tumor cells but also cancer stem cells, tumor blood vessels, and myeloid-derived suppressor cells (MDSCs) [22,23]. Furthermore, the WT1 peptide cancer vaccine has shown potential to convert “cold tumors” (tumors with poor immune cell infiltration) into “hot tumors” (tumors with rich immune cell infiltration), thereby enhancing the effectiveness of the immune response [24].

MDSCs and regulatory T cells (Tregs) contribute to the establishment of a cancer immunosuppressive environment that supports tumor progression and suppresses CTL activity in the tumor microenvironment (TME) and peripheral blood (PB) [[25], [26], [27], [28]]. The three main MDSC subsets are granulocytic MDSCs, monocytic MDSCs (M-MDSCs), and either early-stage or immature MDSCs [[29], [30], [31]]. The presence of these subsets is associated with overall survival (OS) and cancer staging in patients with cancer [32]. In addition, M-MDSCs have been suggested to influence the function of DC-based vaccines by inhibiting DC maturation, antigen uptake, DC migration to lymph nodes, and T-cell proliferation [33]. For these reasons, immunosuppressive cells and TME must be examined to assess the presence of these cells and CTL dysfunction [34], which, in turn, helps determine the appropriate cancer treatment.

In a previous study, a method for manufacturing human platelet lysate (HPL)-induced APCs (HPL-APCs), which could be generated using IFN-α and GM-CSF supplemented with HPL, was developed [35]. HPL-APCs demonstrated a stronger ability to induce antigen-specific CTLs than IFN-DCs without HPL. In our previous phase I clinical trial, gene mutations were identified via a cancer gene panel test on primary tumor tissues, and the safety and tolerability of a personalized HPL-IFN-APC vaccine pulsed with optimized cancer antigen peptides were explored [36]. HPL-APCs were induced from fresh apheresed PBMCs and then cryopreserved until administration in the clinical trial. A total of 2–3 × 10⁷ HPL-APCs were administered biweekly over six doses. This dosage schedule was tolerable and demonstrated effectiveness in stimulating T-cell adaptive immunity.

In another clinical trial with IL-4-DCs, 2.5–5 × 10⁷ cells/session, administered three times biweekly, demonstrated a positive delayed-type hypersensitivity (DTH) skin reaction, antigen-specific T-cell induction, and tumor shrinkage [37].

Therefore, in this pilot study, a three-time-biweekly dosage protocol for the administration of fresh 4 × 10⁷ HPL-APCs loaded with Wilms’ tumor 1 (WT1) antigen-targeted human platelet lysate-induced antigen-presenting cells (WT1-HPL-APCs) was established to reduce physical burden on patients, alleviate cell damage caused by cryopreservation, and evaluate long-lasting acquired T-cell immunity. The safety and tolerability of WT1-HPL-APCs in patients with advanced solid cancers were assessed as the primary endpoints.

2. Methods

2.1. Generation of WT1-HPL-APCs or WT1-IL-4-DCs and in vitro induction of WT1-CTLs

PBMCs were collected via leukapheresis using a Spectra Optia® blood cell separator (Terumo BCT, Inc., Tokyo, Japan). The PBMCs were enriched via density gradient centrifugation using Ficoll-Plaque Premium (Global Life Sciences Solution USA LLC, Marlborough, MA, USA). Then, they were stored until the subsequent processes without cryopreservation at −150°C. HPL-APCs and IL-4-DCs were produced from monocytes, as described in [35,38], respectively. HPL-APCs and IL-4-DCs were pulsed with 10 μg/mL of WT1 peptides (PEPTIDE INSTITUTE, INC., Osaka, Japan) during the maturation phase, as previously described [38]. After 24 h, each cell was collected as a stimulator cell, divided into aliquots, and then cryopreserved. Furthermore, CD8⁺ T cells were separated from HLA-A*24:02-autologous PBMCs using CD8 microbeads (Miltenyi Biotec B.V. & Co. KG, Bergisch Gladbach, Germany) applied as responder cells. The stimulator and responder cells were cocultured in a 96-well U-bottom plate at a ratio of 1:10 in an AIM-V medium (Thermo Fisher Scientific, Inc., Waltham, MA, USA). On days 7, 14, and 21, an HLA-A*24:02-WT1-tetramer assay was conducted to confirm the potential for WT1-CTL induction by WT1-HPL-APCs or WT1-IL-4-DCs, as previously described [35].

2.2. Eligibility criteria for the WT1-HPL-APC pilot study registry

The patients included in this study had one of the following cancers: breast, rectal, pancreatic, stomach, or lung cancer. The inclusion criteria were as follows: (1) aged 20–75 years at the time of informed consent provision; (2) WHO-Eastern Cooperative Oncology Group performance status 0 or 1; (3) cancer types reported to be positive for WT1 antigen; (4) adjuvant therapy following surgical resection; (5) absence of cardiovascular or respiratory diseases that would prevent blood apheresis; (6) absence of organ dysfunction, blood abnormalities, or bleeding tendencies; (7) absence of active infectious diseases; (8) sufficient understanding of the risks and benefits of WT1-HPL-APC administration and ability to provide written informed consent; and (9) nonparticipation in other clinical trials.

2.3. Immunohistochemistry analysis

To examine primary cancer samples, the following antihuman antibodies (Abs) for immunohistochemistry (IHC) were used: WT1 (clone, 6F-H2, Abcam, MA, USA), epithelial membrane antigen (EMA; clone, GP1.4, Leica Biosystems, Nussloch, Germany), and human leukocyte antigen (HLA)-ABC (clone, EMR8-5, HOKUDO Co., Ltd., Sapporo, Japan). Anti-CD4, anti-CD8, and anti-CD68 Abs were provided by ALP Inc. (Ishikawa, Japan). All IHC staining procedures were outsourced to ALP Inc. Images were analyzed at ×100 or ×400 magnifications using the all-in-one inverted fluorescence microscope BZ-X700 or BZ-X800 (KEYENCE Corp., Osaka, Japan).

2.4. Preparation of WT1-HPL-APCs for clinical use

PBMCs were collected via leukapheresis using a Spectra Optia® blood cell separator (Terumo BCT, Inc., Tokyo, Japan). The PBMCs were enriched via density gradient centrifugation using Ficoll-Plaque Premium (Global Life Sciences Solution USA LLC, Marlborough, MA, USA). One portion of the PBMCs was immediately used for subsequent processes without cryopreservation. HPL-APCs were generated from monocytes using 100 ng/mL of recombinant human GM-CSF (Miltenyi Biotec B.V. & Co. KG, Bergisch Gladbach, Germany), 1 μg/mL of PEGylated-IFN-α-2b (PEG-Intron; MSD, Tokyo, Japan; for Case 1) or PEGylated-IFN-α-2a (PEGASYS®; Chugai Pharmaceutical Co., Ltd., Tokyo, Japan; for Cases 2 and 3), and 5% (v/v) GMP-grade UltraGRO^TM-PURE GI (called “HPL” in this study; AventaCell BioMedical Corp., Atlanta, GA, USA) in serum-free medium DCO-K (Shimadzu Diagnostics Corp., Tokyo, Japan) for 3 days. After cell collection and washing, HPL-APCs were matured with 5% (v/v) HPL in DCO-K, supplemented with 100 ng/mL of recombinant human GM-CSF (Miltenyi Biotec B.V. & Co. KG), 1 μg/mL of PEGylated-IFN-α-2b (PEG-Intron®; MSD; for Case 1) or PEGylated-IFN-α-2a (PEGASYS®; Chugai Pharmaceutical Co., Ltd; for Cases 2 and 3), 10 μg/mL of OK-432 (Streptococcal Preparation, Chugai), 10 ng/mL of prostaglandin E2 (PGE2; Kyowa Pharma Chemical Co., Ltd., Toyama, Japan), and 10 μg/mL of WT1 peptides (PEPTIDE INSTITUTE, INC., Osaka, Japan). Two types of WT1 peptides were utilized: the HLA-A*24:02-restricted modified WT1-243 killer peptide (WT1-KP; CYTWNQMNL, residues 235–243) and the HLA-DPB1*05:01-applicable WT1-34 helper peptide (WT1-HP; WAPVLDFAPPGASAYGSL, residues 34–51), which contains killer peptides for HLA-A*02:01 and HLA-A*02:06 [39,40]. After a 16–18-h incubation, sterility and phenotype tests were conducted to ensure the quality of WT1-HPL-APCs. After harvesting, cells were washed twice with normal saline to remove residual culture medium, doublet-depleted with a cell strainer, counted, and resuspended in saline. The vaccine was administered promptly after preparation and shipment, and adverse events were monitored.

2.5. Sterility testing of WT1-HPL-APCs

Each lot was subjected to sterility testing. Endotoxin contamination was assessed using the supernatant derived from WT1-HPL-APCs on culture day 5 with Endosafe® nexgen-PTS^TM (VERITAS Corp., Tokyo, Japan). For endotoxin, the release criterion was <1.0 EU/mL. A mycoplasma test was conducted on the supernatant of WT1-HPL-APCs on culture days 4 and 5 using MycoAlert® (Lonza, Basel, Switzerland) or MycoFinder (Shimadzu Diagnostics Corp.). The releasing criterion was negative for mycoplasma contamination. A general viable bacteria test was conducted to prevent bacterial contamination using the supernatant of HPL-APCs on culture day 4. A portion of the supernatant was applied to the DD checker SCD agar medium (Kyokuto Pharmaceutical Industrial Co., Ltd., Tokyo, Japan) and incubated for ≥20 h at 35°C. The release criterion was the absence of viable bacteria. To confirm WT1-HPL-APC sterility, the supernatant of WT1-HPL-APCs collected on culture day 5 was incubated for 14 days using the direct inoculation method, as per the Japanese Pharmacopoeia. The absence of bacterial or fungal contamination led to a negative result.

2.6. In vitro WT1-CTL induction assay using PBMCs from enrolled patients

WT1-HPL-APCs were manufactured from PBMCs obtained from Cases 1 and 3 with HLA-A*24:02, as described in Section 2.4. To replicate the pilot clinical study protocol, WT1-HPL-APCs were cocultured with autologous CD8⁺ T cells, as described in Section 2.1. The WT1-tetramer analysis was conducted to evaluate the in vitro immunoinducibility of WT1-CTLs by WT1-HPL-APCs.

2.7. Phenotyping of WT1-HPL-APCs

To determine whether WT1-HPL-APCs exhibited the phenotype characteristics of APCs [35], the following fluorescein isothiocyanate (FITC) or phycoerythrin (PE)-conjugated antihuman monoclonal Abs were used as maturation markers on the cell surface for flow cytometry: CD80-FITC (clone L307.4; BD Biosciences, Franklin Lakes, NJ), CD86-PE (clone IT2.2; eBioscience, Inc., San Diego, CA), HLA-ABC-FITC (clone G46-2.6, BD Biosciences), HLA-DR-PE (clone LN3; eBioscience, Inc.), CD40-FITC (clone 5C3; eBioscience, Inc.), and CCR7-PE (clone 150503; R&D System, Minneapolis, MN). The phenotypes of WT1-HPL-APCs on culture day 5 were analyzed using FACS Calibur^TM (BD Biosciences). The measurements were analyzed using FlowJo^TM (ver. 10.10.0; Tree Star, Ashland, OR, USA). The lymphocyte region was removed via forward scatter (FSC) and side scatter (SSC). Live cells were defined as 7-aminoactinomycin D (7-AAD) (Miltenyi Biotec B.V. & Co. KG)-negative cells.

2.8. Detection of cytokine production from WT1-HPL-APCs

Cytokine production was assessed using the supernatant derived from WT1-HPL-APC vaccines with Bio-Plex Pro Cytokine Assays (Bio-Rad Laboratories, Inc., Hercules, CA), as previously described [35]. The IL-6, IL-10, IL-12 (p70), IFN-γ, and TNF-α levels were measured using WT1-HPL-APCs from the enrolled patients and compared with personalized cancer antigen (PCA)- HPL-APC vaccines prepared using the same HPL-APCs [36]. All measurements were performed in duplicate in accordance with the kit protocol. Cytokine values were summarized as the median ± half of the interquartile range (75th–25th percentile).

2.9. Quality criteria and administration of WT1-HPL-APCs

The quality criteria for WT1-HPL-APCs were as follows: cell viability ≥ 70% and total cell number ≥ 4 × 10⁷ cells. The tentative criterion for purity, defined as CD86⁺/HLA-DR⁺ in live cells, was set at ≥70% [36]. WT1-HPL-APCs were harvested and suspended in 2 mL of saline. A total of 4 × 10⁷ WT1-HPL-APCs/session were intradermally and bilaterally injected near the axillary and groin regions. The WT1-HPL-APCs were administered in three sessions biweekly.

2.10. Assessment of safety following WT1-HPL-APC administration

In all patients, adverse events, such as fever, arthralgia, chills, or headache, were monitored following each administration. These events were scored according to the Common Terminology Criteria for Adverse Events (version 5.0-JCOG). The skin reaction at the injection site caused by WT1-HPL-APCs was evaluated 48 h after each vaccination.

2.11. Evaluation of immune response by WT1-Specific CTLs (WT1-CTLs)

HLA phenotyping for HLA-A, HLA-DPB1, and HLA-DRB1 was performed using the WAKFlow HLA typing kit (Wakunaga Pharmaceutical Co., Ltd., Hiroshima, Japan). For immune monitoring, PBMCs were cryopreserved following isolation via Ficoll density gradient centrifugation at each session. To detect WT1-CTLs, PBMCs were stained with PE-conjugated HLA-A* 24:02-modified WT1-tetramer (Medical & Biological Laboratories Co., Ltd., Tokyo, Japan) and FITC-conjugated antihuman CD8 (clone SFCI21Thy2D3, Beckman Coulter, Inc., CA, USA), as previously described [40]. To examine WT1-specific IFN-γ production by WT1-CTLs, enzyme-linked immunospot (ELISpot) assays were conducted, as previously described [40]. The following two WT1 antigen peptides (PEPTIDE INSTITUTE, Inc.) and three HIV antigen peptides were used for the ELISpot assays: 250 μg/mL of WT1-KP, 100 μg/mL of WT1-HP, and 250 or 500 μg/mL of HIV peptides (RYLRDQQLL, SLYNTVATL, and DYVDRFYKTLRAE, synthesized by GenScript Biotech Corp., NJ, USA) as negative controls. In the DTH skin reaction test, following intradermal injection of 20 μg of WT1-KP, WT1-HP, or a negative control (dimethyl sulfoxide) in 100 μL of saline into the patient's forearm, reactions were assessed 48 h after each WT1-HPL-APC administration session in all patients. A positive reaction was defined as an erythema with a maximum diameter ≥ 2 mm [39].

2.12. Measurement of Tregs and M-MDSCs

Tregs and M-MDSCs were evaluated via flow cytometry using the following mouse antihuman Abs: CD3-FITC (clone SK7; BD Biosciences), CD25-PE (clone 2A3; BD Biosciences), CD4-APC-Cy7 (clone SK3; BioLegend, CA, USA), CD127-Brilliant Violet 421 (BV421; clone A019D5; BioLegend), CD45-Brilliant Violet 510 (BV510; clone 2D1; BioLegend), CD14-FITC (clone 61D3; eBioscience), HLA-DR-PE (clone LN3; eBioscience), CD11b-APC-Cy7 (clone ICRF44; eBioscience), and CD33-PE-Cy7 (clone WM53; BD Biosciences). The stained cells were analyzed using FACS Lyric^TM (BD Biosciences), and data were processed using FlowJo^TM (ver10.10.0; Tree Star). The Treg and M-MDSC regions were gated using FSC and SSC. Live cells were defined as 7-AAD (Miltenyi Biotec B.V. & Co. KG)-negative cells, Tregs as CD45⁺CD3⁺CD4⁺CD25⁺CD127^-/low cells [41], and M-MDSCs as CD45⁺CD11b⁺CD33⁺CD14⁺HLA-DR⁻ cells [42]. The gate strategy is illustrated in Fig. S1 for Tregs and Fig. S2 for M-MDSCs.

2.13. Statistical analysis

Fisher's exact test was conducted for the statistical comparisons between the initial administration of WT1-HPL-APCs derived from fresh PBMCs and the second administration of those derived from cryopreserved PBMCs in all three cases. Statistical analyses were conducted using IBM SPSS Statistics version 24 (IBM Japan, Ltd., Tokyo, Japan). A p-value <0.05 was considered significant. The threshold for CD86 and HLA-DR was set at 70% positive cells [36].

3. Results

3.1. Validation of the potency of WT1-HPL-APCs in inducing WT1-CTLs in vitro

To evaluate the ability of WT1-HPL-APCs to induce WT1-CTLs, CD8⁺ T cells were stimulated with WT1-HPL-APCs in vitro every week and cultured for three weeks (Fig. 1). In vitro immune monitoring with the WT1-tetramer assay was conducted using PBMCs from one HLA-A*24:02-positive patient with gastric cancer. WT1-HPL-APCs exhibited higher WT1-CTL induction than IL-4-DCs on day 21 (0.05%, 0.17%, and 0.27% in CD8⁺ T cells only, IL-4-DCs, and WT1-HPL-APCs, respectively; Fig. 1a; 0.9 × 10⁵, 5.9 × 10⁵, and 8.2 × 10⁵ cells in CD8⁺ T cells only, IL-4-DCs, and WT1-HPL-APCs, respectively; Fig. 1b).

Fig. 1.

The potential to induce Wilms' tumor 1 (WT1)-specific CTLs (WT1-CTLs) in vitro, and the administration schedule and manufacturing protocol for WT1-targeted human platelet lysate-induced antigen-presenting cells (WT1-HPL-APCs) in a clinical setting. (a) CD8⁺ T cells were cocultured and stimulated weekly with WT1 peptide-pulsed IL-4-DCs or HPL-APCs, and WT1-CTLs were detected using WT1-tetramer assay. On day 21 (after the third APC stimulation), WT1-CTLs were identified as CD3⁺CD8⁺HLA-A* 24:02 WT1-tetramer⁺ cells. The percentage indicates WT1-tetramer⁺ cells in CD8⁺ T cells. (b) The graph shows the cell number of WT1-CTLs on days 1, 7, 14, and 21. (c) Biweekly WT1-HPL-APC administration and immune monitoring. Clinical records of adverse events, including blood tests, were collected, and immune monitoring was performed using delayed-type hypersensitivity (DTH) skin reaction tests, enzyme-linked immunospot (ELISpot) assays for the detection of IFN-γ production from WT1-specific CD8⁺ T cells, and WT1-tetramer assay for the detection of WT1-specific CD8⁺ T cells. (d) WT1-HPL-APCs manufactured from fresh or cryopreserved peripheral blood mononuclear cells obtained via leukapheresis.

The dosing schedule consisted of three biweekly sessions. WT1-CTL immune response was monitored after WT1-HPL-APC administration (Fig. 1c). WT1-HPL-APCs were manufactured from freshly isolated PBMCs with apheresis for the first administration and cryopreserved PBMCs stored after apheresis for the second and third administrations (Fig. 1d). APCs, which were not cryopreserved, were intradermally injected near the axillary and inguinal regions at a dose of 4 × 10⁷ cells/session.

3.2. Pilot study of WT1-HPL-APCs

A pilot study was conducted to evaluate the safety of three doses of WT1-HPL-APCs in patients with solid tumors and to investigate their potential effectiveness depending on the number of administrations and the impact on the immunosuppressive environment.

Four patients with advanced solid cancers were initially included in the study. However, one patient died of multiorgan failure caused by an infectious disease after providing informed consent. The remaining three patients were enrolled between June 2020 and August 2024. The safety of WT1-HPL-APC vaccination in these patients was evaluated as the primary endpoint. The patients’ characteristics are presented in Table 1.

Table 1.

Characteristics of patients on enrollment at the time of the pilot study.

Case No.	Age (Years)	Sex	Location	Metastases	Stage	Chemotherapy
1	70	F	Pancreas	Liver and Lung	IV	Gemcitabine (GEM) + Nab-paclitaxel (nab-PTX)
2	46	F	Breast	Liver	IV	Eribulin mesylate
3	75	F	Pancreas	Lymph nodes	IV	GEM + nab-PTX

3.3. Detection of WT1 expression in primary cancer for the eligibility criteria

Tumor characteristics were evaluated via IHC on the patient's resected primary cancer tissues. In Case 2, involving a patient diagnosed with breast cancer, the expression levels of WT1, HLA-ABC, and EMA were positive (Fig. 2a). In Case 3, involving a patient diagnosed with pancreatic cancer, the levels of these antigens were also positive (Fig. 2b).

Fig. 2.

Characteristics of primary cancer evaluated via immunohistochemistry (IHC). The hematoxylin and eosin staining and expression levels of WT1, epithelial membrane antigen (EMA), and human leukocyte antigen (HLA)-ABC are illustrated. (a) Breast invasive ductal adenocarcinoma from Case 2. (b) Pancreatic invasive ductal adenocarcinoma from Case 3. Original magnification: ×100 (upper panel) and ×400 (lower panel).

3.4. Safety and clinical outcomes of WT1-HPL-APC vaccination

To ensure the sterility of WT1-HPL-APCs, tests for endotoxin, mycoplasma, and bacteria were conducted, and all lots from patients conformed to the criteria according to the Good Gene, Cellular, and Tissue-based Products Manufacturing Practice in Japan. In Case 1, adverse events such as vagal reflex did not occur during leukapheresis. However, grade 1 skin redness at the injection site was observed (Table 2). Prior to the first administration, the patient's performance status and vital signs were stable, and the complete blood count was within the normal range. No hematological disorder was documented. However, liver function markers, such as alanine aminotransferase, aspartate aminotransferase, and γ-glutamyl transpeptidase, increased from 18, 16, and 19 to 69, 33, and 72 U/L, respectively. The tumor marker cancer antigen (CA) 19-9 increased from 643.4 to 816.1 U/mL. Eight days following the first WT1-HPL-APC administration, bile duct obstruction caused by the progression of pancreatic head cancer with liver metastasis occurred; therefore, vaccination was discontinued in this case before the second administration in accordance with the safety advice of the Committee of Regenerative Medicine.

Table 2.

Adverse events during WT1-HPL-APC administration.

Patient	Adverse events (grade)
Case no.	Skin reaction at the injection site	Fever	Arthralgia	Chills	Headache
1	1	0	0	0	0
2	1	2	1	1	0
3	1	1	0	0	0

The severity of skin reactions at the injection site, fever, arthralgia, chills, and headache was scored based on the Criteria for Adverse Events version 5.0 maintained by the Japan Clinical Oncology Group. Table 2 presents the maximum grades of skin reaction at the injection site observed 48 h following WT1-HPL-APC administration, of fever observed within 48 h of WT1-HPL-APC injection, and of other adverse events during the study. In Case 1, no subsequent administrations were carried out after the first dose due to the discontinuation of this study.

In Case 2, no adverse events, such as vagal reflex, occurred during leukapheresis. However, the patient developed Grade 2 fever within 2 h of administration (Table 2), and an antipyretic analgesic was taken. Grade 1 skin redness at the injection site, arthralgia, and chills occurred within 48 h of administration (Table 2). The CEA and CA15-3 tumor marker levels were 19.4 and 131.4 U/mL before the first administration and 16.9 and 183.8 U/mL 2 weeks after the third administration, respectively. The patient died 8 months following the third administration due to disease progression with metastases.

Case 3 was diagnosed with recurrent pancreatic tail cancer postoperatively. Adverse events, such as vagal reflex, did not occur during leukapheresis. However, the patient developed Grade 1 fever 6 h after administration and skin redness at the injection site (Table 2). Multiple liver metastases, aortic lymph node metastasis, and portal vein infiltration progressed 5 months following the third administration, and the patient died 7 months after the third vaccination due to primary cancer progression with metastases.

3.5. Immunological responses of WT1-CTLs after WT1-HPL-APC vaccination

HLA typing was conducted to determine whether the patient's HLA types were compatible with the WT1 peptides (Table 3). HLA-A*24:02 in Cases 1 and 3 was compatible with WT1-KP, and HLA-A*02:06 in Case 3 was compatible with a killer epitope within WT1-HP. In Case 2, HLA-DPB1*05:01 was also compatible with WT1-HP. However, the HLA-A types in Case 2 were not compatible with WT1-KP. The total WT1-HPL-APC count in Cases 2 and 3 that completed all three sessions was 1.2 × 10⁸ (Table 3).

Table 3.

Immune monitoring for WT1-CTL detection.

Patient	HLA type			Injected WT1-HPL-APCs	Immunological responses
Case no.	A	DPB1	DRB1	Total number of APCs (×107)	ELISpot	WT1-tetramer	DTHc
1	24:02 26:01	02:01 09:01	11:01 15:02	4	NAa	NAa	Negative
2	02:07 31:01	05:01 05:01	08:03 14:03	12	Negative	NAb	Negative
3	02:06 24:02	02:01 04:02	01:01 09:01	12	Negative	Negative	Negative

HLA typing (HLA-A, HLA-DPB1, and HLA-DRB1), the total number of injected WT1-HPL-APCs, and immunological responses after three sessions of WT1-HPL-APC administration. NA, not available.

^aELISpot and WT1-tetramer assays were not conducted due to the discontinuation of this study. In Case 1, only the result of the DTH skin reaction test in the first session is shown.

^bThe WT1-tetramer assay for HLA-A*24:02 was not conducted as the patient's HLA-A type was not 24:02.

^cThe DTH response results induced by WT1-KP and WT1-HP are shown.

Subsequently, to determine whether WT1-CTLs were induced by WT1-HPL-APCs, the presence of WT1-specific CD8⁺ T cells and IFN-γ production from WT1-CTLs exposed to WT1 peptides were examined (Table 3, Fig. S3). ELISpot and WT1-tetramer assays were not conducted in Case 1 as the PBMCs could not be obtained after the third administration. In Case 3, WT1-specific CD8⁺ T cells could not be detected in the WT1-tetramer assay required for the compatible HLA-A*24:02. The WT1-tetramer assay was not conducted in Case 2 due to HLA-A typing incompatibility. In addition, IFN-γ production by WT1-CTLs was not observed in Cases 2 and 3. Furthermore, the DTH skin reaction test with WT1 peptides indicated negative findings 48 h following WT1-HPL-APC administration, and WT1-CTL immune responses were not detected in any of the cases (Table 3). The second and third DTH skin reaction tests were not conducted in Case 1 due to the discontinuation of this study.

3.6. In vitro WT1-CTL inducibility using WT1-HPL-APCs

The potential ability of WT1-HPL-APCs to induce WT1-CTLs in vitro was evaluated after three stimulations. The WT1-CTL induction assay in vitro was performed according to the protocol used to generate WT1-HPL-APCs from PBMCs of enrolled patients with HLA-A*24:02 (Fig. 3a). On day 21, one week after the conclusion of the third WT1-HPL-APC stimulation, the frequency of WT1-CTLs was higher in cocultures containing WT1-HPL-APCs than in cultures of CD8⁺ T cells alone in Case 1 (Fig. 3a; CD8⁺ cells only, 0.05%; WT1-HPL-APC cocultures, 0.11%). The absolute number of CTLs also increased after the third stimulation of WT1-HPL-APCs in Case 1 (Fig. 3b). However, no increase in the frequency or absolute number of WT1-CTLs was observed in Case 3 (Fig. 3a and b).

Fig. 3.

In vitro induction of WT1-CTLs by WT1-HPL-APCs. CD8⁺ T cells were cocultured and stimulated weekly with WT1 peptide-pulsed HPL-APCs, and WT1-CTLs were detected using the WT1-tetramer assay. (a) On days 0 (before the first APC stimulation) and 21 (1 week after the third APC stimulation), WT1-CTLs were identified as CD3⁺/CD8⁺/HLA-A* 24:02 WT1-tetramer⁺ cells. Numbers indicate the ratio of WT1-tetramer⁺ cells in the CD8⁺ T cell population. (b) Graph showing the number of WT1-CTLs on day 21.

3.7. Comparison of the quality of WT1-HPL-APCs between fresh and cryopreserved PBMCs obtained from leukapheresis

Further analysis was conducted to determine whether the cryopreservation of apheresed PBMCs would affect the WT1-HPL-APC phenotype. As no third preparation was performed in Case 1, the phenotype of the third WT1-HPL-APCs in Case 1 was not available for evaluation. The surface molecules defined as maturation markers for APCs are presented in Fig. 4a and b. WT1-HPL-APCs were tentatively defined as phenotypically adequate when ≥70% of the live cells were CD86⁺ and HLA-DR⁺. No significant differences were observed in the expression levels of CD86 and HLA-DR between the cryopreserved and fresh apheresed PBMCs (p = 1.0 for both) (Fig. 4a and b). In all cases, WT1-HPL-APCs derived from cryopreserved PBMCs exhibited lower expression levels of CD80 and CD40 than those derived from fresh apheresed PBMCs (Fig. 4a and b). Furthermore, no clear trends were observed in the expression levels of HLA-ABC and CCR7 between fresh and cryopreserved apheresed PBMCs (Fig. 4a and b). Cell viability ≥ 70% was also defined as a quality criterion for WT1-HPL-APC vaccination. To evaluate the effect of the cryopreservation of apheresed PBMCs on cell viability, WT1-HPL-APCs derived from fresh and cryopreserved PBMCs were compared (Fig. 4c). Consequently, the viability exceeded the predefined criterion in all cases and manufacturing lots, indicating the minimal influence of cryopreservation on cell viability.

Fig. 4.

Maturation profile and cell viability of WT1-HPL-APCs on day 5 of culture, derived from fresh or cryopreserved apheresed PBMCs, for each lot. The expression levels were measured via flow cytometry: (a) red histograms indicate the expression levels of the cells stained with each specific antibody (CD80, CD86, HLA-ABC, HLA-DR, CD40, and CCR7), whereas blue histograms represent the expression levels of the cells stained with the corresponding isotype control antibody. The isotype antibody was used as a negative control. Representative histograms are shown. (b) Expression levels of WT1-HPL-APCs for all administrations. (c) WT1-HPL-APC viability was evaluated using trypan blue to count live and dead cells. In Case 1, the third lot was not assessed due to the discontinuation of this study.

3.8. Cytokine production ability of WT1-HPL-APCs

To evaluate the functional characteristics of HPL-APC vaccines, the secretion of IL-6 and TNF-α, inflammatory cytokines; IL-12 (p70) and IFN-γ, essential for CTL responses [12,13]; and IL-10, immunosuppressive in the culture supernatant of the WT1-HPL-APC vaccine, were measured in all cases (Fig. 5). Furthermore, these cytokine levels were compared with those of the WT1-HPL-APC vaccine, PCA-APC, a PCA-based protocol that has successfully induced PCA-specific CTLs [36]. WT1-HPL-APCs (WT1) induced IL-6 (1472.35 ± 452.42 pg/mL), IL-10 (126.33 ± 17.50 pg/mL), IL-12 (p70) (7.47 ± 0.44 pg/mL), IFN-γ (86.82 ± 12.55 pg/mL), and TNF-α (779.83 ± 161.18 pg/mL). Meanwhile, IL-6 (1617.37 ± 524.28 pg/mL), IL-10 (176.03 ± 14.13 pg/mL), IL-12 (p70) (6.6 ± 0.73 pg/mL), IFN-γ (41.58 ± 14.67 pg/mL), and TNF-α (991.45 ± 160.14 pg/mL) were detected from PCA. The five measured cytokines showed cytokine profiles similar to those of the control vaccine.

Fig. 5.

Cytokine production profile of WT1-HPL-APCs. The levels of IL-6, TNF-α, IL-12 (p70), IFN-γ, and IL-10 were measured via magnetic bead-based multiplex assays. WT1-APC shows the value of each cytokine from WT1-HPL-APCs in Cases 1, 2, and 3. The assay was conducted in duplicate. Each dot represents the average value for each case. WT1 and PCA indicate WT1-HPL-APCs and PCA-APCs, respectively. The bold horizontal bars in the graphs show the median of each parameter.

3.9. TME of primary cancer infiltrated with immune cells before WT1-HPL-APC vaccination

A secondary examination was conducted to determine whether the TME and immune cell populations in the primary cancer tissue influence WT1-CTL induction prior to WT1-HPL-APC vaccination. IHC was conducted to investigate the proportion of immune cells, such as CD4⁺ T cells, CD8⁺ T cells, and macrophages, in the TME of primary cancer tissues (Fig. 6). In Case 2, CD4⁺ T and CD8⁺ T cells infiltrated the outer layer of the cancer ducts, whereas macrophages accumulated inside them (Fig. 6a). By contrast, few CD4⁺ T cells, CD8⁺ T cells, and macrophages in the TME were observed in Case 3 (Fig. 6b).

Fig. 6.

IHC analysis of the tumor microenvironment in primary cancer. Anti-CD4, anti-CD8, and anti-CD68 antibodies were used to detect CD4⁺ T cells, CD8⁺ T cells, and macrophages, respectively. (a) Breast invasive ductal adenocarcinoma in Case 2. (b) Pancreatic invasive ductal adenocarcinoma in Case 3. Original magnification: ×100 (upper panel) and ×400 (lower panel).

To determine whether WT1-HPL-APC vaccination would influence peripheral immunosuppressive cells, the percentages of Tregs and M-MDSCs were determined in the PB (Fig. 7). The percentage of Tregs slightly increased 2 weeks after WT1-HPL-APC vaccination in Cases 2 (8.6%–9.6%) and 3 (6.1%–7.6%) (Fig. 7a and c). Conversely, the percentages of M-MDSCs decreased in Cases 2 (18%–9.3%) and 3 (2.2%–0.011%) (Fig. 7b and c).

Fig. 7.

Comparison of the proportion of immunosuppressive cells in PBMCs before and after the third WT1-HPL-APC vaccination performed via flow cytometry: (a) flow cytometric analysis of regulatory T cells (Tregs) in CD4⁺ T cells; (b) flow cytometric analysis of monocytic myeloid-derived suppressor cells (M-MDSCs) in CD11b⁺ CD33⁺ cells; (c) ratios of Tregs in CD4⁺ T cells and M-MDSCs in CD11b⁺ CD33⁺ cells.

4. Discussion

In this study, we utilized WT1-HPL-APCs rather than WT1-IL-4-DCs, based on our preclinical in vitro evaluation (Fig. 1a and b). We previously reported that HPL-APCs exhibited stronger ability to induce CTLs than IFN-DCs [35]. Furthermore, our previous study demonstrated that 12 × 10⁷ HPL-APCs could have a potency to induce PCA-specific CTLs in a clinical setting [36], the same amount used in this study. It has also been reported that CTLs can be induced by administering 2.5–5 x 10⁷ cells/session three times biweekly [37]. Based on these findings, 4 x 10⁷ WT1-HPL-APCs/session was proposed to enhance WT1-CTL induction over three sessions, aiming to elicit a tertiary immune response as a general protocol of DC vaccination in this study.

In addition, the safety, tolerability, and effectiveness of WT1-HPL-APC administration were explored in patients with advanced solid cancers (Fig. 1c and d, Table 1). The three-time biweekly administration of 4 × 10⁷ WT1-HPL-APCs/session did not cause any severe adverse events during the pilot study (Table 2). Although long-lasting WT1-CTL responses were anticipated from WT1-HPL-APC vaccination, WT1-CTL responses could not be detected in this pilot clinical study (Table 3). The possible reasons may be associated with the dosage schedule (Fig. 1c), WT1 expression in primary cancer (Fig. 2), individual differences and heterogeneity in the ability of WT1-HPL-APCs to induce WT1-CTLs (Fig. 3), expression of cell surface protein on the WT1-HPL-APCs (Fig. 4), cytokine productive ability of the WT1-HPL-APCs (Fig. 5), infiltration of immune cells (Fig. 6), and presence of immunosuppressive cells, such as Tregs and MDSCs, in the TME and blood, which could have influenced WT1-CTL induction in enrolled cases (Fig. 7). Moreover, as this is a safety study involving a small number of cases, the interpretation of the results is limited. This phase I pilot study included three heterogeneous cancer cases, primarily addressing the safety of HPL-APC vaccines that were considered sufficient in terms of the estimated functionality based on the phenotype and cytokine levels [36]. Therefore, further studies involving a larger number of patients with more uniform disease backgrounds are warranted to validate these preliminary observations and examine the therapeutic potential of HPL-APCs.

In the assessment of the adverse reactions of WT1-HPL-APC administration, side effects lower than grade 2, such as skin reactions at the injection site, fever, arthralgia, and chills, occurred within 48 h of WT1-HPL-APC administration (Table 2). These events, such as the flulike reaction, also occur in DC-based cancer immunotherapy [43], and the incidence of these side effects has been reported to be positively correlated with prolonged OS [44]. For these reasons, these mild side effects were considered tolerable and have the potential to prolong OS.

Our prior research using the same HPL-APCs in six sessions (2–3 × 10⁷ cells/session) demonstrated the induction of antigen-specific T cells [36]. However, WT1-CTLs could not be detected in Cases 2 and 3 in Table 3, therefore, the dosing schedule consisted of three biweekly sessions might influence the WT1-CTL levels. In another clinical trial, a three-time biweekly administration of 2.5–5 × 10⁷ cells/session resulted in a positive DTH skin reaction, antigen-specific T-cell induction, and tumor shrinkage [37]. Thus, our study requires further protocol revision for the potential induction of WT1-CTLs.

WT1-HPL-APC therapy targets tumor cells expressing the WT1 antigen on MHC class I, which are attacked by WT1-CTLs. In this study, the expression levels of WT1 and HLA-ABC in the primary cancer tissue were evaluated via IHC. Fig. 2 shows that their expression patterns in Case 2 were quite different from those in Case 3. In Case 3, the low WT1 expression level could be related to either a few clones being stained with anti-WT1 antibody or the undifferentiated state of the tumor cells. A previous study reported that 65% of pancreatic cancers were positive for the anti-WT1 antibody with the 6F-H2 clone, which was used in this study, whereas pancreatic cancers with undifferentiated states were not stained by this antibody [45]. The primary cancer in Case 3 might suggest an undifferentiated state. In a previous study on WT1-pulsed DC-based vaccine therapy, WT1 expression in a pancreatic cancer tissue was positively correlated with the level of acquired immunity [43]. Therefore, the low expression of WT1 may have influenced WT1-CTL induction in Case 3. WT1 antigen expression was also reported in breast cancer [46,47]. Thus, assessing the expression levels of the WT1 antigen in primary cancer seems to be a rational approach to determining the eligibility criteria for targeting WT1 in clinical trials.

We conducted an in vitro autologous WT1-CTL induction assay using WT1-HPL-APCs from enrolled patients harboring HLA-A*24:02 to assess whether WT1-HPL-APCs retained the functional capacity to stimulate antigen-specific T-cells while also validating the HPL-APC manufacturing process. WT1-CTLs could were detected in Case 1, but not in Case 3 (Fig. 3), suggesting that the inducibility of anti-tumor immunity depends on the characteristics and diversity of individual WT1 cancer vaccines as well as patient-specific factors, including the presence of an immunosuppressive environment.

The establishment of quality criteria is crucial for verifying DC-based vaccines [48]; thus, we tentatively proposed a standard for WT1-HPL-APCs, defining them as phenotypically adequate when ≥70% of live cells express both CD86 and HLA-DR. However, the expressions of these markers vary by case and manufacturing lot, as illustrated in Fig. 4b. Previous studies have demonstrated that immunosuppressive cells, including Tregs and MDSCs, reduced CD86 expression on induced APCs in vitro [49,50]. As a similar reduction in CD86 expression was observed in Case 2, we presumed that the presence of immunosuppressive cells, including Tregs and M-MDSCs, in both the TME and PB negatively influenced the quality of WT1-HPL-APCs and induction of WT1-CTLs. M-MDSCs and Tregs were detected in the PB of Case 2 prior to WT1-HPL-APC vaccination (Fig. 7a–c), supporting the possibility that immunosuppressive cells contributed to the reduction in CD86 expression on WT1-HPL-APCs. Furthermore, WT1-HPL-APCs derived from cryopreserved PBMCs exhibited a trend toward reduced CD80 and CD40 expressions in all three cases (Fig. 4b). Previous reports have suggested that cryopreservation and thawing of PBMCs may reduce the proportion of CD80⁺ mature DCs [51]. To optimize vaccine manufacturing using cryopreserved cells, we need to evaluate cell damage during freezing and reduce stress during the thawing process. Specifically, a controlled-rate freezer and thawing at a validated temperature are required. Moreover, the addition of agents, such as TNF-α or TLR ligands, enhances the maturation process of APCs [52,53]. However, given the small sample size, whether the cryopreservation and thawing of PBMCs influenced CD80 and CD40 expressions remains to be elucidated through further detailed studies. Therefore, whether cryopreservation of apheresed PBMCs affects the phenotypes, cytokine production, and CTL-inducing capability of WT1-HPL-APCs remains an important question that should be addressed in future studies involving more patients.

HPL-APC vaccines demonstrate higher IL-6 and TNF-α levels than other cytokines, such as IL-10, IL-12 (p70), and IFN-γ [35]. IL-12 (p70) and IFN-γ, which are important in promoting Th1 differentiation and enhancing CTL responses [12,13], were detected in WT1-HPL-APC vaccines that also exhibited the same IL-6 and TNF-α levels, as previously reported [35] (Fig. 5). In addition, the cytokine levels of WT1-HPL-APC vaccines were similar to those observed in the PCA protocol that successfully induced personalized cancer antigen-specific CTLs [36]. These findings suggest that the WT1-HPL-APC vaccines had characteristics and functionalities that enable the induction of antigen-specific CTLs.

The patients’ immunosuppressive condition caused by advanced metastatic disease may also lead to failed WT1-CTL induction. Previous studies have reported that lymph node metastasis led to an immunosuppressive environment in the lymph nodes [54] and that liver metastasis tended to inhibit CTL induction and DTH reaction [55]. Although WT1-HPL-APCs were injected close to the lymph nodes to facilitate WT1 antigen presentation to the naïve T cells in these nodes, it may be more challenging to induce CTLs with only three sessions of WT1-HPL-APC administration following the previous protocol due to immunosuppressive conditions, such as lymph node or liver metastasis (Table 1).

The potency of the WT1-HPL-APC vaccine to induce WT1-specific CTLs may be influenced by factors other than the immunosuppressive environment, such as the individual variability of vaccines. It has been reported that coadministration of adjuvants, such as OK-432 or Th1 agonists, with IL-4-DC vaccines enhances CTL induction and antitumor responses in vivo [56]. Therefore, it is conceivable that the use of optimized adjuvants with WT1-HPL-APC vaccines could also be expected to ameliorate failed WT1-specific CTL induction. Thus, similar effects could be possibly achieved with the WT1-HPL-APC vaccine. IHC analysis of the TME in primary cancer revealed that immune cells, such as T cells and macrophages, infiltrated the primary cancer prior to WT1-HPL-APC vaccination in Case 2 (Fig. 6a) and were scarcely detected in Case 3 (Fig. 6b). By evaluating circulating Tregs and M-MDSCs in the PB, we also demonstrated that the M-MDSC population decreased after the third administration in Cases 2 and 3. The percentages of Tregs and M-MDSCs before vaccination in Case 2 (Tregs: 8.6% [Fig. 7a and c]; M-MDSCs: 18% [Fig. 7b and c]) were higher than those in healthy donors (Tregs: 6.2%–8.5% [Fig. S4]; M-MDSCs: <1% [Fig. S4]). These findings suggest that the immune cells in the primary cancer of Case 2 are composed of immunosuppressive cells (Fig. 6a), such as M2-type tumor-associated macrophages and Tregs [57,58]. The immunosuppressive cells detected in Case 2 may be resistant to the activation, proliferation, and induction of WT1-CTLs. In Case 3, the percentages of Tregs and M-MDSCs in the PB were nearly the same as those in healthy donors (Fig. S4). Thus, in addition to Tregs and M-MDSCs, the broader immunosuppressive condition may also contribute to interference with WT1-CTL generation in Case 3. Despite the failed WT1-CTL induction, M-MDSC reduction may have ameliorated the recovery of immunosuppressive conditions, such as lymphocyte activation, during the three sessions of WT1-HPL-APC administration, as HPL-APCs have the potential to induce IFN-γ production by antigen-specific CTLs in a clinical setting (Fig. S5). It has been reported that M-MDSCs inhibit the functions of DCs and T cells [33] and contribute to tumor progression [59]. Although M-MDSCs tended to decrease following the third WT1-HPL-APC administration, it remains unclear whether the combined chemotherapy directly influenced this reduction. Certain chemotherapeutic agents, such as gemcitabine (GEM), have been reported to be effective in suppressing MDSCs [60]. GEM can also enhance the suppressive functions of M-MDSCs [61]. In the current study, the ratio of M-MDSC in patients with cancer was higher than that in healthy donors before vaccination, and a decreasing trend in M-MDSCs was observed with the addition of WT1-HPL-APC vaccination during the continuation of chemotherapy (Fig. 7b and c). Therefore, the additive effects of WT1-HPL-APC vaccination may be beneficial in reducing M-MDSCs. Consequently, the selection of chemotherapy or drug therapy to be combined with WT1-HPL-APC administration must be made carefully. Overall, the observed decrease in M-MDSCs in the PB following WT1-HPL-APC administration in two cases, combined with chemotherapy, may help alleviate immune suppression. This suggests the potential to convert a “cold tumor” into a “hot tumor,” enabling the generation of WT1-CTLs through additional administration.

Our findings suggest that, for future clinical applications, it is crucial to evaluate the presence of immunosuppressive cells, such as M-MDSCs and Tregs, prior to patient enrollment. To enhance the efficacy of the WT1-HPL-APC vaccine, its combination with anticancer agents and the use of drugs targeting immunosuppressive cells should be considered. Furthermore, a revised dosing protocol, administration of at least 2 × 10⁷ cells for a minimum of six times, may be more effective than the current regimen [36, Fig. S5]. These improvements may increase the likelihood of inducing WT1-CTLs in future large-scale clinical studies.

5. Conclusions

This pilot study of the WT1-HPL-APC vaccine in patients with advanced solid cancers demonstrated that the vaccine is tolerable, with no severe adverse events observed. In our clinical study, a WT1-CTL response could not be detected after three WT1-HPL-APC administrations, which might be due to individual heterogeneity in the immunoinducibility of WT1-HPL-APCs. However, a decrease in M-MDSCs was found after WT1-HPL-APC vaccination with chemotherapy in two cases, suggesting that WT1-HPL-APC vaccination modulates the immunosuppressive environment. Low CD80 expression on WT1-HPL-APCs due to cryopreserved apheresed PBMCs, as well as the widespread presence of immunosuppressive cells in the PB, appears to influence WT1-CTL induction. Although the administered WT1-HPL-APC vaccine demonstrated sufficient cytokine-producing functionality, its clinical effectiveness could not be determined. The safety of HPL-APC vaccines was confirmed in three patients with different cancer types in this study. Accordingly, further studies are warranted to evaluate the effectiveness of HPL-APCs by both performing an individualized CTL induction assay and validating the quality of HPL-APCs. In the future, improvement of the immunosuppressive condition before WT1-HPL-APC vaccination could lead to the validation of the effective induction of WT1-CTLs in a large-scale clinical trial.

Informed consent statement

Written informed consent was obtained from all participants involved in this study. An accompanying study on HPL-WT1-APC therapy was also conducted after receiving the appropriate informed consent.

Author contributions

Conceptualization, S.S., TJ.Y., YC.L., and S.K.; methodology, S.S. and TJ.Y.; preparation for pilot study, M.T., T.K., K.Y., and H.K.; securing raw material through apheresis sessions, T.T. and S.S.; manufacturing of WT1-HPL-APCs, T.K. and H.K.; investigation, M.T., T.K., K.Y., H.S., and S.K.; formal analysis and validation, M.T., T.K., K.Y., S.Y., TJ.Y., and YC.L.; resources, S.S.; data curation, K.Y. and M.T.; writing—original draft preparation, S.S. and M.T.; writing—review and editing, M.T., T.K., K.Y., S.Y., S.S., TJ.Y., YC.L., S.K., and H.S.; visualization, M.T., T.K., and K.Y.; supervision, S.S. and H.S.; project administration, S.S. and S.K.; funding acquisition, M.T., T.K., and S.S. All authors have read and agreed to the published version of the manuscript.

Institutional review board statement

This study was conducted in accordance with the Declaration of Helsinki and approved by the Kanazawa Medical University Certified Committee for Regenerative Medicine (Class III technologies; approval no. of the committee NB4150006; approval no. PC006) on January 30, 2020. This project was also registered in the Japan Registry of Clinical Trials (project no. jRCTc040200005). Preclinical study in vitro evaluation and an accompanying study to monitor the phenotype of WT1-HPL-APCs and the immunological responses over three sessions and after administration was conducted in accordance with the Declaration of Helsinki and approved by the Ethical Committee of Kanazawa Medical University (approval No. G156) on June 10, 2020.

Data availability statement

All data indicated are available in this article.

Declaration of generative AI and AI-assisted technologies in the manuscript preparation process

During the preparation of this work, the authors used ChatGPT to assist with English language editing. The content was subsequently reviewed and revised by the authors, who take full responsibility for the content of the published article.

Funding

This work was supported by the ALP Institute for Regenerative Medicine Inc.; the Program on Open Innovation Platform with Enterprises, Research Institutes, and Academia of the JST under grant number JPMJOP1722; and Academia of the JSPS KAKENHI under grant numbers JP23K06727, JP25K15878, and JP25K10597.

Declaration of competing interest

S.S., T.K., and M.T. are patent holders for a method for manufacturing human platelet lysate-inducible interferon (IFN)-DCs (PCT/JP/2021/040505). S.S., T.K., and M.T. evaluated DCO-K medium (Shimadzu Diagnostics Corp.) as an HPL-APC induction medium. Kanazawa Medical University has a collaborative investigation agreement with Nissui Pharmaceutical Co., Ltd. (which became a subsidiary of Shimadzu Diagnostics Corp. in 2022). H.S. is the inventor of the WT1 patent (PCT/JP2010/057149 and PCT/JP2006/323827). The funders and H.K. played no role in the study design, data collection, analysis, or interpretation, manuscript writing, or decision to publish the results. This study was conducted based on a joint research agreement between Kanazawa Medical University and Kaohsiung Medical University signed on February 1, 2023.

Appendix A. Supplementary data

The following are the Supplementary data to this article:

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