Over the past 30 years, the therapeutic outcome for pancreatic ductal adenocarcinoma (PDAC) has remained stagnant due to the lack of effective treatments. We performed a genome-wide analysis to identify novel therapeutic targets for PDAC. Our analysis showed that Homo sapiens chromosome 16 open reading frame 74 (C16orf74) was upregulated in most patients with PDAC and associated with poor prognosis. Previously, we demonstrated that C16orf74 interacts with the catalytic subunit alpha of protein phosphatase 3 and plays an important role in PDAC invasion. However, the pathophysiologic function of C16orf74 is still unclear. In this study, through the analysis of C16orf74 interaction, we demonstrate a new strategy to inhibit the growth and invasion of PDAC. C16orf74 exists in the homodimer form under the cell membrane and binds integrin αVβ3 and is also involved in invasion by activating Rho family (Rac1) and MMP2. Considering that this dimeric form was found to be involved in the function of C16orf74, we designed an 11R-DB (dimer block) cell-permeable dominant-negative peptide that inhibits the dimer form of C16orf74. 11R-DB suppressed invasion and proliferation of PDAC cell lines by inhibiting phosphorylation of Akt and mTOR and also by inactivation of MMP2. 11R-DB also showed antitumor effects in an orthotopic xenograft model and peritoneal metastasis model. Thus, this study demonstrates that dimerized C16orf74, present in the cell membrane, is involved in pancreatic cancer invasion and proliferation. In addition, the C16orf74 dimer block cell-permeable peptide (11R-DB) has a potent therapeutic effect on PDAC in vitro and in vivo.

The incidence of pancreatic ductal adenocarcinoma (PDAC) is increasing by about 1% per year, and an estimated 56,770 new cases and 45,750 deaths occurred in the United States in 2019, even today, with progress in therapy, the 5-year combined survival rate of all stages is only 9% (American Cancer Society. Cancer Facts & Figures 2019. Atlanta, GA: American Cancer Society 2019; http://www.cancer.org/research/cancerfactsstatistics/cancer-facts-and-figures-2019). In recent years, FOLFIRINOX (a combination of oxaliplatin, irinotecan, fluorouracil, and leucovorin) and gemcitabine with nab-paclitaxel therapy have been developed and show higher antitumor effect than gemcitabine for patients with metastatic PDAC. Conversely, it has also been reported that various adverse effects appear in the chemotherapy (1–3). In addition, gemcitabine with erlotinib was developed as a molecular target therapy; however, it did not have a significant effect compared with gemcitabine single therapy (only 2 weeks of extension in median overall survival; ref. 4). Therefore, development of new molecular target therapy with strong therapeutic effect on PDAC is essential.

Through a genome-wide cDNA microarray analysis of PDACs, we have previously selected genes that are upregulated highly and commonly in PDAC compared with normal tissues (5). Among the upregulated genes, we have reported a novel gene C16orf74 [Homo sapiens chromosome 16 open reading frame 74 (NM_206967)] as an ideal therapeutic target of PDAC. C16orf74 is frequently overexpressed in PDAC specimens associated with poor prognosis and was found to be essential to growth of PDAC (6). Moreover, other reports have indicated that C16orf74 expression correlated with potential prognostic factor in other types of cancer (7–10). With the functional analysis of C16orf74, previously, we have demonstrated that C16orf74 interacts with the catalytic subunit alpha of protein phosphatase 3 (PPP3CA) as well as the calcineurin (CN) isozyme. This interaction promotes proliferation and invasion of PDAC cells (6, 11). However, the pathophysiologic functional analysis of C16orf74 is unclear.

In this study, we further analyzed the functional characteristics of C16orf74 and found a new strategy to inhibit the growth and invasion of PDAC.

Cell-permeable peptide design

Three cell-permeable peptides were synthesized by Sigma-Aldrich Japan and were purified by preparative reverse-phase high-performance liquid chromatography till they were >80% pure, with the expected amino acid composition and mass spectra. The peptides' amino acid sequences are shown below.

11R-DB; RRRRRRRRRRR-GGG-MGLKMSCLKGFQMCV

11R-C7A_C14A; RRRRRRRRRRR-GGG-MGLKMSALKGFQMAV

11R-7_14AAA; RRRRRRRRRRR-GGG-MGLKMAAAKGFQAAA

Cell lines

The cell lines: PANC-1, MIA PaCa-2, the embryonic kidney, HEK293, and COS7 were purchased from the ATCC. PK-1 and KLM-1 were purchased from RIKEN Bio Resource Center. PK-9 cells were provided by the Cell Resource Center for Biomedical Research, Tohoku University (Sendai, Japan). PCI6 was previously obtained from a patient with PDAC in our laboratory (12). Human umbilical vein endothelial cells (HUVEC) and normal human dermal fibroblasts (NHDF) were purchased from PromoCell. All cells were cultured in appropriate media and were incubated at 37°C with 5% CO2. DMEM (Nacalai Tesque) was used for MIA PaCa-2, PANC-1, NHDF, and COS7. RPMI1640 cell media (Nacalai Tesque) was used for PK-1, PK-9, KLM-1, and PCI6, whereas endothelial cell media (PromoCell) was used for HUVECs. Each medium was supplemented with 10% FBS (Cell Culture Bioscience) and 1% penicillin–streptomycin (Life Technologies).

Quantitative RT-PCR analysis

Total RNA from cells was extracted using an RNeasy Plus Mini Kit (QIAGEN, Hilden, Germany) according to the manufacturer's protocol. cDNA was synthesized using Prime Script RT Master Mix (TAKARA BIO). Using a StepOne Real-time PCR system (Applied Biosystems) and Fast SYBR Green Master Mix (Life Technologies), PCR reactions were performed. Data analysis was performed using the ΔΔCt method, and the housekeeping gene, GAPDH, was used as control. Primer specificity was confirmed by one peak melt curve. All experiments were performed in triplicates for each sample. The primers used in this study are described in Supplementary Table S1.

Western blot analysis

After being washed twice with PBS, cells were harvested in RIPA buffer containing protease inhibitors (aprotinin and PMSF). The protein concentration was standardized by Bradford assay after the cells were homogenized. Proteins were separated by 15% SDS-PAGE and transferred to polyvinylidene difluoride membranes (Millipore) and blocked with 5% skim milk or MaxBlot Solution 1 (MBL) for 1 hour. Membranes were then incubated with the desired primary antibodies for what was being tested. Membranes were probed with the secondary antibodies (1:10,000 dilution) using goat anti-mouse or rabbit IgG. Immunoreactivity was detected with an enhanced chemiluminescence detection system (GE Healthcare). β-Actin was used for equal loading confirmation. Primary antibodies and dilutions are shown in Supplementary Table S2. Secondary antibodies were purchased from Jackson ImmunoResearch.

Native PAGE

The cultured cells were harvested with RIPA buffer without SDS. PAGE was performed similar to as described above but at 4°C and omitting SDS (in all solutions), 2-mercaptoethanol, and sample boiling.

Establishment of PANC-1 cells stably expressing C16orf74-FLAG

PANC-1 cells were plated at a density of 1 × 105 cells/well on 6-well plates and transfected with pCAGGS-C16prf74-3xFLAG using Lipofectamine LTX Reagent and Plus Reagent (Invitrogen), according to the manufacturer's protocol. The medium was then replaced with selection medium containing G418 (800 μg/mL; Invitrogen). The resulting live-cell colonies were plated at a density of 1 cell/well on 96-well plates, and single-cell–derived clones were obtained. Resulting clones were confirmed by immunofluorescence, qRT-PCR, and Western blotting.

Establishment of PK-9 cells stably expressing tdTomato-Luc2

PK-9 cells were transfected with pCSII-CMV-tdTomato-Luc2 (provided by Vascular Biology and Molecular Pathology Graduate School of Dental Medicine, Hokkaido University, Sapporo, Japan), and the clones were selected for tdTomato-Luc expression using 0.1 mg/mL bleomycin. The resulting colonies were plated at a density of 1 cell/well on 96-well plates and single-cell–derived clones were obtained. The expression of luciferase in these cells was confirmed via the in vivo optical imaging showing the quantitative correlation between the cell number and luminescent signal intensity (Promega).

Immunofluorescence

Cells were fixed with 3% paraformaldehyde for 15 minutes at room temperature and permeabilized with 0.1% Triton X-100 in PBS for 4 minutes. To block nonspecific binding of antibodies, cells were incubated in 1% BSA and then further incubated with primary antibodies overnight at 4°C, after which immune complexes were detected by incubation for 1 hour at room temperature in the dark with secondary antibodies (1:250 dilution). FV-10i confocal microscope (Olympus) was used for imaging. The names and dilutions of the antibodies have been listed in Supplementary Table S3.

Immunoprecipitation and Western blotting (pull-down assay)

After transfection 24 hours, cells were harvested with RIPA buffer. Immunoprecipitation was performed with mouse anti-DDDDK(FLAG) tag (FLA-1, MBL) antibody or anti-HA tag (TANA2, MBL) antibody. By incubation with protein A agarose (sc-2001, Santa Cruz Biotechnology), the antibodies were removed, and the wash step was repeated five times. Proteins were extracted with SDS sample buffer and separated on a 4% to 20% gradient SDS page (Bio-Rad). To examine the interaction between flag-tagged proteins and HA-tagged proteins, immune complexes were analyzed by Western blotting. To investigate the interactions between C16orf74 and integrin β3 as well as integrin αV and integrin αIIb, PANC-1 cells were harvested in RIPA buffer and mouse anti-DDDDK (FLAG) immunoprecipitation was performed using a tag antibody. Then, antibodies containing protein A agarose: sc-2001 (Santa Cruz Biotechnology) were collected and washed with PBS five times. Proteins were extracted with SDS sample buffer and separated by 4% to 20% gradient SDS PAGE. Immunocomplexes were analyzed by Western blotting using mouse antibodies (Supplementary Table S2).

Rac1 isolation

Rac1 isolation was performed using PAK-1 PBD agarose beads. PANC-1 cells stably expressing C16orf74 and MOCK (stable expressing empty vector) cells were lysed with MLB (magnesium-containing lysis buffer; 25 mmol/L HEPES pH 7.5, 150 mmol/L NaCl, 1% lagepal CA-630, 10 mmol/L MgCl2, 1 mmol/L EDTA, 10% glycerol, EMD Millipore). PAK-1 PBD agarose (EMD Millipore) was added to each lysate and agitated for 1 hour at 4°C. The beads were collected by centrifugation and the supernatant was discarded. The beads were then washed three times with MLB.

IHC staining

Paraffin-embedded sections were used for IHC staining. Same protocol was followed as our previous study (11). Finally, the sections were counterstained with hematoxylin. Each antibody was shown in Supplementary Table S4.

Cell proliferation assay

The water-soluble tetrazolium salts (WST) cell proliferation assay was performed according to the manufacturer's protocol provided with the cell counting kit-8 (Dojindo). Briefly, cells were seeded on 96-well plates and the culture medium was replaced with various conditioned medium for 24 or 48 hours. At the end of treatment, the WST-8 reagent was added and incubated for another 4 hours. Finally, the plate was directly measured for absorbance at 450 nm.

Matrigel invasion assay

Same protocol was followed as our previous study (11). At least three randomly selected fields were investigated and invaded cells were counted at ×100 magnifications.

Hematogenic invasion assay

A total of 2.5 × 104 HUVECs were seeded on the surface of a BD Matrigel Matrix (BD BioCoat 24-Multiwell Tumor Cell Invasion System) in Endothelial Cell media (PromoCell) to cover the top side of the Transwell inserts (8-μm-pore size), and incubated at 37°C, 5% CO2 for 1 day. After the confluence of HUVECs and monolayer formation, the rehydration solution was removed. A total of 2.5 × 104 stable GFP-expressing C16orf74 stable (C16orf74 stable/GFP) and MOCK (MOCK/GFP) cells were resuspended in 0.5 mL of medium (DMDM:endothelial cell media = 1:1) without FBS and loaded on the top side of the Transwell inserts, and 0.75 mL medium (DMDM:endothelial cell media = 1:1) containing 10% FBS was loaded on the bottom side of the Transwell inserts to induce chemotaxis. The remaining cells on the top side of the Transwell inserts were removed by cell scraping after incubation for 24 hours at 37°C and 5% CO2. The cells permeated through the HUVECs and Transwell inserts were counted by a fluorescence microscope (BZ-9000, KEYENCE). Cells were counted in full visual field (×100) per filter. The results were reported as means of triplicate assays.

Wound-healing assay

Confluent cells were wounded by scraping with a 200 μL pipette tip to analyze cell motility. By phase-contrast microscopy, cell movements were observed at 0, 3, 6, 9, 12, 15, and 18 hours.

Gelatin zymography

Cells were cultured in serum-free medium for 24 hours and then washed twice with PBS. The conditioned medium was clarified by centrifugation at 2,000 × g for 15 minutes. Twenty microliters of 2× nondenaturing loading buffer was mixed with an equal volume of conditioned medium, and separated by 10% SDS-PAGE containing 25 mg/mL gelatin. To remove SDS, gels were washed twice in 2.5% TritonX-100 PBS for 15 minutes at room temperature, and then twice in water, and incubated at 37°C for 24 hours in an incubation buffer (50 mmol/L Tris, 5 mmol/L CaCl2⋅H2O, 1 mmol/L ZnCl2). The resulting gel was stained, fixed with a 50% methanol and 10% acetic acid containing 1.25 mg/mL Coomassie Brilliant Blue for 30 minutes, and destained with 10% methanol and 5% acetic acid. The zymogram was imaged with ChemiDocTMXRS (Bio-Rad) and the intensity of the band corresponding to gelatinase activity was quantified using Image Laboratory software (Bio-Rad).

Construction of the expression vector

The entire coding sequence of each construct was amplified by RT-PCR and the PCR product was inserted into the EcoRI and XhoI sites of pCAGGS to express Flag- or HA-tagged protein (13). RT-PCR primers were shown in Supplementary Table S5.

Pancreatic cancer model mice and in vivo imaging

Female Balb/cA Jcl nu/nu mice were purchased from CLEA Japan. At 6 weeks of age, each of the following models was constructed in the mice under anesthesia induced with ketamine (100 mg/kg) and xylazine (10 mg/kg).

Subcutaneous tumor

PK-9 cells were adjusted to 5 × 106 cells/100 μL Hank balanced salt solution (HBSS, Gibco) containing Matrigel (Corning) and injected subcutaneously in the back using a 27 G needle and 100 μL Hamilton syringe. From 2 weeks after injection of the cancer cells, 11R-DB and a111R-7_14AAA (10 mg/kg, 5 times per week) or PBS only were administered in the subcutaneous tissue near the tumor for 3 weeks.

Orthotopic xenograft model mice

PK-9 cells expressing luciferase (PK-9-tdTomato-luc2) were adjusted to 1 × 106 cells/50 μL HBSS containing Matrigel (Corning) and injected below the serosa of the pancreas tail using a 27 G needle and 100 μL Hamilton syringe (14). One week postinjection, 11R-DB and 11R-7_14AAA (10 mg/kg, 5 times per week) or PBS were administered intraperitoneally for 4 weeks. Using a postintraperitoneal injection of VivoGlo Luciferin in vivo Grade (Promega), bioluminescent imaging was performed with the IVIS Spectrum imaging system (Caliper Life Sciences). Mice were then sacrificed to assess tumor weight and pathologic analysis of tumor and major organs such as, brain, lung, heart, liver, kidney, and spleen.

Peritoneal dissemination model mice

PK-9-tdTomato-luc2 were adjusted to 3 × 106 cells/200 μL HBSS and injected intraperitoneally using a 27 G needle and 1,000 μL syringe. From the day of transplantation of tumor cells until day 5, 11R-DB and 11R-7_14AAA (1 mg/kg, 10 μmol/L) or PBS only were intraperitoneally administered. The mice were analyzed by in vivo imaging on days 3, 7, 14, 21, and 28 after tumor cell implantation and engraftment of the tumor was confirmed.

All animal experiments were conducted according to the guidelines of the Hokkaido University Animal Experiment Implementation Manual and were approved by the Institutional Animal Care and Use Committee of Hokkaido University Graduate School of Medicine (no:17-0011). On the basis of their completion of required animal use and care training, all researchers who performed procedures using live animal were preapproved by the Animal Welfare Committee of Hokkaido University (Sapporo, Japan).

Statistical analysis

The results are shown as the mean ± SEM or SD. Student t test or Mann–Whitney U test was performed to analyze significant differences using JMP version 13 (SAS Institute, Inc.). P < 0.05 was considered statistically significant.

Functional analysis of C16orf74-overexpressed stable transfectant cell line

The morphologic observation of C16orf74 stable transfectant (Red: f actin, Green: paxillin), compared with MOCK stable transfectant demonstrated a strong polarity with cell membrane development in the C16orf74-stable cells (Fig. 1A). From the above observation, we speculated that the Rho family such as lamellipodium and stress fiber would be activated by C16orf74 overexpression. Active Rac1 (Rac1-GTP) was pulled down with GST-PAK-PBD–conjugated agarose beads from extracts of C16orf74 stable and MOCK cells and detected by Immunoblotting. Activated Rac1 was increased in C16orf74-stable cells as compared with MOCK cells (Fig. 1B).

Figure 1.

Functional analysis of C16orf74 overexpressed stable transfectant cell line. A, Observation of cell morphology of C16orf74 stable (A-1) and MOCK cells (A-2) by immunofluorescence (red: F actin; green: paxillin). i, The polarity of the cell was strong, and the direction of cell movement (arrow) was clear. The development of lamellipodium was strong. ii, Filopodia was developing in the direction of cell progression. The development of actin was strong. iii, The array of focal adhesions was irregular, and the development of actin fiber was also weak. iv, Focal adhesion existed in the circumference and no polarity was recognized. B, Pull-down assay for Rac1-GTP. C16orf74 stable showed that expression of Rac1-GTP was increased compared with MOCK. GTPγS, positive control; GDP, negative control. C, WST assay (C16orf74 stable vs. MOCK cells, comparison of 3 cell lines). C16orf74 stable showed an increase in cell proliferation compared with MOCK. D, Matrigel invasion assay (C16orf74 stable vs. MOCK cells, comparison of three cell lines). C16orf74 stable showed a significant increase in invasiveness. E, Wound-healing assay (C16orf74 stable vs. MOCK cells). Cell migration ability was higher in C16orf74 with significant difference (*, P < 0.01). F, Hematogenic invasion assay (C16orf74 stable vs. MOCK cells). C16orf74 stable had significantly more cells infiltrated beyond HUVECs compared with MOCK (**, P < 0.05). G, Western blot analysis for MMP2, MMP9, Akt, p-Akt, ERK, p-ERK, mTOR, and p-mTOR, in C16orf74 stable and MOCK cells. C16orf 74-stable cells showed increased phosphorylation form of Akt. H, Gelatin zymography for MMP2 and MMP9 in C16orf74 stable and MOCK cells. Left, image of gels; right, band quantitative graph (value of C16orf74 based on MOCK = 1). C16orf74-stable cells showed an increase in the amount of activated MMP2 (62 kDa) and a decrease in nonactivated MMP 2 (pro-MMP 2: 72 kDa) compared with MOCK. In addition, MMP9 showed an increase in activity, although it was lower than MMP2.

Figure 1.

Functional analysis of C16orf74 overexpressed stable transfectant cell line. A, Observation of cell morphology of C16orf74 stable (A-1) and MOCK cells (A-2) by immunofluorescence (red: F actin; green: paxillin). i, The polarity of the cell was strong, and the direction of cell movement (arrow) was clear. The development of lamellipodium was strong. ii, Filopodia was developing in the direction of cell progression. The development of actin was strong. iii, The array of focal adhesions was irregular, and the development of actin fiber was also weak. iv, Focal adhesion existed in the circumference and no polarity was recognized. B, Pull-down assay for Rac1-GTP. C16orf74 stable showed that expression of Rac1-GTP was increased compared with MOCK. GTPγS, positive control; GDP, negative control. C, WST assay (C16orf74 stable vs. MOCK cells, comparison of 3 cell lines). C16orf74 stable showed an increase in cell proliferation compared with MOCK. D, Matrigel invasion assay (C16orf74 stable vs. MOCK cells, comparison of three cell lines). C16orf74 stable showed a significant increase in invasiveness. E, Wound-healing assay (C16orf74 stable vs. MOCK cells). Cell migration ability was higher in C16orf74 with significant difference (*, P < 0.01). F, Hematogenic invasion assay (C16orf74 stable vs. MOCK cells). C16orf74 stable had significantly more cells infiltrated beyond HUVECs compared with MOCK (**, P < 0.05). G, Western blot analysis for MMP2, MMP9, Akt, p-Akt, ERK, p-ERK, mTOR, and p-mTOR, in C16orf74 stable and MOCK cells. C16orf 74-stable cells showed increased phosphorylation form of Akt. H, Gelatin zymography for MMP2 and MMP9 in C16orf74 stable and MOCK cells. Left, image of gels; right, band quantitative graph (value of C16orf74 based on MOCK = 1). C16orf74-stable cells showed an increase in the amount of activated MMP2 (62 kDa) and a decrease in nonactivated MMP 2 (pro-MMP 2: 72 kDa) compared with MOCK. In addition, MMP9 showed an increase in activity, although it was lower than MMP2.

Close modal

C16orf74-stable cells demonstrated increased cell proliferation, invasion, migration, and vascular invasion compared with MOCK cells as observed from the results of WST assay (Fig. 1C), Matrigel invasion assay (Fig. 1D), wound-healing assay (Fig. 1E), and hematogenic invasion assay (Fig. 1F). Western blot analysis verified that C16orf74 increased phosphorylation of Akt and expression of MMP2 (Fig. 1G). Therefore, it was suggested that activation of MMP2 and phosphorylation of Akt by C16orf74 may be involved in the increase of cell proliferation and invasiveness. C16orf74-stable cells showed an increase in activated MMP2 and activated MMP9 and a decrease in each of the inactivated form compared with MOCK cells as evidenced by gelatin zymography (Fig. 1H). The expression of C16orf74 in C16orf74 stable and MOCK cell lines by quantitative RT-PCR, Western blot, and immunofluorescence were shown in Supplementary Fig. S1.

C16orf74 forms a homodimer and is localized under the cell membrane

Western blot analysis of the C16orf74 protein revealed multiple bands, resulting in the deduction that C16orf74 form multimers or complexes. From the amino acid sequence of the C16orf74 protein, we predicted a dimer formation at the S-S binding site C7 and C14 (Fig. 2A). Native PAGE showed that two bands were recognized at 60 and 120 kDa (2-fold), suggesting the possibility of dimer formation (Fig. 2B). Cotransfection and pull-down assay showed that C16orf74/FLAG protein and C16orf74/HA protein were directly bound (Fig. 2C). Hence, it was confirmed that C16orf74 exhibits a homodimeric form.

Figure 2.

C16orf74 forms a homodimer and localizes under the cell membrane. A, Amino acid sequence of C16orf74. B, Native PAGE of C16orf74 stable (three cell lines) and MOCK. Because C16orf74 stable showed two bands in native PAGE, C16orf74 was considered to exhibit a homodimer form. Top row, Native PAGE; bottom row, SDS-PAGE; four lanes on the left, SDS and 2-mercaptoethanol (−), Boil (−) at the time of cell lysate collection; four lanes on the right: cell lysate was collected as described earlier. C, Proof of binding of C16orf74/FLAG and C16orf74/HA by pull-down assay. C16orf74/FLAG and C16orf74/HA bind directly (form homodimers). D, Immunofluorescence analysis of the changes in the localization of C16orf74 by administration of 2-hydroxymyristic acid to C16orf74-stable cells. Compared with control (DMSO 0.125 %), in the 2-hydroxymyristic acid administration group, the localization of C16orf74 was changed from cell membrane to cytoplasm diffuse. E, Intracellular localization of C16ord74/WT, C16orf74/G2A, C16orf74/C7A_C14A, C16orf74/Δ2-15, and T44A. C16orf74/WT and C16orf74/T44A localized under the cell membrane of COS7 cells. Conversely, C16ord74/G2A, C16orf74/C7A_C14A, and C16orf74/Δ2-15 could not localize to the cell membrane. F, Pull-down assay of C16orf74/G2A, C16orf74/C7A_C14A, C16orf74/Δ2-15, C16orf74/T44A (with FLAG tag), and C16orf74/WT (with HA tag). C16orf74/WT and C16orf74/T44A formed dimers, but other constructs did not form dimers. G, Schematic representation of the structure of C16orf74. We hypothesized that the first 15 amino acids of the C16orf74 protein is involved in the structure of the protein, dimer formation at disulfide-binding site C7 and C14, and fixation to the cell membrane by G2 that is the N-myristoylation site.

Figure 2.

C16orf74 forms a homodimer and localizes under the cell membrane. A, Amino acid sequence of C16orf74. B, Native PAGE of C16orf74 stable (three cell lines) and MOCK. Because C16orf74 stable showed two bands in native PAGE, C16orf74 was considered to exhibit a homodimer form. Top row, Native PAGE; bottom row, SDS-PAGE; four lanes on the left, SDS and 2-mercaptoethanol (−), Boil (−) at the time of cell lysate collection; four lanes on the right: cell lysate was collected as described earlier. C, Proof of binding of C16orf74/FLAG and C16orf74/HA by pull-down assay. C16orf74/FLAG and C16orf74/HA bind directly (form homodimers). D, Immunofluorescence analysis of the changes in the localization of C16orf74 by administration of 2-hydroxymyristic acid to C16orf74-stable cells. Compared with control (DMSO 0.125 %), in the 2-hydroxymyristic acid administration group, the localization of C16orf74 was changed from cell membrane to cytoplasm diffuse. E, Intracellular localization of C16ord74/WT, C16orf74/G2A, C16orf74/C7A_C14A, C16orf74/Δ2-15, and T44A. C16orf74/WT and C16orf74/T44A localized under the cell membrane of COS7 cells. Conversely, C16ord74/G2A, C16orf74/C7A_C14A, and C16orf74/Δ2-15 could not localize to the cell membrane. F, Pull-down assay of C16orf74/G2A, C16orf74/C7A_C14A, C16orf74/Δ2-15, C16orf74/T44A (with FLAG tag), and C16orf74/WT (with HA tag). C16orf74/WT and C16orf74/T44A formed dimers, but other constructs did not form dimers. G, Schematic representation of the structure of C16orf74. We hypothesized that the first 15 amino acids of the C16orf74 protein is involved in the structure of the protein, dimer formation at disulfide-binding site C7 and C14, and fixation to the cell membrane by G2 that is the N-myristoylation site.

Close modal

We also speculated that the amino acid sequence of G2 may have an important role in C16orf74 localization to the cell membrane as an N-myristoylation site. Administration of 2-hydroxymyristic acid (myristoylation inhibitor) showed that localization of C16orf74 was changed to the cytoplasm (Fig. 2D). Using mutation analysis, the intracellular localization of C16orf74/G2A (mutant of the myristoylation site), C16orf74/C7A_C14A (mutant of the dimer formation site), C16orf74/Δ2-15 (deletion mutant of both the myristoylation and dimer formation site), C16orf74/T44A (mutant of the phosphorylation site that is important to PPP3CA interaction; ref. 6) constructs were examined. C16orf74/WT and C16orf74/T44A localized directly under the cell membrane, oppositely, C16orf74/G2A, C16orf74/C7A_C14A, and C16orf74/Δ2-15 could not localize under the cell membrane (Fig. 2E). To verify dimer formation, pull down assay was performed on each construct (with FLAG tag) and C16orf74/WT (with HA tag). It was found that C16orf74/WT and C16orf74/T44A formed dimers, but other constructs did not form dimers (Fig. 2F).

These results indicate that the first 15 amino acid sequences are considered to be a “structural site” that regulates C16orf7 dimer formation and the localization to cell membrane. In addition, the “functional sites” having a PDIIIT motif that is a PPP3CA-binding site and T44 which is a phosphorylation site (6), are bound to the “structural site” via the -SSSSSS- structure (Fig. 2G).

Suppression of cell proliferation, invasion, and migration by inhibiting homodimers of C16orf74 by dimer-blocking peptides

We hypothesized that inhibition of C16orf74 homodimers would suppress PDAC cell proliferation and invasiveness, and so we designed a homodimer-blocking cell-permeable peptide. 11R-DB (dimer blocking) peptide was designed by a peptide homologous to the first 15 amino acid sequence of C16orf74 linking 11R (cell-permeable peptide signal) with a hydrophilic spacer (-GGG-; Fig. 3A). the negative control peptides for 11R-DB, 11R-7_14AAA peptide (both sides of C7 and C14 were substituted with alanine) and the 11R-C7A_C14A peptide (C7 and C14 were substituted with alanine) were designed (Fig. 3A). The dimer inhibition of C16orf74 using 11R-DB was evaluated by a pull-down assay. 11R-DB was found to inhibit the dimerization of C16orf74 in a dose-dependent manner (Fig. 3B). 11R-7_14 AAA (control peptide) did not inhibit the dimerization of C16orf74 and 11R-C7A_C14A inhibited the dimerization of C16orf74 partially (Fig. 3C).

Figure 3.

Suppression of cell proliferation, invasion, and migration by inhibiting homodimers of C16orf74 by dominant negative peptides. A, Peptide design of 11R-DB (Dimer Block), 11R-C7A_C14A, and 11R-7_14AAA, and the schematic images of the effect of each peptide on dimerized C16orf74. B, Pull-down assay of C16orf74/FLAG and C16orf74/HA with 11R-DB (0–30 μmol/L). The binding of each construct was inhibited in a concentration-dependent manner by 11R-DB. C, Pull-down assay of C16orf74/FLAG and C16orf 74/HA with PBS, 11R-7_14AAA (30 μmol/L), 11R-C7A_C14A (30 μmol/L), and 11R-DB (30 μmol/L). Control (PBS) and 11R-7_14 AAA did not affect dimerization of C16orf74. 11R-DB inhibited each binding, the effect was weaker than 11R-DB, but 11R-C7A_C14A also inhibited binding. D, WST assay for PDAC cell lines and NHDF (normal fibroblast) for each peptide (0–30 μmol/L, 24 hours). E, Matrigel invasion assay of PDAC in presence of the 11R-DB peptide (0, 10, and 20 μmol/L). F, Wound-healing assay for PDAC with 11R-DB peptide (0 or 10 μmol/L). G, Changes in cell morphology after 11R-DB administration (immunofluorescence of C16orf74 stable; green, paxillin; red, f-actin). The localization of the focal adhesion became random (i) or pericytes (ii), and the number of polarity-lost cells were increased. In addition, the form of actin changed randomly. H, Changes in the localization of C16orf74 by 11R-DB administration (immunofluorescence of C16orf74 stable cells; green, C16orf74/FLAG; red, f actin). (a): Control (PBS); (b): 11R-DB. By 11R-DB administration, the localization of C16orf74 moved from the plasma membrane into the cytoplasm. Expression of C16orf74 between cells was strong and remained even after administration of 11R-DB (yellow circle); (c): Actin fiber was broken as seen when using Rho Inhibitor, and development of cell membrane also weakened. NS; no significance; *, P < 0.05; **, P < 0.01.

Figure 3.

Suppression of cell proliferation, invasion, and migration by inhibiting homodimers of C16orf74 by dominant negative peptides. A, Peptide design of 11R-DB (Dimer Block), 11R-C7A_C14A, and 11R-7_14AAA, and the schematic images of the effect of each peptide on dimerized C16orf74. B, Pull-down assay of C16orf74/FLAG and C16orf74/HA with 11R-DB (0–30 μmol/L). The binding of each construct was inhibited in a concentration-dependent manner by 11R-DB. C, Pull-down assay of C16orf74/FLAG and C16orf 74/HA with PBS, 11R-7_14AAA (30 μmol/L), 11R-C7A_C14A (30 μmol/L), and 11R-DB (30 μmol/L). Control (PBS) and 11R-7_14 AAA did not affect dimerization of C16orf74. 11R-DB inhibited each binding, the effect was weaker than 11R-DB, but 11R-C7A_C14A also inhibited binding. D, WST assay for PDAC cell lines and NHDF (normal fibroblast) for each peptide (0–30 μmol/L, 24 hours). E, Matrigel invasion assay of PDAC in presence of the 11R-DB peptide (0, 10, and 20 μmol/L). F, Wound-healing assay for PDAC with 11R-DB peptide (0 or 10 μmol/L). G, Changes in cell morphology after 11R-DB administration (immunofluorescence of C16orf74 stable; green, paxillin; red, f-actin). The localization of the focal adhesion became random (i) or pericytes (ii), and the number of polarity-lost cells were increased. In addition, the form of actin changed randomly. H, Changes in the localization of C16orf74 by 11R-DB administration (immunofluorescence of C16orf74 stable cells; green, C16orf74/FLAG; red, f actin). (a): Control (PBS); (b): 11R-DB. By 11R-DB administration, the localization of C16orf74 moved from the plasma membrane into the cytoplasm. Expression of C16orf74 between cells was strong and remained even after administration of 11R-DB (yellow circle); (c): Actin fiber was broken as seen when using Rho Inhibitor, and development of cell membrane also weakened. NS; no significance; *, P < 0.05; **, P < 0.01.

Close modal

11R-DB showed inhibition of cell growth in C16orf74-overexpressing PDAC cell lines (PK-1 and PK-9) but had a lower effect on C16orf74 low-expressing PDAC cell line and normal cell line (PANC-1, MIA Paca2, and NHDF). 11R-C7A_C14A was weaker than 11R-DB but showed cell growth inhibition correlated with the expression level of C16orf74. 11R-7_14AAA (control peptide) showed no effect on any cell lines (Fig. 3D). The same results were found in the Matrigel invasion assay and wound-healing assay (Fig. 3E and F). The expression of C16orf74 in PDAC or other cell lines by quantitative RT-PCR or Western blot analysis were shown in Supplementary Fig. S2.

The morphologic changes in pancreatic cancer cells under 11R-DB treatment were examined by immunofluorescence with Paxillin (Green: Alexa Fluor 488) and F actin (Red: Alexa Fluor 594). By 11R-DB treatment, the localization of the focal adhesion became scattered or pericytes, and the polarity was lost, and the forms of F-actin changed randomly (Fig. 3G). Similarly, the development of lamellipodium, thought to have developed by Rac1 activated by C16orf74 expression, was attenuated. The cell morphology of C16orf74 stable was changed to same as Mock (in Fig. 1A) under the treatment of 11R-DB. C16orf74 (Green: Alexa Fluor 488) was localized in the cell membrane and was more strongly expressed in the cell-to-cell contact part (Fig. 3H, a); however, after 11R-DB administration, the C16orf74 localization at the cell membrane was lost, and cell–cell contact was attenuated (Fig. 3H, b). The actin fiber was also attenuated as seen when using Rho inhibitor (ref. 15; Fig. 3H, c). The invasiveness of the C16orf74-stable transfectants was also significantly suppressed by 11R-DB (Supplementary Fig. S3). The above findings suggested that 11R-DB inhibits the cytoskeletal change for cancer cell invasion.

C16orf74 binds directly to integrin αVβ3 and controls the PI3K/Akt/mTOR cascade

From C16orf74 localization under the cell membrane and stronger expression in cell-to-cell contact, we speculated that the integrin family and other molecules such as integrin linked kinase (ILK), PDK1, and Paxillin would be involved in C16orf74 signals. The binding between C16orf74 and these molecules was examined by pull-down assays. It was found that integrin β3, ILK, and PPP3CA [shown in previous study (ref. 6) as a control] directly bind to C16orf74 (Fig. 4A). C16orf74 also showed colocalization with integrin β3 by immunofluorescence analysis, in addition to the development of lamellipodium that was found at colocalized site (Fig. 4B). In general, integrin was present as a heterodimer of α-subunit and β-subunit, therefore the combination of integrin α-subunit and β-subunit was determined by pull-down assay. C16orf74 and Integrin αVβ3 were bound together (Fig. 4C). The binding of C16orf74 and Integrin β3 was also inhibited by administration of 11R-DB (Fig. 4D), indicating that dimer formation of C16orf74 is important for binding.

Figure 4.

C16orf74 binds directly to integrin αVβ3 and controls the PI3K/Akt/mTOR cascade. A, Search for molecules that bind C16orf74. Pull-down assay of C16orf74/WT (with FLAG tag) and integrin β1-4 (ITG B1-4), ILK, PPP3CA, PDK1, and paxillin (PXN; with HA tag) revealed that C16orf74 and ITGB3, ILK and PPP3CA strongly bound to each other. B, C16orf74/FLAG and integrin β3/HA were colocalized under the cell membrane of COS7 cells. Development of cell membrane was confirmed at colocalized site. C, The binding of C16orf74, integrin αV, and integrin αIIb was verified by pull-down assay (immunoprecipitated with FLAG antibody and IgG2a as an isotype control) in C16orf74-stable cells (3 types) and MOCK. The degree of binding varied for each cell line. D, Pull-down assay of C16orf74/FLAG and integrin β3/HA with 11R-DB (0–30 μmol/L). The binding of each construct was inhibited in a concentration-dependent manner by 11R-DB. E, Western blotting of Akt, p-Akt, mTOR, p-mTOR, p42/44 MAPK(ERK), p-p42/44 MAPK (p-ERK), MMP2 and MMP9, and β-actin in 30 μmol/L 11R-DB, 11R-C7A_C14A, 11R-7_14AAA, and PBS (control), treated PK-1, PK-9, PANC-1 and MIA PaCA-2 cells. F, Gelatin zymography of PK-1 and PK-9 cells with 11R-DB (0–30 μmol/L). Left, image of gels; right, band quantitative graph. 11R-DB caused a decrease in activated MMP2, but did not significantly affect active MMP9.

Figure 4.

C16orf74 binds directly to integrin αVβ3 and controls the PI3K/Akt/mTOR cascade. A, Search for molecules that bind C16orf74. Pull-down assay of C16orf74/WT (with FLAG tag) and integrin β1-4 (ITG B1-4), ILK, PPP3CA, PDK1, and paxillin (PXN; with HA tag) revealed that C16orf74 and ITGB3, ILK and PPP3CA strongly bound to each other. B, C16orf74/FLAG and integrin β3/HA were colocalized under the cell membrane of COS7 cells. Development of cell membrane was confirmed at colocalized site. C, The binding of C16orf74, integrin αV, and integrin αIIb was verified by pull-down assay (immunoprecipitated with FLAG antibody and IgG2a as an isotype control) in C16orf74-stable cells (3 types) and MOCK. The degree of binding varied for each cell line. D, Pull-down assay of C16orf74/FLAG and integrin β3/HA with 11R-DB (0–30 μmol/L). The binding of each construct was inhibited in a concentration-dependent manner by 11R-DB. E, Western blotting of Akt, p-Akt, mTOR, p-mTOR, p42/44 MAPK(ERK), p-p42/44 MAPK (p-ERK), MMP2 and MMP9, and β-actin in 30 μmol/L 11R-DB, 11R-C7A_C14A, 11R-7_14AAA, and PBS (control), treated PK-1, PK-9, PANC-1 and MIA PaCA-2 cells. F, Gelatin zymography of PK-1 and PK-9 cells with 11R-DB (0–30 μmol/L). Left, image of gels; right, band quantitative graph. 11R-DB caused a decrease in activated MMP2, but did not significantly affect active MMP9.

Close modal

We examined intracellular signaling in PDAC cell lines by administration of 11R-DB (Fig. 4E). It was found that 11R-DB reduces phosphorylation of Akt and mTOR in C16orf74 high expression cell lines PK-1 and PK-9 (quantitative data in Supplementary Fig. S4A and S4B) but does not affect phosphorylation of p42/44 MAPK. In addition, 11R-DB did not affect the signaling of C16orf74 in low expression cell lines PANC-1 and MIA PaCa-2. These results suggest that C16orf74 might be involved in the PI3K/Akt/mTOR cascade, but may not affect the MAPK pathway. In addition, 11R-DB decreased the activation of MMP2, but did not significantly affect MMP9 by zymography (Fig. 4F).

Antitumor effect of 11R-DB peptide on human pancreatic cancer cells grown in nude mice

To assess the antitumor effect of 11R-DB in vivo, treatment of human pancreatic cancer cells, growing subcutaneously and in the pancreas (orthotopic animal model) and in peritoneal metastasis models of nude mice, was done with 11R-DB. In all experiments, PK-9 (C16orf74-overexpressed PDAC cell line) was used. Treatment schedules of each model are shown in Supplementary Fig. S5A and Figs. 5A and 6A. In subcutaneous model, tumor growth was suppressed with significant difference in the 11R-DB administration group (Supplementary Fig. S5B and S5C). In orthotopic model, antitumor effect was examined by in vivo imaging (Fig. 5B) and tumor growth was suppressed with significant difference in the 11R-DB administration group (11R-DB vs. 11R-7_14AAA/PBS: *P < 0.05, 11R-7_14AAA vs PBS: n.s.; Fig. 5C and D). No reduction in weight was observed in the peptide-treated mice after treatment by 11R-DB, and no apparent negative side effects were found to the major vital organs confirmed by hematoxylin and eosin staining (Supplementary Fig. S5D and S5E). 11R-DB suppressed Akt phosphorylation and mTOR phosphorylation in tumor specimen after treatment (Fig. 5E) same as previously shown in vitro assays (Fig. 4E). The Ki-67 index of the tumor specimen was decreased after treatment with 11R-DB (Fig. 5F).

Figure 5.

11R-DB showed antitumor effect on pancreatic tumor model mice. A, Treatment schedule of orthotopic model. B, Representative examples of bioluminescence imaging after tumor cell injection at 5 weeks (4 weeks after initiation of therapy). Representative cases of each groups are shown. C, The graph shows the signal intensities recorded from individual mice for each week and group. The signal intensity at 1 week was set at 1 as a control for each mouse. Each group had 6 mice. Data are presented as mean ± SEM (11R-DB vs. 11R-7_14AAA/PBS: *, P < 0.05, 11R-7_14AAA vs. PBS: n.s.). D, Tumor weights at 5 weeks after tumor cell injection in each group (n = 6; 11R-DB vs. 11R-7_14AAA/PBS: *, P < 0.05, 11R-7_14AAA vs. PBS: n.s.). E, IHC staining of phosphorylated Akt, phosphorylated mTOR, and Ki-67 (MIB-1) in PDAC tumors in vivo. A representative example is shown. F, Ki-67 Index in tumors of each group (PBS, 11R-7_14AAA, and 11R-DB); **, P < 0.05.

Figure 5.

11R-DB showed antitumor effect on pancreatic tumor model mice. A, Treatment schedule of orthotopic model. B, Representative examples of bioluminescence imaging after tumor cell injection at 5 weeks (4 weeks after initiation of therapy). Representative cases of each groups are shown. C, The graph shows the signal intensities recorded from individual mice for each week and group. The signal intensity at 1 week was set at 1 as a control for each mouse. Each group had 6 mice. Data are presented as mean ± SEM (11R-DB vs. 11R-7_14AAA/PBS: *, P < 0.05, 11R-7_14AAA vs. PBS: n.s.). D, Tumor weights at 5 weeks after tumor cell injection in each group (n = 6; 11R-DB vs. 11R-7_14AAA/PBS: *, P < 0.05, 11R-7_14AAA vs. PBS: n.s.). E, IHC staining of phosphorylated Akt, phosphorylated mTOR, and Ki-67 (MIB-1) in PDAC tumors in vivo. A representative example is shown. F, Ki-67 Index in tumors of each group (PBS, 11R-7_14AAA, and 11R-DB); **, P < 0.05.

Close modal
Figure 6.

11R-DB inhibited tumor cell engraftment of pancreatic cancer peritoneal dissemination model mice. A, Treatment schedule of peritoneal dissemination model. B, The graph shows the signal intensities recorded from individual mice for each day and group (n = 5; 11R-DB vs. 11R-7_14AAA/PBS: *P < 0.05, 11R-7_14AAA vs. PBS: n.s.). Data are presented as the mean ± SEM. C, Representative examples of bioluminescence imaging of peritoneal dissemination model mice. D, The number of dissemination nodules at 35 days after tumor cell injection in each group (n = 5; 11R-DB vs. PBS: *, P < 0.05, 11R-DB vs. 11R-7_14AAA **, P < 0.05 11R-7_14AAA vs. PBS: n.s.) E, Representative examples of bioluminescence imaging of peritoneal dissemination nodules (after sacrifice) in the three groups of mice (PBS, 11R-7_14AAA, and 11R-DB).

Figure 6.

11R-DB inhibited tumor cell engraftment of pancreatic cancer peritoneal dissemination model mice. A, Treatment schedule of peritoneal dissemination model. B, The graph shows the signal intensities recorded from individual mice for each day and group (n = 5; 11R-DB vs. 11R-7_14AAA/PBS: *P < 0.05, 11R-7_14AAA vs. PBS: n.s.). Data are presented as the mean ± SEM. C, Representative examples of bioluminescence imaging of peritoneal dissemination model mice. D, The number of dissemination nodules at 35 days after tumor cell injection in each group (n = 5; 11R-DB vs. PBS: *, P < 0.05, 11R-DB vs. 11R-7_14AAA **, P < 0.05 11R-7_14AAA vs. PBS: n.s.) E, Representative examples of bioluminescence imaging of peritoneal dissemination nodules (after sacrifice) in the three groups of mice (PBS, 11R-7_14AAA, and 11R-DB).

Close modal

In peritoneal metastasis model, tumor cell engraftment was significantly suppressed by 11R-DB administration (Fig. 6B and C), and the number of disseminated nodules was suppressed with significant difference in the 11R-DB administration group (11R-DB vs. 11R-7_14AAA/PBS: *, P < 0.05, 11R-7_14AAA vs. PBS: n.s; Fig. 6D and E).

In this study, C16orf74 was found to be localized just beneath the cell membrane and was seen to bind to integrin. From the morphologic observation of C16orf74-stable cell lines, we acknowledged findings that strongly indicated the activation of the Rho family such as lamellipodium and stress fiber (16, 17). An increase in active Rac1 was thought to contribute to cell morphology and invasiveness (17–19). C16orf74 can localize in the cell membrane only when present in a dimeric form and shows direct binding to integrin αVβ3. By administration of 11R-DB, localization and binding of C16orf74 was inhibited. Administration of 11R-DB also caused a decrease in phosphorylation of Akt and mTOR and a decrease in MMP2 activity. Thus, we suggest that 11R-DB inhibited the ILK/PI3K/Akt pathway by blocking the integrin signal. The results of our study suggest that dimerization of C16orf74 is important for binding to integrin αVβ3, and downstream phosphorylation of Akt contribute to activation of MMP2, and eventually invasiveness of pancreatic cancer.

Previous studies showed that C16orf74 interacted to PPP3CA (calcineurin: CN) and was important for proliferation and invasion of PDAC (6, 11). This study demonstrated that C16orf74 might be the molecule that connected the integrin signal and CN signal. C16orf74 transfers signals from the integrin on the cell membrane to the cytoplasmic CN and activates cell proliferation and invasion through the Akt/mTOR pathway. The C16orf74 protein was found to consist of a localization domain for targeting the molecule inside the cell membrane and a functional domain for binding to CN. The localization domain is also important for dimer formation. Therefore, two blocking points were revealed in the development of therapy targeting C16orf74. One is dimer inhibition of C16orf74, which results in signal inhibition from integrin. The other is inhibition of binding between C16orf74 and PPP3CA, that is, signal inhibition to PPP3CA (11). It is also possible to block both, however, the binding site between C16orf74 and PPP3CA has the consensus-binding sequence of nuclear factor of activated T cells; therefore, inhibition of binding has a risk of immunodeficiency by T-cell inhibition (11). The newly discovered dimer inhibition is C16orf74-selective and appears to be extremely useful from the viewpoint of side effects in the development of therapeutic drugs.

The polyarginine signal achieve highly efficient nonspecific uptake of peptides into cells (20). Thus, when DN-C16orf74 is used in a clinic, systemic administration of the peptide may not be able to achieve sufficient concentrations in tumor due to uptake by normal vascular endothelial cells. In this regard, cancer cell–specific cell-permeable peptide signals, which can penetrate only the target cells in vitro and in vivo (21), might be applicable in developing DN-C16orf74 for clinical use. Another important issue is the stability of peptides in the blood. In clinical, peptide drugs such as carperitide (α-human A–type natriuretic peptide) are used in continuous injections, because the half-life of the peptide is very short (22). The clinical application of DN-C16orf74 requires technology that modifies the stabilization of peptides in blood. Another clinical approach for DN-C16orf74 is intraperitoneal administration for patients with PDAC with peritoneal metastases. A phase II clinical trial of intraperitoneal paclitaxel has recently been reported for patients with PDAC with peritoneal metastases (23). In our peritoneal metastasis model, intraperitoneal DN-C16orf74 peptide treatment was extremely effective; therefore, combination therapy can be considered for the peritoneal metastasis of PDAC for clinical trial in the future.

In conclusion, through the significant reduction of Akt and mTOR phosphorylation, DN-C16orf74 inhibited C16orf74 dimerization and suppressed PDAC proliferation in vitro and in vivo. These results suggest that DN-C16orf74 is a potential therapeutic option for patients with PDAC. However, the limitations of our study are (i) the off-target effect of DN-C16orf74 peptides on the binding inhibition of nonspecific proteins is not examined, (ii) intravenous peptide administration is not achieved, (iii) the drug dynamics after the peptide administration is not revealed. Therefore, further studies will be necessary to overcome the above limitations.

No potential conflicts of interest were disclosed.

Conception and design: T. Kushibiki, T. Nakamura, M. Takahashi, Y. Ebihara, T. Shichinohe

Development of methodology: T. Kushibiki, T. Nakamura, K. Inoko

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): T. Kushibiki, T. Nakamura, K. Hontani, K. Inoko, T. Asano, W.-R. Park

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): T. Kushibiki, T. Nakamura, M. Tsuda, K. Inoko, T. Asano, T. Shichinohe, S. Tanaka

Writing, review, and/or revision of the manuscript: T. Kushibiki, T. Nakamura, K. Inoko, K. Okamura, S. Murakami, Y. Kurashima, K. Tanaka, T. Shichinohe, S. Hirano

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): T. Kushibiki, M. Tsuda, K. Inoko, T. Noji, Y. Nakanishi, N. Maishi, K. Sasaki, W.-R. Park, K. Hida

Study supervision: T. Nakamura, M. Tsuda, T. Tsuchikawa, K. Okamura, S. Murakami, T. Noji, T. Shichinohe, S. Hirano

The authors thank Kei Shida and Dr. Yusuke Ohba, and other members of the Department of Gastroenterological Surgery II, Hokkaido University Faculty of Medicine, for helpful discussions. This work was supported by JSPS KAKENHI grant number JP18K08667 (to T. Nakamura).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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Supplementary data