Inhibition of Discoidin Domain Receptor 1 Reduces Collagen-mediated Tumorigenicity in Pancreatic Ductal Adenocarcinoma Free

This article is featured in Highlights of This Issue, p. 2327

The extracellular matrix (ECM) is crucial for tumor progression. Specifically, the amount of ECM deposition correlates with a poor prognosis in cancer patients and ECM-mediated signaling can promote tumor cell survival and invasion (1–4). Fibrillar collagens, major components of tumor-associated ECM, signal through integrins and discoidin domain receptors (DDR). The DDR family includes DDR1 and DDR2, which are closely related collagen-specific receptor tyrosine kinases (RTK) that recognize and bind several types of collagen (5–7). DDR1 and DDR2 are differentially expressed among tissue and cell types: DDR1 expression is predominant in epithelial cells and is found at high levels in the pancreas, brain, lung, kidney, spleen, and placenta (8–11), whereas DDR2 is found in cells of connective tissues that originate from embryonic mesoderm and is expressed highly in skeletal and heart muscle as well as kidney and lung (12–15). DDR1 and DDR2 are functionally important during development. For example, DDR1 participates in organogenesis and DDR2 is critical for bone development and growth (16). DDR1 and DDR2 contribute to key cellular processes, including cell migration, cell proliferation, cell differentiation, and cell survival (17).

Pancreatic ductal adenocarcinoma (PDA) has a collagen-rich microenvironment. Collagen is expressed in PDA by stromal cells, mainly fibroblasts, with a relatively minor but functionally relevant contribution by tumor cells (18, 19). Collagen receptors such as DDR1 have been linked in other tumor types to cellular processes that are prominent in PDA, including phenotypic plasticity [e.g., epithelial-to-mesenchymal transition (EMT)] and chemoresistance. For example, DDR1 can confer chemoresistance and mediate prosurvival signals in breast cancer and lymphoma cell lines (20, 21) and may be involved in the recurrence of certain types of cancer (22). Furthermore, Shintani and colleagues (23) found that collagen I–induced N-cadherin expression (a marker of mesenchymal cells) in human pancreatic cancer cells required activation of DDR1. Moreover, the fact that DDR1 is expressed highly in a variety of tumor types suggests that this collagen-activated RTK may be involved in tumor progression (24–27). We establish here that activation of DDR1 contributes to the tumorigenic properties of PDA and that pharmacologically inhibiting this receptor tyrosine kinase with the selective DDR1 small-molecule 7rh (28–31) slows tumor progression and enhances chemoresponse to standard-of-care PDA regimens.

Human pancreatic cancer cell lines (AsPC-1, PANC-1, and BxPC-3) were purchased from the ATCC and were fingerprinted for validation of authenticity. The murine pancreatic cancer cell line Pan02 (also known as Panc02) was obtained from the NCI (DCTD Tumor Repository). Cells were cultured in DMEM (Invitrogen) or RPMI (Invitrogen) containing 5% FBS and maintained at 37°C in a humidified incubator with 5% CO₂ and 95% air.

Cells were cultured in 6-well tissue culture plates at high density (∼90% confluence) in 2 mL of media with 5% FBS. Uniform scratches were made down the center of each well with a p20 pipette tip, cells were gently washed with PBS to remove the loose cell debris, and drug was added in media containing 5% FBS. 7rh was added in four 4-fold dilutions, to respective wells, and a DMSO control was added to respective wells to demonstrate the vehicle-independent effect. Cells were plated on respective culture conditions and images from the center of each well were taken at 0, 10, 20, and 30 hours. The wound width (μm) was measured using NIS Elements AR 2.30 software. The initial wound width was used to verify consistency in scratches.

Cells were cultured in 6-well tissue culture plates at low density (250 cells per well) in 2 mL of media with 5% FBS with or without 7rh for approximately 1.5–2.0 weeks or until significant colony formation. 7rh was added in four 4-fold dilutions to respective wells, and a DMSO control was added to respective wells to demonstrate the vehicle-independent effect. Cells were then fixed with 10% formalin and stained with crystal violet. Images were analyzed with ImageJ.

Assays were performed with antibodies against: phospho-DDR1 (Tyr792, Cell Signaling Technology; #11994), DDR1 (D1G6, Cell Signaling Technology; #5583), phospho-SRC (Tyr416, Cell Signaling Technology; #2101), SRC (Cell Signaling Technology; #2108), phospho-PYK2 (Tyr402, Cell Signaling Technology; #3291), PYK2 (Cell Signaling Technology; #3292), α-Amylase (D55H10, Cell Signaling Technology; #3796), vimentin (Millipore AB5733), phospho-FAK (Abcam #4803), FAK (Cell Signaling Technology; #3285), PEAK1 (Millipore 09-274) phospho-PEAK1 (Tyr665, Millipore #ABT52), γH2AX (Ser139, Novus Biologicals; #NB100-384), and cleaved caspase-3 (Asp175, Cell Signaling Technology; #9661).

Subconfluent monolayers of cells were lysed, supernatants were recovered by centrifugation at 13,000 rpm, protein concentration was measured, and equal amounts of total protein were separated by SDS-PAGE. Proteins were transferred to polyvinylidene difluoride membranes (Bio-Rad) followed by blockade for 1 hour in 5% milk in TBS-T. The membranes were incubated overnight at 4°C with primary antibodies. Primary antibodies used are detailed above. Membranes were incubated with the corresponding horseradish peroxidase–conjugated secondary antibody (Pierce Biotechnologies) for 1 to 2 hours. Specific bands were detected using the enhanced chemiluminescence reagent (ECL; Perkin Elmer Life Sciences) on autoradiographic film.

Cell lines were lysed in modified radioimmunoprecipitation assay buffer (0.5% deoxycholate, 0.5% SDS, 1% Triton X-100, 10 mmol/L sodium phosphate, pH 7.2, 150 mmol/L sodium chloride) with protease inhibitor (Complete Mini, Roche Life Science). Lysis was performed on serum-starved adherent cells after washing with chilled PBS. Lysates were allowed to rotate at 4°C on a nutator for 1 hour and then vortexed several times before centrifuging at 13,000 rpm for 10 minutes to pellet any insoluble material. Lysates were precleared with protein A/G beads (Thermo Fisher Scientific). We used 200 μg cellular protein in 1 mL lysis buffer per immunoprecipitation reaction. To each sample, 1 μg of the appropriate antibody was added with 50 μL protein A/G bead slurry; each sample was then allowed to rotate overnight at 4°C on a nutator. Immunoprecipitated complexes were washed twice in lysis buffer, boiled in sample buffer, and subjected to SDS-PAGE and Western blot analysis.

Cells were plated 18–24 hours before transfection (1 × 10⁵ cells/well in 6-well dish) at an initial confluence of 60%–80%. TransIT-siQUEST reagent and siRNA complexes were prepared and added according to manufacturer instructions (Mirus Bio, LLC). siRNA complexes were added to the cells at a final siRNA complex concentration of 1 μmol/L. Protein was harvested 72 hours posttransfection for Western blot analysis. siRNA duplexes were purchased from Integrated DNA Technologies. DDR1 duplexes used were (NM_001954 duplexes 1–3):

• Duplex #1 = 5′-GUCUUGUAGCUAGAACUUCUCUAAG-3′,

• 3′-GUCAGAACAUCGAUCUUGAAGAGAUUC-5′;

• Duplex #2: 5′-GCACUAGGCAGGUAAUAAUAAAGGT-3′,

• 3′ GACGUGAUCCGUCCAUUAUUAUUUCCA-5′;

• Duplex #3: 5′-ACACUAAUAUAUGGACCUAGAUUGA-3′,

• 3′-AAUGUGAUUAUAUACCUGGAUCGAACU-5′.

Cell lines were prepared as indicated by the Sircol manual (Sircol Collagen Assay Kit, Oubis Ltd.). Sircol dye reagent was used to saturate the collagen in the supernatant. The colorimetric principle of the Sircol collagen assay is the binding of a dye to collagen. A collagen–dye complex formed and precipitated out, was recovered by centrifugation, eluted with alkali, and measured with a spectrophotometer at 555 nm. The intensity of color measurement was proportional to the collagen concentration in the sample.

All animals were housed in a pathogen-free facility with access to food and water ad libitum. C57BL/6 and NOD-SCID mice were purchased from the UT Southwestern Mouse Breeding Core (Dallas, TX). Kras^LSL-G12D/+; Trp53^LSL-R172H/+; p48^Cre/+ (KPC) mice were generated as described previously (32). Pan02 cells (5 × 10⁵) were injected orthotopically in 6- to 8-week-old C57BL/6 mice as described previously (33, 34). Mice bearing orthotopic Pan02 tumors were used for multiple studies as indicated in Table 1. Eight-week-old NOD SCID mice were injected orthotopically with 1 × 10⁶ AsPC-1 cells as described previously (33, 34). Twenty-seven days after tumor cell injection, mice were randomized to receive treatment as indicated in Table 1. Experiments were approved and performed in accordance with the Institutional Animal Care and Use Committee at UT Southwestern (Dallas, TX). For endpoint studies, experiments were stopped after the designated time post-tumor cell implantation. For survival studies, therapy was maintained until mice attained tumor burden criteria (a loss of >15% body weight, significant reduction of activity, loss of strength, excessive ascites) requiring sacrifice. Plasma from C57BL/6 mice bearing orthotopic Pan02 tumors was collected at the time of sacrifice. The plasma level of albumin, liver transaminases, aspartate transaminase, blood urea nitrogen, creatine, glucose, total bilirubin, and plasma total protein was evaluated by the Mouse Metabolic Phenotyping Core (UT Southwestern, Dallas, TX). For all therapy studies, the person administering therapy was not blinded to the therapy groups. For survival studies, evaluation of health of the animals was determined by the person administering therapy and an independent observer; a decision to sacrifice the animals due to poor performance was determined by consultation between each investigator.

Formalin-fixed tissues were embedded in paraffin and sectioned. Tissue samples from in vivo experiments were stained by Masson Trichrome by the Molecular Pathology Core (UT Southwestern, Dallas, TX). Human (44) and matched PATX (150) TMA samples were scored on a scale of 0 (no stain) to 4 (very high stain) independently by two researchers, and the data were collected and plotted accordingly.

Fluorescent images were captured with a Photometric Coolsnap HQ camera using NIS Elements AR 2.3 Software (Nikon). Color images were obtained with a Nikon Eclipse E600 microscope using a Nikon Digital Dx1200me camera and ACT1 software (Universal Imaging Corporation). Pictures were analyzed using NIS Elements (Nikon). Quantification of IHC was conducted using NIS Elements 3.2 software (Nikon Instruments). All data were analyzed using GraphPad Prism 5.0 software (GraphPad Software, Inc.). Datasets were analyzed by Student t test or ANOVA followed by Dunn posttest or Tukey MCT, multiple comparison test and results were considered as significant at P < 0.05. Results are shown as mean ± SEM.

To characterize the level of collagen-mediated DDR1 signaling in PDA, we determined the expression of phosphorylated DDR1 and phosphorylated PEAK1 in human pancreatic tumor samples with matched patient-derived tumor xenograft (PATX) samples. Primary tumors (44) and PATX samples (150) showed robust activation of DDR1 and PEAK1 (Fig. 1A–C). The overall percentages of staining positivity are shown in Fig. 1C. Furthermore, we examined the expression of active Ddr1, Pyk2, and Peak1, as well as the expression of Muc1 and Sox9 in pancreatic tumors from early (3 month) and later (5 month) stages of the KPC (Kras^LSL-G12D/+; Trp53^LSL-R172H/+; p48^Cre/+) mouse model of PDA (Fig. 1D). The KPC model recapitulates many of the pathologic features seen in human PDA including a dense stromal reaction (ref. 35; Fig. 1E and F). Trichrome analysis revealed robust collagen deposition throughout PDA lesions in KPC mice (Fig. 1F). Ddr1 activation and downstream signaling (Pyk2 and Peak1) were present in early pancreatic intraepithelial (PanIN) lesions as shown by correlative staining with a marker of early PDA lesions, Muc-1. In addition, these effectors were expressed highly throughout the tumor epithelium at the later stage of the model (5-month old KPC) as identified by areas expressing Sox9 (Fig. 1D). Sox9 is expressed in the malignant epithelium and is confined to the duct-like cells; differentiated acinar and endocrine cells do not express Sox9 (36, 37). These data demonstrate that collagen signaling via DDR1 is active in human PDA and mouse models of the disease.

The level of collagen expressed and secreted by AsPC-1 and PANC-1 cells was determined by a Sircol assay (Fig. 2A). AsPC-1 cells expressed a higher level of secreted collagen than PANC-1 cells. The addition of exogenous soluble collagen enhanced the phosphorylation of DDR1, PEAK1, and PYK2 in AsPC-1, PANC-1, and BxPC-3 cells in a time-dependent manner. Consistent with other reports (38), we found that activation of DDR1 resulted in decreased expression of total DDR1 (Fig. 2B). The downstream effectors of DDR1 are ill-defined (16, 17); however, we found that phosphorylation of PYK2 and PEAK1 was induced by collagen robustly in AsPc-1 cells but less so in PANC-1 and BxPC-3 cells. Furthermore, we found that phosphorylated PYK2 and PEAK1, but not integrin αv, coimmunoprecipitated with DDR1 from AsPC-1 cells (Fig. 2C). To further define the contribution of DDR1 to collagen signaling, AsPC-1 cells were stimulated with collagen after siRNA-mediated knockdown of DDR1. Loss of DDR1 expression abrogated the activation of PEAK1 and SRC (Fig. 2D and E), as well as cell migration (Fig. 2F). These data support the premise that collagen-mediated activation of DDR1 induces a signal pathway that includes PEAK1, SRC, and PYK2, which in turn are potentially responsible for collagen-induced chemoresistance (3, 39) and tumor progression.

To demonstrate that DDR1 participates in chemoresponse in PDA, we evaluated the effect of the small-molecule kinase inhibitor 3-(2-(Pyrazolo[1,5-a]pyrimidin-6-yl)-ethynyl) 7rh benzamide (7rh; ref. 28) on collagen-induced signaling in PANC-1 cells. The specificity of 7rh is high for DDR1 versus other related kinases (IC₅₀: DDR1, 6.8 nmol/L; DDR2, 101.4 nmol/L; Bcr-Abl, 355 nmol/L) based on previously published cell-free kinase assays (28). In PANC-1 cells, 7rh inhibited DDR1-mediated signaling induced by soluble collagen (50 μg/mL) in a concentration-dependent manner (Fig. 3A). At pharmacologically relevant concentrations, 7rh inhibited activation of PYK2 and PEAK1, signaling proteins downstream of DDR1 (40). However, 7rh did not affect the activation of focal adhesion kinase (FAK), an effector that has not been previously associated with DDR1-induced signaling (23). Inhibition of the DDR1 signaling with 7rh also reduced cell migration (Fig. 3B) and colony formation (Fig. 3C) in a concentration-dependent manner.

Before initiating the in vivo studies, murine pancreatic (Pan02) cells were investigated for the effect of 7rh on collagen-induced Ddr1 signaling in vitro (Fig. 4A), which demonstrated that 7rh inhibited Ddr1 activation in a dose-dependent manner. Prior pharmacokinetic studies (28) established the in vivo half-life of 7rh to be approximately 12 hours in rats. To determine an appropriate dose for therapy studies, mice bearing established orthotopic Pan02 pancreatic tumors were given a single dose of 0.1, 1, or 10 mg/kg of 7rh via oral gavage (Supplementary Fig. S1). Tumor tissue was collected 12 hours posttreatment and analyzed for Ddr1 activity. We found that 7rh at 1 and 10 mg/kg significantly reduced the phosphorylation of Ddr1 as well as downstream effectors Pyk2 and Peak1, and resulted in an increased apoptotic index (cleaved caspase-3; Supplementary Fig. S1B–S1E) as shown by IHC analysis. After demonstration that 7rh can reduce Ddr1 activity in the tumor microenvironment, a single-agent therapy experiment was performed using a titration of 7rh for 2 weeks. Mice bearing established orthotopic Pan02 tumors were treated with 7rh (3, 10, or 30 mg/kg, 3×/week) via oral gavage (Supplementary Fig. S2). 7rh at 10 and 30 mg/kg resulted in an increase in normal pancreatic tissue as determined by H&E histology and expression of amylase, a marker of normal acinar tissue (Supplementary Fig. S2B and S2C). At these concentrations, 7rh also significantly reduced the level of phosphorylated Ddr1 and Peak1 (Supplementary Fig. S2D and S2E), increased apoptosis (Supplementary Fig. S2F), as well as inhibited cell proliferation noted by the reduction of PCNA levels (Supplementary Fig. S2G). These findings were corroborated by Western blot analysis of tumor lysates that showed a 7rh-dependent reduction of Peak1 phosphorylation (Supplementary Fig. S2H). In addition, 7rh showed no apparent normal tissue toxicity as demonstrated by the maintenance of body weight and the lack of changes in plasma metabolites specific for liver and kidney function (Supplementary Fig. S3A and S3B). Metabolites analyzed included Alb (albumin), ALT (liver transaminases), AST (aspartate transaminase), BUN (blood urea nitrogen), Crea (creatine), Glu (glucose), Tbil (total bilirubin), and Tp (plasma total protein). Next, we performed a single-agent therapy experiment with a fixed concentration of 7rh. Mice bearing established orthotopic Pan02 tumors were treated with 7rh (25 mg/kg, 3×/week; Fig. 4). Therapy was initiated 19 days after tumor cell injection and continued until experiment day 40, at which point animals were sacrificed (Fig. 4B). We found that 7rh significantly reduced primary tumor weight (Fig. 4C). Histologic analysis of pancreata from these animals showed that 7rh slowed the progression of disease (Fig. 4D). This is consistent with increased amylase expression (Fig. 4E) and a significant decrease in Ddr1, Peak1, and Pyk2 activation (Fig. 4F–H) in animals receiving 7rh. This was concordant with enhanced apoptosis (cleaved caspase-3, Fig. 4I) and reduced proliferation (PCNA, Fig. 4J) in the presence of 7rh therapy.

To assess whether 7rh enhanced the efficacy of chemotherapy in vivo, we combined 7rh with the standard-of-care chemotherapy of PDA (gemcitabine and nab-paclitaxel) in a xenograft model of PDA (Fig. 5A). Immunocompromised animals bearing orthotopic AsPC-1 tumors were treated with vehicle, 7rh monotherapy (25 mg/kg, 3×/week), the standard-of-care regimen (chemo: gemcitabine, 15 mg/kg, 2×/week; nab-paclitaxel, 5 mg/kg, 2×/week), or the combination (combo) of 7rh and chemotherapy (Fig. 5A). Therapy was initiated 27 days after tumor cell injection and 3 animals from each cohort were sacrificed on day 28 (one day posttherapy induction) to document tumor burden at the start of therapy (initial group). Each regimen was continued until individual animals became moribund, at which point the moribund animals were sacrificed. The combination of 7rh + chemotherapy significantly enhanced the median overall survival to 98 days, compared with chemotherapy, 7rh, or vehicle at 73, 57, and 54.5 days, respectively. After the median survival was achieved for the combination group, therapy was withdrawn at day 102 to assess the consequence of therapy removal (withdrawn group; Fig. 5B and C and Supplementary Fig. S4). Tumor tissue from each group was analyzed by histology and IHC. Combination therapy resulted in more normal pancreatic tissue (H&E), a significant reduction in collagen signaling (P-DDR1, P-PYK2, P-PEAK1), a reduction in vimentin expression as well as cell proliferation (PCNA), and enhanced apoptosis (cleaved caspase-3) and DNA damage (γH2AX; Fig. 5D–K). Withdrawal of therapy from the combination group resulted in restoration of cell proliferation as well as vimentin expression and collagen signaling to levels similar to that observed in vehicle-treated animals. In addition, we noted that 7rh alone or in combination with chemotherapy reduced trichrome staining, suggesting a reduction in fibrosis (Supplementary Fig. S4A). Tumor weight versus survival days was plotted (Supplementary Fig. S4B) and indicated that therapy with 7rh, chemotherapy, or the combination reduced primary tumor growth compared with treatment with vehicle. Animal weight was monitored throughout the experiment and no therapy-induced changes in body weight were noted (Supplementary Fig. S4C).

To determine whether the therapeutic efficacy of 7rh combinatorial therapy extended to more rigorous in vivo models, we moved to a genetically engineered mouse model (GEMM) of PDA. KPC (LSL-Kras^G12D/+; LSL-Trp53^R172H/+; p48^Cre/+) mice were enrolled into therapy cohorts at 4 months of age (Fig. 6), a time point we have found where greater than 90% of animals have established PDA. Treatment arms were the same as the AsPC-1 xenograft experiment and contained 12 animals/cohort. An additional 9 animals were sacrificed at the start of therapy to document mean tumor burden at the initiation of the experiment. The average tumor burden of these 9 animals was 0.27 g. Treatment with the combination regimen enhanced median of survival to 208 days compared with treatment with chemotherapy, 7rh, or vehicle at 180, 159, and 144 days, respectively (Fig. 6B and C). IHC analyses of tumor tissue harvested at the time of sacrifice demonstrated that inhibition of Ddr1 with 7rh suppressed collagen signaling (P-Ddr1 and P-Peak1), reduced vimentin expression, and cell proliferation (PCNA) while increasing apoptosis (cleaved caspase-3) and DNA damage (γH2ax; Fig. 6D–J). Chemotherapy with gemcitabine and nab-paclitaxel also reduced collagen signaling and vimentin expression as well as decreased the number of PCNA-positive cells. In addition, treatment with 7rh alone, chemotherapy alone, or the combination induced a reduction in trichrome staining (Supplementary Fig. S5A). Tumor weight versus survival days was plotted (Supplementary Fig. S5B) and indicated that therapy with 7rh, conventional chemotherapy, or the combination reduced primary tumor growth compared with treatment with vehicle. Animal weight was not adversely affected by therapy (Supplementary Fig. S5C). These data demonstrate that Ddr1 inhibition can increase the efficacy of standard-of-care chemotherapy in robust preclinical models of PDA.

The goal of this project was to investigate the contribution of collagen-mediated DDR1 signaling to PDA progression. We demonstrated that DDR1 and downstream effectors are expressed and activated in human and mouse PDA. In addition, we found that a novel small-molecule inhibitor, 7rh benzamide (28), effectively abrogated DDR1 signaling thereby reducing liquid colony tumor cell formation and tumor cell migration. Furthermore, we found that 7rh inhibits its target and has significant therapeutic efficacy in vivo at doses that are free from observable tissue toxicity. Finally, 7rh significantly improved the efficacy of standard-of-care chemotherapy in robust mouse models of PDA. Overall these data highlight that collagen signaling through DDR1 is a critical and pharmacologically targetable pathway in PDA.

Physiologic chemoresistance can result from the accumulation of ECM proteins in the tumor microenvironment, a common characteristic of PDA. The dysregulation of ECM-driven signaling can enhance the hostile programs of cancer cells (16). This fibrotic network contributes to the development of a complex tumor microenvironment that promotes PDA development, invasion, metastasis, and resistance to chemotherapy (41). However the ECM-mediated signaling pathways that drive these programs are unclear and the mechanisms of how collagen-mediated DDR1 signaling in particular affects tumor progression require additional investigation.

We previously found that the matricellular protein Sparc (secreted protein acidic and rich in cysteine) reduced collagen I signaling through Ddr1 and that loss of Sparc accelerated PDA progression with a concordant increase in Ddr1 signaling (42). Furthermore, prior reports on the expression of SPARC in pancreatic tumor cells demonstrated that there is a reduction in SPARC expression by promoter hypermethylation in a high frequency of pancreatic tumor cells and other epithelial cancer cells (43, 44). In addition, it was reported that the restoration of SPARC expression enhanced radiosensitivity and chemosensitivity in preclinical models of colon cancer (45) and that SPARC expression enhanced chemoresponse in cancer patients (46, 47). Thus, there was compelling evidence that the loss of tumor cell expression of SPARC correlated with tumor progression and poor chemoresponse. We propose that these observations can be explained by the fact that SPARC inhibits collagen-induced DDR1 activation. This is consistent with reports that collagen signaling is associated with chemoresistance in PDA cell lines (3, 48) and that DDR1 confers resistance to chemotherapy and mediates prosurvival signals (20, 21, 49).

Our studies relied on syngeneic, xenograft, and genetic models of PDA. We chose to utilize Pan02 (also known as Panc02) cells because this cell line grows in C57Bl/6-immunocompetent animals, a useful system to evaluate initial toxicity and efficacy of DDR1 inhibition with 7rh. We used AsPC-1 cells, a commonly used human PDA cell line, because these cells express high levels of endogenous DDR1 activation in vitro and grow robustly in vivo. We also used the KPC model of PDA, which incorporates two common genetic lesions present in human PDA (e.g., KRAS activation and p53 loss). We believe this model is well-suited for endpoint and survival studies as mice develop advanced PDA with 100% penetrance at approximately 3–4 months of age and tumor progression recapitulates many of the characteristics of human PDA (35).

DDR1 is upregulated in fibrotic diseases and contributes to the initiation and progression of fibrosis (50). We observed reduced collagen deposition in tumors from mice treated with 7rh, thus the inhibition of DDR1 might improve response to chemotherapy in a cell-autonomous manner and also improve drug delivery without disrupting the function of cancer-associated fibroblasts. DDR1 inhibition has also been shown to reduce tumorigenicity in multiple tumor models (23, 25, 51, 52). For example, silencing DDR1 by siRNA has been shown to reduce metastatic activity in lung cancer models (27, 51) and enhance chemosensitivity to genotoxic drugs in breast cancer cells (49). In addition, we have recently reported that DDR1 expression and activity correlates with worse outcome in a cohort of gastric cancer patients (31). In this study, 7rh-mediated inhibition of DDR1 in gastric cancer cells reduced tumorigenic characteristics in vitro and tumor growth in vivo. Finally, collagen signaling and DDR1 activity has been linked to epithelial plasticity (23, 40). We found evidence that 7rh induced a more differentiated tumor cell phenotype, which is consistent with improved efficacy of standard chemotherapy.

Several small-molecule inhibitors (imatinib, nilotinib, and dasatinib) that target breakpoint cluster region-Abelson kinase (BCR-ABL) also potently inhibit DDR1/DDR2 activity (53, 54). Thus, the potential activity of imatinib and vinorelbine in a phase I/II trial in metastatic breast cancer patients (55), as well as dasatinib in numerous clinical trials in solid tumors (56), could be due in part to the inhibition of DDRs. Dasatinib in particular has demonstrated promising therapeutic efficacy in lung cancer cells (57) and squamous cell carcinoma patients (58) harboring gain-of-function DDR2 mutations.

Our data suggest that the inhibition of DDR1 can improve the efficacy of standard chemotherapy for pancreatic cancer. Critical questions regarding the contribution of DDR1 to tumor progression remain, including whether the results we have found will extend to other solid tumors (e.g., lung, breast). Furthermore, it is unclear whether responses to DDR1 inhibition will be limited to those tumors that present with extensive stromal deposition or, as suggested by in vitro data with AsPC-1 cells, autocrine stimulation is critical to DDR1 activation on tumor cells. Addressing these questions is now feasible with the development of highly selective DDR1 inhibitors such as 7rh.

R.A. Brekken has ownership interest (including patents) in Tuevol Therapetics. No potential conflicts of interest were disclosed by the other authors.

Conception and design: K.Y. Aguilera, R.A. Brekken

Development of methodology: K.Y. Aguilera, H. Huang

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): K.Y. Aguilera, H. Huang, W. Du, M.M. Hagopian, Z. Wang, S. Hinz, H. Wang

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): K.Y. Aguilera, H. Huang, W. Du, T.H. Hwang, H. Wang, D.H. Castrillon, R.A. Brekken

Writing, review, and/or revision of the manuscript: K.Y. Aguilera, H. Huang, H. Wang, X. Ren, R.A. Brekken

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): K.Y. Aguilera, J.B. Fleming, X. Ren, K. Ding, R.A. Brekken

Study supervision: K.Y. Aguilera, R.A. Brekken

We gratefully acknowledge Dr. Miao Wang for KPC animal tissue, Dr. Hoon Hur for surgical assistance with the orthotopic implantation of Pan02 cells, Dr. Lisa Li for assistance in therapeutic dosing of animals, Dr. James Kim for helpful comments on the manuscript, and Dave Primm for editorial assistance.

This work is supported in part by the NIH (R01CA118240 and U54 CA210181 Project 2, to R.A. Brekken; F31 CA168350, to K.Y. Aguilera), CPRIT (RP160211, to D.H. Castrillon), the National Science Fund for Distinguished Young Scholars (#81425021, to K. Ding), the 100 Distinguished Scientist Award of Guangdong province (Nanyue-Baijie award, to K. Ding), Guangdong province (grant #2016A050502041, to X. Ren), and the Effie Marie Cain Scholarship for Angiogenesis Research (to R.A. Brekken).

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.