Rhabdomyosarcoma (RMS) is a highly aggressive pediatric cancer with features of skeletal muscle differentiation. More than 80% of the high-risk patients ultimately fail to respond to chemotherapy treatment, leading to limited therapeutic options and dismal prognostic rates. The lack of response and subsequent tumor recurrence is driven in part by stem cell–like cells, the tumor subpopulation that is enriched after treatment, and characterized by expression of the AXL receptor tyrosine kinase (AXL). AXL mediates survival, migration, and therapy resistance in several cancer types; however, its function in RMS remains unclear. In this study, we investigated the role of AXL in RMS tumorigenesis, migration, and chemotherapy response, and whether targeting of AXL with small-molecule inhibitors could potentiate the efficacy of chemotherapy. We show that AXL is expressed in a heterogeneous manner in patient-derived xenografts (PDX), primary cultures and cell line models of RMS, consistent with its stem cell–state selectivity. By generating a CRISPR/Cas9 AXL knock-out and overexpressing models, we show that AXL contributes to the migratory phenotype of RMS, but not to chemotherapy resistance. Instead, pharmacologic blockade with the AXL inhibitors bemcentinib (BGB324), cabozantinib and NPS-1034 rapidly killed RMS cells in an AXL-independent manner and augmented the efficacy of the chemotherapeutics vincristine and cyclophosphamide. In vivo administration of the combination of bemcentinib and vincristine exerted strong antitumoral activity in a rapidly progressing PDX mouse model, significantly reducing tumor burden compared with single-agent treatment. Collectively, our data identify bemcentinib as a promising drug to improve chemotherapy efficacy in patients with RMS.

This article is featured in Selected Articles from This Issue, p. 749

Rhabdomyosarcoma (RMS) is the most common pediatric soft tissue sarcoma and a highly aggressive cancer associated with the skeletal muscle lineage (1). The two main RMS subtypes include fusion-negative (FN-RMS) and fusion-positive RMS (FP-RMS), commonly associated to the embryonal, respectively, alveolar histologic subtypes. FP-RMS represents the more aggressive form characterized by characteristic chromosomal translocations between either PAX3 or PAX7 and FOXO1 (PAX3::FOXO1 or PAX7::FOXO1; ref. 2). Despite intense multichemotherapy treatments with vincristine, actinomycin, and cyclophosphamide (3), one-third of localized and two-thirds of patients with metastatic RMS experience relapse, leading to limited therapeutic options and dismal survival rates of up to <5% in patients with FP-RMS (4, 5). One of the main determinants of drug resistance in cancer is intratumoral heterogeneity (6).

To unbiasedly distinguish rare RMS subpopulations within the heterogeneous tumor, we and other have recently employed single-cell analysis, and uncovered the existence of three main cellular states, including: (i) an immature muscle stem cell (MuSC)-like state, (ii) a highly proliferative progenitor-like state, and (iii) a more differentiated muscle-like state (7–9). While conventional chemotherapy regimens effectively deplete the highly proliferative subpopulation, they concurrently lead to the enrichment of immature MuSC-like cells within both FP-RMS and FN-RMS tumors (7, 9). In response to this challenge, a novel therapeutic approach is the “total clonal” strategy, which relies on identifying therapeutic vulnerabilities specific to this tumor compartment. As such, targeting MuSC-like/mesoderm cells with EGFR inhibitors, in combination with chemotherapy, has been shown to improve outcomes in FN-RMS (7). However, it remains unclear whether the same therapeutic benefit extends to FP-RMS. This prior research thus raises the compelling question of whether cotargeting multiple tumor subpopulations could be of therapeutic benefit in FP-RMS.

We previously observed that MuSC-like FP-RMS cells express unique markers, including CD44, CD105, and AXL, which encodes for the homonymous protein (9). AXL is a receptor tyrosine kinase, which belongs to the TAM (TYRO3-AXL-MER) family, and is activated upon binding of its ligand growth arrest–specific protein 6 (GAS6; refs. 10–12). AXL can also be activated in a GAS6-independent manner, for example by oxidative stress, which can lead to AXL homodimerization and autophosphorylation (13). An increasing body of evidence suggests a role for AXL signaling in mediating drug resistance, proliferation, stem cell phenotype, as well as cell migration and invasion in various cancers (14). As such, multiple kinds of AXL-targeting agents are currently in clinical development (15), including: (i) cabozantinib (16), which targets VEGFR, Met, Flt3, c-Kit, and AXL; (ii) NPS-1034 (17), which targets c-Met and AXL; and (iii) bemcentinib (BGB324; R428; ref. 18), the first molecule to have entered clinical trial as an AXL-specific inhibitor, and currently in phase I/II clinical testing for non–small cell lung cancer (NSCLC, NCT02424617) and phase I for acute myeloid leukemia (NCT02488408). Despite a report showing AXL upregulation following IGF-1R–targeted therapy in RMS cell lines (19), a systematic assessment of AXL expression and therapeutic vulnerability in RMS remains uninvestigated.

In this study, we first assessed AXL expression in RMS cell lines, primary cultures, PDX and patient tumors. By using genetic gain- and loss-of-function strategies, we investigated the role of the GAS6-AXL signaling pathway on RMS cell migration, tumorigenesis, and drug response. We show that AXL mediates FP-RMS cell migration, but it does not affect chemotherapy response. We finally assessed the in vitro and in vivo efficacy of pharmacologic AXL inhibition either alone or in combination with chemotherapy, and uncovered a potent AXL-independent effect of bemcentinib as chemosensitizer, providing a new rationale for the clinical evaluation of this molecule in unresponsive RMS patients.

Patient-derived xenografts

All animal experiments were conducted under license of the authorities in compliance with the national laws and regulations and approved by the Zürich canton government (license number ZH013/2021). The patient-derived xenografts (PDX) used in this study were generated from patient biopsy samples collected at St Jude Children's Research Hospital, University Children's Hospital Zurich, Emma Children's Hospital Amsterdam, Institut Curie Paris, and Charité University Hospital Berlin as described previously (9, 20). Patient characteristics and information on the clinical status can be found in Supplementary Table S1. Patient biopsies were first expanded in immunodeficient NOD scid gamma (NSG), Janvier Rj:NMRI-Foxn1nu/nu, or Taconic NOD.Cg-Prkdcscid Il2rgtm1Sug/JicTac mice. Tumors were isolated from mice when reaching a size of 700–1,300 mm3, mechanically minced into smaller pieces using scalpels and retransplanted in secondary recipient mice. To generate single-cell suspensions, PDX tumors were mechanically and enzymatically digested using 200 μg/mL liberase DH (Roche, 5401054001) and 1 mmol/L MgCl2 in 1× HBSS buffer (Sigma, H6648) for 30–60 minutes at 37°C. Cell suspension was filtered through a 70-μm cell strainer, to remove remaining tumor pieces, washed with PBS, and used immediately to produce cultured cells or frozen in freezing medium CryoStor CS10 (StemCell Technologies, #07930).

In vivo drug testing in PDXs

For in vivo drug testing, three million P-1 cells were first expanded in vitro as primary cultures, resuspended in a 1:1 Matrigel (Corning, 354234):Advanced DMEM/F-12 medium (Thermo Fisher Scientific, 12634010) solution, and injected subcutaneously into the right flank of 6- to 10-week-old female NSG mice (n = 24 mice in total). When tumors became palpable (tumor size between 10 and 100 mm3), mice were randomly enrolled into four treatment groups (vehicle-treated control, bemcentinib, vincristine sulfate, or their combination). We excluded n = 2 mice in which no tumor developed, leading to treatment groups of n = 5 mice for vehicle and bemcentinib, and n = 6 for vincristine sulfate and the drug combination. Drugs were dosed as follows, based on a recent publication (21): vincristine sulfate (0.5 mg/kg; ApexBio, A1765) dissolved in PBS and administered by intraperitoneal injection one time a week; bemcentinib (50 mg/kg; SelleckChem, HY-15150) dissolved in double-distilled water with 1% w/w methylcellulose 400 cp (Sigma-Aldrich, M0262), 0.1% Tween-80 (Sigma-Aldrich, P4780), and administered by oral gavage two times a day, five times a week. The cycle was repeated for a total of two weeks. Tumor volume was measured three times a week using a caliper, and mice were euthanized once tumor volume exceeded 1,000 mm3, which was considered the experiment endpoint. During treatment, the mouse weight was monitored daily to ensure no treatment toxicities. Treatment responses were classified according to the Pediatric Preclinical In Vivo Testing Consortium (PIVOT) guidelines.

Primary cultures

To produce primary cultures, PDX tumors were dissociated as described above, and grown on plates coated with Matrigel (Corning, 354234) diluted 1:10 in Advanced DMEM/F-12 medium (Thermo Fisher Scientific, 12634010). Clinical information and culture characteristics for each model can be found in Supplementary Table S1. Cells were cultured in Advanced DMEM/F-12 (Thermo Fisher Scientific, 12634010) medium supplemented with 100 U/mL penicillin/streptomycin (Thermo Fisher, 15140122), 2 mmol/L Glutamax (Life Technologies, 335050–061), 0.75x B-27 (Thermo Fisher Scientific, 17504044), 20 ng/mL bFGF (PeproTech, AF-100–18B), and 20 ng/mL EGF (PeproTech, AF-100–15; “Complete F12” medium) or in Neurobasal medium (Thermo Fisher Scientific) supplemented with 100 U/mL penicillin/streptomycin (Thermo Fisher Scientific, 15140122), 2 mmol/L Glutamax (Life Technologies, 335050–061), 2× B-27 (Thermo Fisher Scientific, 17504044) (“Complete NB” medium). In some cases (see Supplementary Table S1), the medium was supplemented with 20 ng/mL bFGF (PeproTech, AF-100–18B) and 20 ng/mL EGF (PeproTech, AF-100–15). For further passaging, cells were washed with PBS and detached with Accutase (Sigma-Aldrich, A6964) diluted 1:3 in PBS. All RMS primary cultures were regularly tested to ensure no Mycoplasma contamination with the LookOut Mycoplasma PCR Detection Kit (Sigma-Aldrich, MP0035–1KT).

Culture of cell lines

The cell lines Rh41 (RRID: CVCL_2176; provided by Peter Houghton, Greehey Children's Cancer Research Institute, San Antonio, TX), Rh4 (RRID: CVCL_5916; provided by Peter Houghton, Greehey Children's Cancer Research Institute, San Antonio, TX), RMS (ref. 22; provided from J. Shipley, Sarcoma Molecular Pathology, The Institute of Cancer Research), KFR (RRID: CVCL_S637; provided by Jindrich Cinatl, Abteilung für paediatrische Tumor und Virusforschung, Frankfurter Stiftung für krebskranke Kinder, Frankfurt, Germany), and HEK293T (RRID: CVCL_0063) were cultured on uncoated plates in DMEM (Sigma Aldrich, D5671) supplemented with 10% FBS (Life Technologies), 100 U/mL penicillin/streptomycin (Thermo Fisher Scientific, 15140122) and 2 mmol/L Glutamax (Life Technologies, 335050–061). For further passaging, cells were washed with PBS and detached with Trypsin (BioConcept, 5–51F00-I) or Accutase (Sigma-Aldrich, A6964) diluted 1:2 to 1:3 in PBS. All cell lines were authenticated by short tandem repeat analysis (STR) profiling and regularly tested to ensure no Mycoplasma contamination with the LookOut Mycoplasma PCR Detection Kit (Sigma-Aldrich, MP0035–1KT).

Generation of CRISPR/cas9 AXL knockout lines

To generate AXL knockout (KO) lines, we first designed n = 3 guide RNAs against different exons of AXL using CRISPOR (ref. 23; Supplementary Tables S2 and S3), and cloned them into the lentiCRISPRv2 puro expression vector (#98290, RRID:Addgene_98290). We included a control guide targeting the AAVS1 genomic safe-harbor locus (Supplementary Tables S2 and S3). Lentiviral particles containing the desired vectors were produced in HEK293T cells by cotransfecting the cells with second-generation packaging plasmids (psPAX2 #12260 and VSVG #14888; RRID:Addgene_12260, respectively, RRID:Addgene_14888) and the respective transfer plasmids (containing Cas9 and the guide RNAs) using calcium phosphate. After 24 hours, the medium was replaced. Viruses were harvested after additional 48 hours by collecting and concentrating the supernatants with Amicon Ultra centrifugal filter units (Sigma-Aldrich, UFC910024). Rh41 and IC-pPDX-104 cells were infected with the lentiviral particles, and selected with 0.5 ng/μL and, respectively, 0.3 ng/μL puromycin for 5 days.

Generation of AXL-overexpressing lines

To generate AXL overexpressing (OE) lines, we first produced lentiviral particles in HEK293T cells by co-transfecting the cells with a vector with the coding region of AXL (pLX304-AXL; ref. 24), kindly donated by Prof. Yosef Yarden) and second-generation packaging plasmids (psPAX2 #12260 and VSVG #14888, both from Addgene) using calcium phosphate. After 24 hours, the medium was replaced. Viruses were harvested after additional 48 hours by collecting and concentrating the supernatants with Amicon Ultra Centrifugal Filter Units (Sigma-Aldrich, UFC910024). Rh41 and IC-pPDX-104 cells were infected with the lentiviral particles to produce AXL OE lines.

Analysis of AXL gene expression and genetic dependency

To assess AXL expression in patients with RMS, we used a public dataset (25) available on the R2: Genomics Analysis and Visualization Platform (http://r2.amc.nl). To assess AXL expression across cancer cell lines, we downloaded the Expression Public 22Q4+ dataset available on the Cancer Dependency Map (DepMap) online platform (https://depmap.org) containing expression of n = 1,408 cell lines (26, 27).

To assess the genetic dependency of RMS and other cancer cell lines to AXL loss-of-function, we downloaded two datasets available on the Cancer Dependency Map (DepMap) online platform (https://depmap.org; refs. 28–31). The first dataset (CRISPR DepMap Public 22Q4+ screen) contains dependency scores calculated after CRISPR/Cas9 genetic removal of AXL (n = 1,078 cell lines tested, including n = 13 RMS). The second dataset (Achilles+DRIVE+Marcotte RNAi screen) contains dependency scores calculated after AXL gene silencing using RNAi (n = 710 cell lines tested, including n = 7 RMS). The DepMap datasets were plotted in RStudio v1.3.1093 using ggplot2.

Western blotting

Whole-cell lysates were extracted using RIPA lysis buffer (50 mmol/L Tris-HCl pH 7.5, 150 mmol/L NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 1 mmol/L EGTA, 50 mmol/L NaF, 5 mmol/L Na4P2O7, 1 mmol/L Na3VO4, and 10 mmol/L β-glycerol phosphate) in the presence of the cOmplete, Mini, EDTA-free Protease Inhibitor Cocktail (Sigma Aldrich, 11836170001). Protein concentration was measured with Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, 23227) according to the manufacturer's instructions. 5–20 μg of whole-cell lysates were reduced with Laemmli Sample Buffer (Bio-Rad, 161–0747) supplemented with 1:20 DTT. After boiling the samples at 95°C for 5 minutes, proteins were separated using NuPAGE 4%–12%, Bis-Tris, 1.0 mm, Mini Protein Gels (Thermo Fisher Scientific, NP0323BOX) or 4%–20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad, 4561096). Gels were transferred on membranes using Trans-Blot Turbo Transfer System (Bio-Rad, 1704150). After blocking with 5% milk (Carl Roth, T145.3) in TBS/0.05% Tween (Sigma Aldrich, P9416) for 30–60 minutes, membranes were incubated overnight at 4°C or for 2 hours at room temperature with primary antibodies diluted 1:1,000. After three washing steps with TBS/0.05% Tween for 5 minutes, membranes were incubated for 1 hour at room temeprature with HRP-linked secondary antibodies diluted 1:5,000. Finally, after three additional washing steps with TBS/0.05% Tween for 5 minutes, proteins were detected by chemiluminescence using SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Fisher Scientific, 34095) and a ChemiDoc MP imager (Bio-Rad). The following antibodies were used: AXL (Santa Cruz Biotechnology, sc-166269; RRID:AB_2243305), phospho-AXL (Tyr702; Cell Signaling Technology, 5724S; RRID:AB_10544794), phospho-Akt (Ser473; Cell Signaling Technology, 4060L; RRID:AB_2315049), Akt (Cell Signaling Technology, 9272S; RRID:AB_329827), phosphorylated ERK (Thr202/Tyr204; Cell Signaling Technology, 9101S; RRID:AB_331646), ERK1/2 (Cell Signaling Technology, 9102L; RRID:AB_330744), GAPDH (Cell Signaling Technology, 5174S; RRID:AB_10622025), anti-rabbit IgG HRP-linked antibody (Cell Signaling Technology, 7074S; RRID:AB_2099233), anti-mouse IgG HRP-linked antibody (Cell Signaling Technology, 7076S; RRID:AB_330924).

qRT-PCR

Total RNA was extracted using the RNeasy Micro Kit (Qiagen, 74004) from cultured cells and using the RNeasy Mini Kit (Qiagen, 74106) from tumor pieces according to manufacturer's instructions. cDNA was synthesized using the High-Capacity cDNA Reverse Transcription Kit with RNase Inhibitor (Thermo Fisher Scientific, 4374967) modifying the total volume to 10 μL. Specifically, for each sample, a mix consisting of 1.0 μL 10X RT Buffer, 0.4 μL 25XdNTPs Mix (100 mmol/L), 1.0 μL 10X RT Random Primers, 0.5 μL MultiScribeTM Reverse Transcriptase, 0.5 μL RNase inhibitor, 1.6 μL Nuclease-free H2O, and 5 μL Nuclease-Free Water (Thermo Fisher Scientific, AM9937) was incubated in a thermocycler according to manufacturer's instructions. Each sample was analyzed by qRT-PCR using TaqMan Gene Expression Master Mix (Thermo Fisher Scientific, 4369542) and TaqMan gene expression assays. qRT-PCR was performed in technical triplicates for each sample on a 7900HT Fast Real-Time PCR machine. To calculate the relative gene expression of each gene, the 2−ΔΔCt method was used and the quantity of RNA was normalized to the internal control GAPDH. The following TaqMan gene expression assays were used: CD44 (Hs01075864_m1), AXL (Hs01064444_m1), CAV1 (Hs00971716_m1), MYOD1 (Hs00159528_m1), MYOG (Hs01072232_m1), MYH3 (Hs01074230_m1), and GAPDH (Hs02758991_g1).

IHC

PDX tumor pieces fixed in ROTI Histofix 4% (Carl Roth, P087.3) were first embedded with paraffin as formalin-fixed paraffin-embedded (FFPE) tissues, and cut in sections of 2 μm. Sections were stained on the BOND-III Fully Automated Stainer (Leica), and incubated for 30 minutes with primary antibodies against AXL (R&D Systems, AF154; RRID:AB_354852) at a dilution of 1:200. Sections were visualized with a Bond Polymer Refine Detection System (Leica). All sections were counterstained with hematoxylin.

Flow cytometry analysis

To measure AXL, CD44, and CD105 cell surface expression of RMS primary cultures and cell lines, we labeled cells with the following antibodies at a dilution of 1:50: allophycocyanin (APC) AXL (Thermo Fisher Scientific, 17–1087–41; RRID:AB_2723956), FITC anti-human CD44 (BioLegend, 338804; RRID:AB_1501197), and phycoerythrin (PE) anti-CD105 (BioLegend, 800503; RRID:AB_2629655). To gate positive/negative cells, the following isotype controls were used: APC mouse IgG1 (BioLegend, 400122; RRID:AB_326443), FITC mouse IgG1 (BioLegend, 400110; RRID:AB_2861401), and PE mouse IgG1 (BioLegend, 400114; RRID:AB_326435). All FACS analysis were done on FlowJo v10.8 software. Cells were acquired on BD LSRFortessa (BD Biosciences).

In vitro drug treatment

Drugs included vincristine sulfate (ApexBio, A1765–5.1), 4-Hydroperoxycyclophosphamide (4-HC: Niomech, CAS 39800–16–3), bemcentinib (BGB324, Selleckchem, S2841), cabozantinib (Selleckchem, S1119), NPS-1034 (Selleckchem, S7669). To test the sensitivity of AXL inhibitors, cells were plated in 384-well plates at a density of 3,000–4,000 cells/well, equilibrated overnight, and then treated with the inhibitors for 72 hours. For the combination of AXL inhibitors with chemotherapy, cells were plated as described above and pretreated with bemcentinib, cabozantinib, NPS-1034, or DMSO-control for 4 hours, then vincristine sulfate or 4-HC were added for other 72 hours. Cell viability was measured by WST-1 assay according to manufacturer's instructions (Sigma Aldrich, 11644807001), and normalized to DMSO (vehicle)-treated controls. For qRT-PCR analysis, cells were plated in 6-well plates at a concentration of 300,000 cells/well, equilibrated overnight, and then treated with DMSO (vehicle) control or with the drugs for the indicated time. For flow cytometry analysis, cells were plated in 24-well plates at a concentration of 100,000 cells per well, equilibrated overnight, and then treated with DMSO (vehicle) control or with the indicated drugs for 48 hours. IC50 values were determined from the dose–response curves generated using GraphPad Prism (version 9.5.1).

GAS6 stimulation

To analyze GAS6-mediated phosphorylation of AXL, and AKT, we plated cells in 6-well plates, starved them overnight by removing FBS or the growth factors (FGF and EGF) from the medium. The next day, we stimulated cells with the indicated concentration of GAS6 (R&D Systems, 885-GSB-050) for 10 minutes at 37°C, and processed them for Western blot analysis.

Measurement of apoptosis

To measure apoptosis, cells were culture in 6-well plates, treated with the indicated drugs for 48 hours, and stained using the eBioscience Annexin V Apoptosis Detection Kit APC (88–8007–72, Invitrogen) according to the manufacturer's instructions. Cells were acquired on a BD LSRFortessa or BD FACSCelesta (BD Biosciences).

Wound-healing assay

Cell migration assay was performed using culture-inserts 2 well (Ibidi, 80209), where 50,000 cells/well were plated. After incubation for 24 hours at 37°C, the inserts were removed, and cells treated with bemcentinib (0.5 μmol/L or 1 μmol/L) or with GAS6 (400 ng/mL). Cells were imaged by phase contrast microscopy at regular time intervals of 1 hour using the Celloger Mini System (Curiosis). For each condition, 2–3 different regions were imaged from 2 to 4 technical replicates. The wound gap was quantified by ImageJ using the Wound healing macro tool.

Statistical analysis

Data are provided as mean ± SEM from at least n = 3 independent biological replicates experiments, unless stated otherwise. Statistical analyses of differences between groups and IC50 calculations were performed using Prism version 10. Statistical significance was calculated either by the Student t test, ordinary one-way ANOVA or two-way ANOVA, as specified accordingly. Significance is annotated as follows: ns, nonsignificant (P > 0.05); *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; and ****, P ≤ 0.0001.

Data availability

The study did not generate new unique reagents. The PDX models generated in Zürich are available upon request to the corresponding author. The data generated in this study are available within the article and Supplementary Data.

AXL is expressed by the MuSC-like subpopulation of RMS cells

We and others have previously observed that AXL is predominantly expressed by the MuSC-like compartment of FN-RMS and FP-RMS tumors (Fig. 1A; refs. 7, 9); however, its expression and role in rhabdomyosarcomagenesis has not been investigated to date. We assessed total AXL protein and cell surface expression by Western blot analysis (Fig. 1B; Supplementary Fig. S1A), respectively, flow cytometry (Fig. 1C) in a panel of primary cultures and cell lines derived from primary and metastatic sites, diagnostic, and recurrent RMS patients (Supplementary Table S1). We observed cell surface AXL expression in 74% (14/19) of all samples [88% (7/8) of FN-RMS and 64% (7/11) of FP-RMS], and total AXL expression in 61% (11/18) of all samples [89% (8/9) of FN-RMS and 33% (3/9) of FP-RMS], indicating that most AXL localizes at the cell surface membrane of RMS cells (Fig. 1D; Supplementary Table S4). Interestingly, all patients with RMS showed heterogeneity in expression of AXL, with positivity ranging from <1% to 90% of all cells (Fig. 1C), validating the heterogeneity observed at the transcriptomic level (Fig. 1E). FN-RMS showed significantly higher AXL expression and more AXL+ cells compared with FP-RMS samples (Supplementary Fig. S1B), consistent with the higher frequency of MuSC-like cells (7–9); however, in both RMS subtypes, we identified samples with high AXL expression and patients with low or no AXL expression. We further observed a correlation between AXL and CD44 and CD105 expression (Supplementary Fig. S1C), which are markers of the MuSC-like FP-RMS state (9), supporting the MuSC-like selective expression of this protein. We demonstrated the existence of AXL+ cells at the tissue level by performing IHC staining of PDX tumors. PDXs of FP-RMS showed low or no expression of AXL, except for some patients that displayed heterogeneous expression in the cytoplasm, whereas PDXs of FN-RMS generally displayed stronger expression, similarly to what observed in primary cultures sections (Fig. 1F). These findings were validated in patient specimens using a large microarray dataset (25) available on the R2 platform (r2.aml.nl), which confirmed statistically higher AXL mRNA expression in patients with FN-RMS compared with FP-RMS (Fig. 1G).

We next wondered how the expression of AXL in RMS compares with other cancer types for which AXL-directed therapies are currently in clinical trial. We analyzed AXL mRNA expression in n = 1,408 cell lines available on the BROAD DepMap online platform (https://depmap.org; refs. 26, 27). We observed that AXL levels in soft tissue cancers, which include RMS, are comparable to the ones of breast, lung and CNS/brain, for which AXL therapies are currently in clinical trials (Fig. 1H and I). Finally, we compared the genetic dependency of RMS and other cancer lineages to AXL loss-of-function using the CRISPR DepMap Public 22Q4+ screen (n = 1,078 cell lines tested, including n = 13 RMS) or Achilles+DRIVE+Marcotte RNA interference (RNAi) screen (n = 710 cell lines tested, including n = 7 RMS) available on the BROAD DepMap online platform (https://depmap.org; refs. 28–31). We observed no clear dependencies (defined as scores <0.5) across all tumor models upon AXL removal (Supplementary Fig. S1D). Overall, our results indicate that AXL is expressed by a subset of FP-RMS and FN-RMS cells associated with the immature MuSC-like phenotype (Fig. 1J).

GAS6 induces robust activation of AXL in FP-RMS

Having established that AXL is expressed by RMS cells, we sought to better investigate its functional relevance. We focused on FP-RMS, as our previous findings suggest that chemotherapy enriches this tumor type for MuSC-like cells (9). We hypothesized that AXL could be a therapeutic vulnerability of MuSC-like FP-RMS cells, and that cotargeting multiple tumor subpopulations using chemotherapy and MuSC-like inhibitors could be of therapeutic advantage, as shown in FN-RMS tumors (7). We first studied AXL functionality in FP-RMS by genetic gain- and loss-of-function strategies. We selected the primary PDX culture (IC-pPDX-104) and the cell line Rh41 as experimental models, as they displayed the highest AXL expression levels among FP-RMS (Fig. 1B and C). We established CRISPR-Cas9–based AXL knock-out cells using three different sgRNAs, as well as control cells (denoted WT) using a guide targeting the nonessential AAVS1 locus. We selected sgAXL-1 and sgAXL-2 (denoted KO) for follow-up studies, as they demonstrated high knock-out efficiency (Supplementary Fig. S2A). Similarly, we established AXL-overexpressing cells by transduction with a lentiviral construct containing the coding region of AXL (denoted OE; ref. 24). Western blot analysis confirmed lack and overexpression of AXL in KO and OE lines, respectively, compared with WT cells (Fig. 2A; Supplementary Fig. S2A). We did not observe differences in morphologic appearance (Fig. 2B) or proliferation rate if not slightly higher proliferation rates in OE lines (Supplementary Fig. S2B). To functionally verify our model system, we evaluated the ability of GAS6 to activate AXL and its downstream effectors, AKT and ERK (Fig. 2C). We observed robust activation of AXL signaling following treatments with >100 ng/mL GAS6 in WT and OE cells, but not in KO lines (Supplementary Fig. S2C; Fig. 2D). We did not observe any significant activation of the downstream signaling effectors AKT and ERK following GAS6 stimulation (Fig. 2E; Supplementary Fig. S2D). These results demonstrate that AXL activation is modulated in a GAS6-dependent manner in FP-RMS cells.

Bemcentinib, cabozantinib, and NPS-1034 inhibit AXL activation and suppress RMS cell growth in an AXL-independent manner

Several AXL kinase inhibitors are currently being tested in various solid tumors, including lung and breast cancer (12). We therefore investigated the effect of the small-molecule AXL inhibitors bemcentinib, cabozantinib and NPS-1034 on AXL pathway activation in FP-RMS cells. We incubated IC-pPDX-104 and Rh41 cells with increasing concentrations of the AXL inhibitors, and analyzed phosphorylation of AXL (pAXL) and of its downstream effector AKT (pAKT) by Western blot analysis. All the inhibitors effectively blocked pAXL and pAKT signaling at the higher concentrations in IC-pPDX-104 cells, but only cabozantinib reduced pAXL levels in Rh41 cells (Fig. 3A). Cabozantinib demonstrated the highest potency in blocking AXL phosphorylation (Fig. 3B).

We next investigated the therapeutic potential of pharmacological AXL inhibition, by exposing RMS primary PDX cultures and cell lines to bemcentinib, cabozantinib and NPS-1034. All the inhibitors reduced RMS viability in a dose-dependent manner (Fig. 3C ; Supplementary Fig. S3A). Bemcentinib was the most potent inhibitor with half-maximal inhibitory concentration (IC50) values in the low micromolar range (1.2 ± 0.1 μmol/L), compared with cabozantinib (27 ± 9 μmol/L) and NPS-1034 (23 ± 14 μmol/L, Supplementary Table S5). Bemcentinib showed similar IC50 values in the two RMS subtypes (Supplementary Fig. S3B). We observed no correlation between AXL protein expression and in vitro cytotoxicity of the inhibitors, suggesting that the drugs impact cell viability by AXL-independent mechanisms (Supplementary Fig. S3C). To confirm this hypothesis in FP-RMS samples, we measured the effect of the drugs on the viability of WT, KO, and OE IC-pPDX-104 and Rh41 cells. We observed similar dose–response curves, arguing, again, for AXL-independent effects (Supplementary Fig. S3D). Because bemcentinib demonstrated the highest potency in killing RMS cells, we further focused on this inhibitor. We compared the cytotoxicity of bemcentinib upon AXL pathway activation with GAS6 in IC-pPDX-104 cells. Again, we observed no difference in IC50 (Supplementary Fig. S3E), suggesting that neither AXL protein levels nor its activation status influence the effect of the inhibitors. Taken together, our results indicate that bemcentinib inhibits FP-RMS growth by AXL-independent mechanisms.

Bemcentinib sensitizes FP-RMS cells to chemotherapy

We have recently shown that chemotherapy enriches FP-RMS primary cultures, including IC-pPDX-104, in AXL+/CD44+/CD105+/MYOGENIN MuSC-like cells (9). We analyzed the effects of vincristine and 4-Hydroperoxycyclophosphamide (4-HC), the active metabolite of cyclophosphamide, on the cell line Rh41, and confirmed upregulation of AXL gene expression levels and increase in AXL+ cells (Fig. 4A). These results led us to question whether AXL inhibition could enhance cytotoxicity of chemotherapy (Fig. 4B).

To assess whether AXL inhibitors could increase the efficacy of chemotherapy, we pretreated IC-pPDX-104 and Rh41 cells with subtoxic concentrations of bemcentinib, cabozantinib, or NPS-1034 for 4 hours, then added the chemotherapeutics vincristine sulfate or 4-HC for 72 hours, and assessed the cell viability by WST-1 assay. We found that bemcentinib and cabozantinib enhanced the efficacy of both chemotherapeutics by significantly diminishing their IC50 values, whereas combinations with NPS-1034 failed to reach statistical significance (Fig. 4C; Supplementary Fig. S4A).

We next wondered whether pharmacologic AXL inhibition prevents chemotherapy-induced tumor de-differentiation, and therefore enrichment in MuSC-like cells. We treated IC-pPDX-104 cells with bemcentinib, vincristine, or their combination, and measured tumor lineage markers by flow cytometry and qRT-PCR. We observed the already described (9) upregulation of MuSC-like markers AXL, CD44, CAV1 (respectively, AXL+, CD44+, CD105+ cells) upon exposure to vincristine (Supplementary Fig. S4B and S4C). We did not observe a reversed effect in cells treated with bemcentinib or with the drug combination, but we rather found a similar increase in MuSC-like markers (Supplementary Fig. S4B and S4C), indicating that bemcentinib does not prevent chemotherapy-induced lineage shifts.

We wondered whether high AXL levels confer resistance to chemotherapy. We compared the efficacy of vincristine and 4-HC in FP-RMS cells with WT, KO, or OE AXL levels. We did not observe chemotherapy sensitivity in KO lines, nor resistance in OE lines, indicating that AXL does not affect FP-RMS treatment response (Supplementary Fig. S4D). Because AXL levels do not necessarily correlate with AXL pathway activation, we further compared the sensitivity of IC-pPDX-104 cells to vincristine and 4-HC upon AXL stimulation with GAS6. Again, we observed no difference between stimulated and unstimulated cells (Supplementary Fig. S4E), suggesting that neither AXL levels nor its activation status influence chemotherapy response in FP-RMS. Furthermore, we tested whether AXL supports FP-RMS cells in expanding and driving recurrence after treatment. We measured the growth of AXL WT and KO cells following treatments with vincristine, and observed similar recovery rates (Supplementary Fig. S4F), suggesting that AXL does not have a role in FP-RMS drug resistance or recurrence.

Finally, we wished to determine the mechanism by which bemcentinib enhances the cytotoxicity of vincristine. We measured apoptosis in Rh41 and IC-pPDX-104 cells treated with single-agent vincristine or with its combination with bemcentinib, and observed significantly more apoptotic cells in the drug combination (Fig. 4D and E). Overall, these data suggest that AXL by itself does not confer chemoresistance, however, the AXL inhibitors bemcentinib and cabozantinib can augment the efficacy of chemotherapeutics.

AXL contributes to the migratory phenotype of Rh41 cells

In addition to being essential for cancer growth, AXL has been described to promote migration and invasion in numerous cancer types (32, 33). We therefore wondered whether AXL contributes to the migratory phenotype of FP-RMS cells, given the tendency of this cancer type to metastasize to distant organs (2). We utilized the wound-healing assay, a technique that assesses cell migration by measuring the speed at which cells move through a wound field gap, and applied it to Rh41 cells with different AXL levels and/or following exposure to bemcentinib. AXL KO (sgAXL-1) cells showed significantly reduced wound closure rates compared with AXL WT cells (Fig. 5AC), suggesting that AXL contributes to a more migratory phenotype in Rh41 cells. We observed the same trend in Rh41 cells transduced with a second guide against AXL (sgAXL-2), even though it failed to reach statistical significance (Fig. 5B and C). Importantly, AXL OE cells demonstrated a tendency toward accelerated wound closure rates (Fig. 5B and C), confirming a role for this protein in mediating Rh41 cell migration.

We next wondered whether pharmacologic AXL inhibition with bemcentinib phenocopies the effect of its genetic KO on cell migration. Indeed, treatment with bemcentinib at subtoxic concentrations inhibited the wound closure rate of Rh41 cells in a dose-dependent manner (Fig. 5DF). Given the AXL-independent effects of the inhibitor on FP-RMS cell growth inhibition, we tested whether AXL KO would be as responsive as AXL WT cells to bemcentinib treatment. We observed similar inhibition of cell migration in AXL WT and KO (sgAXL-1 and sgAXL-2) cells (Fig. 5G and H), suggesting that the drug inhibits Rh41 migration by AXL-independent mechanisms. Finally, we tested whether GAS6 supplementation accelerates cell migration. We observed no significant increase in migration compared with untreated cells (Fig. 5G and H). Overall, these data suggest that AXL levels but not GAS6-induced activation confer increased migratory capacity to Rh41 cells. This phenotype can be abrogated by treatment with the AXL inhibitor bemcentinib, which acts in an AXL-unspecific manner and thereby inhibits migration of Rh41 cells independently from their AXL levels.

The combination of bemcentinib and vincristine exhibits potent in vivo antitumoral activity

We finally examined the translational in vivo relevance of the combination of vincristine and bemcentinib in a rapidly growing FP-RMS PDX model. We injected IC-pPDX-104 cells subcutaneously, and monitored tumor volume in mice treated with bemcentinib (50 mg/kg daily), vincristine (0.5 mg/kg weekly), or their combination for two weeks. On the basis of the mouse weight change, we observed no treatment-induced toxicity (Supplementary Fig. S5A). Single-agent treatment with bemcentinib did not reduce tumor volume nor prolonged mouse survival (Fig. 6AC; Supplementary Fig. S5B), suggesting that the drug alone does not impair tumor growth. Indeed, all mice treated with bemcentinib experienced progressive disease (PD; Fig. 6D; Supplementary Fig. S5C). On the other hand, single-agent treatment with vincristine significantly reduced tumor volume, and slightly prolonged mouse survival (Fig. 6AC; Supplementary Fig. S5B). Notably, the first dose of vincristine largely decreased tumor volume, whereas the second dose showed weaker effects (Fig. 6A), suggesting that tumors rapidly develop resistance, similarly to clinical settings. The combination of vincristine and bemcentinib not only significantly reduced tumor volume throughout the treatment to a larger extent than single-agent treatments (Fig. 6C), but it also delayed the tumor growth posttreatment (Fig. 6A; Supplementary Fig. S5B), and significantly improved mouse survival (Fig. 6B). Compared with single-agent vincristine, which elicited partial response (PR) in four out of six mice and PD in the other two mice, the combination of vincristine and bemcentinib elicited PR in two mice, PD in one mouse, and complete response (CR) in three out of six mice, with tumors remaining undetected for up to 14 days (Fig. 6D; Supplementary Fig. S5C; Supplementary Table S6). These results indicate that treatment with bemcentinib improves the in vivo sensitivity of FP-RMS PDXs to chemotherapy.

Resistance to therapy is a major challenge in anticancer treatment, and new approaches to overcome it are desperately needed. AXL is a receptor tyrosine kinase with emerging role in mediating resistance to targeted drugs and conventional chemotherapies in several solid cancers (34). AXL kinase inhibitors are attractive therapeutic options, and reached clinical testing for several cancer types, including glioblastoma (NCT03965494), non–small cell lung (NCT03184571), pancreatic (NCT03649321), and breast cancer (NCT03184558; ref. 32). Despite a few studies showing that AXL is expressed in some of the most common soft tissue sarcoma histologic subtypes (35, 36), its expression and therapeutic potential remain unexplored in RMS.

Here, we first evaluated AXL expression and efficacy of three AXL kinase inhibitors in RMS, an aggressive pediatric cancer with limited therapeutic options in relapsed settings. We show that AXL is expressed in a subset (∼60%) of RMS patients, at a frequency higher than what observed in dedifferentiated and pleomorphic liposarcomas (27.7% of all samples) (36), but similar to Ewing sarcoma (72% of all samples; ref. 37). We also show that, within individual RMS samples, AXL expression is heterogeneous, with AXL+ cells ranging from 0% to 90%. Moreover, AXL expression appears higher in FN-RMS than FP-RMS patients. As we and others have previously shown, AXL is predominantly expressed by the immature MuSC-like RMS state, a subpopulation with similarities to mesenchymal stem cells, that is highly abundant in FN-RMS (7, 9). Therefore we attribute the differences in expression between the two RMS subtypes to the difference in MuSC-like cell frequency.

The role of AXL in promoting tumor cell proliferation is debated, with some cancer types, such as squamous cell carcinomas, pancreatic cancers, B-cell chronic lymphocytic leukemia, and glioblastoma (38–41), showing impaired growth upon AXL genetic and therapeutic inhibition, but other cancer types, including ovarian or breast cancer, being unaffected (42, 43). Here, we demonstrate that FP-RMS cells, at least in vitro, are not genetically dependent on AXL, but we uncover an undescribed susceptibility of FP-RMS cells to AXL inhibitors, in particular to bemcentinib. Our data argue for AXL-independent effects of bemcentinib, as the drug inhibited FP-RMS cell growth and migration regardless of their intrinsic AXL expression or activation status. These AXL-independent effects of bemcentinib in RMS align with recent reports in other cancer types, highlighting off-target toxicity effects of the drug (33, 44, 45). As initially proposed by Chen and colleagues (45), and later confirmed by Zdzalik-Bielecka (33) and colleagues, bemcentinib acts as a lysosomotropic compound, impairing the endo-lysosomal compartment and autophagic flux in NSCLC and glioblastoma cells independently of AXL. In FP-RMS, such a mechanism might not only explain the AXL-independent effect of bemcentinib in inhibiting cell proliferation, but also its beneficial combinatorial effect with chemotherapeutic drugs. Considering that bemcentinib is currently in clinical testing as a selective AXL inhibitor, these findings warrant further examination.

Several studies have examined the relationship between AXL expression and chemoresistance in various cancer types (34, 46–49). We and other have previously shown that chemotherapy enriches RMS cells for the mesoderm/MuSC-like cellular state, with consequent upregulation of AXL mRNA and protein expression (7, 9). These findings raise the question of whether AXL mediates drug resistance in RMS. Using a variety of genetic models and pathway modulation with GAS6, we were able to disprove a direct role of AXL in mediating FP-RMS drug resistance. This is in contrast to other cancer types, such as ovarian, NSCLC, or astrocytoma, where AXL knockdown enhanced chemosensitivity by promoting apoptosis (50–52).

Even though AXL expression levels or AXL pathway activation do not mediate chemoresistance in FP-RMS, here we show that bemcentinib greatly increases the in vitro efficacy of the chemotherapeutics vincristine and 4-HC in FP-RMS cells by augmenting chemotherapy-induced apoptosis. Most importantly, we could recapitulate the same responses in vivo using a rapidly growing PDX model of FP-RMS, where the combination of bemcentinib with vincristine significantly decreased tumor burden and prolonged mouse survival. Our findings are in line with studies on ovarian cancer (52), where bemcentinib improved response to paclitaxel and carboplatin, or on mesothelioma, where bemcentinib enhanced the efficacy of cisplatin and pemetrexed (53).

Finally, AXL is a putative driver of invasiveness and migration in several cancer types (12, 54). Here, we show that AXL controls the migration of Rh41 cells, consistent with its primary role in mediating cell adhesion (44), and with its selective expression in the cell adhesion–enriched MuSC-like FP-RMS clusters (9). In addition to genetic AXL KO, treatment with bemcentinib also impaired FP-RMS cell migration, suggesting that the AXL inhibitor may provide dual benefits in FP-RMS, potentiating the cytotoxicity of chemotherapy and blocking cell dissemination. While our findings reveal AXL's involvement in the migratory phenotype of Rh41 cells, technical limitations arising from modified culture conditions hindered our ability to demonstrate a similar effect in the primary culture IC-pPDX-104. Future studies should thus further investigate the AXL-independent mechanisms of action of bemcentinib in additional FP-RMS cell lines. Moreover, these studies should aim at characterizing whether AXL plays a functional role in FN-RMS tumors, known for their high enrichment in MuSC-like AXL+ cells.

No disclosures were reported.

S.G. Danielli: Conceptualization, data curation, formal analysis, funding acquisition, validation, investigation, methodology, writing–original draft, writing–review and editing. J. Wurth: Formal analysis, investigation, visualization. S. Morice: Formal analysis, investigation, visualization, methodology. S. Kisele: Investigation, methodology. D. Surdez: Resources, methodology. O. Delattre: Resources, methodology. P.K. Bode: Resources. M. Wachtel: Conceptualization, supervision, writing–original draft, writing–review and editing. B.W. Schäfer: Conceptualization, resources, supervision, funding acquisition, investigation, writing–original draft, project administration, writing–review and editing.

This work was supported by Cancer League Switzerland KLS-5143–08–2020 (to B.W. Schafer), Childhood Research Foundation Switzerland (to B.W. Schafer), Children's Research Center grant from the University Children's Hospital Zurich 10788_E (to S.G. Danielli), and Sarcoma Foundation of America grant 964047 (to B.W. Schafer). Some illustrations were created with BioRender.com.

Note: Supplementary data for this article are available at Molecular Cancer Therapeutics Online (http://mct.aacrjournals.org/).

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