In squamous cell carcinoma (SCC), tissue invasion by collectively invading cells requires physical forces applied by tumor cells on their surrounding extracellular matrix (ECM). Cancer-related ECM is composed of thick collagen bundles organized by carcinoma-associated fibroblasts (CAF) within the tumor stroma. Here, we show that SCC cell collective invasion is driven by the matrix-dependent mechano-sensitization of EGF signaling in cancer cells. Calcium (Ca2+) was a potent intracellular second messenger that drove actomyosin contractility. Tumor-derived matrix stiffness and EGFR signaling triggered increased intracellular Ca2+ through CaV1.1 expression in SCC cells. Blocking L-type calcium channel expression or activity using Ca2+ channel blockers verapamil and diltiazem reduced SCC cell collective invasion both in vitro and in vivo. These results identify verapamil and diltiazem, two drugs long used in medical care, as novel therapeutic strategies to block the tumor-promoting activity of the tumor niche.

Significance: This work demonstrates that calcium channels blockers verapamil and diltiazem inhibit mechano-sensitization of EGF-dependent cancer cell collective invasion, introducing potential clinical strategies against stromal-dependent collective invasion.

Graphical Abstract:http://cancerres.aacrjournals.org/content/canres/78/18/5229/F1.large.jpg. Cancer Res; 78(18); 5229–42. ©2018 AACR.

Collective cancer cell invasion of the surrounding tumor microenvironment is recurrent in solid cancers, including carcinomas (1) and it occurs independently of epithelial-to-mesenchymal transition (EMT). In fact, collectively invading carcinoma cells retain epithelial characters (2). Recent evidences demonstrate a partial EMT reprograming of a subset of cancer cells at the leading edge of the tumor bulk in head and neck squamous carcinomas (HNSCC; ref. 3). Indeed, collective invasion is a tissue-driven process dictated by extracellular matrix (ECM) remodeling (4, 5), which is orchestrated by carcinoma-associated fibroblasts (CAF). CAFs lead the way to cancer cells by digging tracks within the ECM that SCC cells use to invade (6). Interestingly, CAFs are crucial for track generation, but not necessary for collective invasion once the tracks are created (6).

Track generation by CAFs requires JAK-dependent acto-myosin–driven force generation on collagen fibers (7), which results in thick bundle formation (6), fiber alignment (8), and increased matrix stiffening (9). Accordingly, ECM stiffening is a characteristic of neoplastic lesions (10). ECM rigidity results from increased secretion and assembly of matrix proteins, and collagen cross-linking essentially mediated by lysyl oxidases (LOX) expression by both CAF and tumor cells (11). Consequently, ECM stiffening has been associated with increased metastases and poor clinical outcome in several epithelial cancers due to activation of mechanotransduction-dependent signaling pathways (12). CAF-driven collective invasion also relies on cytokines and growth factor signaling activation in cancer cells (13). From these observations, increasing evidences suggest a connection of mechanotransduction with receptor tyrosine kinase (RTK) signaling pathways. Upon dimerization, RTKs become activated and are involved in integrin-mediated mechanotransduction signaling, which promotes tumor progression (14). Indeed, cell-to-ECM adhesion favors EGFR (also known as ErbB-1/HER1)-dependent tumor growth, and it sensitizes EGFR intracellular signaling in cancer cells (15, 16). EGFR has been reported to be overexpressed, amplified, or mutated in SCC lesions including HNSCC and cutaneous SCC (17). In most cases, EGFR expression is positively correlated with poor patient survival, independent of therapeutic treatments (18, 19).

Ca2+ is one of the most important chemical elements for living organisms and Ca2+ signaling has been linked to a wide variety of physiologic processes. Intracellular calcium entry is tightly regulated in cells by a plethora of Ca2+ channels at the plasma membrane. The L-type calcium channel family is part of the high-voltage activated family of voltage-gated Ca2+ channel (20). They are known to be expressed in excitable cells such as in heart, muscle, and brain organs; however, they have recently been described to be overexpressed in epithelial cancer cells (21). L-type Ca2+ channels, also known as CaV1, are composed of 4 subunits (α1, α2δ, β, and γ), of which α1 is the pore-forming unit and the others are regulatory subunits. The CaV1 α1 subunits (CaV1.1, CaV1.2, CaV1.3, and CaV1.4) are encoded by 4 different genes (CACNA1C, CACNA1D, CACNA1S, and CACNA1F, respectively).

In this study, we used an in vitro model of 3D organotypic coculture and in vivo model to demonstrate the interconnection between EGFR activity and ECM stiffening during collective invasion. We established L-type Ca2+ signaling as a mechanotransducer intracellular second messenger that drives the EGFR and matrix stiffening interplay during cancer cell collective invasion.

Cell culture

Primary human fibroblasts were isolated from foreskin of child and were maintained in DMEM supplemented with 10% FCS and 2 mmol/L glutamine. Human CAFs (gift from E. Sahai, The Francis Crick Institute, London, United Kingdom) isolated from patients with head and neck carcinoma were cultured in DMEM (21969, Gibco) supplemented with 10% FCS, 2 mmol/L glutamine, and insulin-transferrin-selenium (41400-045; Invitrogen). SCC12 cells (gift from E. Sahai, The Francis Crick Institute) were cultured in FAD media (21765, Gibco), supplemented with 10% FCS, 2 mmol/L glutamine, insulin-transferrin-selenium (41400-045, Gibco) and 0.5 μg/mL, hydrocortisone (H-0135; Sigma), hereafter referred to complete media. All cells were grown at 37°C in a humidified 5% CO2 atmosphere. Experiments were performed at passages 3 to 10. Human cell line has been profiled using short tandem repeat and tested for Mycoplasma by PCR (3′ YGC CTG VGT AGT AYR YWC GC and 5′ GCG GTG TGT ACA ARM CCC GA) every 4 months. Hydrogels were purchased from Matrigen and coated with collagen (40 μg/mL, 354249, Corning). Soft condition referred to cells plated on surface of 100 μL of matrix gel for 6-well plates. Matrix gel is composed of 4 mg/mL collagen (354249, Corning), 2 mg/mL Matrigel (354234, Corning), 5x DMEM, HEPES (15630–056, Gibco), and 0.5% FCS DMEM. Stiff conditions correspond to cells cultivated on collagen (40 μg/mL)/Matrigel (20 μg/mL)-coated plates.

Matrix remodeling assay

CAFs (25 × 103) were embedded in 100 μL of matrix gel (22) and seeded in triplicate into 96-well plate. After 1 hour at 37°C, vehicle or inhibitors were added in 100 μL of 0.5% FCS media and changed every 2 days. At day 6, the relative diameter of the well and the gel were measured using ImageJ software (http://rsbweb.nih.gov/ij/). The percentage of gel contraction was calculated using the following formula: 100 × (well diameter – gel diameter)/well diameter.

Traction force microscopy

Contractile forces exerted by clusters of 5 to 10 SCC12 cells on different stiffness gels were assessed by traction force microscopy essentially as described previously (23). Briefly, polyacrylamide substrates with shear moduli of 1 and 50 kPa (PS35-EC-1/PS35-EC-50; Matrigen) were coated with collagen (50 μg/mL) containing fluorescent latex microspheres (18859; Polysciences). SCC12 were plated on fluorescent bead–conjugated discrete stiffness gels and grown for 24 hours. At this time, they were treated with the indicated molecules for 1 hour before traction force measurements. Images of gel surface–conjugated fluorescent beads were acquired for each cluster before and after cell removal using an Axiovert 200M motorized microscope stand (Zeiss) and a ×32 magnification objective. Tractions exerted by clusters of 5 to 10 SCC12 cells were estimated by measuring bead displacement fields, computing corresponding traction fields using Fourier transform traction microscopy, and calculating root-mean-square traction using the PIV (particle image velocity) and TFM (traction force microscopy) package on ImageJ. To measure baseline noise, the same procedure was performed on a cell-free region.

Organotypic invasion assay

CAFs (5 × 105) were embedded in 1 mL of matrix gel (6) composed of 4 mg/mL collagen and 2 mg/mL Matrigel for stiff condition, and 1 mg/mL collagen and 0.5 mg/mL Matrigel for soft condition. After 1 hour at 37°C, 5 × 105 SCC12 were plated on the top of the gel and incubated overnight. The gel was mounted on a metal bridge and fed from underneath with changes of complete media every 2 days. After 6 days, cultures were fixed (4% paraformaldehyde + 0.25% glutaraldehyde in PBS) embedded in paraffin blocks, sectioned, and stained with hematoxylin and eosin. The invasion index was calculated from ImageJ measurement of the total area of SCC cells and the area of noninvading SCC cells, index = average of 1−(noninvading area)/total area of at least four fields separated minimum by 100 μm.

Patient-derived spheroids

Following excision, biopsy samples were directly transferred to freshly prepared 10% FCS, 2 mmol/L glutamine culture medium containing a mixture of antibiotic/antifungal compounds (0.26 μmol/L amphotericin B, ampicillin 0.14 mmol/L, ciprofloxacin 7.54 μmol/L). Fresh tumor tissue samples were mechanically and enzymatically (collagenase 200 μg/mL in PBS; Roche) digested to generate a single-cell suspension. After determination of cell viability using the trypan blue exclusion test, the single-cell suspension was directly processed into spheroids. Briefly, 50 μL droplets containing 25 to 50 × 103 cells were plated onto the underside of a 10-cm culture dish and allowed to form spheroids during 48 hours in a 37°C incubator 48 hours. The spheroids were then embedded in a collagen I/Matrigel gel mix at a concentration of approximately 4 mg/mL collagen and 2 mg/mL Matrigel for stiff condition and 1 mg/mL collagen and 0.5 mg/mL Matrigel for soft condition in 35-mm glass-bottomed cell culture plates (MatTek). The gel was incubated for at least 45 minutes at 37°C with 5% CO2. Then, the gel was covered with complete media. Forty-eight hours later, the spheroids were imaged with an inverted microscope at a magnification of ×4 and ×10. Invasion was quantified using ImageJ.

Generation of patient-derived xenografts

Tumor specimens were obtained at initial surgery (Face and Neck University Institute, Nice, France) from primary diagnosed HNSCC. None of the patients received neoadjuvant chemotherapy and/or radiotherapy. Written informed consent was obtained from each patient and the study was approved by the hospital ethics committee. Patient tumor material was collected in culture medium and partially digested during 1 hour at room temperature in RPMI1640 with 1 mg/mL collagenase IV, 1 mg/mL dispase, and 1 mg/mL hyaluronidase. Approximately 20 to 30 mg tissue fragments in 50% Matrigel were implanted subcutaneously into the flank region of NMRI-nu (RjOrl:NMRI-Foxn1nu/Foxn1nu) mice. The first passage patient-derived xenografts (PDX) were dissociated in a collagenase/dispase mixture and cells were cultured in low serum conditions (2%FBS/F12/DMEM/1× B27) in the presence of 5 ng/mL EGF. Subsequently, 75 × 104 cells in 50% Matrigel were implanted subcutaneously into the flank region of NMRI-nu (RjOrl:NMRI-Foxn1nu/Foxn1nu) mice. One week after tumor engraftment, to avoid any interference with tumor uptake, mice were treated with the corresponding inhibitors. Gefitinib (50 mg/kg/day) was injected with PBS intraperitoneally every 2 days. BAPN (100 mg/kg/day) was dissolved in drinking water. For verapamil and diltiazem treatment (drinking water), low-dose treatment (20 mg/kg/day) started at 4 days postinjection and the final dose (50 mg/kg/day) started at 7 days postinjections. Drug containing drinking water was prepared fresh and changed 3 times a week. The dose in drinking water was determined using average daily water intake (5 mL) and mouse weight. Tumor volume was measured every day from the beginning of the treatment with the following formula: 4/3 × π × (length/2) × [(width/2) × 2].

Study approval

All animal experiments were approved by the local committee of the host institute and by the Institutional Animal Care and Use Committee (CIEPAL AZUR committee, MESR number 2016090714331137) at the University Cote d'Azur, Nice, France. All experimental procedures involving the use of human tissue included the relevant receipt of written informed consent and were approved by Institutional Review Boards. Ethical approval for this study and informed consent conformed to the standards of the Declaration of Helsinki.

Statistical analysis

Cell culture experiments were performed at least three times independently. The number of animals in each group was calculated to measure at least a 20% difference between the means of experimental and control groups with a power of 80% and SD of 10%. The number of patient samples was determined primarily by clinical availability. Histologic analyses of both mouse and human tissue were performed in a blinded fashion. Numerical quantifications for in vitro experiments using cultured cells represent mean ± SD. Numerical quantifications for physiologic experiments using mouse or human sample represent mean ± SEM. Immunoblot images are representative of experiments that have been repeated at least three times. Quantification of immunoblot corresponds to the mean ± SD of the different replicate. Micrographs are representative of experiments in each relevant cohort. Paired samples were compared by a two-tailed Student t test for normally distributed data, whereas Mann–Whitney U nonparametric testing was used for nonnormally distributed data. For comparisons among groups, one-way ANOVA and post hoc Tukey testing was performed. A P value less than 0.05 was considered statistically significant. Correlation analyses were performed by Pearson correlation coefficient calculation. The Mantel–Cox log-rank test was used for statistical comparisons in survival analyses. All statistical analyses have been performed with GraphPad Prism software.

Matrix stiffness sensitizes SCC cells to EGF-dependent collective invasion

Organotypic cell invasion assays were performed using CAFs isolated from human HNSCC biopsies and an SCC cell line (SCC12) that invades collectively (24). We tuned matrix stiffness in vitro, either by modulating collagen concentration (Fig. 1A–D; ref. 10) or by adding ribose to the collagen gels (Fig. 1E and F; ref. 25). On soft collagen gel, collective invasion of SCC12 cells was induced at 10 ng/mL of EGF (Fig. 1A and B), whereas on stiff matrix, 1 ng/mL of EGF was sufficient to promote SCC12 cell collective invasion, but only in the presence of CAFs (Fig. 1C and D). Moreover, although increasing matrix stiffness correlated with increase of the collective invasion index at fixed EGF concentration, the basal level of collective invasion induced by stiff matrices was inhibited by the addition of gefitinib, an FDA-approved EGFR receptor tyrosine kinase inhibitor (RTKi; Fig. 1E and F). Importantly, we demonstrate that EGF does not support the ability of CAFs to physically remodel the collagen gels (Supplementary Fig. S1A). Such remodeling allows creation of proinvasive tracks in the ECM (6) and contributes to increase matrix stiffening (9). Also, the use of CAF-remodeled matrices (24) and addition to SCC12 cells, after CAF removal, demonstrated that SCC12 cells are able to collectively invade within the pre-remodeled matrix only in the presence of EGF (Supplementary Fig. S1B and S1C). Altogether, our results reveal the interplay between CAF-dependent physical remodeling of collagen fibers and EGF-dependent SCC12 cell stimulation during collective invasion.

Figure 1.

Matrix stiffness and EGFR cooperate to promote collective invasion. A, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to EGF in presence or absence of CAF in soft matrices. Median rigidity of soft gel measured by AFM is 0.2345 kPa. Scale bar, 100 μm. B, Quantification of SCC12 cells invasion index shown in A (n = 3; mean + SD). C, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to EGF in the presence or absence of CAF in stiff matrices. Median rigidity of stiff gel measured by AFM is 14.70 kPa. Scale bar, 100 μm. D, Quantification of SCC12 cells invasion index shown in B (n = 3; mean + SD). E, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to different ribose concentration, in the presence or not of EGF and gefitinib. Median rigidity of gels measured by AFM is 0.2345, 1.06, and 1.26 kPa for gels containing 0, 100, and 250 mmol/L ribose, respectively. Scale bar, 100 μm. F, Quantification of SCC12 cells invasion index shown in E (n = 3; mean + SD). G, Heatmap of SCC12 cells traction forces applied on 1 or 50 kPa hydrogels in the presence or absence of EGF (5 ng/mL). H, Quantification of SCC12 cells' traction forces shown in G. Bars correspond to the medians (representative of three independent experiments). I, Immunoblot of p-EGFR and total EGFR in SCC12 cells plated on 1, 12, or 50 kPa hydrogels stimulated EGF for 1 hour. Immunoblot of tubulin shown as control. J, Immunoblot of total EGFR in SCC12 cells plated on 1, 12, or 50 kPa hydrogels for 48 hours. Immunoblot of tubulin shown as control. K, Quantification of EGFR expression in response to substratum rigidity shown in J (fold of induction relative to 1 kPa). L, Histogram of qRT-PCR quantification of EGFR mRNA in SCC12 cells plated on soft and stiff matrices after 48 hours (n = 3; mean +SD). M, Immunoblot of total EGFR, YAP, and TAZ in SCC12 cells plated on stiff matrices following RNAi targeting YAP and TAZ transfection. Immunoblot of tubulin shown as control. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001). A.U., arbitrary unit.

Figure 1.

Matrix stiffness and EGFR cooperate to promote collective invasion. A, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to EGF in presence or absence of CAF in soft matrices. Median rigidity of soft gel measured by AFM is 0.2345 kPa. Scale bar, 100 μm. B, Quantification of SCC12 cells invasion index shown in A (n = 3; mean + SD). C, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to EGF in the presence or absence of CAF in stiff matrices. Median rigidity of stiff gel measured by AFM is 14.70 kPa. Scale bar, 100 μm. D, Quantification of SCC12 cells invasion index shown in B (n = 3; mean + SD). E, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 organotypic culture in response to different ribose concentration, in the presence or not of EGF and gefitinib. Median rigidity of gels measured by AFM is 0.2345, 1.06, and 1.26 kPa for gels containing 0, 100, and 250 mmol/L ribose, respectively. Scale bar, 100 μm. F, Quantification of SCC12 cells invasion index shown in E (n = 3; mean + SD). G, Heatmap of SCC12 cells traction forces applied on 1 or 50 kPa hydrogels in the presence or absence of EGF (5 ng/mL). H, Quantification of SCC12 cells' traction forces shown in G. Bars correspond to the medians (representative of three independent experiments). I, Immunoblot of p-EGFR and total EGFR in SCC12 cells plated on 1, 12, or 50 kPa hydrogels stimulated EGF for 1 hour. Immunoblot of tubulin shown as control. J, Immunoblot of total EGFR in SCC12 cells plated on 1, 12, or 50 kPa hydrogels for 48 hours. Immunoblot of tubulin shown as control. K, Quantification of EGFR expression in response to substratum rigidity shown in J (fold of induction relative to 1 kPa). L, Histogram of qRT-PCR quantification of EGFR mRNA in SCC12 cells plated on soft and stiff matrices after 48 hours (n = 3; mean +SD). M, Immunoblot of total EGFR, YAP, and TAZ in SCC12 cells plated on stiff matrices following RNAi targeting YAP and TAZ transfection. Immunoblot of tubulin shown as control. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001). A.U., arbitrary unit.

Close modal

The Cdc42 small GTPase activity has been mechanistically implicated in the onset of the locomotory forces provided by tumor cells against the tumor microenvironment (26) and in collective SCC cell invasion (6) promoting actomyosin cytoskeletal contractility. Thus, we investigated whether in SCC12 cells Cdc42 activity might be regulated by the matrix stiffness and EGFR interplay. Accordingly, EGF stimulation prompted Cdc42 activity and myosin light chain 2 (MLC2) phosphorylation exclusively when cells were plated on stiff matrix (Supplementary Fig. S1D and S1E). Abrogation of Cdc42 expression in SCC12 cells triggered inhibition of collective cells' invasion in the presence of both EGF and CAFs (Supplementary Fig. S1F–S1H). Locomotory forces applied by SCC12 cells were measured by traction forces microscopy (27), which revealed that EGF triggers increased forces application by SCC12 on the surrounding stiff matrix only (Fig. 1G and H). We thus examined whether matrix stiffness could prime SCC12 cells to EGFR phosphorylation. SCC12 cells were either plated on collagen I–coated hydrogel with controlled stiffness (Fig. 1I) or embedded within a soft and stiff matrix (Supplementary Fig. S1I). In both conditions, a strong enhancement of EGFR phosphorylation depending on the substrate stiffness was detected in the presence of EGF (Fig. 1I; Supplementary Fig. S1I). Interestingly, increased EGFR expression at protein and mRNA levels was noted on stiff substrates in the absence of EGF (Fig. 1J–L). Moreover, inhibition of the mechano-responsive transcription factors YAP and TAZ abrogated the stiffness-dependent EGFR regulation (Fig. 1M). Together, these results support the notion that collective cancer cell invasion is triggered by mechanotransduction-dependent EGFR signaling, which leads to locomotory forces' traction onto the surrounding substrate.

Interplay between matrix stiffness and EGFR activity promotes collective invasion in vivo

We next investigated whether ECM stiffness and EGFR signaling also cooperate to promote SCC collective invasion in vivo. We subcutaneously engrafted HNSCC tumor cells isolated from patients into the flanks of nude mice. One week later, to avoid any interference with tumor uptake, mice were treated with either gefitinib or BAPN, a LOX inhibitor, or both in combination. Such in vivo analyses revealed that both gefitinib and BAPN treatments alone or in combination block tumor cell collective invasion (Fig. 2A and B). AFM quantification of tumor rigidity confirmed the inhibitory effect of BAPN on ECM stiffening in vivo (Fig. 2C). IHC analyses by Picrosirius Red coloration and polarized light visualization of the tumors confirmed ECM remodeling and collagen bundle formation in the tumor stroma that both were inhibited in BAPN-treated mice (Fig. 2D–F). In these tumors, EGFR expression was significantly reduced, which indicated that tumor cells increased EGFR expression in response to ECM stiffening (Fig. 2D–F). A strong reduction of the tumor volume was also noticed in the treated mice compared with control (Supplementary Fig. S2A).

Figure 2.

Matrix stiffness and EGFR cooperate to promote collective invasion in vivo. A, Representative hematoxylin and eosin pictures of human patient–derived HNSCC tumors showing collective invasion in control, BAPN, gefitinib or BAPN, and gefitinib-treated group mice. Scale bar, 200 μm. B, Quantification of tumors presenting invading cell cohorts shown in A (n = 6 tumors/group). C, Atomic force microscopy quantification of tissue rigidity (in Pascal) from tumor shown in A. Median quantification was performed on 3 to 5 anatomic regions from frozen sections of three independent tumors for each group. Outliners data points above 2,500 Pa were removed for quantification. D, Representative picture of EGFR immunostaining and Sirius Red–stained collagen bundles observed under polarized light of PDX. Scale bar, 200 μm. E, Quantification of EGFR expression shown in D [Quick Score (QS) method, n = 2 for 3 individual PDXs; mean + SEM]. F, Quantification of fibrilar collagen shown in D. (n = 2 for 3 individual PDXs; mean + SEM). G, Representative pictures of human patient–derived HNSCC multicellular spheroids embedded in soft or stiff matrices in the presence or absence of gefitinib (5 μmol/L). Scale bar, 200 μm. H, Quantification of HNSCC multicellular spheroid invasion shown in G (n = 5 for 5 individual patients; bars correspond to mean). I, Representative picture of EGFR expression (measured by the Quick Score method) and collagen fibrilar deposition in a cohort of 48 patients with HNSCC (scale bar, 200 μm), classify between low (QS ≤ 8), medium (8 < QS ≤ 12), and high (12 < QS) EGFR expression. J, Relative number of patient biopsies (shown in I) classified between EGFR QS and Sirius Red intensity. White numbers represent the real number of biopsies per conditions. A Pearson correlation was performed. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; ***, P < 0.001). A.U., arbitrary unit.

Figure 2.

Matrix stiffness and EGFR cooperate to promote collective invasion in vivo. A, Representative hematoxylin and eosin pictures of human patient–derived HNSCC tumors showing collective invasion in control, BAPN, gefitinib or BAPN, and gefitinib-treated group mice. Scale bar, 200 μm. B, Quantification of tumors presenting invading cell cohorts shown in A (n = 6 tumors/group). C, Atomic force microscopy quantification of tissue rigidity (in Pascal) from tumor shown in A. Median quantification was performed on 3 to 5 anatomic regions from frozen sections of three independent tumors for each group. Outliners data points above 2,500 Pa were removed for quantification. D, Representative picture of EGFR immunostaining and Sirius Red–stained collagen bundles observed under polarized light of PDX. Scale bar, 200 μm. E, Quantification of EGFR expression shown in D [Quick Score (QS) method, n = 2 for 3 individual PDXs; mean + SEM]. F, Quantification of fibrilar collagen shown in D. (n = 2 for 3 individual PDXs; mean + SEM). G, Representative pictures of human patient–derived HNSCC multicellular spheroids embedded in soft or stiff matrices in the presence or absence of gefitinib (5 μmol/L). Scale bar, 200 μm. H, Quantification of HNSCC multicellular spheroid invasion shown in G (n = 5 for 5 individual patients; bars correspond to mean). I, Representative picture of EGFR expression (measured by the Quick Score method) and collagen fibrilar deposition in a cohort of 48 patients with HNSCC (scale bar, 200 μm), classify between low (QS ≤ 8), medium (8 < QS ≤ 12), and high (12 < QS) EGFR expression. J, Relative number of patient biopsies (shown in I) classified between EGFR QS and Sirius Red intensity. White numbers represent the real number of biopsies per conditions. A Pearson correlation was performed. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; ***, P < 0.001). A.U., arbitrary unit.

Close modal

The crucial role of matrix stiffening and EGF cooperation in tumor invasion was further assessed using a patient-derived multicellular spheroid model of HNSCC cells invasion in vitro. Spheroids composed of cells isolated from fresh human HNSCC tumor biopsies were embedded in either soft or stiff collagen-rich matrices. Assessment of tumor cell invasion demonstrated that stiff matrix promotes cancer cell invasion dependent on EGFR activity in all the tested patient biopsies (Fig. 2G and H). These experimental data were finally corroborated by IHC analyses of 48 human HNSCC biopsies. EGFR was overexpressed in tumor cells specifically at the tumor–stroma interface, and a positive correlation between cross-linked collagen bundles and EGFR expression was established by Picrosirus Red coloration visualized under polarized light (Fig. 2I and J). Together, these results support the notion that tumor stiff extracellular matrix mechano-sensitizes SCC cells to the EGFR-dependent collective invasion both in vitro and in vivo.

L-type calcium channel CaV1.1 triggers matrix stiffness and EGFR interplay for collective invasion

Identification of the molecular mechanisms driving tumor cell collective invasion downstream of ECM stiffness and EGFR signalization would provide valuable insight in tumor cell biology. In addition, it might lead to the discovery of novel molecular targets for therapeutic approaches in patients with SCC. Invasion assays on stiff matrices in the presence of CAFs and EGF were thus performed to screen for FDA-approved pharmacologic compounds able to inhibit SCC12 cell collective invasion. The cell toxicity of 380 chemical compounds was evaluated on SCC12 cells and CAFs after 4-day exposure to 10 μmol/L final concentration of each single drug. Sixty-seven molecules that inhibited at least 40% of the cell mitochondrial activity were discarded (Supplementary Fig. S3A; Supplementary Table S1). The remaining 313 compounds targeting kinases, phosphatases, ion channels, nuclear receptors, epigenetic, and the Wnt signaling were further screened to assess their capacity to block SCC12 cell collective invasion in vitro. A total of 48 small molecules, which inhibited SCC12 cell collective invasion by at least 50% compared with control DMSO, were identified (Fig. 3A; Supplementary Table S2). To distinguish between molecules that specifically block SCC cell invasion but not the CAF proinvasive activity, ECM remodeling assays based on matrix contraction were performed using CAF cells alone (6). Thirty-four of the 48 selected small molecules displayed no significant blocking activity of CAF proinvasive ECM remodeling (Supplementary Fig. S3B; Supplementary Table S3). A Metascape analysis of all the known molecular target genes for the remaining 34 compounds revealed that 14 of them are calcium (Ca2+) channel inhibitors (Supplementary Table S4). Accordingly, Ca2+ channel signaling was the best candidate pathway to be targeted to inhibit SCC12 cell collective invasion in vitro (Fig. 3B). Because 9 of them are inhibitors of L-type Ca2+ channels, the role of this family of Ca2+ channel in SCC invasion was further investigated.

Figure 3.

The L-type calcium channel CaV1.1 triggers EGF-dependent SCC cell collective invasion. A, Quantification of organotypic invasion assay screening, performed with SCC12 cells in the presence of HNCAF, EGF (10 ng/mL), and stiff matrix, with all nontoxic selected molecules (10 μmol/L final concentration) classified by libraries (n = 1; mean + SD of 4 pictures). Dotted bar represents the threshold used to select molecules that efficiently blocked SCC12 invasion (50% of inhibition). B, Metascape analysis (using Kegg pathways enrichment) of the targeted genes triggered by the 34 inhibitors identified for blocking SCC12 invasion but not CAF matrix remodeling. C, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 invasion assays in the presence or not of diltiazem (30 μmol/L) and verapamil (15 μmol/L). Scale bar, 100 μm (n = 3; mean + SD). D, Quantification of SCC12 cells' organotypic invasion index shown in C. E, Hematoxylin and eosin coloration of paraffin-embedded sections of organotypic culture of SCC12 cells transduced with two different siRNA against CaV1.1 or siRNA against luciferase for control. Scale bar, 100 μm. F, Quantification of the invasion index of SCC12 cells shown in E (n = 3; mean + SD). G, Immunoblot of CaV1.1 in SCC12 cells corresponding to the assay presented in G. Tubulin was used as control. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; ***, P < 0.001). A.U., arbitrary unit.

Figure 3.

The L-type calcium channel CaV1.1 triggers EGF-dependent SCC cell collective invasion. A, Quantification of organotypic invasion assay screening, performed with SCC12 cells in the presence of HNCAF, EGF (10 ng/mL), and stiff matrix, with all nontoxic selected molecules (10 μmol/L final concentration) classified by libraries (n = 1; mean + SD of 4 pictures). Dotted bar represents the threshold used to select molecules that efficiently blocked SCC12 invasion (50% of inhibition). B, Metascape analysis (using Kegg pathways enrichment) of the targeted genes triggered by the 34 inhibitors identified for blocking SCC12 invasion but not CAF matrix remodeling. C, Hematoxylin and eosin coloration of paraffin-embedded sections of SCC12 invasion assays in the presence or not of diltiazem (30 μmol/L) and verapamil (15 μmol/L). Scale bar, 100 μm (n = 3; mean + SD). D, Quantification of SCC12 cells' organotypic invasion index shown in C. E, Hematoxylin and eosin coloration of paraffin-embedded sections of organotypic culture of SCC12 cells transduced with two different siRNA against CaV1.1 or siRNA against luciferase for control. Scale bar, 100 μm. F, Quantification of the invasion index of SCC12 cells shown in E (n = 3; mean + SD). G, Immunoblot of CaV1.1 in SCC12 cells corresponding to the assay presented in G. Tubulin was used as control. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; ***, P < 0.001). A.U., arbitrary unit.

Close modal

To validate the possible role of L-type Ca2+ channels in SCC12 cell collective invasion, two CaV1 calcium channel blockers (CCB) were selected: the phenylalkylamine verapamil and the nondihydropiridine diltiazem. Both drugs are administered to patients as cardiac antiarrhythmic or antihypertensive therapeutic agents. At experimental level, these two compounds efficiently inhibit SCC12 cell collective invasion in a dose-dependent manner (Fig. 3C and D; Supplementary Fig. S3C), without affecting CAF-dependent proinvasive ECM remodeling (Supplementary Fig. S3D). To identify the specific isoform of the L-type Ca2+ channels mediating SCC12 cell collective invasion, expression of each α1 subunit was depleted in SCC12 cells by a SMARTpool RNAi approach. CaV1.1 was specifically identified (Supplementary Fig. S3E and S3F) and validated using two independent RNAi sequences (Fig. 3E–G). Collectively, these results reveal the L-type Ca2+ channel CaV1.1 as a critical regulatory element of SCC12 cell collective invasion downstream of ECM stiffness and EGFR signalization in vitro.

Interplay between matrix stiffness and EGFR signaling regulates CaV1.1 expression and induces calcium entry in epithelial cancer cells

So far, the molecular mechanisms leading to L-type Ca2+ channel expression in epithelial cancer cells are unknown. We hypothesized that CaV1.1 expression is regulated by the interplay between matrix stiffness and EGFR signaling. Accordingly, expression of CaV1.1 protein was induced only when SCC12 cells were plated on stiff matrix in the presence of EGF. In contrast, CaV1.1 expression was undetectable even at high EGF concentration when cells were plated on soft matrix (Fig. 4A–D). Moreover, upregulation of CaV1.1 occurred at the mRNA level in tumor cells plated on stiff matrix in the presence of EGF (Fig. 4E). These observations suggest a complex mechanism of regulation governing CaV1.1 expression in epithelial cancer cells. We next explored whether CaV1.1 calcium channel in SCC12 is functional. The Ca2+ flux in SCC12 was thus quantified by conducting Fura 2 (fluorescent Ca2+ probe) experiments in response to matrix stiffness. Ca2+ entry was significantly increased in SCC12 cells grown on stiff matrix compared with those seeded on soft counterparts upon EGF stimulation. Moreover, the steady state of intracellular Ca2+ concentration resulted independent from ECM stiffness, but it reached maximal responses when EGF was applied to the cells grown on stiff matrix (Fig. 4F and G; Supplementary Fig. S4A and S4B). In addition, blocking of CaV1.1 activity using both verapamil and diltiazem reduced Ca2+ entry to rates comparable with those observed in cells plated on soft matrix (Fig. 4F and G). Specifically, the role of CaV1.1 was confirmed by RNAi-mediated knockdown. Indeed, loss of CaV1.1 expression recovered a Ca2+ entry in the cytoplasm at levels similar to those induced by EGF in cells grown on soft matrix (Supplementary Fig. S4C and S4D). These results demonstrated the presence of a functional CaV1.1 L-type calcium channel, whose expression is upregulated by EGFR signaling in SCC12 plated on stiff matrix, resulting in increased Ca2+ entry in the cytosol.

Figure 4.

EGFR signaling and matrix stiffness regulate CaV1.1 expression and activity. A, Representative immunoblot of CaV1.1 in SCC12 cells plated 3 days on soft or stiff matrices in the presence or absence of EGF. Tubulin was used as control. B, Quantification of CaV1.1 relative to tubulin in three immunoblots performed in the same conditions as in A (n = 3; mean + SD). C, Representative confocal merged images of CaV1.1 and DAPI staining in SCC12 plated for 3 days on soft or stiff matrices in the presence or absence of EGF (5 ng/mL). Scale bar, 40 μm. D, Quantification of CaV1.1 staining from experiment shown in C. Bars represent the mean and each dot the relative fluorescence of CaV1.1 in a SCC12 cell (n = 30 cells at least). A.U., arbitrary unit. E, mRNA fold of induction measured by qPCR of CaV1.1 (CACNA1S) in SCC12 cells cultivated on soft matrix for 3 days and then plated on stiff matrix for 3 hours (n = 3; mean + SD). F, Ca2+ entry in SCC12 cells plated for 3 days on soft or stiff matrices, stimulated by 5 ng/mL of EGF during the experiment, in the presence or absence of verapamil (15 μmol/L) or diltiazem (30 μmol/L) for 30 minutes before the experiment. Representative data of three independent experiments. G, Representation of Ca2+ level measured at the end of the experiment shown in C. Bars correspond to the mean and each dot represents Ca2+ measurement in a cell (n = 30 cells at least). H, Pictures of CaV1.1 and EGFR immunostaining [measured by the Quick Score (QS) method] and collagen fibrilar deposition in a cohort of 48 patients with HNSCC (scale bar, 200 μm) classify between low (QS ≤ 8), medium (8 < QS ≤ 12), and high (12 < QS) CaV1.1 expression. I, Relative number of patient biopsies (shown in H) classified between CaV1.1 and EGFR QS. J, Relative number of patient biopsies (shown in H) classified between CaV1.1 and Sirius Red intensity. White numbers represent the real number of biopsies per conditions. A Pearson correlation was performed. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

Figure 4.

EGFR signaling and matrix stiffness regulate CaV1.1 expression and activity. A, Representative immunoblot of CaV1.1 in SCC12 cells plated 3 days on soft or stiff matrices in the presence or absence of EGF. Tubulin was used as control. B, Quantification of CaV1.1 relative to tubulin in three immunoblots performed in the same conditions as in A (n = 3; mean + SD). C, Representative confocal merged images of CaV1.1 and DAPI staining in SCC12 plated for 3 days on soft or stiff matrices in the presence or absence of EGF (5 ng/mL). Scale bar, 40 μm. D, Quantification of CaV1.1 staining from experiment shown in C. Bars represent the mean and each dot the relative fluorescence of CaV1.1 in a SCC12 cell (n = 30 cells at least). A.U., arbitrary unit. E, mRNA fold of induction measured by qPCR of CaV1.1 (CACNA1S) in SCC12 cells cultivated on soft matrix for 3 days and then plated on stiff matrix for 3 hours (n = 3; mean + SD). F, Ca2+ entry in SCC12 cells plated for 3 days on soft or stiff matrices, stimulated by 5 ng/mL of EGF during the experiment, in the presence or absence of verapamil (15 μmol/L) or diltiazem (30 μmol/L) for 30 minutes before the experiment. Representative data of three independent experiments. G, Representation of Ca2+ level measured at the end of the experiment shown in C. Bars correspond to the mean and each dot represents Ca2+ measurement in a cell (n = 30 cells at least). H, Pictures of CaV1.1 and EGFR immunostaining [measured by the Quick Score (QS) method] and collagen fibrilar deposition in a cohort of 48 patients with HNSCC (scale bar, 200 μm) classify between low (QS ≤ 8), medium (8 < QS ≤ 12), and high (12 < QS) CaV1.1 expression. I, Relative number of patient biopsies (shown in H) classified between CaV1.1 and EGFR QS. J, Relative number of patient biopsies (shown in H) classified between CaV1.1 and Sirius Red intensity. White numbers represent the real number of biopsies per conditions. A Pearson correlation was performed. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

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CaV1.1 expression was then evaluated in tumor biopsies of the engrafted HNSCC tumor mice by immunostaining. We disclosed the presence of CaV1.1 in the tissue area positive for EGFR staining. Interestingly, in mice treated with BAPN and gefitinib, either alone or in combination, CaV1.1 staining in the tumor tissue was significantly reduced (Supplementary Fig. S4E and S4F). Finally, CaV1.1 expression in sections from the 48 human HNSCC tumors was found to correlate with EGFR expression and collagen cross-linking (Fig. 4H–J). These results show that in human tumor tissue sections, expression of the CaV1.1 L-type Ca2+ channel is essentially located in the cells at the interface with the tumor ECM.

Cav1-dependent calcium signaling mediates actomyosin contractility in response to ECM stiffening

We found that inhibition of CaV1.1 expression suppresses SCC12 cell collective invasion promoted by the cooperation between EGFR and ECM stiffness (Fig. 3). We thus investigated the role of CaV1.1 in the actomyosin contractility. The role of CaV1.1 and Ca2+ in actin cytoskeleton organization of SCC12 cells was first explored. A cortical actin organization was correlated with increased actin-rich microspikes formation in response to EGF and matrix stiffness cooperation (Fig. 5A and B), which was abolished by the addition of verapamil and diltiazem (Fig. 5A and B). We next determined whether Ca2+ prompted locomotory forces induced by EGF in SCC12 cells on stiff matrix. The increased forces applied by SCC12 cells stimulated with EGF that were observed in the control were reduced by the addition of diltiazem or verapamil (Fig. 5C and D) concomitantly with the abrogation of both Cdc42-GTP loading and MLC2 phosphorylation induced by EGF (Fig. 5E–H). Overall, these data show that intracellular Ca2+ entry through L-type calcium channels in SCC12 cells mediates actomyosin contractility and Cdc42 activity in epithelial cancer cells, leading to increased locomotory forces and collective invasion.

Figure 5.

L-type-dependent Ca2+ entry regulates actomyosin contractility in SCC cells. A, Representative confocal merged images of phalloidin and DAPI staining in SCC12 cells plated 3 days on soft or stiff condition, with or without EGF (5 ng/mL), diltiazem (30 μmol/L), or verapamil (15 μmol/L). Scale bar, 20 μm. B, Quantification of the number of filopodia per cells represented in A. Bars correspond to the mean and each dot represents the number of filopodia in one cell (n = 30 cells at least; representative experiment of three independent experiments). C, Heatmap of SCC12 cells traction forces applied on 50 kPa hydrogels in the presence or absence of EGF (5 ng/mL), diltiazem (30 μmol/L), or verapamil (15 μmol/L). D, Quantification of SCC12 cells' traction forces corresponding to G. Bars correspond to the medians (representative data of three independent experiments). E, Immunoblot of Cdc42-GTP pull-down assay in SCC12 cells plated 3 days on soft or stiff matrices and stimulated 5 minutes with 5 ng/mL EGF in presence or absence of diltiazem or verapamil. Cdc42 total and tubulin were used as control. F, Quantification of Cdc42-GTP/Cdc42 from three independent experiments performed in the same conditions as in E. G, Immunoblot of p-MLC2 in SCC12 cells plated in the same conditions as in E. MLC2 and tubulin were used as control. H, Quantification of p-MLC2/tubulin of three independent experiments performed as in G. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

Figure 5.

L-type-dependent Ca2+ entry regulates actomyosin contractility in SCC cells. A, Representative confocal merged images of phalloidin and DAPI staining in SCC12 cells plated 3 days on soft or stiff condition, with or without EGF (5 ng/mL), diltiazem (30 μmol/L), or verapamil (15 μmol/L). Scale bar, 20 μm. B, Quantification of the number of filopodia per cells represented in A. Bars correspond to the mean and each dot represents the number of filopodia in one cell (n = 30 cells at least; representative experiment of three independent experiments). C, Heatmap of SCC12 cells traction forces applied on 50 kPa hydrogels in the presence or absence of EGF (5 ng/mL), diltiazem (30 μmol/L), or verapamil (15 μmol/L). D, Quantification of SCC12 cells' traction forces corresponding to G. Bars correspond to the medians (representative data of three independent experiments). E, Immunoblot of Cdc42-GTP pull-down assay in SCC12 cells plated 3 days on soft or stiff matrices and stimulated 5 minutes with 5 ng/mL EGF in presence or absence of diltiazem or verapamil. Cdc42 total and tubulin were used as control. F, Quantification of Cdc42-GTP/Cdc42 from three independent experiments performed in the same conditions as in E. G, Immunoblot of p-MLC2 in SCC12 cells plated in the same conditions as in E. MLC2 and tubulin were used as control. H, Quantification of p-MLC2/tubulin of three independent experiments performed as in G. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

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Diltiazem and verapamil prevent SCC collective invasion in vivo

Inhibition of Ca2+ signaling prevents SCC cell invasion in vitro, which implies that counteracting Ca2+ influx in cancer cells may lead to inhibition of tumor cell invasion in vivo. To verify this possibility, we determined whether inhibition of the L-type–dependent Ca2+ influx influenced human HNSCC tumor cell invasion both in vitro and in vivo. HNSCC multicellular spheroids were embedded in stiff matrix gel in culture media supplemented with EGF to assess their invasion capacity. Addition of verapamil or diltiazem to the patient-derived spheroid cultures drastically reduced the invasiveness of the tumor cells (Fig. 6A and B). Then, the anti-invasive potential of both diltiazem and verapamil was tested in vivo using human HNSCC patient cells subcutaneously engrafted on nude mice. Four days after injection, the mice were pretreated with verapamil and diltiazem in drinking water at low doses (20 mg/kg/day). Administration of full inhibitory doses started 3 days later (7 days postinjection). Tumor growth was significantly reduced in the treated mice compared with the controls (Supplementary Fig. S5A). Immunohistologic analyses of the tumors demonstrated that both verapamil- and diltiazem-treated mice presented less collectively invading tumor cells compared with the untreated control mice (Fig. 6C and D). In control tumors, presence of stromal αSMA–positive cells correlated with robust MLC2 phosphorylation staining. In contrast, tumors developed in verapamil- or diltiazem-treated mice presented reduced levels of pMLC2 (Fig. 6E–G). Both verapamil and diltiazem, therefore, display a strong antitumor activity toward the development of invasive human tumors in vivo.

Figure 6.

Diltiazem and verapamil prevent HNSCC tumor cell collective invasion. A, Representative pictures of human patient–derived HNSCC multicellular spheroid embedded in stiff matrices in the presence or absence of diltiazem (30 μmol/L) or verapamil (15 μmol/L) for 5 days. Scale bar, 200 μm. B, Quantification of HNSCC multicellular spheroid invasion shown in A (n = 5 for 5 individual patients; bars correspond to mean). A.U., arbitrary unit. C, Representative hematoxylin and eosin pictures of human patient–derived HNSCC tumors showing collective invasion in control, diltiazem, or verapamil-treated group mice. Scale bar, 200 μm. D, Quantification of tumors presenting invading cell cohorts shown in C (n = 6 tumors/group). E, Representative confocal images of αSMA, p-MLC2, and DAPI staining in PDX from control mice, verapamil or diltiazem-treated mice. Scale bar, 50 μm. F, Quantification of αSMA staining shown in E. G, Quantification of p-MLC2 staining shown in E. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

Figure 6.

Diltiazem and verapamil prevent HNSCC tumor cell collective invasion. A, Representative pictures of human patient–derived HNSCC multicellular spheroid embedded in stiff matrices in the presence or absence of diltiazem (30 μmol/L) or verapamil (15 μmol/L) for 5 days. Scale bar, 200 μm. B, Quantification of HNSCC multicellular spheroid invasion shown in A (n = 5 for 5 individual patients; bars correspond to mean). A.U., arbitrary unit. C, Representative hematoxylin and eosin pictures of human patient–derived HNSCC tumors showing collective invasion in control, diltiazem, or verapamil-treated group mice. Scale bar, 200 μm. D, Quantification of tumors presenting invading cell cohorts shown in C (n = 6 tumors/group). E, Representative confocal images of αSMA, p-MLC2, and DAPI staining in PDX from control mice, verapamil or diltiazem-treated mice. Scale bar, 50 μm. F, Quantification of αSMA staining shown in E. G, Quantification of p-MLC2 staining shown in E. For all data, paired samples were compared by two-tailed Student t test, whereas one-way ANOVA and post hoc Tukey tests were used for group comparisons (NS, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001).

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Fibroblast activation in cancer leads to the emergence of carcinoma-associated fibroblasts that produce a variety of different procarcinogenic components, which results in pleiotropic actions on the tumor cells (28). CAFs support each step of the neoplastic development from tumor initiation to metastatic spreading (29, 30). CAFs lead the way of collectively invading epithelial carcinoma cells by digging tracks within the ECM that tumor cells use to invade (6). Thus, cancer cell invasion and tumor progression rely on complex communication networks that involve a variety of noncancerous cells and noncellular components found within the tumor microenvironment. Most studies agree that CAF-enriched tumor stroma supports tumor progression and invasion, either through secretion of carcinogenic molecules (28) or physical matrix organization and stiffening (8, 10). Here, we uncover a novel cross-talk mechanism by which matrix stiffening mechano-sensitizes human SCC cells to EGFR-driven invasiveness. We show that SCC cells, in response to matrix stiffening, increase EGFR expression, thus sensitizing carcinoma cells to EGFR phosphorylation, which results in acto-myosin contractility and collective invasion. Modulation of matrix stiffening was obtained by increasing collagen I concentration (Fig. 1A–D), which also results in microscale matrix parameter modification that critically regulates cell invasion, such as pore size, fiber architecture, local material deformability, and out-side-in signaling pathways in cancer cells (31). Thus, to confirm the critical role of matrix rigidity specifically, we conducted key experiments in collagen I matrices of different rigidity using both ribose-dependent collagen glycation and hydrogels (25), which confirmed that collective invasion is driven by the mechanical regulation of EGFR activity (Fig. 1E–J). Altogether, this work provides evidence that mechanical matrix property regulates EGFR signaling and collective invasion of squamous cell carcinoma.

Blocking EGFR activity, using the FDA-approved RTKi gefitinib, results in inhibition of both SCC12 cell collective invasion and tumor development in vivo. However, it is admitted that EGFR-targeting therapies fail to improve the patients' survival (32). It has been noted that more that 40% of HNSCC lesions present SNP in EGFR gene, which leads to EGFR-targeted therapy resistance (33). Moreover, blocking EGFR induces EMT and CAF activation in HNSCC tumors (34). Nevertheless, the EGFR signaling pathways remain interesting pharmaceutical targets because of the high level of EGFR amplification and genetic mutations found in patients with cancer (35).

Our work aimed to target the molecular mechanisms downstream the EGFR signaling in cancer cell collective invasion. Screening for small molecules in organotypic culture assays identified the L-type CCBs verapamil and diltiazem as potent cancer invasion inhibitors. Further experiments highlighted the role of Cav1.1 L-type channel isoform downstream of the cooperation between EGFR and stiff ECM in SCC cells. Overall, our screening approach led to the identification of 48 inhibitors that block SCC12 cell collective invasion that represented 16% of the total library. Among the 48 identified compounds that actively inhibit SCC12 cell invasion, 14 of them (29.2%) target the CAF proinvasive activity, and 34 (70.8%) the SCC12 cells. Among these latter, 14 (41.2%) were calcium channel inhibitors, and 9 (26.5%) of them specifically target L-types Ca2+ channels. Undoubtedly, deregulated Ca2+ homeostasis is observed in various disorders, including cancers (20, 36). In solid cancers, Ca2+ signaling has been linked to metastasis formation by controlling several fundamental aspects of cancer cell behavior, such as EMT (37, 38), migration (39), angiogenesis, and intravasation (40). Alterations in the expression of specific Ca2+ channels have been reported in several cancer types (41, 42), including HNSCC and colon (39, 43, 44), where it sustains Ca2+-sensitive oncogenic pathways. We identified the CaV1.1 Ca2+ channel as a downstream signaling molecule of the EGFR-dependent tumor cell collective invasion in HNSCC carcinomas. We also show a preferential CaV1.1 expression in cells at the edge with the matrix in human tumor samples biopsies. Because other L-type Ca2+ channels are expressed in epithelial cancer cells (39, 44), the broad spectrum of action of verapamil and diltiazem reinforces their potential therapeutic use in epithelial cancers.

This work reinforces the crucial role of the Cdc42 small GTPase activity in SCC12 cell collective invasion. We have previously shown that a Cdc42/MRCK signaling pathway, and not the RhoA/ROCK signaling pathway, drives actomyosin contractility in SCC12 cells (6). Here, we propose that increased substratum rigidity in tumor tissue favors EGFR tyrosine kinase activity and leads to a Ca2+-dependent regulation of the Cdc42 small GTPase activity in cancer cells. Such a signaling route seems crucial for the capacity of the cells to generate and apply forces to the substrata, which allows tumor cell collective strands to invade. Interestingly, the myosin light chain kinase (MLCK), a fundamental regulator of actomyosin contractility in cells (45), does not play a role in SCC12 cell collective invasion; however, it regulates actomyosin–dependent ECM remodeling in CAF cells (Supplementary Tables S2 and S3). Such a differential regulation observed in tumor and stromal cells demonstrates that additional comprehensive work must be addressed to identify a single molecular target for therapeutic purpose that could block actomyosin activity in both cell types.

The major ligands of EGFR are EGF, heparin binding-EGF, TGFα, amphiregulin, epiregulin, epigen, and betacellulin (46). Tumor-associated macrophages are a prominent source of EGF in the tumor microenvironment, and their presence is associated with poor prognosis in multiple cancers (47). Although our work specifically focuses on the downstream activation of the EGFR in SCC, we can speculate that other EGFR ligands may be implicated in this process. Also, it has been reported that adhesion to ECM could lead to activation of several growth factors, including EGFR, in the absence of ligand (48, 49), which might explain that addition of gefitinib inhibits the basal level of collective invasion we observed in response to stiffness alone.

Verapamil and diltiazem are known to inhibit multidrug resistance pumps, implicated in chemoresistance (50). We provide evidence that treatment with either verapamil or diltiazem reduces SCC tumor invasion in vitro, in vivo, and in patient primary cells. These properties of the two drugs are novel, despite the fact that verapamil and diltiazem have been used in medical care for decades. Indeed, the clinical relationship between carcinoma progression and nondihydropyridine CCBs in patients was never reported so far. However, verapamil was tested in patients in combination with traditional chemotherapy by intra-arterial infusion at high concentrations. This protocol improved the treatment efficacy without any side effects with positive effects on the patients' survival and quality of life (51–53), whereas the EGFR-targeted therapies induced severe side effects, including cutaneous (54) or pulmonary toxicity (55).

In conclusion, our work demonstrates that targeting the main downstream second messenger of EGFR signaling in cancer results in inhibition of tumor cell collective invasion both in vitro and in vivo. In particular, targeting L-type Ca2+ channels in SCC lesions could open novel therapeutic strategies to block the tumor-promoting activity of the tumor niche with reduced risks of resistance to treatments.

No potential conflicts of interest were disclosed.

Conception and design: E.M. Grasset, T. Bertero, I. Bourget, M. Lecacheur, C. Gaggioli

Development of methodology: E.M. Grasset, I. Bourget, P. Hofman, C. Duranton, C. Gaggioli

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): E.M. Grasset, T. Bertero, A. Bozec, J. Friard, I. Bourget, S. Pisano, M. Lecacheur, C. Bailleux, A. Emelyanov, M. Ilie, P. Hofman, G. Meneguzzi, C. Duranton, D.V. Bulavin, C. Gaggioli

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): E.M. Grasset, T. Bertero, S. Pisano, M. Lecacheur, C. Duranton, C. Gaggioli

Writing, review, and/or revision of the manuscript: E.M. Grasset, P. Hofman, G. Meneguzzi, C. Duranton, C. Gaggioli

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): E.M. Grasset, I. Bourget, M. Maiel, D.V. Bulavin, C. Gaggioli

Study supervision: C. Gaggioli

The authors acknowledge the IRCAN's Molecular and Cellular Core Imaging Facility (PICMI), the Histology core facility, the IRCAN Animal core facility, and the Genomic core facility. The IRCAN facilities were supported financially by the Région PACA, Canceropole PACA, the EEC ERC program, and the “Conseil Général 06.” We thank E. Selva (Hospital-Integrated Tumor Biobank, Pasteur Hospital, Nice, France) for providing the tumor samples. We thank Dr. E. Boulter for his help on the locomotory forces quantification. This work was supported by the French Government (National Research Agency, ANR) through the « Investments for the Future » LABEX SIGNALIFE: the Program reference # ANR-11-LABX-0028-01 (to G. Meneguzzi). This work was supported by grants obtained by C. Gaggioli from ARC “PJA20171206434,” FRM “DEQ20180339183,” ANR “ANR-14-RARE-0004-02,” Foundation Silab Jean Paufique, and the Debra UK foundation. E.M. Grasset is the recipient of an LNCC and ARC fellowships.

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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