Matrix stiffness regulates epithelial-mesenchymal transition via cytoskeletal remodeling and MRTF-A translocation in osteosarcoma cells

Journal of the Mechanical Behavior of Biomedical Materials 90 (2019) 226–238

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Matrix stiffness regulates epithelial-mesenchymal transition via cytoskeletal remodeling and MRTF-A translocation in osteosarcoma cells ⁎

Jun Dai1, Liang Qin1, Yan Chen, Huan Wang, Guanlin Lin, Xiao Li, Hui Liao , Huang Fang

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Department of Orthopedics, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, 1095 Jiefang Avenue, Qiaokou District, Wuhan 430030, China

A R T I C LE I N FO

A B S T R A C T

Keywords: Matrix stiffness Epithelial–mesenchymal transition Cytoskeleton Myocardin-related transcription factor A Osteosarcoma

Matrix stiffness is known to alter cellular behaviors in various biological contexts. Previous investigations have shown that epithelial–mesenchymal transition (EMT) promotes the progression and invasion of tumor. Mechanical signaling is identified as a regulator of EMT. However, the molecular mechanisms underlying the influence exerted by matrix stiffness on EMT in osteosarcoma remains largely unknown. Using polyacrylamide hydrogel model, we investigate the effects of matrix stiffness on EMT and migration in osteosarcoma. Our data indicates that high matrix stiffness regulates cell morphology and promotes EMT and migration in osteosarcoma MG63 cell line in vitro. Notably, matrix stiffness promotes polymerization of actin and nuclear accumulation of myocardin-related transcription factor A (MRTF-A). Furthermore, inhibiting MRTF-A by CCG 203971 significantly reduces EMT and migration on rigid gels. These data suggest that matrix stiffness of the tumor microenvironment actively regulate osteosarcoma EMT and migration through cytoskeletal remodeling and translocation of MRTF-A, which may contribute to cancer progression.

1. Introduction Osteosarcoma is considered as one of the most common non-hematologic and primary malignancy of bone with peak incidence in pediatric patients (Reed et al., 2017; Torre et al., 2015), accounting for 20–40% of all bone cancers (Valery et al., 2015). Thanks to establishment of standard treatment paradigm of neoadjuvant chemotherapy combined with surgery, the 5-year survival rate has increased from 10% to 70% (Harrison et al., 2018; Yang et al., 2018). However, osteosarcoma prognosis remains poor and its recurrence usually leads to death due to metastases (Brown et al., 2017). To provide new cancer therapy and improved clinical outcomes, research efforts should be promoted to deepen our understanding of osteosarcoma initiation and progression from the molecular mechanistic level. Microenvironment surrounding tumor cells is crucial for cell fate and behavior. From the standpoint of a cell, the term “microenvironment” can be categorized into chemical environment, surface chemistry and physical environment, cell-cell interactions and cell-extracellular matrix interactions. As a vital constituent of local microenvironment, matrix stiffness is closely associated with the initiation and development of tumor (DuFort et al., 2011). Due to the deposition and remodeling of ECM, surface tension and stiffness are significantly

increased in various solid tumors. Such alternation is believed to promote tumor cell proliferation and migration (Lampi and Reinhart-King, 2018; Miyazawa et al., 2018; Wei et al., 2017). Such a process is particularly pronounced in liver cancer. The rigid matrix of liver cancer tissue can be attributed from fibrosis forces modification of cell contractility to adapt to environment and to maintain tensional homeostasis, which subsequently exerts on intracellular signaling leading to malignant transformation (Affo et al., 2017; Saneyasu et al., 2016). Moreover, similar changes can be observed in breast cancer that tumorigenesis is associated with collagen crosslinking extent and ECM stiffening. In addition, when the stiffness increases in the tumor microenvironment, cancer-associated fibroblasts can be activated, leading to progression of breast cancer (Katara et al., 2018; Wang et al., 2017). Through transmembrane proteins with multicomponent adhesion sites, matrix stiffness can be converted from an extracellular mechanical signal to an intracellular biological signal to regulate cell behavior. (Lantoine et al., 2016; Muller and Pompe, 2016; Trappmann et al., 2012). More specifically, ECM stiffness can interact with cytoskeleton through forming focal adhesions and regulating various downstream signaling pathways. A number of studies have been conducted to unveil the relationship between mechanotransduction and physiological/pathophysiological status of cells. One of these studies demonstrated that

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Corresponding authors. E-mail addresses: [email protected] (H. Liao), [email protected] (H. Fang). 1 Jun Dai and Liang Qin contributed equally to this work. https://doi.org/10.1016/j.jmbbm.2018.10.012 Received 16 May 2018; Received in revised form 2 October 2018; Accepted 3 October 2018 Available online 16 October 2018 1751-6161/ © 2018 Elsevier Ltd. All rights reserved.

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the β-catenin expression was upregulated by biomechanical changes in uterine fibroids (Ko et al., 2018). Another study suggested that modification of stiffness cues, the arterial and venous differentiation of endothelial progenitor cells could be controlled via the Ras/Mek pathway (Xue et al., 2017). Furthermore, as known downstream transducers of the Hippo pathway, YAP/TAZ co-regulators also possess important effects in connecting mechanical stimuli with gene transcription (Dupont, 2016; Rammensee et al., 2017). However, few studies were reported regarding specific mechanisms of mechanotransduction in osteosarcoma. Therefore, further investigation on disease mechanism, in particular initiation and progression of osteosarcoma, is still necessary to warrant a more detailed understanding. The epithelial–mesenchymal transition (EMT) acts as a crucial process in tumor progression and metastasis, as well as in fibrosis (Li et al., 2018). Epithelial cells can acquire characteristics of mesenchymal cells, including the loss of cell–cell and cell–basement membrane associations and changes in cell morphology and the expression of many proteins (Lopez-Novoa and Nieto, 2009). More specific, a monolayer cuboidal shaped epithelial cells with polarity transition to an elongated spindle-shaped mesenchymal cell with ability of migration (Bhowmick et al., 2001). Previous research have demonstrated that matrix stiffness plays important role in driving EMT in mammary and kidney epithelial cells via soluble factor induction and intercellular signaling cascades, such as TGF-β1, YAP and TAZ (De Craene and Berx, 2013). However, the key factor that matrix stiffness controlling induction of EMT processes in osteosarcoma cell remains unclear. Myocardin-related transcription factor A (MRTF-A), initially found in an acute megakaryoblastic leukemia study of a chromosomal translocation, accounts as a transcriptional coactivator of serum-response factor (SRF) (Kalita et al., 2012). The subcellular localization and activity of MRTF-A is mediated by the remodeling of the actin cytoskeleton, while the arrangement of actin can be controlled by ECM stiffness. After the process of nuclear translocation, MRTF-A combined with CArG-box in the promoters of genes to activate transcription. This process can be triggered by TGF-β and Rho signaling pathway (Morita et al., 2007). Meanwhile, as a specific targeted small molecule inhibitor, CCG 203971 exerts blocking effect on translocation of MRTF-A through binding of the N-terminal basic domain (Sisson et al., 2015). The advantages of CCG-203971 over the original compound CCG 1423 are a significant reduction in cytotoxicity, with increased potency and selectivity. Although previous study demonstrated that the subcellular localization of MRTF-A is an important factor in mediating EMT (O'Connor et al., 2015), few relevant report on osteosarcoma has be found yet. Owing to these research gaps, we hypothesize that matrix stiffness can regulate EMT process in osteosarcoma cells via MRTF-A translocation. Our results suggested that matrix stiffness plays a crucial role in promoting upregulation of mesenchymal markers, cell morphology and reorganization of the actin cytoskeleton. Besides, MRTF-A is a key regulator of matrix rigidity controlled EMT. Thus, it is important for studies cellular mechanotransduction in osteosarcoma, suggesting a role for MRTF-A in sensing mechanics and tuning EMT activity.

purchased from Proteintech Group (Wuhan, Hubei, China). Horseradish peroxidase (HRP)-conjugated secondary antibodies were acquired from Jackson ImmunoResearch Laboratories (West Grove, PA, USA). 2.2. Fabrication of polyacrylamide hydrogels and compression test According to a previously established protocol, polyacrylamide hydrogels of diversity stiffness were prepared by adjusting the relative concentrations of acrylamide (monomer) and bis-acrylamide (crosslinker) (Sigma-Aldrich). Firstly, acrylamide and bis-acrylamide were mixed with the corresponding proportions respectively according to the desired difference of stiffness. This solution was sterilized by filtration with 0.22 µm filter. Dissolved oxygen in solution was removed by vacuum. Then, solution containing 0.1 M NaOH on coverslips was dried on a heating plate. About 200 µL 3-aminopropyltrimethoxysilane (APES; Sigma-Aldrich) was added on the surface of coverslip for 5 min and fully rinsed with distilled H2O. Then coverslips were soaked in 0.5% glutaraldehyde (Sigma-Aldrich) for 30 min. After drying in a fume hood, amino-silanated coverslips (25 mm diameter for western blotting and 12 mm diameter for immunofluorescence) were prepared. Glass slides were treated with dichlorodimethylsilane (DCDMS) for 5 min before rinsing with distilled H2O, resulting in chloro-silanated glass slides. Finally, TEMED (Sigma-Aldrich) and 10% ammonium persulfate (Sigma-Aldrich) were added to the mixed acrylamide and bis-acrylamide solutions for polymerization. Before the solution was polymerized, it (100 µL) was transferred onto prepared chloro-silanated glass slide and covered by amino-silanated coverslips. After polymerization completed in about 20 min, hydrogels were removed from slides and cover slips for further evaluation. To facilitate cell adhesion, protein coating was performed on the surface of hydrogels. About 100 µL sufosuccinimidyl-6-(4'-azido-2'-nitrophenylamino)-hexanoate (sulfo-SANPAH; Sigma-Aldrich) was smeared to the surface of hydrogels which was exposed to UV light at 365 nm for 10 min. Resultant gels were then rinsed with 50 mM HEPES buffer, incubated at 37 ° overnight with 10 µg/mL solution of Rat Tail Collagen I (Thermo Fisher Scientific), and sterilized under UV light for 20 min. Previous study demonstrated that the stiffness of the hydrogel could not be affect by protein coating (Chen et al., 2010). Here we applied an Instron Universal Testing Machine (Instron Series 5500 Load Frames) to perform compression test of freshly prepared hydrogels. Elastic modulus represented the measurement scale of an object to resist elastic deformation, which only related to chemical composition of the material and consistent with Hooke's law. The calculation formula of the elastic modulus (E) is known as E = F / A , where F is the force exerted △L / L on hydrogel under compression, A is the actual cross-sectional area which equals the area of the cross-section perpendicular to the applied force, ΔL is the amount by which the height of hydrogel changes, L is the original height of hydrogel. Acrylamide and bis-acrylamide solution were mixed at various proportions (Table 1) with addition of TEMED and APS before pouring into a 24-well plate for polymerization. The hydrogels was removed after solidification. Diameter and height of this columnar hydrogel were measured to be 18 mm and 10 mm, respectively (Figs. 2A, 2D and 2G). To conduct pressure test, hydrogels placed in the center of the test platform were compressed at a constant rate of 0.5 mm/min to obtain the stress–strain curve (Fig. 1). Elastic modulus of hydrogels can be determined by the slope of the first 10% linear

2. Materials and methods 2.1. Reagents and antibodies CCG 203971 was purchased from Tocris Bioscience (Bristol, United Kingdom). Before each use, CCG 203971 was freshly dissolved in dimethyl sulfoxide (DMSO) (vehicle). Calcein AM (ab141420), primary antibodies against MRTF-A (ab49311), E-cadherin (ab76055), vimentin (ab8978), snail (ab180714) and fibronectin (ab2413), and goat antirabbit second antibody Alexa Fluor 594 were obtained from Abcam (Cambridge, United Kingdom). Fluorescein isothiocyanate labeled peptide from Amanita phalloidin was obtained from Sigma Aldrich (St. Louis, MO, USA). Primary antibody against MMP-9 (10375–2-AP) was

Table 1 Preparation of polyacrylamide hydrogels with various stiffness.

Acrylamide (40%) (mL) Bis-acrylamide (2%) (mL) Water (mL)

227

Soft

Medium

Rigid

0.75 1.5 7.75

2 1.32 6.68

2.5 1.5 6

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circularity equals 1.0, whereas 0.0 indicates an irregular polygon. In brief, images were converted into 8-bit pattern, and subjected to adjustment of appropriate of brightness/contrast and threshold. Then reversed images were created using “invert” function. Parameter A and P were then obtained by “analyze particles” function. Then the mean value of A in each group was calculated and the results greater than 1.5 times the mean value were excluded. For each group, 3 areas and at least 80 cells were analyzed. 2.6. Immunofluorescence Cells on glass slides or on hydrogels substrate were fixed in 4% paraformaldehyde at room temperature for 20 min and permeabilized using 0.5% Triton X-100 (Solarbio, Shanghai, China) for 10 min. Fixed cells were blocked with 5% bovine serum albumin (Sigma-Aldrich) in PBS-0.1% Triton X-100 for 30 min, followed by overnight incubation at 4 ° with rabbit polyclonal primary antibody for MRTF-A. After rinsing with PBS, the cells were incubated with goat anti-rabbit second antibody Alexa Fluor 594 at room temperature for 1 h in the dark. Then, for staining of the actin cytoskeleton, the cells incubated with phalloidin at 37 °C for 1 h as a counterstain. Finally, the cells were stained with DAPI. Images were collected by using FV1000 laser scanning confocal microscopy equipped with an Olympus Fluoview FV1000 system (Olympus, Tokyo, Japan) or using an EVOS FL Auto Imaging Systems (Thermo Fisher Scientific).

Fig. 1. Schematic diagram describing the process of pressure test.

region of stress–strain curve (Pilipchuk et al., 2013), which in this case 10% of measured hydrogel height (10%*10 mm = 1 mm) was employed as endpoint of stress test (Figs. 2B, 2E and 2H). Mean elastic modulus of at least five hydrogels were measured for each stiffness condition. 2.3. Cell culture The cell lines MG63 were obtained from the Chinese Academy of Sciences (Shanghai, China). MG63 is an established human osteosarcoma cell lines, which derives from a juxtacortical osteosarcoma diagnosed in the distal diaphysis of the left femur of a 14 year old male (Billiau et al., 1977). MG63 cells exhibit osteoblast phenotypic characteristics and it has the ability to retain a differentiated phenotype under culturing conditions. Not only MG63 have PTH-unresponsive adenylate cyclase, but also contain a specific lα,25-dihydroxyvitamin D3 receptor (Czekanska et al., 2012). Meanwhile, MG63 cells show fast proliferation and is therefore widely used in research of osteosarcoma (Decker et al., 1999). The cells were maintained in DMEM/F12 medium, which was supplemented with 10% heat-inactivated fetal bovine serum, 100 U/mL penicillin and 100 µg/mL streptomycin and incubated at 37 °C in a water-saturated atmosphere with 5% CO2. MG63 cells were plated in each specimen at a density of 5000 cells/cm2. Prior to any treatment, cells were grown to about 70–80% confluence.

2.7. Western blotting Western blotting was used to determine MRTF-A and EMT protein expression in cells of different groups. After the incubation period, cell protein was extracted. RIPA buffer (100 µL) supplemented with 1 mM phenylmethanesulfonyl fluoride solution (PMSF) and a phosphotransferase inhibitor (Boster Biotechnology) were transferred onto a sheet of parafilm. Then the coverslips were pinched by curved forceps and placed (cell-side down) on the top of droplets for each group. Cells were thus incubated with the lysis buffer for 3 min before protein lysates being transferred to a micro centrifuge tube. Protein concentration was determined with the BCA Protein Assay Kit (Boster Biotechnology). Samples were loaded onto and separated by 8% or 10% SDS-PAGE gel electrophoresis, which was then transferred to a polyvinylidene difluoride membrane (Millipore, Billerica, MA, USA). The membranes were blocked with 5% BSA in TBST (Tris-buffered saline with 0.1% Tween-20) for 1 h at room temperature and then incubated with an appropriate amount of primary antibodies overnight at 4 °C. The Blots were washed in TBST and incubated with HRP-conjugated secondary antibodies in 5% non-fat milk for 1 h at room temperature. Then, the signals were detected with ECL (Thermo Fisher Scientific). Protein levels were normalized with GAPDH and laminB1. The bands were semiquantified through densitometry using Image J software.

2.4. Treatment of CCG 203971 MG63 cells were seeded on polyacrylamide hydrogels of rigid substrate. In particular, cells were seeded in serum-reduced media (DMEM/ F12 supplemented with 0.5% FBS and 100 U/mL penicillin and 100 µg/ mL streptomycin) and incubated for 24 h. Medium was then replaced by fresh serum-reduced media containing 10 µM CCG 203971 or an equivalent volume of DMSO vehicle control. After treatment for 24 h, cells were harvested for analysis.

2.8. Wound healing assay by culture insert Cell migration assay was performed in a culture-insert assay. In accordance with the manufacturer's instructions (ibidi 80206, Martinsried, Germany), culture inserts composed of two chambers and separated by a 500 µm distance, were placed onto the glass slides or hydrogels substrate by using sterile micro forceps. Cell suspensions were prepared and counted with cell counter at a concentration of 5 * 105 cells/mL in 10% FBS medium, and then 70 µL was placed into each chamber of culture insert and incubated at 37 °C and 5% CO2. After cell attachment was observed, the culture inserts were removed and medium was replaced by serum-free media (For CCG203971 treated group and control group, the medium was replaced by FBS free media containing 10 µM CCG 203971 and DMSO respectively). All cell groups on glass or hydrogel coverslips were incubated at 37 °C in 5% CO2 for 24 h and media was not changed during this time. For each

2.5. Calcein AM staining and measurement of cell circularity To visualize cell morphology and quantify cell circularity，cells were stained with calcein AM staining according to the manufacturer's instructions (Abcam, Cambridge, United Kingdom). Adherent MG63 cells were washed by PBS, and then incubated with 1 µM calcein AM at 37 °C for 30 min, protected from light. Images were captured by an EVOS FL Auto Imaging Systems (Thermo Fisher Scientific). To calculate cell circularity depending on the various stiffness of polyacrylamide substrates, images were processed based on image J software. Circularity was defined as C = 4 π (A/P2), hereinto A is cell area and P is cell perimeter. It means a perfect circle when the value of 228

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Fig. 2. Elastic modulus measured by Instron Universal Testing Machine. (A, D and G) were anteroposterior and lateral views of three kinds of hydrogels. (B, E and H) were graphs of stress against strain in different stiffness, red box indicated the first 10% linear region. (C, F and I) were slopes of the line, respectively. (J) showed the stiffness range of hydrogels (mean ± S.D.). Soft, medium and rigid were to be 2.10 ± 0.25, 21.92 ± 1.10 and 33.65 ± 1.50 kPa, respectively.

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abundance in stiff substrate and control groups (Fig. 3D). The results were consistent with the previous studies that the cells adopted a spindle-shape morphology and exhibited the organization of filamentous actin on the rigid substrate hydrogels (Sanz-Ramos et al., 2013; Trappmann et al., 2012). The actin cytoskeleton mediated the change of cell morphology by regulating SRF transcriptional activity (Connelly et al., 2010). SRF has been documented to interact with co-factor MRTF-A, while transcriptional activity of MRTF-A can be regulated by association with G-actin (a monomeric actin form of F-actin) (Miralles et al., 2003). Immunofluorescence and western blotting were performed to determine subcellular location of MRFT-A expression in various substrate stiffness. Our results demonstrated that MRTF-A predominantly expressed and distributed in nuclei of MG63 cells with increasing matrix stiffness, whereas almost little nuclear accumulation of MRTF-A was observed on soft substrate (Fig. 3D and Fig. 4). Consistent with previously published studies, our data suggest that subcellular localization of MRTF-A could be greatly affected by extracellular matrix stiffness through the assembly of actin. In addition, this was the first time to show matrix stiffness could regulate nuclear translocation of MRTF-A in osteosarcoma cells.

group, image was obtained before and after experiment using an EVOS FL Imaging Systems (Thermo Fisher Scientific), and analyzed with Image J software determine would closure. Each experiment was performed at least three times independently. 2.9. Statistical analysis Prism 6 software (Version 6.0; GraphPad Software Inc., CA, USA) was utilized for statistical analysis and generation of graphs. At least three experimental replicates were performed for all studies. Data were reported as mean ± standard deviation. Student's t-test and one-way ANOVA were used to analyze the differences among groups. For all tests, p < 0.05 was considered as statistical significance and p values were denoted as follows: * p < 0.05, * * p < 0.01, * ** p < 0.001 and * ** * p < 0.0001. 3. Results 3.1. Tunable polyacrylamide hydrogels were established An extremely crucial step in this study was to establish the hydrogel system with suitable gradient stiffness. The mechanical properties of hydrogel substrates could readily be tuned by altering the concentration of cross-links in the gel. Here three different relative density of acrylamide and bis-acrylamide were adopted to provide a spectrum of material stiffness for further evaluation. The compression test by Instron Universal Testing Machine was conducted to confirm stiffness range and stability of hydrogel. According to formula E = F / A , elastic △L / L modulus (E) could be positively correlated with slope (F/ΔL) which was tabulated from Figs. 2C, 2F and 2I. Our results showed that stiffness ranging from soft, medium and rigid were to be 2.10 ± 0.25, 21.92 ± 1.10 and 33.65 ± 1.50 kPa (mean ± SD), respectively (Fig. 2J). This data suggested high consistency and reproducibility in measuring elastic modulus using Instron Universal Testing Machine.

3.4. Matrix stiffness promoted the epithelial–mesenchymal transition The EMT process not only results in significant changes in cell morphology, but also involves down-regulation of the epithelial marker E-cadherin and activation of the mesenchymal marker vimentin and fibronectin. As a critical indicator of EMT, E-cadherin can be found at the junctions of cells, and transcriptional expression of E-cadherin was in turn regulated by snail. Detection of aforementioned proteins would provide evidence to determine whether matrix stiffness could actually promote EMT process. As demonstrated in Fig. 5, western blotting results showed that increased of E-cadherin and decreased of vimentin, fibronectin and snail in cells on soft substrate, compared to rigid and control groups. Whereas, decreased E-cadherin and increased vimentin, fibronectin and snail were observed in medium, rigid and control groups. Therefore, this data clearly indicated that increase in matrix stiffness facilitated formation of mesenchymal phenotype during EMT, which play a significant role in the process of tumor metastasis.

3.2. Extracellular matrix stiffness regulated cell morphology in vitro The morphology of cells undergoes significant changes during the EMT process. Therefore, precise assessment of cell morphology is essential in our study. To determine whether matrix stiffness could affect cell shape, specifically circularity, calcein AM staining was employed for visualizing cell boundaries. With optical microscopy, dramatic distinction in cell morphology and organization of MG63 cells were observed when seeded on various substrates including soft, medium and rigid hydrogels and glass slides (Control). Cells on soft substrate appeared to be rounded and organized in rounded cell clusters, while cells on medium, rigid and control group tend to be elongated and scattered (Fig. 3A). Such visual observation was further supported by the quantitative measurement of cell perimeter and surface area with Image J and ultimately circularity (Fig. 3B). As shown in Fig. 3C, unlike most of groups with circularity value below 0.5, the soft group exhibited a much rounded cell shape with circularity of 0.75 ± 0.12 (closer to 1.0), suggesting beneficial effect of soft substrate to improve cell circularity. This data suggested that morphology of cell, in particular circularity was significantly affected by extracellular matrix stiffness.

3.5. CCG 203971 prohibited nuclear translocation of MRTF and EMT process To investigate whether MRTF-A nuclear translocation was essential for EMT, MG63 cells were treated with CCG 203971 on the most rigid hydrogel substrate which illustrated abundant accumulation of MRTF-A in nuclear. Likewise, we first determined the cell morphology and circularity, and then the localization of MRTF-A and protein levels of EMT. Cells exposed to CCG 203971 appeared smaller and rounder compared to DMSO treated group (Figs. 6A, 6B and 6C). In addition, nuclear translocalization of MRTF-A was effectively blocked by CCG 203971 (Figs. 6D and 6E). Western blotting also revealed down-regulated expression of mesenchymal markers, albeit restoration of E-cadherin protein level was not detected CCG 203971 treated group (Fig. 7). These data further strengthens the evidence linking MRTF-A with EMT in osteosarcoma cells. 3.6. Matrix stiffness facilitated migration of MG63 cells

3.3. Extracellular matrix stiffness regulated arrangement of actin cytoskeleton and nuclear translocation of MRTF-A

As the next step, a wound-healing assay was performed to observe the effect of matrix stiffness and CCG 203971 during cell migration. After 24 h incubation, MG63 cells on soft substrate showed healing delay in comparison to other groups, and as expected, control group suggested obvious healing effect as seen by decreased gap area remaining (Figs. 8A and 8B). Additionally, CCG 203971 exhibited inhibitory effect on cell migration behaviors, as shown by less MG63 cells

F-Actin was visualized via fluorescein isothiocyanate labeled phalloidin, thus we could observe whether matrix stiffness effects actin cytoskeletal structure. Although formation of F-actin were not observed in both soft and medium groups, cell morphology in medium group seemed to be more spreading than that in soft group. Unlike soft and medium groups, actin filaments or stress fibers could be visualized in 230

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Fig. 3. Osteosarcoma cells on glass slides (control) and hydrogel's substrate (soft, medium and rigid). (A) Cell's morphology on various stiffness substrate under light microscopy. Scale Bar= 50 µm. (B) Calcein AM staining revealed the outline of each cell. Scale Bar= 50 µm. (C) Circularity of each group was calculated (mean ± S.D.). (D) Confocal images showed differences in actin organization and MRTF-A localization. Scale Bar= 50 µm.

stiffness in high elastic modulus, abundance MMP-9 expression was observed, consistent with wound-healing assay. In accordance with these results, we concluded that increasing matrix stiffness promoted cell migration, in which could be effectively blocked by RhoA pathway

in the gap region (Figs. 8C and 8D). Since MMP 9 could be involved in the migration of osteosarcoma cells (Wu et al., 2013), it was of our interests to explore the link between MMP expression and migration in our cell lines. Again, our results in Fig. 8E suggested that when matrix 231

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Fig. 4. (A) Western blotting analysis of MRTF-A. Values were normalized to GAPDH and laminB1. (B, C and D) Histogram of relative protein expression in total protein, cytoplasmic protein and nucleic protein. Data were presented of three independent experiments. *p < 0.05, * *p < 0.01, * **p < 0.001 and * ** *p < 0.0001.

remodeling of the cytoskeleton, and ultimately affected EMT process of osteosarcoma cells as manifested in the reduction of epithelial marker E-cadherin on the rigid matrix. The levels of expression of the mesenchymal markers vimentin, fibronectin, and the invasion index MMP 9 were increased, whereas those on the soft substrate were opposite to those described above. In addition, the mesenchymal phenotype of osteosarcoma cells can be significantly inhibited by blocking the nuclear import of MRTF-A using drugs. Our results suggest that targeting MRTF-A by molecular-mechanical signaling pathways may be a potentially effective method for the treatment of bone tumors. The cells in our body can sense and respond to various mechanical stimuli from the surrounding three-dimensional environment, and can convert these stimuli into biochemical signals resulting in a series of downstream effects. This process is defined as mechanical signal transduction (mechanotransduction)(Ingber, 2006). Mechanical signal transduction has been widely reported in the regulation of cell and extracellular structures, which play an important role in fostering development and maintaining homeostasis of organs and tissues. However, when such as process shows defective, it might lead to occurrence and development of pathological conditions, ranging from muscular

inhibitor CCG 203971. 4. Discussion Tumor tissues possess an organ-like structures and exhibit substantial heterogeneity between cellular and non-cellular components within their microenvironment. Numerous studies have approved that microenvironment plays a critical role in the occurrence and progression of tumors (Calvo and Sahai, 2011; Joyce and Pollard, 2009). Thorough studies in tumor microenvironment-specific molecular and cellular interactions should draw tremendous attention in cancer research, since these precious understanding not only contribute to clarify tumor pathogenesis but also warrant personalized and/or targeted therapy, to a great extent. Although the hypothesis, design and significance of our study was intended to resonate with such a mainstream tone in cancer research, we discovered a unique relationship between substrate stiffness and cell morphology with entailed underlying molecular mechanisms. One of the most important mechanism found in our study accounts for that matrix stiffness regulated nuclear translocation of MRTF-A, 232

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Fig. 5. Western blotting analysis of EMT markers for osteosarcoma cells under different stiffness. (A) Expression of EMT protein in four groups. Values were normalized to GAPDH. (B, C, D and E) Histogram of relative protein expression. Data were presented of three independent experiments. * p < 0.05, * * p < 0.01, * ** p < 0.001 and * ** * p < 0.0001.

freshly prepared hydrogel, we showed that our preparation protocol was highly feasible and reproducible to yield hydrogels with consistent mechanical properties as in literature (Tse and Engler, 2010). Studies have shown that different proteins possess various biological effects on cells. Although we adopted type I collagen throughout the study, we found that osteosarcoma cells coated with type I collagen and laminin showed different cell morphology (higher circularity and more cluster) in the soft conditions (Supporting Information Fig. 3) (Choi et al., 2013; Rehfeldt et al., 2007). The most common subtype of osteosarcoma (approximately 60%) is an osteoblast subtype characterized by an osteoid matrix in which lesions are mainly composed of a large number of extracellular matrix mineralization components secreted by transformed osteoblasts. (Ragland et al., 2002). Studies have demonstrated that cell culture within microenvironment with mineralized materials can promote phenotypic change of osteosarcoma cells (Rubio et al., 2014). In addition, TGFβ1-induced EMT process of mammary epithelial cells could also be promoted when cultured on rigid substrates mimicking breast cancer (O'Connor et al., 2015). Similar results have also been reported in pancreatic and prostate cancer studies (Haage and Schneider, 2014; Prauzner-Bechcicki et al., 2015). These reported clued urged us to examine specific protein markers of osteosarcoma cells under various substrate stiffness. As shown in Fig. 5, we detected that rigid substrate promoted the malignant process of EMT, and elevated MMP9 expression when increasing substrate stiffness, further fortifying invasive nature of these cells. In fact, the dramatic difference between soft group and medium/rigid groups also signified that matrix stiffness might regulate cellular behavior within a certain range of substrate mechanical stiffness.

dystrophies, cardiomyopathies to cancer. (Jaalouk and Lammerding, 2009). In particular, physical properties of extracellular matrix (ECM), particularly stiffness or elasticity, widely involved in various cell behaviors such as adhesion, spreading, migration, proliferation, differentiation, and apoptosis. Thanks to previous research efforts, we are able to quantitatively describe different levels of stiffness of various tissues and organs in our human bodies, e.g. brain (0.1–1 kPa), pancreas (1.2 kPa), muscle (8–17 kPa) and precalcified bone tissue (25–40 kPa). It was of great interests that these numbers are not identical in the same type of tissue depending upon specific situation of tissue-of-interests (e.g. physiological or pathological states) (Lv et al., 2015). To elaborate on this topic, we adopted polyacrylamide hydrogels (PAAMs) as in vitro biomaterials to create various substrate. First, PAAMs have been widely used as a substrate for culture because of their adjustable stiffness properties by changing chemical equivalents (ratios) between acrylamide and methylene bisacrylamide. To be specific, increasing the ratio of either acrylamide or bis-acrylamide would lead to a higher elastic modulus after hydrogel polymerization (Shih et al., 2011). Previous study demonstrated that human tissue displayed a variety of stiffness to fulfill corresponding physiological needs (Handorf et al., 2015), while PAAMs can be prepared to simulate different stiffness of organ/tissues in human body. Moreover, PAAMs possess advantageous mechanical properties such as restored normal elastic modulus after loading. It is also transparent material with great optical properties, compatible for fluorescence microscopy. Due to the biologically inert nature, PAAMs hydrogel is not conducive to cell adhesion, however, this problem can be simply addressed via surface functionalizing with appropriate proteins before use (Wouters et al., 2016). Using Instron Universal Testing Machine for elastic modulus measurement of

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Fig. 6. Exposure of osteosarcoma cells to CCG 203971 and DMSO on rigid hydrogel's substrate. (A) Cell's morphology under light microscopy. Scale Bar= 50 µm. (B) Calcein AM staining revealed the outline of each cell. Scale Bar= 50 µm. (C) Circularity of each group was calculated (mean ± S.D.). (D) Fluorescence images exhibited differences in MRTF-A localization under treatment. Scale Bar= 50 µm. (E) Western blotting analysis of MRTF-A. Values were normalized to GAPDH and laminB1. (F and G) Histogram of relative protein expression in total protein and nucleic protein, respectively. Data were presented of three independent experiments. *p < 0.05, * *p < 0.01, * **p < 0.001 and * ** *p < 0.0001.

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Fig. 7. Western blotting analysis of EMT markers for osteosarcoma cells under CCG 203971 treatment. (A) Expression of EMT protein in two groups. Values were normalized to GAPDH. (B, C, D and E) Histogram of relative protein expression. Data were presented of three independent experiments. *p < 0.05, * *p < 0.01, * **p < 0.001 and * ** *p < 0.0001.

cytoskeleton were reorganized accompanied by cell spreading and elongation (Fig. 3). Meanwhile the EMT-related processes were initiated, which reduced the expression of epithelial markers and increased the expression of mesenchymal markers (Fig. 5). Studies confirm the identification of MRTFs as master regulators of EMT (Gasparics and Sebe, 2018). This is consistent with our observations above. Our study was further validated using CCG 203971. We found that the expression levels of the mesenchymal markers vimentin, fibronectin, transcription factor snail, and invasion index MMP 9 were significantly reduced in CCG 203971 treated groups (Figs. 7 and 8E), revealing important role of MRTF-A during EMT. It was also noteworthy that the epithelial marker E-cadherin did not increase when MRTF-A was blocked. This might be due to the loss of cell-to-cell contact, which had an effect on the expression of E-cadherin. Even if the nuclear localization of MRTF was blocked, it could not form an effective connection between cells. Therefore, it is necessary that future effort to illustrate underlying mechanisms controlling E-cadherin expression. Fibronectin (FN) is an extracellular protein that known to modulate cell-matrix interactions, which may affect molecular alteration of osteosarcoma responsible for its aggressive clinical behavior (Na et al., 2012). On one hand, cells are constantly synthesizing, breaking down, and rearranging ECM components to alter ECM composition. On the other hand, ECM could in turn regulate cell behavior. Imbalance in the synthesis and metabolism of ECM can affect matrix stiffness through deposits of large amounts of protein. In our study, we found fibronectin more expression with increasing ECM stiffness. Therefore, we might speculated that increasing ECM stiffness triggers integrin aggregation and activation of extracellular signal-regulated kinase (ERK), resulting

Cell morphological changes also reflects dynamic remodeling process of cytoskeleton. In other words, as stiffness of ECM increases, number of integrins linked to matrix-specific ligand changes, and assembly of receptor molecules such as focal adhesion kinase (FAK), vinculin, and paxillin can be initiated to assist cell-to-matrix interactions. Focal adhesion (FA), a large protein complex, accounts for an anchoring site of cytoskeleton and corresponding mechanical signal transduction. Under rigid matrix conditions, cells are linked to extracellular via integrin-based adhesions. Actin filaments can then organize into stress fibers to mediate cytoskeletal tension, morphology, and downstream events. On soft substrates, such reaction might be inhibited (Steward and Kelly, 2015; Zaidel-Bar et al., 2007) due to the blocking of integrin-ECM binding. Specifically, with the recruitment and phosphorylation of FA proteins, activated Rho GTPases can promote assembly of actin filaments and formation of actin stress fibers through Rho-associated kinase (ROCK) and mammalian Diaphanous-related formins (mDia), allowing the monomeric G-actin to polymerize into Factin. Through dynamic recombination of F-actin (Parreno et al., 2014; Posern and Treisman, 2006), cytoskeleton can activate the SRF/MRTFA axis to regulate gene expression. To be specific, MRTF-A, originally formed a reversible complex with G-actin, could also dissociate from the complex and translocate from cytoplasm to nucleus to participate in transcriptional activation upon was association with SRF (Huveneers and Danen, 2009; Young and Copeland, 2010) (Fig. 9). We clearly detected nuclear translocation process of MRTF-A, which was control by matrix stiffness (Fig. 3D and Fig. 4). Consistent with literature findings, there was no formation of F-actin be observed when MRTF-A was present in cytoplasm, while MRTF-A translocated into the nucleus, 235

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Fig. 8. Wound healing assay was performed with culture-insert. (A and C) Photomicrograph of initial linear defect compared with the same gap at end of study (24 H) for each experimental group. A showed the phase contrast images of cells on glass slides(control) and hydrogel's substrate(soft, medium and rigid). C showed the phase contrast images of cells on rigid substrate. (B and D) Index of migration under different conditions. (E) Protein expression of MMP9. Values were normalized to GAPDH. (F and G) Histogram of relative protein expression. All data were presented of three independent experiments. *p < 0.05, * *p < 0.01, * **p < 0.001 and * ** *p < 0.0001.

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Fig. 9. Graphical summary of the EMT regulatory mechanism of matrix stiffness in osteosarcoma cells.

Appendix A. Supplementary material

in elevated Rho-mediated cell contractility, and that continual increase in cell tension will increase ECM protein by affecting downstream signaling molecules. Secreted ECM further increases stiffness, resulting in a positive feedback loop. However, questions regarding which specific ECM proteins play a dominant role in matrix stiffness and how to accurately target this to intervention this feedback process remain to be studied. In conclusion, our study manifested that extracellular matrix stiffness plays a significant role in the EMT process by affecting reorganization of cytoskeleton and subcellular localization of MRTF-A in osteosarcoma cells. These data would be highly valuable for comprehensive understanding of osteosarcoma progression, discovering targeted interventions to conquer mechanosensing of tumor cells, and ultimately effective cancer therapy.

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.jmbbm.2018.10.012. References Affo, S., Yu, L.X., Schwabe, R.F., 2017. The role of cancer-associated fibroblasts and fibrosis in liver cancer. Annu. Rev. Pathol. 12, 153–186. Bhowmick, N.A., Ghiassi, M., Bakin, A., Aakre, M., Lundquist, C.A., Engel, M.E., Arteaga, C.L., Moses, H.L., 2001. Transforming growth factor-beta1 mediates epithelial to mesenchymal transdifferentiation through a RhoA-dependent mechanism. Mol. Biol. Cell 12, 27–36. Billiau, A., Edy, V.G., Heremans, H., Van Damme, J., Desmyter, J., Georgiades, J.A., De Somer, P., 1977. Human interferon: mass production in a newly established cell line, MG-63. Antimicrob. Agents Chemother. 12, 11–15. Brown, H.K., Tellez-Gabriel, M., Heymann, D., 2017. Cancer stem cells in osteosarcoma. Cancer Lett. 386, 189–195. Calvo, F., Sahai, E., 2011. Cell communication networks in cancer invasion. Curr. Opin. Cell Biol. 23, 621–629. Chen, W.L., Likhitpanichkul, M., Ho, A., Simmons, C.A., 2010. Integration of statistical modeling and high-content microscopy to systematically investigate cell-substrate interactions. Biomaterials 31, 2489–2497. Choi, J.S., Kim, E.Y., Kim, M.J., Giegengack, M., Khan, F.A., Khang, G., Soker, S., 2013. In vitro evaluation of the interactions between human corneal endothelial cells and extracellular matrix proteins. Biomed. Mater. 8, 014108. Connelly, J.T., Gautrot, J.E., Trappmann, B., Tan, D.W., Donati, G., Huck, W.T., Watt, F.M., 2010. Actin and serum response factor transduce physical cues from the microenvironment to regulate epidermal stem cell fate decisions. Nat. Cell Biol. 12,

Acknowledgement None.

Conflict of interest The authors declare that they have no conflict of interest. 237

Journal of the Mechanical Behavior of Biomedical Materials 90 (2019) 226–238

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Influence of crosslinking on the stiffness and degradation of dermis-derived hydrogels. J. Biomed. Mater. Res. Part A 101, 2883–2895. Posern, G., Treisman, R., 2006. Actin' together: serum response factor, its cofactors and the link to signal transduction. Trends Cell Biol. 16, 588–596. Prauzner-Bechcicki, S., Raczkowska, J., Madej, E., Pabijan, J., Lukes, J., Sepitka, J., Rysz, J., Awsiuk, K., Bernasik, A., Budkowski, A., Lekka, M., 2015. PDMS substrate stiffness affects the morphology and growth profiles of cancerous prostate and melanoma cells. J. Mech. Behav. Biomed. Mater. 41, 13–22. Ragland, B.D., Bell, W.C., Lopez, R.R., Siegal, G.P., 2002. Cytogenetics and molecular biology of osteosarcoma. Lab. Investig.; a J. Tech. Methods Pathol. 82, 365–373. Rammensee, S., Kang, M.S., Georgiou, K., Kumar, S., Schaffer, D.V., 2017. Dynamics of mechanosensitive neural stem cell differentiation. Stem Cells 35, 497–506. Reed, D.R., Hayashi, M., Wagner, L., Binitie, O., Steppan, D.A., Brohl, A.S., Shinohara, E.T., Bridge, J.A., Loeb, D.M., Borinstein, S.C., Isakoff, M.S., 2017. Treatment pathway of bone sarcoma in children, adolescents, and young adults. Cancer 123, 2206–2218. Rehfeldt, F., Engler, A.J., Eckhardt, A., Ahmed, F., Discher, D.E., 2007. Cell responses to the mechanochemical microenvironment–implications for regenerative medicine and drug delivery. Adv. Drug Deliv. Rev. 59, 1329–1339. Rubio, R., Abarrategi, A., Garcia-Castro, J., Martinez-Cruzado, L., Suarez, C., Tornin, J., Santos, L., Astudillo, A., Colmenero, I., Mulero, F., Rosu-Myles, M., Menendez, P., Rodriguez, R., 2014. Bone environment is essential for osteosarcoma development from transformed mesenchymal stem cells. Stem Cells 32, 1136–1148. Saneyasu, T., Akhtar, R., Sakai, T., 2016. Molecular Cues Guiding Matrix Stiffness in Liver Fibrosis. BioMed. Res. Int. 2016, 2646212. Sanz-Ramos, P., Mora, G., Vicente-Pascual, M., Ochoa, I., Alcaine, C., Moreno, R., Doblare, M., Izal-Azcarate, I., 2013. Response of sheep chondrocytes to changes in substrate stiffness from 2 to 20 Pa: effect of cell passaging. Connect. Tissue Res. 54, 159–166. Shih, Y.R., Tseng, K.F., Lai, H.Y., Lin, C.H., Lee, O.K., 2011. Matrix stiffness regulation of integrin-mediated mechanotransduction during osteogenic differentiation of human mesenchymal stem cells. J. Bone Mineral. Res. : Off. J. Am. Soc. Bone Mineral. Res. 26, 730–738. Sisson, T.H., Ajayi, I.O., Subbotina, N., Dodi, A.E., Rodansky, E.S., Chibucos, L.N., Kim, K.K., Keshamouni, V.G., White, E.S., Zhou, Y., Higgins, P.D., Larsen, S.D., Neubig, R.R., Horowitz, J.C., 2015. Inhibition of myocardin-related transcription factor/ serum response factor signaling decreases lung fibrosis and promotes mesenchymal cell apoptosis. Am. J. Pathol. 185, 969–986. Steward, A.J., Kelly, D.J., 2015. Mechanical regulation of mesenchymal stem cell differentiation. J. Anat. 227, 717–731. Torre, L.A., Bray, F., Siegel, R.L., Ferlay, J., Lortet-Tieulent, J., Jemal, A., 2015. Global cancer statistics, 2012. CA: a Cancer J. Clin. 65, 87–108. Trappmann, B., Gautrot, J.E., Connelly, J.T., Strange, D.G., Li, Y., Oyen, M.L., Cohen Stuart, M.A., Boehm, H., Li, B., Vogel, V., Spatz, J.P., Watt, F.M., Huck, W.T., 2012. Extracellular-matrix tethering regulates stem-cell fate. Nat. Mater. 11, 642–649. Tse, J.R., Engler, A.J., 2010. Preparation of hydrogel substrates with tunable mechanical properties. Current protocols in cell biology Chapter 10, Unit 10 16. Valery, P.C., Laversanne, M., Bray, F., 2015. Bone cancer incidence by morphological subtype: a global assessment. Cancer causes Control : CCC 26, 1127–1139. Wang, Y., Gong, T., Zhang, Z.R., Fu, Y., 2017. Matrix stiffness differentially regulates cellular uptake behavior of nanoparticles in two breast cancer cell lines. ACS Appl. Mater. Interfaces 9, 25915–25928. Wei, B., Zhou, X., Liang, C., Zheng, X., Lei, P., Fang, J., Han, X., Wang, L., Qi, C., Wei, H., 2017. Human colorectal cancer progression correlates with LOX-induced ECM stiffening. Int. J. Biol. Sci. 13, 1450–1457. Wouters, O.Y., Ploeger, D.T., van Putten, S.M., Bank, R.A., 2016. 3,4-Dihydroxy-L-phenylalanine as a novel Covalent linker of extracellular matrix proteins to polyacrylamide hydrogels with a Tunable stiffness. tissue engineering. Part C., Methods. Wu, C.L., Tsai, H.C., Chen, Z.W., Wu, C.M., Li, T.M., Fong, Y.C., Tang, C.H., 2013. Ras activation mediates WISP-1-induced increases in cell motility and matrix metalloproteinase expression in human osteosarcoma. Cell. Signal. 25, 2812–2822. Xue, C., Zhang, T., Xie, X., Zhang, Q., Zhang, S., Zhu, B., Lin, Y., Cai, X., 2017. Substrate stiffness regulates arterial-venous differentiation of endothelial progenitor cells via the Ras/Mek pathway. Biochim. Et. Biophys. Acta 1864, 1799–1808. Yang, Y., Han, L., He, Z., Li, X., Yang, S., Yang, J., Zhang, Y., Li, D., Yang, Y., Yang, Z., 2018. Advances in limb salvage treatment of osteosarcoma. J. Bone Oncol. 10, 36–40. Young, K.G., Copeland, J.W., 2010. Formins in cell signaling. Biochim. Et. Biophys. Acta 1803, 183–190. Zaidel-Bar, R., Itzkovitz, S., Ma'ayan, A., Iyengar, R., Geiger, B., 2007. Functional atlas of the integrin adhesome. Nat. Cell Biol. 9, 858–867.

711–718. Czekanska, E.M., Stoddart, M.J., Richards, R.G., Hayes, J.S., 2012. In search of an osteoblast cell model for in vitro research. Eur. Cells Mater. 24, 1–17. De Craene, B., Berx, G., 2013. Regulatory networks defining EMT during cancer initiation and progression. Nat. Rev. Cancer 13, 97–110. Decker, S., Winkelmann, W., Nies, B., van Valen, F., 1999. Cytotoxic effect of methotrexate and its solvent on osteosarcoma cells in vitro. J. Bone Jt. Surg. Br. Vol. 81, 545–551. DuFort, C.C., Paszek, M.J., Weaver, V.M., 2011. Balancing forces: architectural control of mechanotransduction. Nat. Rev. Mol. Cell Biol. 12, 308–319. Dupont, S., 2016. Role of YAP/TAZ in cell-matrix adhesion-mediated signalling and mechanotransduction. Exp. Cell Res. 343, 42–53. Gasparics, A., Sebe, A., 2018. MRTFs- master regulators of EMT. Developmental dynamics : an official publication of the American Association of Anatomists 247, pp. 396-404. Haage, A., Schneider, I.C., 2014. Cellular contractility and extracellular matrix stiffness regulate matrix metalloproteinase activity in pancreatic cancer cells. FASEB J. : Off. Publ. Fed. Am. Soc. Exp. Biol. 28, 3589–3599. Handorf, A.M., Zhou, Y., Halanski, M.A., Li, W.J., 2015. Tissue stiffness dictates development, homeostasis, and disease progression. Organogenesis 11, 1–15. Harrison, D.J., Geller, D.S., Gill, J.D., Lewis, V.O., Gorlick, R., 2018. Current and future therapeutic approaches for osteosarcoma. Expert Rev. Anticancer Ther. 18, 39–50. Huveneers, S., Danen, E.H., 2009. Adhesion signaling - crosstalk between integrins, Src and Rho. J. Cell Sci. 122, 1059–1069. Ingber, D.E., 2006. Cellular mechanotransduction: putting all the pieces together again. FASEB J. : Off. Publ. Fed. Am. Soc. Exp. Biol. 20, 811–827. Jaalouk, D.E., Lammerding, J., 2009. Mechanotransduction gone awry. Nat. Rev. Mol. Cell Biol. 10, 63–73. Joyce, J.A., Pollard, J.W., 2009. Microenvironmental regulation of metastasis. Nat. Rev. Cancer 9, 239–252. Kalita, K., Kuzniewska, B., Kaczmarek, L., 2012. MKLs: co-factors of serum response factor (SRF) in neuronal responses. Int. J. Biochem. Cell Biol. 44, 1444–1447. Katara, G.K., Kulshrestha, A., Mao, L., Wang, X., Sahoo, M., Ibrahim, S., Pamarthy, S., Suzue, K., Shekhawat, G.S., Gilman-Sachs, A., Beaman, K.D., 2018. Mammary epithelium-specific inactivation of V-ATPase reduces stiffness of extracellular matrix and enhances metastasis of breast cancer. Mol. Oncol. 12, 208–223. Ko, Y.A., Jamaluddin, M.F.B., Adebayo, M., Bajwa, P., Scott, R.J., Dharmarajan, A.M., Nahar, P., Tanwar, P.S., 2018. Extracellular matrix (ECM) activates beta-catenin signaling in uterine fibroids. Reproduction 155, 61–71. Lampi, M.C., Reinhart-King, C.A., 2018. Targeting extracellular matrix stiffness to attenuate disease: from molecular mechanisms to clinical trials. Sci. Transl. Med. 10. Lantoine, J., Grevesse, T., Villers, A., Delhaye, G., Mestdagh, C., Versaevel, M., Mohammed, D., Bruyere, C., Alaimo, L., Lacour, S.P., Ris, L., Gabriele, S., 2016. Matrix stiffness modulates formation and activity of neuronal networks of controlled architectures. Biomaterials 89, 14–24. Li, L., Su, N., Zhou, T., Zheng, D., Wang, Z., Chen, H., Yuan, S., Li, W., 2018. Mixed lineage kinase ZAK promotes epithelial-mesenchymal transition in cancer progression. Cell death Dis. 9, 143. Lopez-Novoa, J.M., Nieto, M.A., 2009. Inflammation and EMT: an alliance towards organ fibrosis and cancer progression. EMBO Mol. Med. 1, 303–314. Lv, H., Li, L., Sun, M., Zhang, Y., Chen, L., Rong, Y., Li, Y., 2015. Mechanism of regulation of stem cell differentiation by matrix stiffness. Stem Cell Res. Ther. 6, 103. Miralles, F., Posern, G., Zaromytidou, A.I., Treisman, R., 2003. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113, 329–342. Miyazawa, A., Ito, S., Asano, S., Tanaka, I., Sato, M., Kondo, M., Hasegawa, Y., 2018. Regulation of PD-L1 expression by matrix stiffness in lung cancer cells. Biochem. Biophys. Res. Commun. 495, 2344–2349. Morita, T., Mayanagi, T., Sobue, K., 2007. Dual roles of myocardin-related transcription factors in epithelial mesenchymal transition via slug induction and actin remodeling. J. Cell Biol. 179, 1027–1042. Muller, C., Pompe, T., 2016. Distinct impacts of substrate elasticity and ligand affinity on traction force evolution. Soft Matter 12, 272–280. Na, K.Y., Bacchini, P., Bertoni, F., Kim, Y.W., Park, Y.K., 2012. Syndecan-4 and fibronectin in osteosarcoma. Pathology 44, 325–330. O'Connor, J.W., Riley, P.N., Nalluri, S.M., Ashar, P.K., Gomez, E.W., 2015. Matrix Rigidity Mediates TGFbeta1-Induced Epithelial-Myofibroblast Transition by Controlling Cytoskeletal Organization and MRTF-A Localization. J. Cell. Physiol. 230, 1829–1839. Parreno, J., Raju, S., Niaki, M.N., Andrejevic, K., Jiang, A., Delve, E., Kandel, R., 2014. Expression of type I collagen and tenascin C is regulated by actin polymerization through MRTF in dedifferentiated chondrocytes. FEBS Lett. 588, 3677–3684. Pilipchuk, S.P., Vaicik, M.K., Larson, J.C., Gazyakan, E., Cheng, M.H., Brey, E.M., 2013.

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