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. 2018 Apr 16;233(9):7320–7332. doi: 10.1002/jcp.26563

Tyrosine kinase receptor c‐ros‐oncogene 1 inhibition alleviates aberrant bone formation of TWIST‐1 haploinsufficient calvarial cells from Saethre–Chotzen syndrome patients

Esther Camp 1,2, Peter J Anderson 2,3, Andrew C W Zannettino 2,4, Carlotta A Glackin 5, Stan Gronthos 1,2,✉
PMCID: PMC13482113  PMID: 29663378

Abstract

Saethre–Chotzen syndrome (SCS), associated with TWIST‐1 mutations, is characterized by premature fusion of cranial sutures. TWIST‐1 haploinsufficiency, leads to alterations in suture mesenchyme cellular gene expression patterns, resulting in aberrant osteogenesis and craniosynostosis. We analyzed the expression of the TWIST‐1 target, Tyrosine kinase receptor c‐ros‐oncogene 1 (C‐ROS‐1) in TWIST‐1 haploinsufficient calvarial cells derived from SCS patients and calvaria of Twist‐1del/+ mutant mice and found it to be highly expressed when compared to TWIST‐1 wild‐type controls. Knock‐down of C‐ROS‐1 expression in TWIST‐1 haploinsufficient calvarial cells derived from SCS patients was associated with decreased capacity for osteogenic differentiation in vitro. Furthermore, treatment of human SCS calvarial cells with the tyrosine kinase chemical inhibitor, Crizotinib, resulted in reduced C‐ROS‐1 activity and the osteogenic potential of human SCS calvarial cells with minor effects on cell viability or proliferation. Cultured human SCS calvarial cells treated with Crizotinib exhibited a dose‐dependent decrease in alkaline phosphatase activity and mineral deposition, with an associated decrease in expression levels of Runt‐related transcription factor 2 and OSTEOPONTIN, with reduced PI3K/Akt signalling in vitro. Furthermore, Crizotinib treatment resulted in reduced BMP‐2 mediated bone formation potential of whole Twist‐1del/+ mutant mouse calvaria organotypic cultures. Collectively, these results suggest that C‐ROS‐1 promotes osteogenic differentiation of TWIST‐1 haploinsufficient calvarial osteogenic progenitor cells. Furthermore, the aberrant osteogenic potential of these cells is inhibited by the reduction of C‐ROS‐1. Therefore, targeting C‐ROS‐1 with a pharmacological agent, such as Crizotinib, may serve as a novel therapeutic strategy to alleviate craniosynostosis associated with aberrant TWIST‐1 function.

Keywords: C‐ROS‐1, calvarial cells, osteoblasts, Saethre–Chotzen, TWIST‐1, Twist‐1 del/+− heterozygous mice

C‐ROS‐1 is a downstream mediator of aberrant bone formation in TWIST‐1 haploinsufficient cranial bone cells. Pharmacological targeting of C‐ROS‐1 activity could help control cranial pre‐fusion in Saethre–Chotzen syndrome patients with TWIST‐1 mutations.

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1. INTRODUCTION

Craniosynostosis is a medical condition affecting infants, which describes partial or complete premature fusion of the sutures. These are the fibrous connections between the bones of the calvaria which bridge the two osteogenic fronts (Wilkie, 1997). Sutures normally remain patent by maintaining active bone growth without fusion in young children, allowing the skull to expand to accommodate the growing brain (Opperman, 2000; Rice & Rice, 2008). Plagiocephaly, ocular hypertelorism, proptosis, beaking of the nose and midface hypoplasia are the common cranio‐facial features of craniosynostosis. It can also lead to an increase in intracranial pressure that can result in optic atrophy, blindness and developmental delay (Kimonis, Hoffman, Panchal, & Boyadjiev, 2007). In addition, many neurological studies show that children with just single suture craniosynostosis have an increased risk of developing cognitive, language, and motor skill difficulties during infancy and childhood (Knight, Anderson, Spencer‐Smith, & Da Costa, 2014). Craniosynostosis can occur as an isolated event resulting in nonsyndromic craniosynostosis, or it can occur due to a genetic anomaly in conjunction with other anomalies in well‐defined patterns that make up clinically recognized syndromes. Saethre–Chotzen syndrome (SCS) is the second most prevalent syndromic craniosynostosis occurring in approximately 1/25,000–1/50,000 births. It usually involves unilateral or bilateral coronal synostosis and mild limb deformities (brachydactyly and cutaneous syndactyly) (Anderson, Hall, Evans, Hayward, & Jones, 1996; Anderson et al., 1997). It is autosomal‐dominantly inherited and is induced by loss‐of‐function mutations of TWIST‐1 (Howard et al., 1997). There are approximately 100 different heterozygous mutations of TWIST‐1, all of which result in TWIST‐1 haploinsufficiency and varying phenotypes of SCS (el Ghouzzi et al., 1997; Elanko et al., 2001; Howard et al., 1997; Johnson et al., 1998). The clinical phenotypes of SCS vary profoundly, ranging from isolated unicoronal craniosynostosis to the extreme manifestation of multiple suture involvement.

The TWIST‐1 gene encodes a basic helix‐loop‐helix (bHLH) transcription factor that mediates skeletal and head mesodermal tissue development (Chen & Behringer, 1995). Mouse, Twist‐1 homozygous mutant embryos die before birth, while, Twist‐1del/+ heterozygous embryos survive post birth and have craniofacial and limb abnormalities similar to the characteristics observed in SCS patients, including premature fusion of the coronal suture (Behr, Longaker, & Quarto, 2011; Carver, Oram, & Gridley, 2002; Chen & Behringer, 1995; Yoshida et al., 2005). Post embryonic development, TWIST‐1 is expressed in number of adult mesenchymal stem cell (MSC)‐like populations including bone marrow derived stromal cells (BMSC), cranial mesenchyme cells within the cranial sutures during calvarial and suture development (Johnson, Wilkie, & Morriss‐Kay, 2000; Menicanin, Bartold, Zannettino, & Gronthos, 2010; Rice et al., 2000, 2003; Yoshida et al., 2005) and primary osteoblastic cells derived from newborn mouse calvariae (Murray, Winters, Gazit, Kahn, & Murray, 1992). We have previously demonstrated that calvarial cells isolated from SCS patients have reduced TWIST‐1 expression compared to cells from healthy aged matched controls and exhibit an increased capacity to form mineralized nodules in vitro under osteogenic conditions (Cakouros et al., 2012). Moreover, TWIST‐1 haploinsufficient SCS calvarial cells exhibit alterations in gene expression patterns, leading to decreased osteogenic progenitor proliferation and increased osteogenesis (Yousfi, Lasmoles, & Marie, 2002; Yousfi, Lasmoles, Lomri, Delannoy, & Marie, 2001). The cause of the premature bony bridging between apposed skull plates in SCS patients results from the loss of TWIST‐1, leading to an increase in the osteogenic potential of osteoprogenitor calvarial cells and accelerated bone growth (Yousfi et al., 2001, 2002; Guenou, Kaabeche, Dufour, Miraoui, & Marie, 2006). However, the molecular factors, which regulate suture fusion and the process of suture fusion in humans is still not fully understood.

A recent study has identified Tyrosine kinase receptor c‐ros‐oncogene 1 (C‐ROS‐1) to be a novel TWIST‐1 target (Camp, Anderson, Zannettino, & Gronthos, 2017). C‐ROS‐1 belongs to the sevenless subfamily of tyrosine kinase insulin receptors (Chen, Heller, Poon, Kang, & Wang, 1991) and encodes for a tyrosine kinase receptor, which consists of a glycoprotein‐rich extracellular domain, a transmembrane domain, and an intracellular tyrosine kinase (PT kinase) domain (Acquaviva, Wong, & Charest, 2009). C‐ROS‐1 is temporally and spatially expressed in epithelial cells of the developing and adult lungs, kidneys, heart, intestine, and epididymis (Sonnenberg‐Riethmacher, Riethmacher, Gödecke, & Birchmeier, 1996; Sonnenberg, Walter, Bladt, & Birchmeier, 1991; Tessarollo et al., 1992), in BMSC and in pre‐osteoblast calvarial cells (Camp et al., 2017). Expression of C‐ROS‐1 increases in BMSC during osteogenic differentiation; however, it is down‐regulated in TWIST‐1 over‐expressing cells (Camp et al., 2017), which have a reduced osteogenic differentiation potential in vitro and in vivo (Isenmann et al., 2009). Furthermore, chromatin immunoprecipitation analysis demonstrated that TWIST‐1 directly binds to two E‐box motif regulatory regions on the C‐ROS‐1 promoter during osteogenic differentiation (Camp et al., 2017). Knock‐down of C‐ROS‐1 in BMSC and pre‐osteoblast calvarial cells resulted in a decreased capacity for osteogenic differentiation in vitro (Camp et al., 2017). In the present study, we examined the expression profile of C‐ROS‐1 in TWIST‐1 haploinsufficient calvarial cells and examined whether C‐ROS‐1 inhibition had an effect on the osteogenic potential of these cells. We identified that C‐ROS‐1 is elevated in TWIST‐1 haploinsufficient calvarial cells compared to TWIST‐1 wild‐type cells, during osteogenic differentiation and within the coronal suture. Furthermore, inhibition of C‐ROS‐1 expression and C‐ROS‐1 activity reduced the osteogenic potential of TWIST‐1 haploinsuffient calvarial cells in vitro and reduced bone formation in calvaria organotypic cultures.

2. MATERIALS AND METHODS

2.1. Isolation of human cranial cells

Human parietal bone calvarial cells were derived from the cranium of donors with informed consent by the Human Ethics Committee of the Women's and Children's Hospital, South Australia (Approval# REC1033/06/2019). Cells were cultured as previously described (Isenmann et al., 2009).

2.2. Osteogenic differentiation and osteogenesis assays

Cells were cultured in osteogenic inductive media (αMEM supplemented with 5% fetal calf serum, 2 mM l‐glutamine, 1 mM sodium pyruvate, 10 mM HEPES buffer, 50 U/mL penicillin, 50 μg/ml streptomycin, 1 × 10−7 M dexamethasone, 100 μM l‐ascorbate‐2‐phosphate, and 2.6 mM KH2PO4 (Sigma–Aldrich, Inc., Sydney, NSW, Australia) for 7 days, 2 weeks, or 3 weeks. Media was changed twice a week. Alkaline phosphatase (ALK) staining was performed using Phosphatase Alkaline kit (Cat#86R‐1KT, Sigma–Aldrich Inc) following manufacturer's instructions. Mineralized bone matrix formation was identified with Alizarin red S staining (Cat#A5533, Sigma–Aldrich, Inc.) of mineral deposits. Extracellular matrix‐bound calcium was measured in triplicate samples and normalized to DNA content per well as previously described (Isenmann et al., 2009).

2.3. siRNA knock‐down and Crizotinib

Cells were seeded at 2.2 × 104 cells per well (24‐well plate) the day before transfection of siRNAs. Sequence‐specific siRNA against C‐ROS‐1 (s12113) or negative siRNA#1 control (Ambion/Life Technologies, Mulgrave, VIC, Australia) were used at 15 pmol with Lipofectamine™ RNAiMAX Transfection Reagent (Cat#13778150, Thermo Fisher Scientific) to achieve a 70–80% knockdown of transcript levels as previously described (Camp et al., 2017). The procedure was performed following manufacturer's instructions with a 72 hr incubation period before adding osteogenic differentiation media. Crizotinib (PF‐2341066, Xalkori, Pfizer, Inc. New York, NY) was reconstituted in dimethyl sulfoxide (DMSO) and stored at −80 °C. Cells were treated with Crizotinib at 50–2 μM or DMSO (0.1%).

2.4. RNA extractions, cDNA synthesis, and real time polymerase chain reaction analysis

The anterior segment of the calvaria of day (P) 10 wild‐type and Twist‐1del/+ mice were retrieved and snap frozen in liquid nitrogen. Total RNA from cultured human calvarial cells and mouse calvaria fragments was isolated using TRIzol Reagent (Cat#15596026, Invitrogen/Thermo Fisher Scientific. Waltham, MA) according to the manufacturer's instructions. Generation of cDNA and real‐time analysis was performed as previously described in triplicate (Camp et al., 2017). Primer sets used in this study (GeneWorks Pty Ltd, Thebarton, SA, Australia) were: human β‐actin (Accession#: M28424.1 Fwd:5′‐gatcattgctcctcctgagc‐3′; Rev:5′‐gtcatagtccgcctagaagcat‐3′), Runx2 (Accession#:NM_001015051.3 Fwd:5′‐gtggacgaggcaagagtttca‐3′; Rev:5′‐catcaagcttctgtctgtgcc‐3′), OPN (Accession#:J04765.1 Fwd:5′‐ acatccagtaccctgatgctacag‐3′; Rev:5′‐gtgggtttcagcactctggt‐3′), human C‐ROS‐1 (Accession#:NM_002944.2 Fwd:5′‐atgcagacctaccaactgctc‐3′; Rev:5′‐gggcttgaccacataggacg‐3′), Twist‐1 (Accession#:Y10871.1 Fwd;5′‐tcttacgaggagctgcagacgca‐3′; Rev:5′‐atcttggagtccagctcgtcgct‐3′), mouse β‐actin (Accession#:AY618569.1 Fwd:5′‐ttgctgacaggatgcagaag‐3′; Rev:5′‐aagggtgtaaaacgcagctc‐3′), mouse C‐ROS‐1 (Accession #NM_011282.2 Fwd:5′‐caagtvaaggcctccaggag‐3′; Rev:5′‐gggtgggtaattccatgcca‐3′).

2.5. Cell growth

Cells were seeded into 96‐well plates (2.5 × 103) in the presence of Crizotinib (500, 1, or 2 μM). Proliferating cells were assessed at the indicated time points using Cell proliferation Elisa, bromodeoxyuridine (BrdU) colometric kit (Roche Products Pty Limited. Sydney NSW Australia) as per manufacturer's instructions. Absorbance was read at 450 nM on an iMark microplate absorbance reader (Bio‐Rad Laboratories, Hercules, CA).

2.6. Cell viability assay

Cells were cultured in the presence of DMSO (0.1%) or Crizotinib (500, 1, or 2 mM) for 7 days. Cells then were stained with annexin V–488 (Cat#:A13202, Invitrogen/Thermo Fisher Scientific) and 7‐aminoactinomycin (7‐AAD, Cat#A1310, Invitrogen/Thermo Fisher Scientific) as per the manufacturers’ instructions and analyzed immediately by flow cytometry (BD LSRFortessa X20 Analyser, Becton Dickinson Biosciences, Franklin Lakes, NJ). Cells were incubated for 10 min in either DMSO (100%) or ethanol (80%) and were used as positive controls.

2.7. Calvaria organ cultures

Calvaria bone organ cultures established from 4‐day‐old Twist‐1del/+ mice were cultured in the presence of rhBMP‐2 (50 ng/mL, Cat#:PHC7145, Thermo Fisher Scientific) and vehicle control (0.1% DMSO), rhBMP‐2 and Crizotinib at 500 nM or rhBMP‐2 and Crizotinib at 1 mM for 10 days as previously described (Garrett, 2003; Mohammad, Chirgwin, & Guise, 2008). Calvaria were then fixed in 10% formalin for 6 hr, decalcified overnight (14% EDTA, pH 7.2), and embedded in paraffin. Sections (5 mm) were cut and stained with hematoxylin and eosin. The area of new bone formed relative to the length of calvaria bone specimen analyzed was quantitated using OsteoMeasure XP Advanced Bone Histomorphometry ver. 3.3.02 (OsteoMetrics, Inc., Decatur, GA) software on an Olympus BX53Microscope (Olympus, Notting Hill, VIC, Australia).

2.8. Micro–computed tomography

Three‐dimensional (3D) microarchitecture of day 10 mouse skulls was evaluated using micro–computed tomography (micro‐CT; SkyScan 1076 X‐ray Micro‐CT; Bruker microCT, Kontich, Belgium). Skulls were scanned at 9 mm resolution, AI 0.5 mm filter, excitation 5890 ms, voltage 48 kV, current 159 μA, rotation step 0.4, and 1‐frame averaging. Reconstruction of the original scan data was performed using NRecon Reconstruction (64‐bit, ver. 1.6.10; Bruker microCT) with a smoothing of 1, ring artifact of 10, and beam hardening of 30%. Reconstructed .bmp files were realigned in Data Viewer (64‐bit, ver.1.5.2; Bruker microCT). A region of interest (ROI) was selected as defined by approximately 166 slices down from the top/surface of the calvaria (at the anterior region of the sphenoid bone) to a defined region of the anterior part of the calvaria where the coronal sutures are located. ROI was subjected to 2D and 3D morphometric parameters calculated by a Comprehensive Tex Archive Network CT‐analyzer (CTAn, ver. 1.15 and CTVol, ver.2.3: 32‐bit version XP, Vista,Win‐7 2D/3D processing, analysis, visualization; Bruker microCT) in 2D or 3D based on a surface rendered volume model in 2D or 3D space.

2.9. Western blot analysis

Whole cell lysates were prepared from cells cultured in 9.5 cm2 wells in triplicate in 300 μl of lysis buffer per well (50 mM Tris/HCL pH 7.4; 1 mM EDTA pH 8.0; 100 mM NaCl; 1% NP‐40; 10 mM NaF; 0.5 mM PMSF; 5 mM sodium vanadate; 10 mM sodium pyrophosphate; 1× complete protease inhibitor) (Roche Applied Science Roche Diagnostics GmbH, Mannheim, Germany). The lysates were incubated on ice for 30 min and centrifuged for 15 min at 16,000g at 4 °C. Protein concentrations were measured using the RCDC protein assay kit (Bio‐Rad Laboratories). Western blot analysis was performed as described (Camp et al., 2017). The membranes were probed overnight at 4 °C with the following primary antibodies diluted in 5% ECL membrane blocking agent (Cat#:RPN2125, GE Healthcare Australia Parramatta, NSW, Australia); anti‐rabbit Runx‐2 (M‐70, Cat#:SC‐10758, 1:500 dilution, RRID:AB_2184247, Santa Cruz Biotechnology, Inc. Dallas, TX) anti‐rabbit SHP90α/β (H‐114, Cat#:SC7947 1:1000 dilution, RRID: AB_2121235, Santa Cruz Biotechnology, Inc.), anti‐rabbit C‐ROS‐1 (Cat#:ab5512, 1:350 dilution, RRID: AB_304948, Abcam, Cambridge, MA), anti‐rabbit phosphorylated C‐ROS‐1 (Tyr2274, Cat#:3078, 1:400 dilution, RRID:AB_2180473, Cell Signaling Technology, Danvers, MA), anti‐mouse beta‐actin (Cat#:8H10D10, 1:1000 dilution, RRID: AB_2242334, Cell Signaling Technology), anti‐rabbit Akt (Cat#:9272S, 1/500 dilution, RRID: AB_329827, Cell Signaling Technology), anti‐mouse phosphorylated Akt‐Ser473 (Cat#:4051S, 1/500 dilution, RRID: AB_331158, Cell Signaling Technology), anti‐rabbit phosphorylated Akt‐Thr308 (Cat#:9275L, 1/500 dilution, RRID: AB_329828, Cell Signaling Technology), anti‐rabbit PI3K kinase (Cat#:3011, 1:1000 dilution, RRID: AB_10831524, Cell Signaling Technology). Blots were washed three times with TBS/0.1% Tween 20, then incubated for 1 hr at room temperature with the corresponding secondary detection antibodies, Secondary detection was performed using anti‐Rabbit‐Alkaline phosphatase conjugate (AP) (Cat#:ab97048, 1:2500 dilution, RRID:AB_10680574, Abcam) and anti‐Mouse‐Alkaline phosphatase conjugate (AP) (Cat#:AP124A, 1:2500 dilution, RRID:AB_92456, Merk Life Science, Bayswater, VIC, Australia) antibodies. After washing off secondary antibody, the blots were developed with enhanced chemifluorescence (ECF) substrate (GE Healthcare UK Ltd., Buckinghamshire, England).

2.10. Immunohistochemical analysis

For immunohistochemical analysis, calvaria were isolated from day 10 wild‐type and Twist‐1del/+ mice, fixed in 10% formalin for 24 hr, decalcified overnight (14% EDTA, pH 7.2), and embedded in paraffin. Transverse sections (5 μm) were cut and deparaffinized in xylene, rehydrated in decreasing concentrations of ethanol. Following this, the sections were subjected to antigen retrieval by heating in citrate buffer (pH 6) for 20 min at 95 °C and then treated with 3% hydrogen peroxide for 20 min at room temperature in order to block endogenous peroxidase activity. The slides were then blocked with 5% normal goat serum (Cat#:AB7481, Abcam) and incubated overnight with an anti‐ROS1 antibody (Cat#:ab5512, 1:500 dilution, RRID: AB_304948, Abcam) at 4 °C. Subsequent to washing in phosphate‐buffered saline (PBS), the sections were incubated with biotinylated secondary antibodies for 40 min (Cat#:BA1000, 1/250 dilution, Vector Laboratories, Burlingame), washed in PBS and then incubated with Steptavidin peroxidase conjugate (1:100 dilution, Thermo Fisher Scientific) for 1 hr. Staining was performed using a DAB kit (3,3‐Diaminbenzidine, Dako/Aligent, Santa Clara, CA) following manufacturer's instructions. The slides were counterstained with Haematoxylin and then dehydrated and mounted. Negative controls, which were established by replacing the primary antibody with rabbit IgG (Cat#:I5006, Sigma–Aldrich), showed no immunoreactivity.

2.11. Statistics

Experiments were performed in triplicates. Data analysis was carried out using Microsoft GraphPad Prism 5 (GraphPad Software, RRID:CR_002798, http://www.graphpad.com/), which was used for the generation of graphs and calculation of statistical significance. Statistical differences (*) of p < 0.05 between samples are shown.

3. RESULTS

3.1. TWIST‐1loss of function mutant calvarial cells derived from patients with SCS exhibit high expression of C‐ROS‐1

Calvarial cells were obtained from four SCS affected patients and three aged matched unaffected control individuals. In our cohort of SCS patients, three donors carried a variant that results in premature truncation or absence of the protein (single point deletions and microdeletion) expressed by the affected TWIST‐1 allele. The fourth donor carried a heterozygous in‐frame duplication of seven amino acids in the affected TWIST‐1 allele. All four SCS patient donors had TWIST‐1 haploinsufficiency and exhibited prematurely fused skull bones along the coronal suture (Figure 1a). In keeping with previous studies, we observed that SCS calvarial cells from all four donors exhibited reduced TWIST‐1 gene expression (Figure 1b) and increased mineral production when cultured under osteogenic inductive conditions, compared to normal aged matched control calvarial cells (Figures 1c and 1d).

Figure 1.

Figure 1

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TWIST‐1 haploinsufficient SCS calvarial cells exhibit an increased osteogenic capacity and express higher levels of C‐ROS‐1 during osteogenic differentiation when compared to TWIST‐1 wild‐type calvarial cells. (a) Five‐month‐old child with Saethre‐Chotzen syndrome (SCS). Frontal CT scan of child with bulging forehead and fused coronal sutures (arrows) on both sides of the skull. The sagittal suture is still unfused (star). (b) Real‐time PCR analysis for TWIST‐1 expression in TWIST‐1 wild‐type (WT) and TWIST‐1 mutant SCS patient calvarial cells. Graph represents mean ± S.E.M TWIST‐1 expression relative to β‐actin (*p < 0.05, student's t‐test, n = 3 WT and n = 4 SCS donors). (c–e) TWIST‐1 WT and TWIST‐1 mutant SCS calvarial cells were cultured in osteogenic inductive media for 3 weeks. (c) Representative images depicting mineral deposits stained with Alizarin red, scale bar = 200μm. (d) Mean ± S.E.M levels of extracellular calcium quantitated using the Calcium Arsenazo III reagent and normalised to total DNA content per well (*p < 0.05, student's t‐test, n = 3 WT, n = 4 SCS). (e) C‐ROS‐1 expression was assessed using quantitative real‐time PCR in TWIST‐1 WT and TWIST‐1 mutant SCS calvarial cells cultured under growth media (CM) or osteogenic inductive media (OI) for 3 weeks. Raw data obtained from CM and OI conditions were normalized relative to β‐actin and expressed as fold change. Graph represents mean (±S.E.M) gene expression as a fold change between CM and OI conditions (*p < 0.05, unpaired student's t‐test, n = 3 WT, n = 4 SCS). (f) Representative Western blot analysis of C‐ROS‐1 protein levels in TWIST‐1 WT and TWIST‐1 mutant SCS cells following osteogenic induction (n = 3 WT, n = 4 SCS). HSP90 protein was used as loading control

To determine the relative gene expression levels of C‐ROS‐1, osteoprogenitor calvarial cells obtained from SCS patients and aged‐matched controls were cultured under osteogenic induction conditions. During osteogenic differentiation C‐ROS‐1 gene expression levels increased in control cells by logFC = 0.6054 (SEM ± 0.45) with significantly higher increases in SCS cells by logFC = 4.918 SEM ± 1.718 (Figure 1e). Confirmatory studies showed that C‐ROS‐1 protein levels were higher in TWIST‐1 haploinsufficient SCS osteoblasts compared to control calvarial osteoblasts as assessed by Western blot analysis (Figure 1f).

3.2. Twist‐1 del/+ heterozygous mice express high levels of C‐Ros‐1 in the coronal suture and have a greater calvaria bone volume

Previous histological studies investigating suture closure in Twist‐1del/+ heterozygous mice have reported that the coronal sutures fuse between postnatal (P) day 9 (P9) and day 15 (P15) (Behr et al., 2011; Carver et al., 2002). Using 3‐dimensional microcomputer tomography (µCT) scanning (Skyscan1174, SAHMRI Bioresources) the cranium of Twist‐1del/+ mutant mice were examined during growth at P10. We observed that 40% of Twist‐1del/+ mutant animals demonstrated some degree of coronal suture fusion (Figure 2a) while no wild‐type animals exhibited coronal suture fusion (data not shown). Moreover, bone volume was greater in this area of the cranium in Twist‐1del/+ heterozygous mice compared to wild type littermates (Figure 2b), regardless of whether premature fusion of the coronal sutures was observed or not. Analysis of C‐ros‐1 gene expression levels in samples isolated from the coronal sutures revealed a significant increase in C‐ros‐1 transcript expression levels in Twist‐1del/+ heterozygous mice, compared to wild‐type controls (Figure 2c). This corresponded to an increase in C‐ROS‐1 protein levels observed in the coronal sutures of Twist‐1del/+ heterozygous mice at P10, when compared to C‐ROS‐1 levels in the coronal sutures of wild‐type mice, as assessed by immunohistochemistry (Figure 2d). These results suggest that the aberrant increase of C‐ROS‐1 within the coronal suture region of Twist‐1del/+ heterozygous mice may play a role in facilitating the craniosynostosis phenotype, at least during calvaria development at P10.

Figure 2.

Figure 2

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Twist‐1del/+ heterozygous mice have premature fusion of the coronal suture by P10 and express high levels of C‐ROS‐1 in the coronal suture. (a) A representative cross‐section CTAn image generated from µCT analysis of the region of interest (ROI) encompassing the coronal suture of P10 Twist‐1del/+ mutant mouse calvaria (i) and a representative 3D CTVol image of the ROI (ii). An open coronal suture on one side of the calvaria is shown (arrow) and fusing/fused coronal suture (asterix) on the opposite side of the calvaria. (b) Graph represents the mean (±S.E.M) bone volume from the ROI analysis performed using CTAn on Twist‐1 wild‐type (WT) and Twist‐1del/+ mutant P10 calvaria (*p < 0.05, student's t‐test, n = 5 mice per genotype). (c) C‐Ros‐ 1 expression was assessed using quantitative real‐time PCR in Twist‐1 WT and Twist‐1del/+ P10 cranial calvarial cells (*p < 0.05, student's t‐test, n = 4). (d) Representative photomicrographs of C‐Ros‐1 immunohistochemistry on Twist‐1 WT and Twist‐1del/+ mutant calvaria sections depicting mesenchyme (arrows) within an unfused coronal suture

3.3. C‐ROS‐1 promotes the osteogenic potential of SCS derived calvarial cells

C‐ROS‐1 has previously been shown to promote calvarial cell osteogenic differentiation (Camp et al., 2017). To determine the effects of knock‐down of C‐ROS‐1 in TWIST‐1 haploinsufficient SCS derived calvarial cells during osteogenesis, cells were transfected with scrambled siRNA control or siRNA targeting C‐ROS‐1 as previously described (Camp et al., 2017). Following osteogenic inductive culture, we observed a reduction in C‐ROS‐1 gene and protein expression levels in the presence of C‐ROS‐1 siRNA compared to scrambled siRNA controls, as determined by qPCR and Western blot analysis, respectively (Figures 3a and 3b). Furthermore, siRNA‐mediated knockdown of C‐ROS‐1 lead to a significant reduction in the formation of Alizarin red positive mineral deposits (Figure 3c), corresponding to a decrease in levels of extracellular calcium (Figure 3d), when compared to scramble siRNA controls. In agreement with these findings, siRNA knockdown of C‐ROS‐1 in SCS cells resulted in a significant reduction in the expression of the osteogenic markers, Runt‐related transcription factor 2 (RUNX2) and OSTEOPONTIN (OPN), when compared to scramble siRNA controls (Figure 3e).

Figure 3.

Figure 3

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Knock‐down of C‐ROS‐1 inhibits the osteogenic potential of TWIST‐1 haploinsufficient cranial cells from SCS patients (a) C‐ROS‐1 gene expression was assessed using quantitative real‐time PCR in TWIST‐1 haploinsufficient calvarial cells from SCS patients transfected with scrambled siRNA (siScram) or an siRNA targeting c‐ros‐1 (siRos‐1) and cultured under osteogenic conditions. Graph shows relative mean C‐ROS‐1 gene expression mean (±S.E.M) relative to β‐actin. (*p < 0.05, unpaired student's t‐test, n = 4 SCS donors). (b) Representative Western blot analysis to detect C‐ROS‐1 protein levels in siScram control and c‐ros‐1 siRNA transfected TWIST‐1 haploinsufficient cells. HSP90 antibody was used as a control for protein loading. (c) Representative images depicting mineral deposits stained with Alizarin red, scale bar = 200 μm. (d) Mean ± S.E.M levels of extracellular calcium quantitated using the Calcium Arsenazo III reagent and normalised to total DNA content per well (*p < 0.05, student's t‐test, n = 4 donors). (e) Real‐time PCR analysis of cranial cells derived from SCS patients transfected with either siScram control or siRos‐1 and cultured under osteogenic inducing media. Graph represents mean ± S.E.M of RUNX2 and OPN expression relative to β‐actin (*p < 0.05, student's t‐test, n = 4 donors)

3.4. Crizotinib is a C‐ROS‐1 inhibitor that can inhibit alkaline phosphatase production in TWIST‐1 haploinsufficient calvarial cells without affecting cell proliferation or viability

TWIST‐1 haploinsufficient SCS calvarial cells were cultured under osteogenic conditions in 0.1% DMSO (vehicle control) or in the presence of increasing concentrations (50–2 μM) of the C‐ROS‐1 inhibitor, Crizotinib, for seven days. Alkaline phosphatase staining, an early marker of osteogenesis and osteoblast function in vitro, was visibly reduced in cells treated with 500–2 μM of Crizotinib, compared to control (Figure 4a). We next examined the effect of these concentrations of Crizotinib on cell proliferation using bromodeoxyuridine (BrdU) incorporation. Treatment of cells with 500–1 μM of Crizotinib for 4 or 7 days had no effect on proliferation compared to untreated cells. However, treatment with 2 μM resulted in a reduction in cell proliferation at both time points (Figure 4b). Moreover, to assess the toxicity of Crizotinib, Annexin V/7AAD staining and flow cytometric analysis was performed DMSO and Crizotinib treated TWIST‐1 haploinsufficient calvarial cells. Treatment of cells with 500nM of Crizotinib did not alter the percentage of apoptotic or necrotic cells when compared to control (Figures 4c and 4d). However, there was a statistically significant increase in the percentage of early apoptotic cells when treated with Crizotinib at 2 μM (Annexin V positive cells) and an increase in the percentage of late apoptotic cells (Annexin V and 7AAD positive cells) when treated with 1–2 μM of Crizotinib compared to control. Treatment of cells with Crizotinib at 1 and 2 μM of did not alter the percentage of necrotic cells (7AAD positive cells) when compared to control (Figures 4c and 4d).

Figure 4.

Figure 4

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Effects of Crizotinib on TWIST‐1 haploinsufficient calvarial cell alkaline phosphatase activity, proliferation and viability. (a) TWIST‐1 haploinsufficient calvarial cells were cultured in osteogenic inducing conditions and treated with DMSO or Crizotinib at increasing concentrations (5–2 μM) for 7 days. (a) Representative images depicting positive alkaline phosphatase staining scale bar = 200μm. (b) TWIST‐1 haploinsufficient calvarial cells were left untreated (UT) or treated with DMSO or Crizotinib at 500, 1, or 2 μM. Graph represents the mean ± S.E.M proliferation rates of cranial cells cultured for 4 days (i) or 7 days (ii) as measured by bromodeoxyuridine (BrdU) incorporation (*p < 0.05, One‐way Anova with Tukey post‐hoc multiple comparison test, comparison to mean of untreated cells, n = 4). (c and d) Flow cytometric analysis of vehicle control and Crizotinib treated cells double stained with Annexin V and 7AAD. (c) Representative histograms of flow cytometric data. (d) Graphs represent mean percentage ± S.E.M of early apoptotic gated cells stained with Annexin V (i), late apoptotic gated cells stained with Annexin V and 7AAD (ii) and necrotic gated cells stained with 7AAD (iii) (*p < 0.05, One‐way Anova with Tukey post‐hoc multiple comparison test, comparison to mean of control, n = 4 donors)

3.5. Inhibition of C‐ROS‐1 activity by crizotinib inhibits the osteogenic potential of TWIST‐1 haploinsufficient human cranial cells and mouse calvariae

Osteoprogenitor TWIST‐1 haploinsufficient calvarial cells derived from SCS patients were cultured under osteogenic inducing conditions and treated with DMSO or Crizotinib at 500 or 1 μM. Western blot analysis demonstrated Crizotinib treatment at 500 or 1 μM had little effect on C‐ROS‐1 receptor levels yet reduced the level of phosphorylated C‐ROS‐1 and the osteogenic marker, RUNX2, compared to cells treated with DMSO vehicle control (Figure 5a). TWIST‐1 haploinsufficiency is associated with increased PI3K/Akt signalling in mutant osteoblasts from SCS patients. This mediates an increase in mutant osteoblast proliferation and an increase in overall bone growth compared to control (Guenou et al., 2006). Western blot analysis demonstrated Crizotinib treatment at 500 or 1 μM had little effect on total PI3K and Akt levels, yet reduced the level of the PI3K target proteins; phosphorylated Akt (Thr308) and phosphorylated Akt (Ser473) when compared to cells treated with DMSO vehicle control (Figure 5b). Furthermore, treatment with Crizotinib at 500 or 1 μM resulted in significant reduction in the formation of Alizarin red positive mineral (Figure 5c), corresponding to a decrease in levels of extracellular calcium (Figure 5d), when compared to with DMSO controls. However, there was no difference in calcium levels between cells, which were treated with either 500 or 1 μM of Crizotinib. Treatment of cells with Crizotinib at 500 or 1 μM also significantly reduced the expression levels of the osteogenic marker genes, RUNX2 and OPN when compared to treatment with DMSO alone (Figures 5a and 5e).

Figure 5.

Figure 5

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Crizotinib mediated reduction of C‐ROS‐1 activity and inhibition of osteogenesis in vitro and in a calvaria organotypic culture system. TWIST‐1 haploinsufficient calvarial cells were treated with DMSO or Crizotinib at 500 or 1 μM and cultured under osteogenic inducing conditions. (a and b) Representative Western blot analysis of protein lysates prepared for immunoblotting. Membranes were probed with total or phosphorylation‐specific antibodies as indicated. HSP90 protein levels (a) and β‐actin protein levels (b) were used as protein loading controls. (c) Representative images depicting mineral deposit stained with Alizarin red, scale bar = 200μm. (d) Mean ± S.E.M levels of extracellular calcium quantitated using the Calcium Arsenazo III reagent and normalised to total DNA content per well (*p < 0.05, One‐way Anova with Tukey post‐hoc multiple comparison test, n = 4 donors). (e) Real‐time PCR analysis. Graph represents mean ± S.E.M of RUNX2 and OPN gene expression relative to β‐actin (*p < 0.05, One‐way Anova with Tukey post‐hoc multiple comparison test, n = 4 donors). (f) Twist‐1del/+ heterozygous mouse neonatal calvaria were cultured in recombinant human BMP‐2 (50 ng/ml) in the presence of DMSO or Crizotinib at 500 or 1 μM. Representative photomicrographs of treated calvaria sections stained with hematoxylin and eosin illustrate the extent of bone formation induced. (g) Total bone area was quantitated morphometrically. Data is expressed as the mean ± S.E.M of two independent experiments each performed using three calvaria (*p < 0.05, One‐way Anova with Tukey post‐hoc multiple comparison test)

The effect of Crizotinib on bone formation was further examined using a calvaria organotypic culture system. The murine calvaria bone explant culture model has been used extensively to identify compounds that are anabolic for bone formation (Garrett, 2003; Garrett et al., 2003; Martin et al., 2010). Calvaria explant cultures were treated with recombinant BMP‐2 (50 ng/mL) for 10 days, to stimulate bone formation, in the presence of DSMO or Crizotinib at 500 or 1 μM. Calvaria treated with Crizotinib at 500 or 1 μM demonstrated a decrease in bone formation compared with calvaria treated with DMSO (Figures 5f and 5g). There was no difference in total bone formed between calvaria treated with 500 or 1 μM of Crizotinib. Collectively, these findings indicate that reduction of the aberrant C‐ROS‐1 levels found in TWIST‐1 haploinsufficient osteoprogenitor calvarial cells can reduce the osteogenic potential of these cells.

4. DISCUSSION

Our study has identified a novel mechanism responsible for regulating aberrant bone formation rates in TWIST‐1 haploinsufficient osteogenic progenitor calvarial cells derived from SCS patients. During development, Twist‐1 expression is greatest at sites of immature cranial mesenchyme cells within the cranial sutures (Connerney, Leshem, Muentener, Mercado, & Spicer, 2006; Johnson et al., 2000; Rice et al., 2000; Yoshida et al., 2005), where Twist‐1del/+ mutant mice display diminished Gli‐1+ cells and premature maturation within the suture mesenchyme, leading to craniosynostosis (Zhao et al., 2015). Previous studies have reported that TWIST‐1 maintains pre‐osteogenic stromal populations, and enhances their proliferation potential and life span, whilst inhibiting chondrogenesis and osteogenesis in vitro and in vivo (Cakouros et al., 2015, 2012; Isenmann et al., 2009). This highlights the importance of TWIST‐1 in suture biogenesis and in maintaining the suture as a growth centre. Previous studies have reported various molecular targets linked to TWIST‐1 function including; BMP, FGF, FGFR2, and Notch signalling (Connerney et al., 2008, 2006; Guenou, Kaabeche, Mee, & Marie, 2005; Hayashi et al., 2007; Howard et al., 1997; Johnson et al., 2000; Rice et al., 2000, 2003; Wilkie, 1997; Yen, Ting, & Maxson, 2010) and EphA4 expression in sutures (Ting et al., 2009). Recently we have identified C‐ROS‐1, a novel TWIST‐1 target, which promotes bone formation by BMSC and cranial osteoprogenitor cells (Camp et al., 2017). Previous studies of heterozygous and homozygous mutations of the C‐ROS‐1 gene (loss of function mutations) in mouse, showed no gross phenotypic deformities, but did report infertility in male homozygotes (Sonnenberg‐Riethmacher et al., 1996). However, in mice, the fusion of the interfrontal suture is complete at P15 (Sahar, Longaker, & Quarto, 2005), while all other sutures remain unfused throughout postnatal life (Bradley, Levine, Blewett, et al., 1996; Bradley, Levine, Roth, McCarthy, & Longaker, 1996). Therefore, latent fusion of the interfrontal suture and any aberrant defects in the remaining sutures may not result in an obvious cranial phenotype in these mice.

The present study found that TWIST‐1 haploinsufficient SCS calvarial cells expressed greater levels of C‐ROS‐1 in vitro compared to calvarial cells derived from unaffected individuals. Similarly, Twist‐1del/+ calvarial mouse cells expressed higher levels of C‐Ros‐1 than Twist‐1 wild‐type cells in vitro and in vivo. Functional studies demonstrated that siRNA mediated C‐ROS‐1 knockdown in TWIST‐1 haploinsufficient SCS calvarial cells lead to decreased levels of mineral formation and reduced expression of the early pre‐osteoblast marker RUNX2 and the osteoblast associated marker OPN, when compared to the siRNA scramble controls. These results suggest that C‐ROS‐1 plays an important role in suture mesenchyme osteogenic differentiation, and that C‐ROS‐1 levels are an important mediator of the effects of TWIST‐1 mutations on premature bone growth in sutures.

In this study we further investigated whether pharmacological targeting of C‐ROS‐1 activity in TWIST‐1 haploinsufficient calvarial cells could inhibit their osteogenic potential. Previous studies have reported that cancers with chromosomal rearrangements of the gene encoding C‐ROS‐1, are susceptible to the therapeutic tyrosine kinase inhibitor, Crizotinib (Li et al., 2017; Shaw & Solomon, 2015). Crizotinib is a small molecule tyrosine kinase inhibitor of C‐ROS‐1 (Bergethon et al., 2012), anaplastic lymphoma kinase (ALK) (Christensen et al., 2007) and tyrosine‐protein kinase Met (C‐MET) (Zou et al., 2007) receptor tyrosine kinase and belongs to the 3‐benzyloxy‐2‐aminopyridine series of kinase inhibitors (Timofeevski et al., 2009). Crizotinib acts by binding to the adenosinetriphosphate (ATP)‐binding site, thereby preventing binding of ATP and subsequent phosphorylation, which is required for activation of the enzyme. Our study showed that Crizotinib treatment of TWIST‐1 haploinsufficient SCS calvarial cells caused a dose‐dependent decrease in alkaline phosphatase activity, mineral deposition rates and bone gene expression levels. Importantly, Crizotinib at the inhibitory concentrations of 500 and 1 μM showed little or no affect on cell proliferation and viability. Analysis of 500 and 1 μM Crizotinib treated TWIST‐1 haploinsufficient calvarial cells showed a decrease in C‐ROS‐1 activity, via reduced phosphorylated C‐ROS‐1, and RUNX2 protein levels, with no affect on total C‐ROS‐1 receptor protein levels. Western blot analysis also demonstrated that Crizotinib treatment at 500 or 1 μM had little effect on total PI3K and Akt levels yet resulted in reduced PI3K/Akt signalling demonstrated by reduced phosphorylated Akt protein levels when compared to cells treated with DMSO vehicle control. In bone, PI3K signalling has been shown to control osteoblast differentiation and survival (Golden & Insogna, 2004). In addition, Akt signalling has been shown to be a crucial regulator of osteoblasts by promoting their differentiation and survival (Maruyama, Mirando, Deng, & Hsu, 2010; Mukherjee and Rotwein, 2009). Reduced Akt has been shown in mesenchymal stem cells to reduce osteoblast differentiation (Mukherjee & Rotwein, 2009). Moreover, a decrease in Akt signalling in mouse calvarial cells leads to osteoblast dysfunction due to decreased transcriptional activity of RUNX2 (Maruyama et al., 2010). Thus, treatment of cells from SCS patients with Crizotinib at 500 and 1 μM reduced C‐ROS‐1 activity and PI3K/Akt signalling resulting in; reduced alkaline phosphatase activity, decreased RUNX2 and OPN expression and reduced formation of Alizarin red positive mineral deposits, corresponding to a decrease in levels of extracellular calcium Moreover, Crizotinib treatment significantly inhibited BMP‐2 mediated osteogenesis in an organotypic culture system using calvaria derived from Twist‐1del/+ mice It has been previously demonstrated that the IGF‐induced PI3‐kinase‐Akt signalling cascade is essential for BMP2‐activated osteoblast differentiation and maturation, bone development and growth (Mukherjee & Rotwein, 2009). Thus, Crizotinib treatment may be able to inhibit BMP‐2 mediated osteogenesis and bone growth in TWIST‐1 haploinsufficient calvaria by reducing PI3K/Akt signalling. Collectively, these findings indicate that reduction of the aberrant C‐ROS‐1 levels found in TWIST‐1 haploinsufficient calvarial osteogenic progenitor cells can reduce the osteogenic potential of these cells in vitro and in an explant calvaria model. Furthermore, treatment of TWIST‐1 haploinsufficient calvarial cells with Crizotinibat concentrations of 500 and 1 μM was effective at reducing C‐ROS‐1 activity without affecting total C‐ROS‐1 protein levels and cell proliferation.

Crizotinib is currently the most effective C‐ROS‐1 tyrosine kinase inhibitor, which is approved by the therapeutic and goods administration (TGA) for the treatment of small lung carcinomas that express high levels of C‐ROS‐1. However, Crizotinib not only inhibits C‐ROS‐1 but also inhibits the receptor tyrosine kinases ALK and c‐MET. ALK is phylogenetically related to C‐ROS however, there is no known function of ALK signalling in mesenchymal stem cell osteogenic differentiation. Moreover, inhibition of C‐MET signalling can increase mineral formation in bone cell lines (Shibasaki et al., 2015) in contrast to our findings which found that C‐ROS‐1 knockdown or Crizotinib treatment decreased bone formation in BMSC and calvarial cells (Camp et al., 2017). Other studies have reported that C‐ROS‐1 high expressing cancers are often refractory to C‐MET inhibitors, indicating that these signalling pathways are not dependent (Aljohani et al., 2015).

Currently, the only treatment option available for SCS patients with craniosynostosis is major invasive cranial surgery, which involves the excision of the affected sutures and repositioning the skull bones. As dysmorphic growth may continue while the child develops, further surgical intervention may be required to ensure optimal cognitive and skeletal development. These procedures are potentially life threatening and traumatic for the children and their families, requiring substantial hospitalization and resources. The development of new approaches with therapeutic agents to prevent or minimize fusion could have a profound impact on an affected child's management resulting in reduced cranial surgery. Identification of TWIST‐1 regulated downstream molecular mechanisms involved in suture biogenesis may lead to the development of new approaches to prevent or minimize premature suture fusion. In conclusion, given that treatment with Crizotinib was well tolerated by TWIST‐1 haploinsufficient calvarial cells, this approach may be a feasible non‐surgical therapeutic strategy that could be used to treat craniosynostosis in SCS patients in the future.

ACKNOWLEDGMENTS

This work was supported in‐part by the National Health and Medical Research Project Grant APP1120989 and fellowship APP1042677, and the ACMFF.

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