Tissue and Cell 47 (2015) 115–121
Contents lists available at ScienceDirect
Tissue and Cell journal homepage: www.elsevier.com/locate/tice
Effects of matrix metalloproteinase 13 on vascular smooth muscle cells migration via Akt–ERK dependent pathway Sung Won Yang a,1 , Leejin Lim b,1 , Sujin Ju a , Dong-Hyun Choi c , Heesang Song b,d,∗ a
Department of Opthalmology, Chosun University School of Medicine, Gwangju, Republic of Korea Department of Biomaterials, Chosun University, Gwangju, Republic of Korea Department of Cardiology, Chosun University School of Medicine, Gwangju, Republic of Korea d Department of Biochemistry and Molecular Biology, Chosun University School of Medicine, Gwangju, Republic of Korea b c
a r t i c l e
i n f o
Article history: Received 2 June 2014 Received in revised form 12 December 2014 Accepted 13 December 2014 Available online 18 December 2014 Keywords: Smooth muscle cells Migration PDGF Matrix metalloproteinase 13
a b s t r a c t Migration of vascular smooth muscle cells (VSMCs) is an early event of atherosclerosis, which is mediated mainly by matrix metalloproteinase (MMP) 2 and 9. Because MMP13 is associated with tumor cells migration, we hypothesized that MMP13 participates in VSMC migration induced by certain stimuli such as platelet-derived growth factor (PDGF) and angiotensin II (Ang II). We found that the mRNA level of MMP13 in rat aortic smooth muscle cells (RAoSMCs) was increased by both PDGF and Ang II. We observed the significant decrease of migration in PDGF- or Ang II-treated RAoSMCs by MMP13 specific inhibitor treatment. Silencing of MMP13 by a specific small interfering RNA (siRNA) significantly decreased expression of the active form of MMP13, which is followed by the decreased migration of PDGFor Ang II-treated RAoSMCs. Interestingly, we observed synergistic inhibitory effects on migration by treatment with MMP2 and 13 or MMP9 and 13 inhibitors compared with that in single treatments. Moreover, we found that cordycepin, a known inhibitor of VSMC migration, caused significant downregulation of MMP2, 9, and 13 expression in PDGF-treated RAoSMCs. We further show that the level of phosphorylated extracellular signal-regulated kinase (ERK) was significantly decreased in LRP1-silenced cells, suggesting that ERK is a potential mediator of LRP1-regulated MMP2 and MMP9 expression in U87 cells. Together, our data strongly suggest that MMP13 involves VSMCs migration via an Akt and ERK-dependent regulation. © 2014 Elsevier Ltd. All rights reserved.
1. Introduction Increased proliferation and migration of vascular smooth muscle cells (VSMCs) are critical events in the pathophysiology of several prominent cardiovascular disease states such as atherosclerosis and restenosis (Owens, 1995; Schwartz et al., 1990). Following migration, VSMCs proliferate in the intima and secrete several extracellular matrix proteins and proteases that form atheromatous plaques under the influence of stimulatory cytokines (Jiang et al., 2006, 2008). Many reports have demonstrated that degradation of the extracellular matrix and basement membrane by proteases, such as matrix metalloproteinases (MMPs) is critical for
∗ Corresponding author at: Department of Biochemistry and Molecular Biology, Chosun University School of Medicine, 309 Pilmundaero, Gwangju 501-759, Republic of Korea. Tel.: +82 62 230 6290; fax: +82 62 226 4165. E-mail address: [email protected] (H. Song). 1 These authors contributed equally to this work. http://dx.doi.org/10.1016/j.tice.2014.12.004 0040-8166/© 2014 Elsevier Ltd. All rights reserved.
atherogenesis initiated by VSMCs (Newby, 2007, 2012). Most studies have focused on the role of MMP2 and 9 during atherogenesis (Dey and Lincoln, 2012; Gurjar et al., 1999; Newby, 2006) and have shown that initiation of atherogenesis can be controlled by downregulation of MMP2 and MMP9 expressions and/or inhibition of their activities (Chien et al., 2012; Gurjar et al., 1999; Karki et al., 2013). MMP13 (collagenase-3) was originally cloned from a human breast tumor cDNA library (Freije et al., 1994). MMP13 has activity against not only types I and III collagen but also uniquely type II collagen predominating in cartilage (Knauper et al., 1996a; Mitchell et al., 1996). Indeed MMP13 is highly overexpressed in chondrocytes and synovial cells in rheumatoid arthritis and osteoarthritis (Inada et al., 2004). In addition to association with bone and joint pathology, MMP13 has been reported to be elevated in various cancers (Acuff et al., 2006). MMP13 was reported to affect the levels of a functionally diverse range of proteins associated with the promotion of tumor growth, angiogenesis and metastasis (Neutzner et al., 2007). In fact, there are many reports indicating that MMP13 plays a critical role in cancer cell migration and invasion
116
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
(Meierjohann et al., 2010; Pan et al., 2011; Yu et al., 2011). The metastasis of esophageal squamous cell carcinoma induced by MUC1, one of the transmembrane mucins, was reported to be mediated by upregulation of MMP13 expression (Ye et al., 2011). Non-tumor endothelial cell migration is also mediated by MMP13 via nitric oxide activation (Lopez-Rivera et al., 2005). After the expression of MMP13 was indicated in human abdominal aortic aneurysms and vascular smooth muscle cells in culture (Mao et al., 1999), many evidence for association of MMP13 with vascular pathology have been reported. Indeed, the level of expression of MMP13 is related to the rate of progression in atherosclerosis, which is associated with MMP13 promoter polymorphism (Halpert et al., 1996; Sukhova et al., 1999; Yoon et al., 2002). Specific inhibition of MMP13 by siRNA increased collagen accumulation in atherosclerotic plaques, which may lead to resistance to rupture (Quillard et al., 2011). However, the role of MMP13 in vascular smooth muscle cells still remains to be elucidated in the atherogenic event. Therefore, we hypothesized that MMP13 has an effect on the migration of VSMCs. Because platelet-derived growth factor (PDGF) and angiotensin II (Ang II) are known to be important stimuli for the migration of VSMCs and atherogenesis (Cheng et al., 2012; Jiang et al., 2008), we investigated the role of MMP13 in PDGF- or Ang II-stimulated VSMCs migration. 2. Materials and methods 2.1. Cell culture and reagents Rat aortic SMCs (RAoSMCs) were isolated from 6- to 8-weekold Sprague-Dawley rats. Their thoracic aortas were removed and transferred on ice to serum-free Dulbecco’s modified Eagle medium (DMEM). The aorta was freed from connective tissue, transferred to a Petri dish containing 5 ml of an enzyme dissociation mixture consisting of DMEM containing 1 mg/ml collagenase type I (Sigma) and 0.5 mg/ml elastase (Worthington), and incubated for 30 min at 37 ◦ C. Then, the aorta was transferred to DMEM and the adventitia was stripped off with forceps under a stereomicroscope. The aorta was transferred to a plastic tube containing 5 ml of the enzyme dissociation mixture and incubated for 2 h at 37 ◦ C. The suspension was centrifuged (300 × g for 10 min) and then resuspended in DMEM with 10% fetal bovine serum (FBS). The cells were cultured in a humidified atmosphere with 5% CO2 , which was confirmed their homogeneity using immunocytochemistry with smooth muscle cell-specific ␣SMA (alpha-Smooth Muscle Actin) and SM-MHC (Smooth Muscle-Myosin Heavy Chain) and endothelial cell-specific vWF (von Willebrand Factor). PDGF, Ang II, cordycepin, and signal mediators inhibitors were obtained from Sigma. PI3K-specific inhibitor LY294002, Akt specific inhibitor, and ERK-specific inhibitor U0126 were also obtained from Sigma. MMP inhibitors, OA-Hy for MMP2, inhibitor I for MMP9, and pyrimidine dicarboxamide for MMP13, were obtained from Calbiochem (Cat. No. are 444244, 444278, and 444283, respectively). 2.2. MMP13 siRNA and transfection Single-stranded, rat MMP13-specific sense and antisense RNA oligonucleotides were synthesized by Bioneer, and doublestranded RNA molecules were generated according to the manufacturer’s instructions. Three different siRNA molecules for MMP13 were tested and their sequences were as follows: #1, GACAUCAUGAGAAAACCAAtt, #2, CUGUGAACAAGCUUCAGUAtt, #3, CACACUGAUAGAGGACACAtt. For transfection, RAoSMCs were grown to 80% confluence, and then transfected with MMP13 siRNA using siLentfect (Bio-Rad) according to the manufacturer’s instructions. After 48 h of transfection, the cells were treated with the indicated concentrations of PDGF or Ang II.
2.3. Proliferation assay Cells on 96-well plates were transfected with 20 or 100 nM of MMP13 siRNA #1 using siLentfect, then the BrdU proliferation assay was performed after 24 h using the Cell proliferation colorimetric ELISA system (Promega) according to the manufacturer’s instructions. 2.4. Migration assay Cell migration was examined using a three-dimensional Boyden chamber assay and a two-dimensional wound healing assay. Boyden chamber assays were carried out in 6.5-mm diameter transwell chambers with a pore size of 8.0 m. Cells (4 × 104 cells in 400 l) were resuspended in serum-free DMEM, and placed in the upper compartment of transwell chambers coated with 1% gelatin on the lower surface. The lower compartment was filled with 600 l of the DMEM migration medium containing 1% FBS and 5 ng/ml PDGF. After incubation for 16 h at 37 ◦ C, the membrane were fixed and stained. Then non-migrated cells on the upper compartment were removed using cotton swab, and the number of cells on the lower surface in five random fields were counted at 200× magnification. For the wound healing assay, a rectangular lesion was created using a cell scraper, and then the cells were rinsed twice with serum free DMEM and cultured in DMEM containing 10% FBS. After the indicated times, three randomly selected fields at the lesion border were imaged using a CCD camera (Olympus) attached to an inverted microscope. In each field, the distances from the margin of the lesion to the 10 most migrated cells were measured, and the mean value of the distances was taken as the mobility of the cells in each culture dish. 2.5. Zymography Aliquots of control and test media were electrophoresed on a 10% sodium dodecyl sulfate (SDS)-polyacrylamide gel containing 0.8% gelatin. The gel was washed twice with 2.5% Triton X-100 to remove SDS for 30 min, washed with deionized water for 1 h, rinsed with 50 mM Tris–HCl, pH 7.5, and then incubated overnight at room temperature with the development buffer (50 mM Tris–HCl, pH 7.5, 5 mM CaCl2 , 1 M ZnCl2 , 0.02% sodium azide, and 1% Triton X-100). The zymographic activities were revealed by staining with 1% Coomassie Blue and quantified using laser densitometry of the corresponding bands in the linear response of the gelatin zymogram. 2.6. Western blotting RAoSMCs were lysed in lysis buffer (phosphate-buffered saline containing 1% Triton X-100, protease inhibitor cocktail, and 1 mM phenylmethylsulfonyl fluoride). Protein concentrations were determined using a Bradford Protein Assay Kit. Equal quantities of protein were separated by 10% SDS-polyacrylamide gel electrophoresis and transferred to a polyvinylidene difluoride membrane. The membranes were incubated with anti-MMP2, -9, and -13 or anti-actin antibodies and then a horseradish peroxidase-conjugated secondary antibody. Immunoreactive proteins were detected using an ECL system (Intron). ImageJ software (National Institutes of Health; Bethesda, MD) was used for quantification. 2.7. Quantitative real-time PCR analysis Quantitative RT-PCR was carried out using SYBR Green. Total RNA isolated using Trizol was reverse transcribed to cDNA with
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
a 1st Strand cDNA Synthesis kit (Nanohelix). Quantitative realtime PCR was performed using a qPCR kit (Nanohelix). cDNA was amplified to detect the mRNA levels of several MMPs. GAPDH was used as an internal control. The relative expression levels were quantified and analyzed using Rotor-Gene 6 software (Corbettresearch). Three independent experiments were performed to analyze the relative gene expression, and each sample was tested in triplicate. Data were analyzed using the cycle threshold (CT) method. The amount of target mRNA (2−CT ) was normalized to the endogenous GAPDH reference (CT) and relative to the amount of the target mRNA in control samples, which was set at 1.0. 2.8. Statistical analysis All quantified data represent the mean of at least triplicate samples. Error bars represent standard deviations of the mean. Statistical significance was determined using Student’s t-test, and P < 0.05 was considered significant. 3. Results 3.1. Inhibition of MMP13 activity results in a reduction of PDGFor Ang II-induced RAoSMCs migration First, we confirmed the identity of RAoSMCs by immunocytochemistry. Over 95% of the RAoSMCs expressed smooth muscle cell-specific ␣SMA and SM-MHC, and endothelial cellspecific vWF was not expressed (data not shown). Further, we confirmed that treatment of RAoSMCs with PDGF or Ang II increased RAoSMCs migration in a dose-dependent manner using both two-dimensional wound healing and three-dimensional Boyden chamber assays (data not shown). In order to examine whether isoform-specific MMP activity is required for RAoSMCs migration, we tested the effect of specific inhibitors, OA-Hy for MMP2, inhibitor I for MMP9, and pyrimidine dicarboxamide for MMP13. We showed that each inhibitor has successfully inhibited the corresponding MMPs activity using zymographic analysis without any significant cross-inhibition for other isoform of MMPs (Fig. 1). Fig. 2 shows that 5 ng/ml PDGF or 200 nM Ang II increased RAoSMCs migration significantly in both two- and threedimensional assays. Because MMP2 and MMP9 are major mediators of RAoSMCs migration, we compared the effects of MMP activity inhibition using specific inhibitors on RAoSMCs migration. As expected, treatment with an MMP2-specific inhibitor decreased both PDGF- and Ang II-induced two-dimensional RAoSMCs migration by ∼50% and ∼70%, respectively, compared with that in
Fig. 1. Isoform-specific MMP inhibition with inhibitors. Zymographic analysis were performed with cultured media of RAoSMCs in the presence of 5 M OA-Hy, 10 nM inhibitor I, or 20 nM pyrimidine dicarboxamide for MMP2, MMP9, or MMP13, respectively.
117
untreated controls. Treatment with an MMP9-specific inhibitor also showed a significant reduction in PDGF- or Ang II-induced RAoSMCs migration. Interestingly, treatment with an MMP13specific inhibitor resulted in significant reductions of both PDGFand Ang II-induced two-dimensional RAoSMCs migration by ∼20% and ∼30%, respectively (Fig. 2A), even though the degree of reduction was lower than those of MMP2 and MMP9. We also performed three-dimensional Boyden chamber assays to confirm these results. As shown in Fig. 2B, treatment with the MMP13specific inhibitor reduced both PDGF- and Ang II-induced RAoSMCs migration by ∼20% and ∼30%, respectively, which were almost the same as those in the two-dimensional assay. In particular, the reduction of Ang II-induced RAoSMCs migration by MMP2, 9, or 13 was almost the same as that in the three-dimensional assay. Furthermore, to investigate the role of MMP13 in MMP2- and MMP9-mediated RAoSMCs migration, we evaluated the effects of co-inhibition of MMP13 activity with MMP2 or MMP9 activities on RAoSMCs migration using the three-dimensional Boyden chamber assay. As shown in Fig. 2C, no significant synergistic effect was observed by co-inhibition of MMP2 and MMP13 activities in PDGFinduced RAoSMCs migration compared with that of the MMP2 inhibitor alone. However, we observed a significant reduction by co-inhibition of MMP9 and MMP13 activities in PDGF-induced RAoSMCs migration compared with that of the MMP9 inhibitor alone. We also observed obvious synergistic effects of co-inhibition in Ang II-induced RAoSMCs migration. Co-inhibition with MMP13 showed a ∼20% increase in the reduction of Ang II-induced cell migration compared with that of the MMP2 or MMP9 inhibitors alone. 3.2. MMP13 silencing decreases PDGF- and Ang II-induced RAoSMCs migration To determine the specific role of MMP13 in RAoSMCs migration, we knocked down MMP13 expression using an MMP13-specific siRNA. We designed three double-stranded, 21-nucleotide siRNAs with 30 TT dinucleotide overhangs against the coding sequence of rat MMP13 (NM 133530). None of the siRNAs shared homology with exons of other known rat genes. The three siRNAs were screened for their effect on MMP13 protein levels using Western blot in order to indicate the most effective silencer for further analysis. As shown in Fig. 3A, MMP13 siRNAs #2 and #3 had no significant effect on MMP13 expression. However, MMP13 siRNA #1 silenced Ang II-induced MMP13 expression by more than 80%, suggesting that MMP13 siRNA #1 could modulate MMP13 expression. The reduction in MMP13 expression levels was maintained up to 72 h after transfection with MMP13 siRNA #1 (data not shown). We confirmed that mRNA expression level of MMP13 was also significantly decreased in MMP13 siRNA-treated RAoSMCs (Fig. 3B). Additionally, we demonstrated the specificity of siRNA #1 for MMP13 by showing no cross-reactivity with MMP2 and 9 (Fig. 3A and B). We further observed that MMP13 secreted into media was significantly decreased by ∼50% in zymographic analysis (Fig. 3C). To investigate whether MMP13 silencing affected RAoSMCs migration, we conducted three-dimensional cell migration assays using transwell chambers. We found that knockdown of MMP13 in PDGF- or Ang II-treated RAoSMCs decreased cell migration by ∼40% and ∼20%, respectively, compared with that in negative siRNA-treated controls (Fig. 3D). In addition, we examined the effect of MMP13 silencing on the survival and proliferative activity of RAoSMCs, because a decrease in survival or proliferation after MMP13 silencing may have contributed to the observed decrease in RAoSMCs migration. We found that MMP13 silencing did not significantly decrease the proliferation rate of RAoSMCs compared with that of control cells (Fig. 3E).
118
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
Fig. 2. Effect of MMP13 inhibition on RAoSMCs migration. (A) Two-dimensional migration assays were conducted with RAoSMCs using a modified wound healing assay in the presence of 5 M OA-Hy, 10 nM inhibitor I, or 20 nM pyrimidine dicarboxamide. The migrated distance during the designated period was measured. Scale bar, 200 m. (B) Three-dimensional migration assays were performed with RAoSMCs using Boyden chambers in the presence of designated MMP inhibitors above. (C) Three-dimensional migration assays were conducted in co-treatment of pyrimidine dicarboxamide with OA-Hy or inhibitor I in RAoSMCs. Columns, average of triple determinations; bars, SD. *P < 0.05 or **P < 0.01 as compared with control.
3.3. MMP13 is downregulated by a known RAoSMCs migration inhibitor, cordycepin Cordycepin is a type of nucleoside analog isolated from Cordyceps militaris, which inhibits VSMC migration via downregulation of MMP2 and MMP9 (Chang et al., 2008; Tuli et al., 2013). Therefore, we used cordycepin to confirm whether MMP13 plays a role in VSMC migration. Similar to previous reports, treatment with PDGF increased intracellular MMP2 and MMP9 expression levels as well as their secreted levels in the medium, which were significantly decreased by cordycepin in a dose-dependent manner. Interestingly, we observed a similar trend in MMP13 expression levels. The secreted level of MMP13 was ∼2-fold higher than that of the untreated control but no significant difference was observed in cell lysates. Treatment with cordycepin decreased MMP13 expression by up to 50% in both the media and cell lysates (Fig. 4A and B). Furthermore, the increased mRNA levels of MMP13 by PDGF treatment were inhibited by cordycepin (Fig. 4C). In addition to the result that silencing MMP13 significantly decreased the migration
of RAoSMCs (Fig. 3D), these results indicate that MMP13 might be one of mediators in VSMCs migration.
3.4. Akt and ERK are potential mediators of MMP13 expression The signal transduction pathways that modulate the activity of MMP transcription factors are highly diverse. Many reports have demonstrated that mitogen-activated protein kinase (MAPK) signal transduction pathways, including p38, ERK, and JNK, are well known mediators that stimulate or inhibit MMP expression depending on the cell types (Johansson et al., 2000; Nakai et al., 2013). To understand the mechanism by which stimuli regulate MMP13 expression in VSMCs, we analyzed the effects of signal mediator inhibitors on MMP13 expression. We found that an Akt or ERK-specific inhibitors, but not PI3K inhibitor, significantly decreased MMP13 expression levels (Fig. 5). These results indicate that Akt and ERK are likely upstream mediators that regulate MMP13 expression and functions in cell migration of VSMCs.
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
119
Fig. 3. MMP13 silencing in RAoSMCs. RAoSMCs were transfected with MMP13 siRNA or control siRNA using siLentFect. After 24 h transfection, cells and media were harvested for Western blotting, RNA extraction, zymographic analysis, or Boyden chamber assays. (A) MMP13 silencing by three different MMP13 siRNAs in RAoSMCs. The same amounts of lysates were analyzed by Western blotting using an antibody against MMP13. Anti-actin blot was used as a loading control. (B) mRNA levels of MMP13 were quantified by real-time PCR after treatment control or MMP13 siRNAs. (C) Extracellular activity of MMP13 after treatment control or MMP13 siRNAs was analyzed using zymography. (D) RAoSMCs migration after treatment control or MMP13 siRNAs was analyzed using Boyden chamber assays. Columns, average of triple determinations; bars, SD. *P < 0.05 or **P < 0.01 as compared with Neg. siRNA-treated control.
4. Discussion In this study, we showed that MMP13 silencing led to a significant decrease in PDGF- and Ang II-induced RAoSMCs migration, which is a key event in the early progression of atherosclerosis (Doran et al., 2008), and provided evidence that MMP13 participated in this process. Although many reports demonstrate that MMP13 is involved in the migration and invasion of various tumor and non-tumor cell types, including esophageal squamous cell carcinoma, cutaneous squamous cell carcinoma, and endothelial cells (Lopez-Rivera et al., 2005; Xu et al., 2012; Ye et al., 2011), the role of MMP13 in migratory VSMCs has been unclear. Our finding that Ang II treatment resulted in a significant increase of MMP13 expression in RAoSMCs is similar to a previous report in which Ang II induced production of MMP3 and MMP13 via the AT1 receptor in osteoblasts (Nakai et al., 2013). In contrast, Tarin et al. showed that a lack of NO inhibits MMP13 and increases ICAM-1 levels in atherosclerosis, leading to worsen atherosclerosis by increasing monocyte adhesion to endothelial cells (Tarin et al., 2009). Therefore, because Ang II is known to decrease the bioavailability of NO, one could hypothesize that an increase in Ang II would result in decreased MMP13. However, it has been shown that NO regulated MMP13 release during wound repair (Lizarbe et al., 2008). The disparity of MMP13 expression and role among these studies may reflect the diverse characteristics of the experimental systems including cell types, stimuli, and signaling pathways
that participate in cellular migration. Nevertheless, we found that PDGF treatment resulted in a significant increase of transcriptional and translational MMP13 levels, indicating that MMP13 may be a mediator of RAoSMCs migration in addition to MMP2 and MMP9. This finding is supported by the fact that inhibition of MMP13 activity led to a significant decrease of RAoSMCs migration and co-inhibition of MMP13 activity with MMP2 or MMP9 showed synergistic effects on the reduction of RAoSMCs migration. We also found that silencing of MMP13 markedly inhibited the transcriptional and translational levels of MMP13 induced by PDGF or Ang II, leading to a decrease in RAoSMCs migration. Although ∼80% of MMP13 expression was silenced by siRNA, the migration rates of RAoSMCs induced by PDGF or Ang II were reduced by only ∼40% and ∼20%, respectively. A potential explanation for this discrepancy is that MMP2 and MMP9 were still functional in the MMP13silenced RAoSMCs. In addition, other modulators may regulate MMP expression and RAoSMCs migration independent of MMP13. Our observation that silencing or inhibition of MMP13 successfully decreased cell migration of PDGF- or Ang II-treated RAoSMCs strongly suggests that MMP13 expression levels are positively correlated with RAoSMCs migration rates. Indeed, these results are coincided with the previous report that B16F1 melanoma grafts displayed reduced tumor growth and strongly decreased metastasis and angiogenesis in a MMP13−/− mouse model compared to wildtype mice (Zigrino et al., 2009). Our results provide strong evidence that reducing MMP13 expression and function may protect
120
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
Fig. 4. Altered MMP13 expression by cordycepin. (A) Altered MMPs expression by cordycepin was evaluated in media and (B) cell lysates of 5 ng PDGF-treated RAoSMCs using Western blot. (C) The effects of cordycepin on mRNA levels of MMP13 were quantified by real-time PCR in PDGF-induced RAoSMCs. Columns, average of triple determinations; bars, SD. **P < 0.01 as compared with PDGF-treated control.
Fig. 5. Effects of inhibitors on MMP13 expression. The resultant MMP13 expressions in the absence or presence of 5 M LY294002, 2 M Akt inhibitor, or 5 M U0126 were revealed using Western blot. Columns, average of triple determinations; bars, SD. **P < 0.01 as compared with non-treated control.
against early atherogenesis, particularly VSMC migration. Additionally, because MT1-MMP (MMP14) is a known activator of both MMP2 and 13 (Knauper et al., 1996b, 2002), further investigation will be needed to reveal the relationship of MT1-MMP or MMP2 and MMP13 in atherogenesis. In addition, further studies aimed at identifying the precise signal transduction pathways and transcription factors that mediate MMP13 expression may provide a molecular understanding of the roles of MMP13 in atherosclerosis.
Acknowledgement This was supported by grants from the Chosun University Medical Institute, 2012. References Acuff, H.B., Carter, K.J., Fingleton, B., Gorden, D.L., Matrisian, L.M., 2006. Matrix metalloproteinase-9 from bone marrow-derived cells contributes to survival
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121 but not growth of tumor cells in the lung microenvironment. Cancer Res. 66, 259–266. Chang, W., Lim, S., Song, H., Song, B.W., Kim, H.J., Cha, M.J., Sung, J.M., Kim, T.W., Hwang, K.C., 2008. Cordycepin inhibits vascular smooth muscle cell proliferation. Eur. J. Pharmacol. 597, 64–69. Cheng, C., Haasdijk, R.A., Tempel, D., den Dekker, W.K., Chrifi, I., Blonden, L.A., van de Kamp, E.H., de Boer, M., Burgisser, P.E., Noorderloos, A., Rens, J.A., Ten Hagen, T.L., Duckers, H.J., 2012. PDGF-induced migration of vascular smooth muscle cells is inhibited by heme oxygenase-1 via VEGFR2 upregulation and subsequent assembly of inactive VEGFR2/PDGFRbeta heterodimers. Arterioscler. Thromb. Vasc. Biol. 32, 1289–1298. Chien, Y.C., Sheu, M.J., Wu, C.H., Lin, W.H., Chen, Y.Y., Cheng, P.L., Cheng, H.C., 2012. A Chinese herbal formula “Gan-Lu-Yin” suppresses vascular smooth muscle cell migration by inhibiting matrix metalloproteinase-2/9 through the PI3K/AKT and ERK signaling pathways. BMC Complement Altern. Med. 12, 137. Dey, N.B., Lincoln, T.M., 2012. Possible involvement of cyclic-GMP-dependent protein kinase on matrix metalloproteinase-2 expression in rat aortic smooth muscle cells. Mol. Cell. Biochem. 368, 27–35. Doran, A.C., Meller, N., McNamara, C.A., 2008. Role of smooth muscle cells in the initiation and early progression of atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 28, 812–819. Freije, J.M., Diez-Itza, I., Balbin, M., Sanchez, L.M., Blasco, R., Tolivia, J., Lopez-Otin, C., 1994. Molecular cloning and expression of collagenase-3, a novel human matrix metalloproteinase produced by breast carcinomas. J. Biol. Chem. 269, 16766–16773. Gurjar, M.V., Sharma, R.V., Bhalla, R.C., 1999. eNOS gene transfer inhibits smooth muscle cell migration and MMP-2 and MMP-9 activity. Arterioscler. Thromb. Vasc. Biol. 19, 2871–2877. Halpert, I., Sires, U.I., Roby, J.D., Potter-Perigo, S., Wight, T.N., Shapiro, S.D., Welgus, H.G., Wickline, S.A., Parks, W.C., 1996. Matrilysin is expressed by lipid-laden macrophages at sites of potential rupture in atherosclerotic lesions and localizes to areas of versican deposition, a proteoglycan substrate for the enzyme. Proc. Natl. Acad. Sci. U. S. A. 93, 9748–9753. Inada, M., Wang, Y., Byrne, M.H., Rahman, M.U., Miyaura, C., Lopez-Otin, C., Krane, S.M., 2004. Critical roles for collagenase-3 (MMP13) in development of growth plate cartilage and in endochondral ossification. Proc. Natl. Acad. Sci. U. S. A. 101, 17192–17197. Jiang, M., Bujo, H., Ohwaki, K., Unoki, H., Yamazaki, H., Kanaki, T., Shibasaki, M., Azuma, K., Harigaya, K., Schneider, W.J., Saito, Y., 2008. Ang II-stimulated migration of vascular smooth muscle cells is dependent on LR11 in mice. J. Clin. Invest. 118, 2733–2746. Jiang, M., Bujo, H., Zhu, Y., Yamazaki, H., Hirayama, S., Kanaki, T., Shibasaki, M., Takahashi, K., Schneider, W.J., Saito, Y., 2006. Pitavastatin attenuates the PDGF-induced LR11/uPA receptor-mediated migration of smooth muscle cells. Biochem. Biophys. Res. Commun. 348, 1367–1377. Johansson, N., Ala-aho, R., Uitto, V., Grenman, R., Fusenig, N.E., Lopez-Otin, C., Kahari, V.M., 2000. Expression of collagenase-3 (MMP-13) and collagenase-1 (MMP1) by transformed keratinocytes is dependent on the activity of p38 mitogenactivated protein kinase. J. Cell Sci. 113 (Pt 2), 227–235. Karki, R., Jeon, E.R., Kim, D.W., 2013. Nelumbo nucifera leaf extract inhibits neointimal hyperplasia through modulation of smooth muscle cell proliferation and migration. Nutrition 29, 268–275. Knauper, V., Bailey, L., Worley, J.R., Soloway, P., Patterson, M.L., Murphy, G., 2002. Cellular activation of proMMP-13 by MT1-MMP depends on the C-terminal domain of MMP-13. FEBS Lett. 532, 127–130. Knauper, V., Lopez-Otin, C., Smith, B., Knight, G., Murphy, G., 1996a. Biochemical characterization of human collagenase-3. J. Biol. Chem. 271, 1544–1550. Knauper, V., Will, H., Lopez-Otin, C., Smith, B., Atkinson, S.J., Stanton, H., Hembry, R.M., Murphy, G., 1996b. Cellular mechanisms for human procollagenase3 (MMP-13) activation. Evidence that MT1-MMP (MMP-14) and gelatinase A (MMP-2) are able to generate active enzyme. J. Biol. Chem. 271, 17124–17131. Lizarbe, T.R., Garcia-Rama, C., Tarin, C., Saura, M., Calvo, E., Lopez, J.A., Lopez-Otin, C., Folgueras, A.R., Lamas, S., Zaragoza, C., 2008. Nitric oxide elicits functional MMP13 protein-tyrosine nitration during wound repair. FASEB J. 22, 3207–3215.
121
Lopez-Rivera, E., Lizarbe, T.R., Martinez-Moreno, M., Lopez-Novoa, J.M., RodriguezBarbero, A., Rodrigo, J., Fernandez, A.P., Alvarez-Barrientos, A., Lamas, S., Zaragoza, C., 2005. Matrix metalloproteinase 13 mediates nitric oxide activation of endothelial cell migration. Proc. Natl. Acad. Sci. U. S. A. 102, 3685–3690. Mao, D., Lee, J.K., VanVickle, S.J., Thompson, R.W., 1999. Expression of collagenase-3 (MMP-13) in human abdominal aortic aneurysms and vascular smooth muscle cells in culture. Biochem. Biophys. Res. Commun. 261, 904–910. Meierjohann, S., Hufnagel, A., Wende, E., Kleinschmidt, M.A., Wolf, K., Friedl, P., Gaubatz, S., Schartl, M., 2010. MMP13 mediates cell cycle progression in melanocytes and melanoma cells: in vitro studies of migration and proliferation. Mol. Cancer 9, 201. Mitchell, P.G., Magna, H.A., Reeves, L.M., Lopresti-Morrow, L.L., Yocum, S.A., Rosner, P.J., Geoghegan, K.F., Hambor, J.E., 1996. Cloning, expression, and type II collagenolytic activity of matrix metalloproteinase-13 from human osteoarthritic cartilage. J. Clin. Invest. 97, 761–768. Nakai, K., Kawato, T., Morita, T., Iinuma, T., Kamio, N., Zhao, N., Maeno, M., 2013. Angiotensin II induces the production of MMP-3 and MMP-13 through the MAPK signaling pathways via the AT(1) receptor in osteoblasts. Biochimie 95, 922–933. Neutzner, M., Lopez, T., Feng, X., Bergmann-Leitner, E.S., Leitner, W.W., Udey, M.C., 2007. MFG-E8/lactadherin promotes tumor growth in an angiogenesisdependent transgenic mouse model of multistage carcinogenesis. Cancer Res. 67, 6777–6785. Newby, A.C., 2006. Matrix metalloproteinases regulate migration, proliferation, and death of vascular smooth muscle cells by degrading matrix and non-matrix substrates. Cardiovasc. Res. 69, 614–624. Newby, A.C., 2007. Metalloproteinases and vulnerable atherosclerotic plaques. Trends Cardiovasc. Med. 17, 253–258. Newby, A.C., 2012. Matrix metalloproteinase inhibition therapy for vascular diseases. Vasc. Pharmacol. 56, 232–244. Owens, G.K., 1995. Regulation of differentiation of vascular smooth muscle cells. Physiol. Rev. 75, 487–517. Pan, X., Mao, X., Cheng, T., Zhang, X., 2011. Macrophage migration inhibitory factor: a regulator of MMP13 and inflammation in titanium particles-stimulated air pouch in vivo. Mol. Cell. Biochem. 357, 313–321. Quillard, T., Tesmenitsky, Y., Croce, K., Travers, R., Shvartz, E., Koskinas, K.C., Sukhova, G.K., Aikawa, E., Aikawa, M., Libby, P., 2011. Selective inhibition of matrix metalloproteinase-13 increases collagen content of established mouse atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 31, 2464–2472. Schwartz, S.M., Heimark, R.L., Majesky, M.W., 1990. Developmental mechanisms underlying pathology of arteries. Physiol. Rev. 70, 1177–1209. Sukhova, G.K., Schonbeck, U., Rabkin, E., Schoen, F.J., Poole, A.R., Billinghurst, R.C., Libby, P., 1999. Evidence for increased collagenolysis by interstitial collagenases1 and -3 in vulnerable human atheromatous plaques. Circulation 99, 2503–2509. Tarin, C., Gomez, M., Calvo, E., Lopez, J.A., Zaragoza, C., 2009. Endothelial nitric oxide deficiency reduces MMP-13-mediated cleavage of ICAM-1 in vascular endothelium: a role in atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 29, 27–32. Tuli, H.S., Sharma, A.K., Sandhu, S.S., Kashyap, D., 2013. Cordycepin: a bioactive metabolite with therapeutic potential. Life Sci. 93, 863–869. Xu, N., Zhang, L., Meisgen, F., Harada, M., Heilborn, J., Homey, B., Grander, D., Stahle, M., Sonkoly, E., Pivarcsi, A., 2012. MicroRNA-125b down-regulates matrix metallopeptidase 13 and inhibits cutaneous squamous cell carcinoma cell proliferation, migration, and invasion. J. Biol. Chem. 287, 29899–29908. Ye, Q., Yan, Z., Liao, X., Li, Y., Yang, J., Sun, J., Kawano, T., Wang, X., Cao, Z., Wang, Z., Huang, L., 2011. MUC1 induces metastasis in esophageal squamous cell carcinoma by upregulating matrix metalloproteinase 13. Lab. Invest. 91, 778–787. Yoon, S., Kuivaniemi, H., Gatalica, Z., Olson, J.M., Buttice, G., Ye, S., Norris, B.A., Malcom, G.T., Strong, J.P., Tromp, G., 2002. MMP13 promoter polymorphism is associated with atherosclerosis in the abdominal aorta of young black males. Matrix Biol. 21, 487–498. Yu, T., Wu, Y., Helman, J.I., Wen, Y., Wang, C., Li, L., 2011. CXCR4 promotes oral squamous cell carcinoma migration and invasion through inducing expression of MMP-9 and MMP-13 via the ERK signaling pathway. Mol. Cancer Res. 9, 161–172. Zigrino, P., Kuhn, I., Bauerle, T., Zamek, J., Fox, J.W., Neumann, S., Licht, A., SchorppKistner, M., Angel, P., Mauch, C., 2009. Stromal expression of MMP-13 is required for melanoma invasion and metastasis. J. Invest. Dermatol. 129, 2686–2693.
Contents lists available at ScienceDirect
Tissue and Cell journal homepage: www.elsevier.com/locate/tice
Effects of matrix metalloproteinase 13 on vascular smooth muscle cells migration via Akt–ERK dependent pathway Sung Won Yang a,1 , Leejin Lim b,1 , Sujin Ju a , Dong-Hyun Choi c , Heesang Song b,d,∗ a
Department of Opthalmology, Chosun University School of Medicine, Gwangju, Republic of Korea Department of Biomaterials, Chosun University, Gwangju, Republic of Korea Department of Cardiology, Chosun University School of Medicine, Gwangju, Republic of Korea d Department of Biochemistry and Molecular Biology, Chosun University School of Medicine, Gwangju, Republic of Korea b c
a r t i c l e
i n f o
Article history: Received 2 June 2014 Received in revised form 12 December 2014 Accepted 13 December 2014 Available online 18 December 2014 Keywords: Smooth muscle cells Migration PDGF Matrix metalloproteinase 13
a b s t r a c t Migration of vascular smooth muscle cells (VSMCs) is an early event of atherosclerosis, which is mediated mainly by matrix metalloproteinase (MMP) 2 and 9. Because MMP13 is associated with tumor cells migration, we hypothesized that MMP13 participates in VSMC migration induced by certain stimuli such as platelet-derived growth factor (PDGF) and angiotensin II (Ang II). We found that the mRNA level of MMP13 in rat aortic smooth muscle cells (RAoSMCs) was increased by both PDGF and Ang II. We observed the significant decrease of migration in PDGF- or Ang II-treated RAoSMCs by MMP13 specific inhibitor treatment. Silencing of MMP13 by a specific small interfering RNA (siRNA) significantly decreased expression of the active form of MMP13, which is followed by the decreased migration of PDGFor Ang II-treated RAoSMCs. Interestingly, we observed synergistic inhibitory effects on migration by treatment with MMP2 and 13 or MMP9 and 13 inhibitors compared with that in single treatments. Moreover, we found that cordycepin, a known inhibitor of VSMC migration, caused significant downregulation of MMP2, 9, and 13 expression in PDGF-treated RAoSMCs. We further show that the level of phosphorylated extracellular signal-regulated kinase (ERK) was significantly decreased in LRP1-silenced cells, suggesting that ERK is a potential mediator of LRP1-regulated MMP2 and MMP9 expression in U87 cells. Together, our data strongly suggest that MMP13 involves VSMCs migration via an Akt and ERK-dependent regulation. © 2014 Elsevier Ltd. All rights reserved.
1. Introduction Increased proliferation and migration of vascular smooth muscle cells (VSMCs) are critical events in the pathophysiology of several prominent cardiovascular disease states such as atherosclerosis and restenosis (Owens, 1995; Schwartz et al., 1990). Following migration, VSMCs proliferate in the intima and secrete several extracellular matrix proteins and proteases that form atheromatous plaques under the influence of stimulatory cytokines (Jiang et al., 2006, 2008). Many reports have demonstrated that degradation of the extracellular matrix and basement membrane by proteases, such as matrix metalloproteinases (MMPs) is critical for
∗ Corresponding author at: Department of Biochemistry and Molecular Biology, Chosun University School of Medicine, 309 Pilmundaero, Gwangju 501-759, Republic of Korea. Tel.: +82 62 230 6290; fax: +82 62 226 4165. E-mail address: [email protected] (H. Song). 1 These authors contributed equally to this work. http://dx.doi.org/10.1016/j.tice.2014.12.004 0040-8166/© 2014 Elsevier Ltd. All rights reserved.
atherogenesis initiated by VSMCs (Newby, 2007, 2012). Most studies have focused on the role of MMP2 and 9 during atherogenesis (Dey and Lincoln, 2012; Gurjar et al., 1999; Newby, 2006) and have shown that initiation of atherogenesis can be controlled by downregulation of MMP2 and MMP9 expressions and/or inhibition of their activities (Chien et al., 2012; Gurjar et al., 1999; Karki et al., 2013). MMP13 (collagenase-3) was originally cloned from a human breast tumor cDNA library (Freije et al., 1994). MMP13 has activity against not only types I and III collagen but also uniquely type II collagen predominating in cartilage (Knauper et al., 1996a; Mitchell et al., 1996). Indeed MMP13 is highly overexpressed in chondrocytes and synovial cells in rheumatoid arthritis and osteoarthritis (Inada et al., 2004). In addition to association with bone and joint pathology, MMP13 has been reported to be elevated in various cancers (Acuff et al., 2006). MMP13 was reported to affect the levels of a functionally diverse range of proteins associated with the promotion of tumor growth, angiogenesis and metastasis (Neutzner et al., 2007). In fact, there are many reports indicating that MMP13 plays a critical role in cancer cell migration and invasion
116
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
(Meierjohann et al., 2010; Pan et al., 2011; Yu et al., 2011). The metastasis of esophageal squamous cell carcinoma induced by MUC1, one of the transmembrane mucins, was reported to be mediated by upregulation of MMP13 expression (Ye et al., 2011). Non-tumor endothelial cell migration is also mediated by MMP13 via nitric oxide activation (Lopez-Rivera et al., 2005). After the expression of MMP13 was indicated in human abdominal aortic aneurysms and vascular smooth muscle cells in culture (Mao et al., 1999), many evidence for association of MMP13 with vascular pathology have been reported. Indeed, the level of expression of MMP13 is related to the rate of progression in atherosclerosis, which is associated with MMP13 promoter polymorphism (Halpert et al., 1996; Sukhova et al., 1999; Yoon et al., 2002). Specific inhibition of MMP13 by siRNA increased collagen accumulation in atherosclerotic plaques, which may lead to resistance to rupture (Quillard et al., 2011). However, the role of MMP13 in vascular smooth muscle cells still remains to be elucidated in the atherogenic event. Therefore, we hypothesized that MMP13 has an effect on the migration of VSMCs. Because platelet-derived growth factor (PDGF) and angiotensin II (Ang II) are known to be important stimuli for the migration of VSMCs and atherogenesis (Cheng et al., 2012; Jiang et al., 2008), we investigated the role of MMP13 in PDGF- or Ang II-stimulated VSMCs migration. 2. Materials and methods 2.1. Cell culture and reagents Rat aortic SMCs (RAoSMCs) were isolated from 6- to 8-weekold Sprague-Dawley rats. Their thoracic aortas were removed and transferred on ice to serum-free Dulbecco’s modified Eagle medium (DMEM). The aorta was freed from connective tissue, transferred to a Petri dish containing 5 ml of an enzyme dissociation mixture consisting of DMEM containing 1 mg/ml collagenase type I (Sigma) and 0.5 mg/ml elastase (Worthington), and incubated for 30 min at 37 ◦ C. Then, the aorta was transferred to DMEM and the adventitia was stripped off with forceps under a stereomicroscope. The aorta was transferred to a plastic tube containing 5 ml of the enzyme dissociation mixture and incubated for 2 h at 37 ◦ C. The suspension was centrifuged (300 × g for 10 min) and then resuspended in DMEM with 10% fetal bovine serum (FBS). The cells were cultured in a humidified atmosphere with 5% CO2 , which was confirmed their homogeneity using immunocytochemistry with smooth muscle cell-specific ␣SMA (alpha-Smooth Muscle Actin) and SM-MHC (Smooth Muscle-Myosin Heavy Chain) and endothelial cell-specific vWF (von Willebrand Factor). PDGF, Ang II, cordycepin, and signal mediators inhibitors were obtained from Sigma. PI3K-specific inhibitor LY294002, Akt specific inhibitor, and ERK-specific inhibitor U0126 were also obtained from Sigma. MMP inhibitors, OA-Hy for MMP2, inhibitor I for MMP9, and pyrimidine dicarboxamide for MMP13, were obtained from Calbiochem (Cat. No. are 444244, 444278, and 444283, respectively). 2.2. MMP13 siRNA and transfection Single-stranded, rat MMP13-specific sense and antisense RNA oligonucleotides were synthesized by Bioneer, and doublestranded RNA molecules were generated according to the manufacturer’s instructions. Three different siRNA molecules for MMP13 were tested and their sequences were as follows: #1, GACAUCAUGAGAAAACCAAtt, #2, CUGUGAACAAGCUUCAGUAtt, #3, CACACUGAUAGAGGACACAtt. For transfection, RAoSMCs were grown to 80% confluence, and then transfected with MMP13 siRNA using siLentfect (Bio-Rad) according to the manufacturer’s instructions. After 48 h of transfection, the cells were treated with the indicated concentrations of PDGF or Ang II.
2.3. Proliferation assay Cells on 96-well plates were transfected with 20 or 100 nM of MMP13 siRNA #1 using siLentfect, then the BrdU proliferation assay was performed after 24 h using the Cell proliferation colorimetric ELISA system (Promega) according to the manufacturer’s instructions. 2.4. Migration assay Cell migration was examined using a three-dimensional Boyden chamber assay and a two-dimensional wound healing assay. Boyden chamber assays were carried out in 6.5-mm diameter transwell chambers with a pore size of 8.0 m. Cells (4 × 104 cells in 400 l) were resuspended in serum-free DMEM, and placed in the upper compartment of transwell chambers coated with 1% gelatin on the lower surface. The lower compartment was filled with 600 l of the DMEM migration medium containing 1% FBS and 5 ng/ml PDGF. After incubation for 16 h at 37 ◦ C, the membrane were fixed and stained. Then non-migrated cells on the upper compartment were removed using cotton swab, and the number of cells on the lower surface in five random fields were counted at 200× magnification. For the wound healing assay, a rectangular lesion was created using a cell scraper, and then the cells were rinsed twice with serum free DMEM and cultured in DMEM containing 10% FBS. After the indicated times, three randomly selected fields at the lesion border were imaged using a CCD camera (Olympus) attached to an inverted microscope. In each field, the distances from the margin of the lesion to the 10 most migrated cells were measured, and the mean value of the distances was taken as the mobility of the cells in each culture dish. 2.5. Zymography Aliquots of control and test media were electrophoresed on a 10% sodium dodecyl sulfate (SDS)-polyacrylamide gel containing 0.8% gelatin. The gel was washed twice with 2.5% Triton X-100 to remove SDS for 30 min, washed with deionized water for 1 h, rinsed with 50 mM Tris–HCl, pH 7.5, and then incubated overnight at room temperature with the development buffer (50 mM Tris–HCl, pH 7.5, 5 mM CaCl2 , 1 M ZnCl2 , 0.02% sodium azide, and 1% Triton X-100). The zymographic activities were revealed by staining with 1% Coomassie Blue and quantified using laser densitometry of the corresponding bands in the linear response of the gelatin zymogram. 2.6. Western blotting RAoSMCs were lysed in lysis buffer (phosphate-buffered saline containing 1% Triton X-100, protease inhibitor cocktail, and 1 mM phenylmethylsulfonyl fluoride). Protein concentrations were determined using a Bradford Protein Assay Kit. Equal quantities of protein were separated by 10% SDS-polyacrylamide gel electrophoresis and transferred to a polyvinylidene difluoride membrane. The membranes were incubated with anti-MMP2, -9, and -13 or anti-actin antibodies and then a horseradish peroxidase-conjugated secondary antibody. Immunoreactive proteins were detected using an ECL system (Intron). ImageJ software (National Institutes of Health; Bethesda, MD) was used for quantification. 2.7. Quantitative real-time PCR analysis Quantitative RT-PCR was carried out using SYBR Green. Total RNA isolated using Trizol was reverse transcribed to cDNA with
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
a 1st Strand cDNA Synthesis kit (Nanohelix). Quantitative realtime PCR was performed using a qPCR kit (Nanohelix). cDNA was amplified to detect the mRNA levels of several MMPs. GAPDH was used as an internal control. The relative expression levels were quantified and analyzed using Rotor-Gene 6 software (Corbettresearch). Three independent experiments were performed to analyze the relative gene expression, and each sample was tested in triplicate. Data were analyzed using the cycle threshold (CT) method. The amount of target mRNA (2−CT ) was normalized to the endogenous GAPDH reference (CT) and relative to the amount of the target mRNA in control samples, which was set at 1.0. 2.8. Statistical analysis All quantified data represent the mean of at least triplicate samples. Error bars represent standard deviations of the mean. Statistical significance was determined using Student’s t-test, and P < 0.05 was considered significant. 3. Results 3.1. Inhibition of MMP13 activity results in a reduction of PDGFor Ang II-induced RAoSMCs migration First, we confirmed the identity of RAoSMCs by immunocytochemistry. Over 95% of the RAoSMCs expressed smooth muscle cell-specific ␣SMA and SM-MHC, and endothelial cellspecific vWF was not expressed (data not shown). Further, we confirmed that treatment of RAoSMCs with PDGF or Ang II increased RAoSMCs migration in a dose-dependent manner using both two-dimensional wound healing and three-dimensional Boyden chamber assays (data not shown). In order to examine whether isoform-specific MMP activity is required for RAoSMCs migration, we tested the effect of specific inhibitors, OA-Hy for MMP2, inhibitor I for MMP9, and pyrimidine dicarboxamide for MMP13. We showed that each inhibitor has successfully inhibited the corresponding MMPs activity using zymographic analysis without any significant cross-inhibition for other isoform of MMPs (Fig. 1). Fig. 2 shows that 5 ng/ml PDGF or 200 nM Ang II increased RAoSMCs migration significantly in both two- and threedimensional assays. Because MMP2 and MMP9 are major mediators of RAoSMCs migration, we compared the effects of MMP activity inhibition using specific inhibitors on RAoSMCs migration. As expected, treatment with an MMP2-specific inhibitor decreased both PDGF- and Ang II-induced two-dimensional RAoSMCs migration by ∼50% and ∼70%, respectively, compared with that in
Fig. 1. Isoform-specific MMP inhibition with inhibitors. Zymographic analysis were performed with cultured media of RAoSMCs in the presence of 5 M OA-Hy, 10 nM inhibitor I, or 20 nM pyrimidine dicarboxamide for MMP2, MMP9, or MMP13, respectively.
117
untreated controls. Treatment with an MMP9-specific inhibitor also showed a significant reduction in PDGF- or Ang II-induced RAoSMCs migration. Interestingly, treatment with an MMP13specific inhibitor resulted in significant reductions of both PDGFand Ang II-induced two-dimensional RAoSMCs migration by ∼20% and ∼30%, respectively (Fig. 2A), even though the degree of reduction was lower than those of MMP2 and MMP9. We also performed three-dimensional Boyden chamber assays to confirm these results. As shown in Fig. 2B, treatment with the MMP13specific inhibitor reduced both PDGF- and Ang II-induced RAoSMCs migration by ∼20% and ∼30%, respectively, which were almost the same as those in the two-dimensional assay. In particular, the reduction of Ang II-induced RAoSMCs migration by MMP2, 9, or 13 was almost the same as that in the three-dimensional assay. Furthermore, to investigate the role of MMP13 in MMP2- and MMP9-mediated RAoSMCs migration, we evaluated the effects of co-inhibition of MMP13 activity with MMP2 or MMP9 activities on RAoSMCs migration using the three-dimensional Boyden chamber assay. As shown in Fig. 2C, no significant synergistic effect was observed by co-inhibition of MMP2 and MMP13 activities in PDGFinduced RAoSMCs migration compared with that of the MMP2 inhibitor alone. However, we observed a significant reduction by co-inhibition of MMP9 and MMP13 activities in PDGF-induced RAoSMCs migration compared with that of the MMP9 inhibitor alone. We also observed obvious synergistic effects of co-inhibition in Ang II-induced RAoSMCs migration. Co-inhibition with MMP13 showed a ∼20% increase in the reduction of Ang II-induced cell migration compared with that of the MMP2 or MMP9 inhibitors alone. 3.2. MMP13 silencing decreases PDGF- and Ang II-induced RAoSMCs migration To determine the specific role of MMP13 in RAoSMCs migration, we knocked down MMP13 expression using an MMP13-specific siRNA. We designed three double-stranded, 21-nucleotide siRNAs with 30 TT dinucleotide overhangs against the coding sequence of rat MMP13 (NM 133530). None of the siRNAs shared homology with exons of other known rat genes. The three siRNAs were screened for their effect on MMP13 protein levels using Western blot in order to indicate the most effective silencer for further analysis. As shown in Fig. 3A, MMP13 siRNAs #2 and #3 had no significant effect on MMP13 expression. However, MMP13 siRNA #1 silenced Ang II-induced MMP13 expression by more than 80%, suggesting that MMP13 siRNA #1 could modulate MMP13 expression. The reduction in MMP13 expression levels was maintained up to 72 h after transfection with MMP13 siRNA #1 (data not shown). We confirmed that mRNA expression level of MMP13 was also significantly decreased in MMP13 siRNA-treated RAoSMCs (Fig. 3B). Additionally, we demonstrated the specificity of siRNA #1 for MMP13 by showing no cross-reactivity with MMP2 and 9 (Fig. 3A and B). We further observed that MMP13 secreted into media was significantly decreased by ∼50% in zymographic analysis (Fig. 3C). To investigate whether MMP13 silencing affected RAoSMCs migration, we conducted three-dimensional cell migration assays using transwell chambers. We found that knockdown of MMP13 in PDGF- or Ang II-treated RAoSMCs decreased cell migration by ∼40% and ∼20%, respectively, compared with that in negative siRNA-treated controls (Fig. 3D). In addition, we examined the effect of MMP13 silencing on the survival and proliferative activity of RAoSMCs, because a decrease in survival or proliferation after MMP13 silencing may have contributed to the observed decrease in RAoSMCs migration. We found that MMP13 silencing did not significantly decrease the proliferation rate of RAoSMCs compared with that of control cells (Fig. 3E).
118
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
Fig. 2. Effect of MMP13 inhibition on RAoSMCs migration. (A) Two-dimensional migration assays were conducted with RAoSMCs using a modified wound healing assay in the presence of 5 M OA-Hy, 10 nM inhibitor I, or 20 nM pyrimidine dicarboxamide. The migrated distance during the designated period was measured. Scale bar, 200 m. (B) Three-dimensional migration assays were performed with RAoSMCs using Boyden chambers in the presence of designated MMP inhibitors above. (C) Three-dimensional migration assays were conducted in co-treatment of pyrimidine dicarboxamide with OA-Hy or inhibitor I in RAoSMCs. Columns, average of triple determinations; bars, SD. *P < 0.05 or **P < 0.01 as compared with control.
3.3. MMP13 is downregulated by a known RAoSMCs migration inhibitor, cordycepin Cordycepin is a type of nucleoside analog isolated from Cordyceps militaris, which inhibits VSMC migration via downregulation of MMP2 and MMP9 (Chang et al., 2008; Tuli et al., 2013). Therefore, we used cordycepin to confirm whether MMP13 plays a role in VSMC migration. Similar to previous reports, treatment with PDGF increased intracellular MMP2 and MMP9 expression levels as well as their secreted levels in the medium, which were significantly decreased by cordycepin in a dose-dependent manner. Interestingly, we observed a similar trend in MMP13 expression levels. The secreted level of MMP13 was ∼2-fold higher than that of the untreated control but no significant difference was observed in cell lysates. Treatment with cordycepin decreased MMP13 expression by up to 50% in both the media and cell lysates (Fig. 4A and B). Furthermore, the increased mRNA levels of MMP13 by PDGF treatment were inhibited by cordycepin (Fig. 4C). In addition to the result that silencing MMP13 significantly decreased the migration
of RAoSMCs (Fig. 3D), these results indicate that MMP13 might be one of mediators in VSMCs migration.
3.4. Akt and ERK are potential mediators of MMP13 expression The signal transduction pathways that modulate the activity of MMP transcription factors are highly diverse. Many reports have demonstrated that mitogen-activated protein kinase (MAPK) signal transduction pathways, including p38, ERK, and JNK, are well known mediators that stimulate or inhibit MMP expression depending on the cell types (Johansson et al., 2000; Nakai et al., 2013). To understand the mechanism by which stimuli regulate MMP13 expression in VSMCs, we analyzed the effects of signal mediator inhibitors on MMP13 expression. We found that an Akt or ERK-specific inhibitors, but not PI3K inhibitor, significantly decreased MMP13 expression levels (Fig. 5). These results indicate that Akt and ERK are likely upstream mediators that regulate MMP13 expression and functions in cell migration of VSMCs.
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
119
Fig. 3. MMP13 silencing in RAoSMCs. RAoSMCs were transfected with MMP13 siRNA or control siRNA using siLentFect. After 24 h transfection, cells and media were harvested for Western blotting, RNA extraction, zymographic analysis, or Boyden chamber assays. (A) MMP13 silencing by three different MMP13 siRNAs in RAoSMCs. The same amounts of lysates were analyzed by Western blotting using an antibody against MMP13. Anti-actin blot was used as a loading control. (B) mRNA levels of MMP13 were quantified by real-time PCR after treatment control or MMP13 siRNAs. (C) Extracellular activity of MMP13 after treatment control or MMP13 siRNAs was analyzed using zymography. (D) RAoSMCs migration after treatment control or MMP13 siRNAs was analyzed using Boyden chamber assays. Columns, average of triple determinations; bars, SD. *P < 0.05 or **P < 0.01 as compared with Neg. siRNA-treated control.
4. Discussion In this study, we showed that MMP13 silencing led to a significant decrease in PDGF- and Ang II-induced RAoSMCs migration, which is a key event in the early progression of atherosclerosis (Doran et al., 2008), and provided evidence that MMP13 participated in this process. Although many reports demonstrate that MMP13 is involved in the migration and invasion of various tumor and non-tumor cell types, including esophageal squamous cell carcinoma, cutaneous squamous cell carcinoma, and endothelial cells (Lopez-Rivera et al., 2005; Xu et al., 2012; Ye et al., 2011), the role of MMP13 in migratory VSMCs has been unclear. Our finding that Ang II treatment resulted in a significant increase of MMP13 expression in RAoSMCs is similar to a previous report in which Ang II induced production of MMP3 and MMP13 via the AT1 receptor in osteoblasts (Nakai et al., 2013). In contrast, Tarin et al. showed that a lack of NO inhibits MMP13 and increases ICAM-1 levels in atherosclerosis, leading to worsen atherosclerosis by increasing monocyte adhesion to endothelial cells (Tarin et al., 2009). Therefore, because Ang II is known to decrease the bioavailability of NO, one could hypothesize that an increase in Ang II would result in decreased MMP13. However, it has been shown that NO regulated MMP13 release during wound repair (Lizarbe et al., 2008). The disparity of MMP13 expression and role among these studies may reflect the diverse characteristics of the experimental systems including cell types, stimuli, and signaling pathways
that participate in cellular migration. Nevertheless, we found that PDGF treatment resulted in a significant increase of transcriptional and translational MMP13 levels, indicating that MMP13 may be a mediator of RAoSMCs migration in addition to MMP2 and MMP9. This finding is supported by the fact that inhibition of MMP13 activity led to a significant decrease of RAoSMCs migration and co-inhibition of MMP13 activity with MMP2 or MMP9 showed synergistic effects on the reduction of RAoSMCs migration. We also found that silencing of MMP13 markedly inhibited the transcriptional and translational levels of MMP13 induced by PDGF or Ang II, leading to a decrease in RAoSMCs migration. Although ∼80% of MMP13 expression was silenced by siRNA, the migration rates of RAoSMCs induced by PDGF or Ang II were reduced by only ∼40% and ∼20%, respectively. A potential explanation for this discrepancy is that MMP2 and MMP9 were still functional in the MMP13silenced RAoSMCs. In addition, other modulators may regulate MMP expression and RAoSMCs migration independent of MMP13. Our observation that silencing or inhibition of MMP13 successfully decreased cell migration of PDGF- or Ang II-treated RAoSMCs strongly suggests that MMP13 expression levels are positively correlated with RAoSMCs migration rates. Indeed, these results are coincided with the previous report that B16F1 melanoma grafts displayed reduced tumor growth and strongly decreased metastasis and angiogenesis in a MMP13−/− mouse model compared to wildtype mice (Zigrino et al., 2009). Our results provide strong evidence that reducing MMP13 expression and function may protect
120
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121
Fig. 4. Altered MMP13 expression by cordycepin. (A) Altered MMPs expression by cordycepin was evaluated in media and (B) cell lysates of 5 ng PDGF-treated RAoSMCs using Western blot. (C) The effects of cordycepin on mRNA levels of MMP13 were quantified by real-time PCR in PDGF-induced RAoSMCs. Columns, average of triple determinations; bars, SD. **P < 0.01 as compared with PDGF-treated control.
Fig. 5. Effects of inhibitors on MMP13 expression. The resultant MMP13 expressions in the absence or presence of 5 M LY294002, 2 M Akt inhibitor, or 5 M U0126 were revealed using Western blot. Columns, average of triple determinations; bars, SD. **P < 0.01 as compared with non-treated control.
against early atherogenesis, particularly VSMC migration. Additionally, because MT1-MMP (MMP14) is a known activator of both MMP2 and 13 (Knauper et al., 1996b, 2002), further investigation will be needed to reveal the relationship of MT1-MMP or MMP2 and MMP13 in atherogenesis. In addition, further studies aimed at identifying the precise signal transduction pathways and transcription factors that mediate MMP13 expression may provide a molecular understanding of the roles of MMP13 in atherosclerosis.
Acknowledgement This was supported by grants from the Chosun University Medical Institute, 2012. References Acuff, H.B., Carter, K.J., Fingleton, B., Gorden, D.L., Matrisian, L.M., 2006. Matrix metalloproteinase-9 from bone marrow-derived cells contributes to survival
S.W. Yang et al. / Tissue and Cell 47 (2015) 115–121 but not growth of tumor cells in the lung microenvironment. Cancer Res. 66, 259–266. Chang, W., Lim, S., Song, H., Song, B.W., Kim, H.J., Cha, M.J., Sung, J.M., Kim, T.W., Hwang, K.C., 2008. Cordycepin inhibits vascular smooth muscle cell proliferation. Eur. J. Pharmacol. 597, 64–69. Cheng, C., Haasdijk, R.A., Tempel, D., den Dekker, W.K., Chrifi, I., Blonden, L.A., van de Kamp, E.H., de Boer, M., Burgisser, P.E., Noorderloos, A., Rens, J.A., Ten Hagen, T.L., Duckers, H.J., 2012. PDGF-induced migration of vascular smooth muscle cells is inhibited by heme oxygenase-1 via VEGFR2 upregulation and subsequent assembly of inactive VEGFR2/PDGFRbeta heterodimers. Arterioscler. Thromb. Vasc. Biol. 32, 1289–1298. Chien, Y.C., Sheu, M.J., Wu, C.H., Lin, W.H., Chen, Y.Y., Cheng, P.L., Cheng, H.C., 2012. A Chinese herbal formula “Gan-Lu-Yin” suppresses vascular smooth muscle cell migration by inhibiting matrix metalloproteinase-2/9 through the PI3K/AKT and ERK signaling pathways. BMC Complement Altern. Med. 12, 137. Dey, N.B., Lincoln, T.M., 2012. Possible involvement of cyclic-GMP-dependent protein kinase on matrix metalloproteinase-2 expression in rat aortic smooth muscle cells. Mol. Cell. Biochem. 368, 27–35. Doran, A.C., Meller, N., McNamara, C.A., 2008. Role of smooth muscle cells in the initiation and early progression of atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 28, 812–819. Freije, J.M., Diez-Itza, I., Balbin, M., Sanchez, L.M., Blasco, R., Tolivia, J., Lopez-Otin, C., 1994. Molecular cloning and expression of collagenase-3, a novel human matrix metalloproteinase produced by breast carcinomas. J. Biol. Chem. 269, 16766–16773. Gurjar, M.V., Sharma, R.V., Bhalla, R.C., 1999. eNOS gene transfer inhibits smooth muscle cell migration and MMP-2 and MMP-9 activity. Arterioscler. Thromb. Vasc. Biol. 19, 2871–2877. Halpert, I., Sires, U.I., Roby, J.D., Potter-Perigo, S., Wight, T.N., Shapiro, S.D., Welgus, H.G., Wickline, S.A., Parks, W.C., 1996. Matrilysin is expressed by lipid-laden macrophages at sites of potential rupture in atherosclerotic lesions and localizes to areas of versican deposition, a proteoglycan substrate for the enzyme. Proc. Natl. Acad. Sci. U. S. A. 93, 9748–9753. Inada, M., Wang, Y., Byrne, M.H., Rahman, M.U., Miyaura, C., Lopez-Otin, C., Krane, S.M., 2004. Critical roles for collagenase-3 (MMP13) in development of growth plate cartilage and in endochondral ossification. Proc. Natl. Acad. Sci. U. S. A. 101, 17192–17197. Jiang, M., Bujo, H., Ohwaki, K., Unoki, H., Yamazaki, H., Kanaki, T., Shibasaki, M., Azuma, K., Harigaya, K., Schneider, W.J., Saito, Y., 2008. Ang II-stimulated migration of vascular smooth muscle cells is dependent on LR11 in mice. J. Clin. Invest. 118, 2733–2746. Jiang, M., Bujo, H., Zhu, Y., Yamazaki, H., Hirayama, S., Kanaki, T., Shibasaki, M., Takahashi, K., Schneider, W.J., Saito, Y., 2006. Pitavastatin attenuates the PDGF-induced LR11/uPA receptor-mediated migration of smooth muscle cells. Biochem. Biophys. Res. Commun. 348, 1367–1377. Johansson, N., Ala-aho, R., Uitto, V., Grenman, R., Fusenig, N.E., Lopez-Otin, C., Kahari, V.M., 2000. Expression of collagenase-3 (MMP-13) and collagenase-1 (MMP1) by transformed keratinocytes is dependent on the activity of p38 mitogenactivated protein kinase. J. Cell Sci. 113 (Pt 2), 227–235. Karki, R., Jeon, E.R., Kim, D.W., 2013. Nelumbo nucifera leaf extract inhibits neointimal hyperplasia through modulation of smooth muscle cell proliferation and migration. Nutrition 29, 268–275. Knauper, V., Bailey, L., Worley, J.R., Soloway, P., Patterson, M.L., Murphy, G., 2002. Cellular activation of proMMP-13 by MT1-MMP depends on the C-terminal domain of MMP-13. FEBS Lett. 532, 127–130. Knauper, V., Lopez-Otin, C., Smith, B., Knight, G., Murphy, G., 1996a. Biochemical characterization of human collagenase-3. J. Biol. Chem. 271, 1544–1550. Knauper, V., Will, H., Lopez-Otin, C., Smith, B., Atkinson, S.J., Stanton, H., Hembry, R.M., Murphy, G., 1996b. Cellular mechanisms for human procollagenase3 (MMP-13) activation. Evidence that MT1-MMP (MMP-14) and gelatinase A (MMP-2) are able to generate active enzyme. J. Biol. Chem. 271, 17124–17131. Lizarbe, T.R., Garcia-Rama, C., Tarin, C., Saura, M., Calvo, E., Lopez, J.A., Lopez-Otin, C., Folgueras, A.R., Lamas, S., Zaragoza, C., 2008. Nitric oxide elicits functional MMP13 protein-tyrosine nitration during wound repair. FASEB J. 22, 3207–3215.
121
Lopez-Rivera, E., Lizarbe, T.R., Martinez-Moreno, M., Lopez-Novoa, J.M., RodriguezBarbero, A., Rodrigo, J., Fernandez, A.P., Alvarez-Barrientos, A., Lamas, S., Zaragoza, C., 2005. Matrix metalloproteinase 13 mediates nitric oxide activation of endothelial cell migration. Proc. Natl. Acad. Sci. U. S. A. 102, 3685–3690. Mao, D., Lee, J.K., VanVickle, S.J., Thompson, R.W., 1999. Expression of collagenase-3 (MMP-13) in human abdominal aortic aneurysms and vascular smooth muscle cells in culture. Biochem. Biophys. Res. Commun. 261, 904–910. Meierjohann, S., Hufnagel, A., Wende, E., Kleinschmidt, M.A., Wolf, K., Friedl, P., Gaubatz, S., Schartl, M., 2010. MMP13 mediates cell cycle progression in melanocytes and melanoma cells: in vitro studies of migration and proliferation. Mol. Cancer 9, 201. Mitchell, P.G., Magna, H.A., Reeves, L.M., Lopresti-Morrow, L.L., Yocum, S.A., Rosner, P.J., Geoghegan, K.F., Hambor, J.E., 1996. Cloning, expression, and type II collagenolytic activity of matrix metalloproteinase-13 from human osteoarthritic cartilage. J. Clin. Invest. 97, 761–768. Nakai, K., Kawato, T., Morita, T., Iinuma, T., Kamio, N., Zhao, N., Maeno, M., 2013. Angiotensin II induces the production of MMP-3 and MMP-13 through the MAPK signaling pathways via the AT(1) receptor in osteoblasts. Biochimie 95, 922–933. Neutzner, M., Lopez, T., Feng, X., Bergmann-Leitner, E.S., Leitner, W.W., Udey, M.C., 2007. MFG-E8/lactadherin promotes tumor growth in an angiogenesisdependent transgenic mouse model of multistage carcinogenesis. Cancer Res. 67, 6777–6785. Newby, A.C., 2006. Matrix metalloproteinases regulate migration, proliferation, and death of vascular smooth muscle cells by degrading matrix and non-matrix substrates. Cardiovasc. Res. 69, 614–624. Newby, A.C., 2007. Metalloproteinases and vulnerable atherosclerotic plaques. Trends Cardiovasc. Med. 17, 253–258. Newby, A.C., 2012. Matrix metalloproteinase inhibition therapy for vascular diseases. Vasc. Pharmacol. 56, 232–244. Owens, G.K., 1995. Regulation of differentiation of vascular smooth muscle cells. Physiol. Rev. 75, 487–517. Pan, X., Mao, X., Cheng, T., Zhang, X., 2011. Macrophage migration inhibitory factor: a regulator of MMP13 and inflammation in titanium particles-stimulated air pouch in vivo. Mol. Cell. Biochem. 357, 313–321. Quillard, T., Tesmenitsky, Y., Croce, K., Travers, R., Shvartz, E., Koskinas, K.C., Sukhova, G.K., Aikawa, E., Aikawa, M., Libby, P., 2011. Selective inhibition of matrix metalloproteinase-13 increases collagen content of established mouse atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 31, 2464–2472. Schwartz, S.M., Heimark, R.L., Majesky, M.W., 1990. Developmental mechanisms underlying pathology of arteries. Physiol. Rev. 70, 1177–1209. Sukhova, G.K., Schonbeck, U., Rabkin, E., Schoen, F.J., Poole, A.R., Billinghurst, R.C., Libby, P., 1999. Evidence for increased collagenolysis by interstitial collagenases1 and -3 in vulnerable human atheromatous plaques. Circulation 99, 2503–2509. Tarin, C., Gomez, M., Calvo, E., Lopez, J.A., Zaragoza, C., 2009. Endothelial nitric oxide deficiency reduces MMP-13-mediated cleavage of ICAM-1 in vascular endothelium: a role in atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 29, 27–32. Tuli, H.S., Sharma, A.K., Sandhu, S.S., Kashyap, D., 2013. Cordycepin: a bioactive metabolite with therapeutic potential. Life Sci. 93, 863–869. Xu, N., Zhang, L., Meisgen, F., Harada, M., Heilborn, J., Homey, B., Grander, D., Stahle, M., Sonkoly, E., Pivarcsi, A., 2012. MicroRNA-125b down-regulates matrix metallopeptidase 13 and inhibits cutaneous squamous cell carcinoma cell proliferation, migration, and invasion. J. Biol. Chem. 287, 29899–29908. Ye, Q., Yan, Z., Liao, X., Li, Y., Yang, J., Sun, J., Kawano, T., Wang, X., Cao, Z., Wang, Z., Huang, L., 2011. MUC1 induces metastasis in esophageal squamous cell carcinoma by upregulating matrix metalloproteinase 13. Lab. Invest. 91, 778–787. Yoon, S., Kuivaniemi, H., Gatalica, Z., Olson, J.M., Buttice, G., Ye, S., Norris, B.A., Malcom, G.T., Strong, J.P., Tromp, G., 2002. MMP13 promoter polymorphism is associated with atherosclerosis in the abdominal aorta of young black males. Matrix Biol. 21, 487–498. Yu, T., Wu, Y., Helman, J.I., Wen, Y., Wang, C., Li, L., 2011. CXCR4 promotes oral squamous cell carcinoma migration and invasion through inducing expression of MMP-9 and MMP-13 via the ERK signaling pathway. Mol. Cancer Res. 9, 161–172. Zigrino, P., Kuhn, I., Bauerle, T., Zamek, J., Fox, J.W., Neumann, S., Licht, A., SchorppKistner, M., Angel, P., Mauch, C., 2009. Stromal expression of MMP-13 is required for melanoma invasion and metastasis. J. Invest. Dermatol. 129, 2686–2693.
