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Am J Physiol Heart Circ Physiol 288: H3001-H3005, 2005. First published February 4, 2005; doi:10.1152/ajpheart.01002.2004
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Podosome-mediated matrix resorption and cell motility in vascular smooth muscle cells

Gerald Burgstaller1 and Mario Gimona1,2

1Institute of Molecular Biology, Department of Cell Biology, Austrian Academy of Sciences, Salzburg, Austria; and 2Consorzio Mario Negri Sud, Department of Cell Biology and Oncology, Santa Maria Imbaro, Italy

Submitted 1 October 2004 ; accepted in final form 29 January 2005


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The migration of vascular smooth muscle cells (VSMCs) is a principal factor for the development and progression of vascular diseases. In addition, phenotypic alteration from the contractile (differentiated) to the synthetic (dedifferentiated) state and a proteolytic process in the form of extra cellular matrix degradation are necessary for SMC invasion. The actual mechanism leading to the focal degradation of basement membrane matrix components and, hence, SMC migration within the tissue itself is, however, unclear. In response to phorbol ester [phorbol-12,13-dibutyrate (PDBu)], VSMCs in culture form podosomes, dynamic organelles critical for cell adhesion and substrate degradation that are typically found in invasive cells and cells that cross tissue boundaries. Here, we show that PDBu-stimulated VSMCs resorb the extracellular matrix at the sites of podosomes. Podosome formation correlates with an increased polarization of VSMCs on fibronectin- or collagen-coated flexible substrates in addition to a concomitant induction of cell motility. VSMCs embedded in reconstituted basement membrane support adopt the typical spindle-shaped morphology of differentiated SMCs in vivo and, after PDBu treatment, form peripheral lamellipodia and podosomes around their matrix-contacting surface. Our findings demonstrate that podosome formation is the potential mechanism underlying the ability of VSMCs to traverse the surrounding basement membrane and escape the barrier of the tunica media in vascular diseases.

podosomes; matrix resoption; smooth muscle; motility

ATHEROGENESIS causes smooth muscle cells (SMCs) to change their phenotype, which is accompanied by a loss of contractile elements and the acquisition of replicative and migratory potential (5). The ultimate consequence of this phenotypic modulation is the invasion of the intimal layer of the artery, inappropriate replication within the arterial wall, and the deposition of fibrotic connective tissue (27). Whereas the concept that SMCs need to remove the barrier formed by the basal lamina [primarily via the elevated production and secretion of matrix metalloproteases (MMPs)] before engaging in motile activities has been established earlier (12, 22), no definite answers to the question of how this activity would be applied in a spatially controlled manner have been supplied.

We have demonstrated recently that the podosomes formed in rat aortic vascular SMCs (VSMCs) are structurally and with respect to their molecular composition similar to the podosomes of monocytes (13). Podosomes are unique actin-rich adhesion structures found in macrophages and osteoclasts (for a review, see Ref. 16). Strikingly, cancer cells also develop podosomes when stimulated with oncogenic agents or activated Src kinases and might employ these adhesive structures to migrate to new sites in the body (2, 3). Thus podosome development not only significantly contributes to the regulation of bone strength but also to the frequency of metastasis.

Differentiated SMCs appear to retain a substantial level of plasticity, and cultured SMCs can readily change their gene expression pattern to a nonmuscle type within 48 h (10, 28). The formation of podosomes in response to a tumor-promoting agent like phorbol ester may represent a reversible dedifferentiation process that could occur in vivo under specific conditions. Our previous studies (4, 11, 13) imply that podosome formation, induced by the tumor-promoting phorbol ester phorbol-12,13-dibutyrate (PDBu), may indeed activate the matrix-resorbing and motility-stimulating potential of VSMCs in vitro.

Here, we investigated the potential of vascular smooth muscle podosomes to mediate cell motility and resorption of extracellular matrix (ECM) components. Our findings shed new light on the potential correlation between smooth muscle phenotypic plasticity and dedifferentiation and the development of malignant alterations of smooth muscle such as atherosclerosis.


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Cell culture, transfection, and immunofluorescence microscopy. A7r5 rat smooth muscle cells (American Type Culture Collection) were grown in low-glucose (1,000 mg/l) DMEM without phenol red supplemented with 10% FBS (PFA) and penicillin-streptomycin (GIBCO-BRL) at 37°C and 5% CO2. For transient expression, cells were transfected using Superfect (Qiagen) at 70% confluence, essentially as described (14). Cells were replated onto 15-mm coverslips 16 h posttransfection and prepared for immunofluorescence microscopy after an additional 48 h. Cells were washed three times in PBS, extracted in 3.7% paraformaldehyde (PFA)-0.3% Triton X-100 in PBS for 5 min, and fixed in 3.7% PFA (Merck) in PBS for 30 min. Fluorescence images were recorded on a Zeiss Axioscop equipped with an Axiocam driven by the manufacturer's software package (Zeiss) using a x63 oil immersion lens.

Confocal microscopy and volume rendering. Stacks of optical sections (z-step = 0.1 µm) were captured using a confocal spinning disk system (QLC100 confocal head from Visitech) mounted on a Zeiss Axiovert 100M motorized microscope (Zeiss) equipped with a x63 objective, numerical aperture 1.4, at exposure times of 200–1,000 ms using 458- and 568-nm laser excitation. Images were acquired using a 512 x 512-pixel MicroMax camera (Princeton Instruments) driven by IPLab version 3.5.5 software (Scanalytics) running on a Macintosh G4 computer. Collected confocal stacks were processed using the freeware ImageJ 1.32 software (http://rsb.info.nih.gov/ij/) running on a Macintosh G4 computer. For volume rendering of confocal stacks, we used the software plugin VolumeJ 1.7 developed by M.D. Abramoff (http://bij.isi.uu.nl/) for ImageJ 1.32.

Flexible substrates. Flexible substrates were prepared according to the protocol of Wang (32) (http://ylwang.umassmed.edu/protocol/mc/pasubs.htm). For our experiments, we used stiff matrix (final acrylamide-bisacrylamide concentration 8:0.1%, exhibiting a Young's modulus of >75 kN/m2). Flexible substrates were coated and cross-linked with fibronectin (FN; Sigma Aldrich) at a final concentration of 50 µg/ml in PBS or collagen type I (Vitrogen) at a final concentration of 200 µg/ml in water. The crosslinking agent Sulfo-Sanpah was purchased from Pierce.

ECM degradation assay. ECM degradation assays were performed essentially as described in detail by Baldassarre et al. (2). Briefly, 15-mm coverslips were coated with Alexa488 (Molecular Probes)-labeled FN (Sigma Aldrich) at a final concentration of 50 µg/ml (in 2 M urea) in PBS and incubated for 30 min at room temperature. Cells were plated and treated as described above. For podosome stimulation, 1 µM PDBu was added to cells in complete growth medium (10% FCS in DMEM without phenol red) for 30 min at 37°C. Cells were fixed in 3.7% PFA 30–60 min after the addition of PDBu, and the cells were processed for immunofluorescence microscopy as described above. Images of cells with prominent podosomes were analyzed for matrix degradation using individual single-channel images and resulting overlay images from multichannel immunofluorescence microscopy imaged with the Axiovision software (Zeiss). All experiments were repeated at least five times.


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VSMC migration is greatly enhanced by the production of MMPs (8, 9, 12, 18), and rat SMCs require active MMP-2 for the invasion of basement membrane support in vitro (8, 24). Invadopodia of metastasizing cancer cells and podosomes of osteoclasts serve a role in coordinating the spatial secretion of these enzymes (2, 3, 16). As a first attempt to validate the importance of podosome formation for smooth muscle pathophysiology, we therefore determined the matrix-resorbing potential of cultured VSMCs. A7r5 rat VSMCs stained with fluorescent phalloidin to visualize F-actin-rich structures were allowed to grow on glass coverslips coated with Alexa488-conjugated FN. Under normal serum conditions, the rounded cells showed substantial rearrangement of the underlying FN matrix in the cell center, due to the normal contractile activity of the A7r5 cells (Fig. 1A, a–c). After the application of PDBu, the cells locally degraded the FN matrix at and around podosomes, footprinting the dynamic appearance of podosomes during the time from PDBu application to the point of fixation (Fig. 1A, d–f). This finding signifies that matrix degradation in VSMCs has a local constraint and is confined to podosomes (Fig. 1B). This result is in agreement with the findings that invadopodia of cancer cells are directly responsible for matrix degradation in vitro (2, 7) and suggests that a similar mechanism may be functioning in VSMCs upon dedifferentiation and phenotypic modulation. Remarkably, both cancer cells and PDBU-stimulated VSMCs also employ p190RhoGAP for the formation of focal matrix degradation structures (4, 21, 23).



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  		Fig. 1. A: vascular smooth muscle cells (VSMCs) remodel and resorb extracellular matrix. a–c: Cells on fluorescently labeled fibronectin (FN)-coated glass support display a prominent actin stress fiber network. Contractile forces lead to rearrangement of the matrix in the cell center but not in the cell periphery. Note the almost homogenous fluorescent matrix in the area of actin stress fibers arranged perpendicularly to the cell axis (bracket). d–f: Peripheral podosomes form after 30 min of phorbol-12,13-dibutyrate (PDBu) treatment. Induced cytoskeletal rearrangement and increased contractility cause a further rearrangement of the matrix in the cell center in addition to the degradation at regions of podosome formation. B: enlarged images of cells shown in A, d–f. Matrix removal is evident in regions of podosome formation. Beginning degradation by the formation podosomes (arrows) and regions of completed matrix degradation where podosomes have already been disassembled (arrowheads) can be identified. The actin cytoskeleton was visualized by Alexa568 phalloidin. Bar = 20 µm.

 
The degradation of the matrix appeared restricted to the cell periphery and the leading lamella in migrating VSMCs, coinciding with the region of podosome formation, but was never observed in the perinuclear region, as is generally seen for invadopodia of cancer cells (2). Degradation of the matrix was also clearly discernable from the rearrangement of the matrix, which is observed in the cell center. As seen in Fig. 1A, only those actin filaments directed toward the cell center contributed to the matrix rearrangement, whereas filaments lying perpendicular to the cell axis exerted no contractile forces and caused no matrix reorganization. Strikingly, in our experimental cell system, the majority of podosomes were induced already 30–60 min after the addition of PDBu. Hence, our assay for matrix degradation was established within the same time frame, whereas the commonly used time for matrix degradation assays for cancer cells (and using cross-linked gelatin as a substrate) averages 15 h (2). This narrow time window is also way below what can be envisaged for the de novo synthesis, maturation, and secretion of MMPs. A possible explanation for this phenomenon may be the storage of MMPs in the vicinity of focal adhesion sites where podosome formation is initiated, but experimental evidence for such a scenario is currently missing. Also, the VSMCs used herein were unable to degrade FN or gelatin that had been chemically cross-linked with glutaraldehyde (not shown). It has been noted before by Baldassarre et al. (2) that such insoluble matrix is only sufficiently dissolved by highly aggressive cancer cells, which form invadopodia that remain stationary for several hours. Finally, our assays using fluorescently labeled collagen did not demonstrate consistent degradation, albeit some degradation in the vicinity of podosomes was detected (data not shown).

The commonly observed morphology of an A7r5 cell on glass is characterized by a round shape, prominent actin stress fibers, and large focal adhesions (Fig. 2A). No change in phenotype was observed when the glass support was coated with cross-linked FN or type 1 collagen. Although PDBu induced podosome formation in VSMCs on glass, it failed to trigger cell polarization and net-directional movement (not shown). The morphological differences between primary VSMCs and A7r5 cells are thus not solely explained by the type of matrix support provided. Recently, "cytoskeletal prestress" (26) and mechanical stiffness properties of the ECM have been brought into the discussion of VSMC regulation. It has also been noted previously that the cellular response to physical parameters such as substrate stiffness or flexibility may represent a key mechanism in the interactions with the cell's surrounding environment (17, 25). Thus it was important to investigate how flexible substrates, which would allow a higher level of contractility to be built up in the cells, would impact on the general morphology. When allowed to spread on flexible polyacrylamide substrates coated with either FN or type 1 collagen, A7r5 VSMCs rapidly adopted an elongated shape (Fig. 2A). This morphology was retained after PDBu stimulation, and the cells now featured additionally a pronounced leading edge with active lamellipodia and a trailing edge structure, indicative of directional cell motility. Podosome formation was mainly confined to the region of active actin cytoskeleton remodeling in the region of lamellipodia formation.



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  		Fig. 2. Phenotypic modulation and induction of cell motility in VSMCs on flexible substrates. A, 1: VSMCs on glass exhibit a round shape. A, 2: on collagen type 1 (Coll-1)-coated flexible polyacrylamide substrates (Flex + Coll-1), the cells change their phenotype to an elongated appearance. A, 3: addition of PDBu induces further cell polarization and podosome formation in the region of the leading edge. Bars = 10 µm. B: live cell phase-contrast imaging on FN-coated flexible substrates (Flex + FN). 1: 50-min PDBu; 2: 92-min PDBu; 3: 102-min PDBu; 4: 150-min PDBu. Polarized cells display directional movement upon the addition of PDBu and continue to form transient podosomes in the leading lamella (arrows in the inset in 2). See also the supplementary video sequence (http://ajpheart.physiology.org/cgi/content/full/01002.2004/DC1).

 
Wang and colleagues (17) concluded that changes in tissue rigidity and strain could play an important controlling role in a number of normal and pathological processes involving cell locomotion. Live phase-contrast video imaging of cells on FN-coated flexible substrate confirmed that, in addition to driving podosome formation, cells treated with phorbol ester developed increased cell polarization and motility (Fig. 2B). Identical results were obtained when flexible substrates were coated with type 1 collagen instead of FN.

These results strongly indicated that the differentiated SMC phenotype is not only influenced by the type of ECM but also by the ability to exert forces and to undergo contraction. In contrast to fibroblasts or epithelial cells, SMCs in vivo are contacting the basement membrane not only at the ventral surface but are rather encapsulated by basal membrane. Aguilera et al. (1) have provided the first study that directly correlated the degradation of basement membrane with the migration and proliferation of VSMCs. Along the same lines, it was shown that the basement membrane can inhibit both proliferation and migration of VSMCs (22). More specifically, the basement membrane has been suggested to both provide a physical barrier (or "cage") for migration and, in addition, supplies a "break" on signaling processes. Removal of these constraints to movement and the formation of productive cell-matrix interactions are essential for migration. VSMCs lose their basement membrane before migrating to the neointima, and the basement membrane has been shown to inhibit phenotypic changes of VSMCs, and inhibitors of MMPs inhibit VSMC migration. Thus MMP-mediated loss of basement membrane appears tightly related to the induction of cell migration, whereas VSMC proliferation appears not to require basement membrane degradation via MMPs. Thus it has been concluded that proliferating cells might use other proteases to degrade the surrounding basement membrane (or specific components of it).

The resulting consequence of these considerations was to place VSMCs into a three-dimensional matrix support, which mimics the natural environment of the surrounding basal lamina in vessel tissue. When VSMCs are embedded in reconstituted basement membrane matrix (Matrigel), they rapidly establish a network of spindle-shaped cells (Fig. 3A). A similar end-to-end alignment of elongated spindle-shaped cells with a tubelike morphology, tapered ends, and a central nucleus, and forming interconnected strands or cords, has been observed previously (29) in freshly isolated aorta SMCs embedded in Matrigel. The actin filaments of VSMCs in Matrigel are much thinner and less prominent compared with those of cells grown on a rigid glass surface (Fig. 3B). They align along the longitudinal axis, in the periphery of the cells (insets in Fig. 3b). Within 30 min after the addition of PDBu, these spindle-shaped cells also develop numerous ruffle-like structures at the cell periphery and, in addition, podosome-like complexes, which emanate from the cell periphery (inset in Fig. 3C). Volume rendering of the three-dimensional reconstructions from a stack of confocal images reveals the typical cone shape of these actin-rich clusters (Fig. 3D).



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  		Fig. 3. Spindle-shaped VSMCs form podosomes in three-dimensional basement membrane support. A: networks of interconnected cells formed in Matrigel as seen by phase-contrast microscopy. The elongated cells connect nodal points of less expanded cells. B, left: confocal fluorescence image of a VSMC in Matrigel stained with phalloidin to visualize F-actin. B, right: actin bundles are arranged along the longitudinal axis of the spindle-shaped VSMCs. Insets 1–4, selected cross sections of a resliced confocal stack. The positions are indicated. For better visibility of the fragile actin filaments (seen as black dots), the contrast in the images of the insets was inverted using Adobe Photoshop. C: stimulation with PDBu induces podosome formation also in cells embedded in soft, flexible extracellular matrix support. Individual podosomes can be seen in the enlarged area. D: volume rendering reveals the characteristic cone shape of the podosomes shown in C. The selected images are rotated sequentially by 60° from one image to the next.

 
Podosomes and invadopodia play an elementary role in many different aspects of cellular invasion of tissues. Substrate degradation of osteoclasts and tumor cells is confined to punctate areas corresponding in both size and location to the appearance of podosomes (3, 6, 19). This work provides explanations for the complex mechanisms underlying SMC invasion in the course of the development of atherosclerotic lesions and restenosis. Phenotypic plasticity of SMCs paired with the rapid and transient formation of podosomes supports the concept that matrix degradation at these specialized sites, and the concomitant engagement of the molecular machinery, initiating actin-based cell motility, drive tissue invasion in this cell type. Strikingly, phenotypic plasticity (31), SMC migration, and podosome formation (11) are suppressed by normal levels of calponin h1 expression via the ability of calponin to regulate actin filament turnover (15). The most recent identification of podosomes in epithelial (30) and endothelial cells (20) further points toward a more common use of equivalent structures in a variety of pathophysiological conditions.

Clearly, there are two important factors that contribute to the formation of an atherosclerotic plaque in the vasculature. Besides the discussed migratory behavior of VSMCs undergoing phenotypic modulation, an increased proliferative activity is required to mediate growth of the plaque structure. As shown by Aguilera et al. (1), the requirement of these two processes with respect to the degradation of the basement membrane are quite divergent and may thus represent independent yet complementary processes. These observations are in good agreement with our present observations as we could not observe the induction of proliferative activity before the formation of podosomes (not shown); however, the short time window of a few hours used in this study may have been insufficient to draw clear conclusions on this issue.

Taken together, the present study highlights the potential of cultured immortalized VSMCs to degrade the ECM before or during engagement of cellular motility. Confirmation of these data in smooth muscle tissue will be the challenging next step. Experiments aiming at the confirmation of these data in smooth muscle tissue and primary SMCs are currently underways.


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This study was supported in part by grants from the Austrian Science Foundation and by Marie Curie Excellence Grant MEXT-CT-2003-002573 of the European Union.


   ACKNOWLEDGMENTS
 
We thank Ulrike Tischler for expert technical assistance and Dr. Roberto Buccione (CMNS, Italy) for insightful suggestions regarding matrix degradation experiments.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. Gimona, Consorzio Mario Negri Sud, Dept. of Cell Biology and Oncology, Unit of Actin Cytoskeleton Regulation, Via Nazionale 8a, I-66030 Santa Maria Imbaro, Italy (E-mail: gimona{at}dcbo.negrisud.it)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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