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Vol. 19, Issue 5, 2051-2058, May 2008
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Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, TX 75390-9039
Submitted September 18, 2007;
Revised February 6, 2008;
Accepted February 19, 2008
Monitoring Editor: Yu-Li Wang
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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1-mm gap between paired dermal equivalents in floating, nested collagen matrices. Our findings demonstrate that when fibroblasts interact with collagen matrices, tractional force exerted by the cells can couple to matrix translocation as well as to cell migration. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Mitchison and Cramer, 1996; Galbraith and Sheetz, 1997; Beningo et al., 2001; Ridley et al., 2003). The extensive body of research underlying the multistep model is based on studies with tissue cells on extracellular matrix (ECM)-coated planar surfaces including rigid materials such as glass or plastic coverslips as well as flexible materials such as polyacrylamide. On ECM-coated planar surfaces, cells can modulate their cytoskeletal function and adhesion strength in response to surface mechanics (Balaban et al., 2001; Discher et al., 2005; Ingber, 2006; Vogel and Sheetz, 2006). However, adsorbed or covalently attached ECM molecules tend to be held in register with each other with little capacity to undergo cell-mediated mechanical and molecular reorganization.
Type 1 collagen is the major protein component of fibrous connective tissues. These connective tissues provide mechanical support and frameworks throughout the body, and fibroblasts are the cell type primarily responsible for their biosynthesis and remodeling. Three-dimensional (3D) matrices prepared with type I collagen exhibit mechanical properties that resemble connective tissue (Barocas et al., 1995; Wakatsuki et al., 2000; Roeder et al., 2002; Silver et al., 2002; Ahlfors and Billiar, 2007). Unlike ECM-coated material surfaces, fibroblasts can mechanically remodel collagen matrices both locally and globally (Brown et al., 1998; Tomasek et al., 2002; Grinnell, 2003; Petroll, 2004; Tranquillo, 1999. Such mechanical remodeling of connective tissue ECM is believed to be important for tissue homeostasis (Silver et al., 2002; Wiig et al., 2003; Goldsmith et al., 2004; Langevin et al., 2004), aging (Varani et al., 2004), repair (Tonnesen et al., 2000; Tomasek et al., 2002; Grinnell, 2003), fibrosis (Eckes et al., 2000; Desmouliere et al., 2005), and tumorigenesis (Beacham and Cukierman, 2005; Gaggioli et al., 2007; Yamada and Cukierman, 2007).
Cells interacting with 3D collagen matrices exhibit distinct patterns of cell signaling (Cukierman et al., 2002; Wozniak et al., 2003; Beningo et al., 2004; Rhee et al., 2007) and increased plasticity of cell migration (Sahai and Marshall, 2003; Shreiber et al., 2003; Friedl, 2004; Even-Ram and Yamada, 2005; Zaman et al., 2006; Wolf et al., 2007). We have been studying human foreskin fibroblast migration in nested collagen matrices (Grinnell et al., 2006). To prepare nested matrices, contracted collagen matrices known as dermal equivalents (Bell et al., 1979) are polymerized within cell-free outer matrices. Cells can migrate from the dermal equivalents into the outer matrices. Platelet-derived growth factor (PDGF) is unique among growth factors in its capacity to promote human fibroblast migration in nested collagen matrices, whereas sphingosine-1-phosphate acts as an inhibitor of migration (Jiang et al., 2008).
Time-lapse microscopic observations of fibroblasts migrating in nested collagen matrices showed that in addition to cell migration, collagen fibril flow occurred in the outer matrix toward the dermal equivalent boundary. Features of this flow suggested that it depends on the same cell motile machinery normally used by cells for migration. Collagen fibril flow was capable of producing large-scale tissue translocation as shown by closure of a 1-mm gap between paired dermal equivalents in floating, nested collagen matrices. Our findings demonstrate that tractional force exerted by fibroblasts in collagen matrices can couple differentially to cell migration or matrix translocation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell Culture and Preparation of Nested Collagen Matrices
Early passage human foreskin fibroblasts (hTERT-immortalized; Rhee et al., 2007) were cultured in DMEM supplemented with 10% FBS at 37°C in a 5% CO2 humidified incubator. Cell migration in nested collagen matrices was performed as described previously (Figure 1A; Grinnell et al., 2006). Briefly, dermal equivalents were formed by contraction for 6–48 h of 200-µl collagen matrices (1.5 mg/ml collagen, 2 x 105 cells/matrix). Subsequently, dermal equivalents were embedded in 200 µl outer collagen matrices containing 1.5 mg/ml collagen (Figure 1B). Nested matrices were incubated in DMEM (or CO2-independent DMEM for time-lapse studies) containing 5 mg/ml BSA and 50 ng/ml PDGF or other growth factors as indicated.
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14-mm inner diameter). Paired nested matrices contained two dermal equivalents that were placed 1 mm apart on a 60-µl cushion of outer matrix and then covered with the remaining 140 µl of outer matrix collagen solution. Overall appearance of nested collagen matrices was recorded using an Epson 4870 photo scanner (Epson America, Long Beach, CA).
Time-Lapse Microscopy
Nested collagen matrices in 24-well culture dishes were placed in an environmental chamber at 37°C, and time-lapse microscopy was carried out using a Zeiss Axiovert 200M inverted microscope equipped with an A-PLAN 10x/0.25 PH1 Zeiss objective (Carl Zeiss MicroImaging, Thornwood, NY) and a Hamamatsu model Orca 285 CCD camera (Hamamatsu, Bridgewater, NJ). Z stack images were acquired at 20-µm steps and 5–10-min intervals using Openlab 4.02 (Improvision, Lexington, MA) software. The Z stack range was 800 µm to ensure complete coverage of the range of cell movement, which typically occurred within 100–200 µm. Each Z plane was reconstructed into a single 2D video. For quantitative analysis of cell migration and collagen translocation, the image field was calibrated with a micrometer slide, and Openlab software was used to measure displacements. In some experiments, reorganization of the collagen matrix was analyzed by visualizing the distribution of 3- or 6-µm beads embedded in the matrix. In initial experiments, we used beads coated with BSA. In later experiments, we added beads directly as supplied by the manufacturer, which was simpler because a sonication step was not required, and no differences were observed compared with albumin-coated beads.
Immunofluorescence Microscopy
Preparation of samples for actin staining with Alexa Fluor 594–conjugated phalloidin and PI was carried out as previously described (Grinnell et al., 2006). Images of PI-stained cells used for quantification of cell migration were collected with a Nikon Elipse 400 fluorescence microscope (Melville, NY) and 10x/0.45 Nikon Plan Apo infinity corrected objective using a Photometrics SenSys CCD camera (Tucson, AZ) and MetaVue acquisition and imaging software (Molecular Devices, Menlo Park, CA). The cell migration index was calculated by counting the average number of cells that had migrated out of dermal equivalents in four 10x microscopic fields selected arbitrarily. Each field included the border of the dermal equivalent (detected by dark field microscopy) and the furthest moving cells (detected by nuclear staining with PI).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Migrating cells all were localized within the outer collagen matrix. No direct interaction occurred between migrating cells and the underlying culture dish on which nested collagen matrices were restrained.
Collagen Translocation in Nested Collagen Matrices
Figure 2A presents representative, phase-contrast images from a video (Sup_1.mov) in which individual cells can be seen migrating from the dermal equivalent into the outer matrix. Migrating cells typically had leading dendritic extensions that were branched. Some of these extensions increased in size and became stabilized during migration; others regressed. In addition to cell migration, collagen translocation occurred in the outer matrix toward the dermal equivalent interface. Collagen movement can be appreciated by noting the position of matrix deformations (Figure 2A, asterisks). In the video, collagen translocation has the appearance of collagen fibril flow.
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The decrease in velocity of collagen translocation might have been a feature of the experimental system independent of the time when cell migration began. To test this possibility, studies were carried out using nested collagen matrices prepared with dermal equivalents that had been contracted for 48 h. With 48-h dermal equivalents, a longer lag phase precedes cell migration compared with the 6-h dermal equivalents used for the experiments described in Figures 1 and 2. Figure 3A shows representative images from the beginning and end of a time-lapse video (Sup_2.mov). During the 15-h incubation, little cell migration across the interface occurred. As can be seen in the video, collagen flow took place during this period. Polystyrene beads (3µm) in the collagen help visualize collagen flow. Quantitative measurements (Figure 3B) demonstrate that collagen translocation in the absence of cell migration reached a higher velocity than observed in Figure 2 and declined later.
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). Figure 4 shows that blebbistatin, which blocks myosin II activity, completely prevented both cell migration and collagen translocation. Y27632, which blocks Rho kinase activity, partially inhibited cell migration and collagen translocation. These findings were consistent with the idea that cells use the same motile machinery for collagen translocation as for cell migration.
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–4 all were made using nested collagen matrices restrained on culture dish surfaces. We and others have shown that fibroblast physiology and morphology differ markedly when fibroblasts interact with collagen matrices floating in culture medium versus restrained on culture dishes (Grinnell, 2003). Therefore, we compared cell migration with nested collagen matrices that were restrained on culture dishes or floating in culture medium. Figure 5A shows fluorescence images of samples stained by PI, and Figure 5B presents quantification of the results. If nested collagen matrices were floating in medium, then the cell migration index was reduced markedly compared with restrained conditions.
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1 mm apart in outer matrices. Figure 6A shows the experimental design and presents immunofluorescence images taken at the edge and gap regions of the paired matrices after culture for 24 h. Figure 6B shows quantification of the findings. With restrained, paired nested matrices, cell migration occurred to a similar extent around the edges of the dermal equivalents and in the gap between. With floating, paired nested matrices, migration was limited almost entirely to the gap region in between. Taken together, the findings in Figures 5 and 6 suggested that fibroblast migration in collagen matrices depended on the ability of the matrix to resist tractional force exerted by the cells. Resistance could be provided by restraint of nested matrices on culture surfaces or by the opposition between paired dermal equivalents if the nested matrices were floating in culture medium.
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). Additional experiments were carried out to compare the effects of these agonists on gap closure with paired, nested collagen matrices. Figure 8C shows that neither LPA- nor FBS-stimulated gap closure. Also, no gap closure occurred if paired, nested matrices were restrained. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Grinnell, 2003). Remodeling also has been studied by observing local changes in collagen organization in response to cell motile activity (Roy et al., 1997; Tamariz and Grinnell, 2002) and cell migration (Gaggioli et al., 2007; Wolf et al., 2007). In addition, if fibroblast explants are embedded in collagen matrices, then collagen fibrils in between the explants become aligned into linear tracks (Stopak and Harris, 1982; Sawhney and Howard, 2002). In the current article, we describe another aspect of mechanically remodeling of collagen matrices: collagen fibril flow and large-scale tissue translocation. In the case of floating nested matrices, collagen fibril flow was sufficient to permit closure of a 1-mm gap between paired dermal equivalents.
Under some circumstances, collagen flow has been reported to occur within collagen matrices independent from tissue cells (Newman et al., 1985). The collagen flow we detected in the current studies required fibroblasts and appeared to utilize the same cell motile machinery involved in cell migration. Interfering with actin or inhibiting myosin II or Rho kinase inhibited cell migration and collagen translocation. In addition, the growth factor specificity of collagen flow was similar to cell migration. Gap closure in floating, paired nested matrices took place in the presence of PDGF but not LPA or FBS. Previously, we showed that PDGF but not LPA or FBS stimulates human fibroblast migration in nested collagen matrices (Grinnell et al., 2006). Serum frequently has been used as an agonist to study cell migration, and PDGF is believed to be the major promigratory factor for fibroblasts in serum (Li et al., 2004; Gao et al., 2005). However, serum also contains the lipid growth factor sphingosine-1-phosphate (Eichholtz et al., 1993; Yatomi et al., 1997). S1P recently was shown to be a potent inhibitor of human fibroblast migration in collagen matrices and to reduce the promigratory activity of serum (Jiang et al., 2008).
Evidence regarding the regulatory role that Rho kinase plays in cell migration is complex. Blocking Rho kinase activity can inhibit or stimulate cell migration depending on cell type and experimental conditions (Riento and Ridley, 2003; Totsukawa et al., 2004). Besides for collagen fibril flow and cell migration, Rho kinase activity also is required for PDGF-stimulated fibroblast-collagen matrix contraction (Rhee and Grinnell, 2006). Human fibroblasts in collagen lack detectable Rho activation upon PDGF stimulation (Grinnell et al., 2003). Therefore, basal rather than agonist-stimulated Rho kinase probably is required for cell migration and contraction, perhaps by maintaining basal levels of myosin light chain phosphorylation (Abe et al., 2003; Knock et al., 2008).
Collagen fibril flow not only appeared to depend on the same cell motile machinery as cell migration, but also tended to occur reciprocally with migration. That is, collagen translocation was greatest before cells began to migrate in the outer matrices. Using more contracted dermal equivalents to extend the period before cell migration begins increased the period of collagen flow. Why collagen flow eventually decreased under the latter conditions even in the absence of cell migration remains to be determined. In the case of floating nested matrices, we observed decreased cell migration and increased collagen flow. Collagen flow in paired nested collagen matrices resulted in dermal equivalent gap closure if the matrices were floating.
Why there is a lag phase before cell migration begins, and why the length of the lag phase increases along with the time of dermal equivalent contraction are questions yet to be resolved. One possibility is that during the lag phase, changes occur in the cells or matrix required for fibroblasts to move from across the interface between the dermal equivalent whose collagen density can be as high as 25 mg/ml collagen (Ahlfors and Billiar, 2007) into the outer collagen matrix composed of 1.5 mg/ml collagen. On collagen-coated planar surfaces, cells tend not to move from stiffer to softer materials (Lo et al., 2000). Whatever the final explanation, we believe that the studies with restrained compared to floating nested matrices indicate an important role for tension. Fibroblasts in floating collagen matrices have fewer stress fibers and focal adhesions than cells in restrained matrices (Grinnell, 2003). Based on the finding that cellular stress fibers and focal adhesions are indicators of cell tension (Singer et al., 1984; Burridge et al., 1988; Balaban et al., 2001; Galbraith et al., 2002), fibroblasts are less able to develop tension in floating compared with restrained matrices. Given that tension plays a positive role in cell migration on planar surfaces (Tucker et al., 1985; Kolega, 1986; Beloussov et al., 2000; Lo et al., 2000; Wang et al., 2001; Raeber et al., 2007), the decreased ability of cells in floating nested collagen matrices to develop tension (unless the dermal equivalents are paired) provides a reasonable explanation for the lack of migration.
Figure 9 offers what we believe to be an attractive hypothesis to account for our overall observations. If the collagen matrix can resist cellular tractional force, then the cells can move. If the matrix cannot resist cellular traction force, then the matrix moves. Likewise, when tissue cells attempt to spread on a silicone oil surface, if the viscosity of the oil is not sufficient to resist the pull of cell extensions, then the oil flows and the cells remain round (Harris, 1973).
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). Two different mechanisms for mechanical force generation were implicated. Based on studies with splinted wounds, cellular force was believed to be generated from within the granulation tissue (Abercrombie et al., 1960), which now is known to represent myofibroblast contraction (Tomasek et al., 2002). Other studies showed that removal of granulation tissue did not decrease the rate of wound closure (Grillo et al., 1957; Gross et al., 1995), and fibroblasts migrating at the wound margins were suggested to provide the mechanical force for closure albeit by an unknown mechanism. We suggest that tractional forces exerted by fibroblasts attempting to migrate at the wound edge could cause inward movement of the connective tissue matrix, depending on potential mobilization of the surrounding tissue (cf. Peacock, 1984), a cell-dependent version of what surgeons call "mechanical creep" (Johnson et al., 1993; Wilhelmi et al., 1998).
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Frederick Grinnell (frederick.grinnell{at}utsouthwestern.edu)
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