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Am J Physiol Cell Physiol 290: C1640-C1650, 2006. First published January 11, 2006; doi:10.1152/ajpcell.00455.2005
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EXTRACELLULAR MATRIX, CELL INTERACTIONS

Intrinsic mechanical properties of the extracellular matrix affect the behavior of pre-osteoblastic MC3T3-E1 cells

Chirag B. Khatiwala,1 Shelly R. Peyton,1 and Andrew J. Putnam1,2

1Department of Chemical Engineering and Materials Science; and 2Department of Biomedical Engineering, The Henry Samueli School of Engineering, University of California, Irvine, Irvine, California

Submitted 7 September 2005 ; accepted in final form 8 January 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Mechanical cues present in the ECM have been hypothesized to provide instructive signals that dictate cell behavior. We probed this hypothesis in osteoblastic cells by culturing MC3T3-E1 cells on the surface of type I collagen-modified hydrogels with tunable mechanical properties and assessed their proliferation, migration, and differentiation. On gels functionalized with a low type I collagen density, MC3T3-E1 cells cultured on polystyrene proliferated twice as fast as those cultured on the softest substrate. Quantitative time-lapse video microscopic analysis revealed random motility speeds were significantly retarded on the softest substrate (0.25 ± 0.01 µm/min), in contrast to maximum speeds on polystyrene substrates (0.42 ± 0.04 µm/min). On gels functionalized with a high type I collagen density, migration speed exhibited a biphasic dependence on ECM compliance, with maximum speeds (0.34 ± 0.02 µm/min) observed on gels of intermediate stiffness, whereas minimum speeds (0.24 ± 0.03 µm/min) occurred on both the softest and most rigid (i.e., polystyrene) substrates. Immature focal contacts and a poorly organized actin cytoskeleton were observed in cells cultured on the softest substrates, whereas those on more rigid substrates assembled mature focal adhesions and robust actin stress fibers. In parallel, focal adhesion kinase (FAK) activity (assessed by detecting pY397-FAK) was influenced by compliance, with maximal activity occurring in cells cultured on polystyrene. Finally, mineral deposition by the MC3T3-E1 cells was also affected by ECM compliance, leading to the conclusion that altering ECM mechanical properties may influence a variety of MC3T3-E1 cell functions, and perhaps ultimately, their differentiated phenotype.

bone; focal adhesion kinase; mechanotransduction; cytoskeleton; integrins

ONCE THOUGHT TO PROVIDE only structural support to tissues by acting as a scaffold to which cells bind, the ECM is now known to contain critical biochemical information that regulates virtually all cell functions, including adhesion, spreading, migration, proliferation, survival, and differentiation. Most of these functions depend on integrins, a class of heterodimeric cell surface receptors composed of {alpha}- and beta-subunits that are responsible for the majority of adhesive interactions between a cell and the ECM (14). After adhesion to the ECM, integrins cluster in the plane of the cell membrane and integrate a number of structural and signaling proteins on the cytoplasmic surface of the cell membrane. They can activate various protein tyrosine kinases [e.g., focal adhesion kinase (FAK), Src, Abl], serine-threonine kinases (e.g., ERK, other MAPK family members, PKC), GTPases of the Rho family (RhoA, Rac, and Cdc42), and phosphoinositide lipid mediators (5, 13, 38). In addition, integrin attachment has been determined to be critical for optimal activation of growth factor-induced mitogenic pathways (39).

Integrins also provide a direct physical link between the ECM and the underlying cytoskeleton. This linkage is bidirectional, reciprocal, and dynamic (37), which implies that externally applied mechanical forces can be transduced directly from the ECM to the underlying cytoskeleton, imparting changes in cytoskeletal assembly and organization (16). It also implies that cell-generated forces can be transmitted across integrins to the supporting ECM. A striking example occurs when cells cultured on thin silicon rubber substrates visibly wrinkle the substrates as a result of cell-based tractional forces (12). The ECM's ability to resist these cell-based tractional forces appears to be a critical factor in controlling cell shape, which can regulate the balance between cell growth, differentiation, and death in some cell types (4, 6, 10, 17, 28).

In the context of bone, Julius Wolff (19, 42) first recognized the importance of mechanical forces in the late 1800s by proposing that mechanical stresses play a critical role in normal bone development and adaptation. Recent efforts to engineer functional bone replacements have documented that both applied strain and fluid shear stresses influence bone development and remodeling (40), strengthening the premise of Wolff's law. However, the influence of local stresses in the microenvironment on osteoblast behavior remains unclear. Recent evidence from our group and others suggests that the mechanical characteristics of the ECM (i.e., its stiffness or compliance) provide vital instructional cues to migrating smooth muscle cells (8, 32, 43) and fibroblasts (26, 30) and can even induce cells to migrate in a directional fashion from softer substrates to stiffer substrates, but not vice-à-versa. This new form of directional migration has been dubbed "mechanotaxis" or "durotaxis."

In this study, we studied the influence of ECM mechanics on cells of the osteoblast lineage, using the pre-osteoblastic MC3T3-E1 cell line as a model. By exploiting polyacrylamide substrates covalently functionalized with uniform densities of type I collagen, we have found that tuning ECM compliance modulates the migratory potential of these pre-osteoblastic cells. On surfaces covalently functionalized with low-density type I collagen, MC3T3-E1 cell migration speeds increased as compliance decreased, with maximum speeds achieved on rigid polystyrene control surfaces. By contrast, when gels were functionalized with high-density type I collagen, migration speed was found to depend on ECM compliance in a biphasic manner, in agreement with our recent findings that ECM compliance governs the migration speed of smooth muscle cells in a biphasic fashion (32). Parallel studies have revealed that ECM compliance influences MC3T3-E1 proliferation, cytoskeletal and focal adhesion assembly, the activity of FAK, and differentiation independently of changes in cell spreading.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture and drug treatment. Pre-osteoblastic MC3T3-E1 cells [subclone 4; American Type Culture Collection (ATCC), Manassas, VA] were maintained and expanded in {alpha}-MEM (GIBCO-BRL/Invitrogen, Carlsbad, CA) supplemented with 10% FBS (ATCC), 100 U/ml penicillin, and 0.1 mg/ml streptomycin (both from GIBCO-BRL) at 37°C and 5% CO2. MC3T3-E1 cells between passages 3 and 9 were used in all studies. For migration experiments, cells were cultured in {alpha}-MEM containing a reduced amount of serum (2% FBS). In some cases, microtubules (MTs) were depolymerized using nocodazole {methyl [5-(2-thienylcarbonyl)-1H-benzimidazol-2-yl]carbamate; Sigma, St. Louis, MO} by first dissolving the lyophilized powder in DMSO to yield a stock solution of 1 mg/ml, and subsequently diluting the drug in {alpha}-MEM culture medium with 2% FBS to obtain a final concentration of 10 µg/ml.

Fabrication and functionalization of polyacrylamide substrates. Model substrates with tunable mechanical properties were fabricated from polyacrylamide by adapting published methods (2, 30, 32). Briefly, stock solutions of 40% acrylamide and 2% N,N'-methylene-bis-acrylamide (Bio-Rad Laboratories, Hercules, CA) were combined to yield a final concentration of 8% acrylamide and varying concentrations of bis-acrylamide (0.1–0.3%) in 10 mM HEPES, pH 8.5. Following the addition of 10% ammonium persulfate and the free radical stabilizer TEMED [1,2-di-(dimethylamino) ethane] (both from Bio-Rad), the resulting solution was immediately syringe filtered (0.22-µm filter), poured between sterile glass plates separated by 0.7-mm spacers, and allowed to polymerize for at least 30 min. The use of the spacers provided uniformly thick hydrogels to ensure that attached cells were not influenced by the stiffness and/or rigidity of the glass or polystyrene surface underneath the gels. Following polymerization, the plates were separated and gel disks were punched from the polymerized material using a -in.- or -in.-diameter steel punch (McMaster-Carr, Atlanta, GA) and placed into multiwell culture plates for subsequent assays.

Gels were functionalized to support cell adhesion by covalently attaching type I collagen (Cohesion, Palo Alto, CA) as described (2). Briefly, N-sulfosuccinimidyl-6-(4'-azido-2'-nitrophenylamino) hexanoate (sulfo-SANPAH; Pierce Biotechnology, Rockford, IL), a UV light-sensitive heterobifunctional cross-linker, was covalently attached to the otherwise inert polyacrylamide surface via its azide functionality. A solution of 0.5 mM sulfo-SANPAH prepared in 50 mM HEPES, pH 8.5, and 0.5% DMSO was added to cover the gel surface and then induced to react with the acrylamide surface by exposing the gels to UV light (Blak-Ray; UVP, Cambridge, UK) at 365 nm for 15 min. Next, the gels were washed once with 50 mM HEPES, pH 8.5, and then photoactivation was repeated. The hydrogels were washed three times with 50 mM HEPES to remove excess reagent. A solution of type I collagen (either 10 µg/ml or 100 µg/ml) in PBS was added to the substrate and allowed to react overnight at 4°C. ECM-derivatized hydrogels were washed repeatedly with PBS before use. For rigid controls, type I collagen was adsorbed passively to the surface of polystyrene dishes or glass slides by being diluted in a COFormula-HCO3– buffer (in mM: 15 Na2CO3 and 35 NaHCO3, pH 9.4) as previously described (35).

Characterization of polyacrylamide gel mechanical properties. The bulk macroscopic elastic moduli (Young's moduli) of the polyacrylamide hydrogels were quantified using an MTS Synergie 100 (MTS Systems, Eden Prairie, MN) with 10 N load/cell as previously described (32). Briefly, hydrogels were formed into a "dog bone" shape that conformed to ASTM D638 standards (type V) by pouring the solution into a Teflon mold. Following polymerization, gels were hydrated in PBS for 30 min before testing. The hydrogels were then attached to the MTS load cell and fixed head using transparency film and a cyanoacrylate ester-based adhesive and subjected to a uniform deformation rate of 1 mm/min and a data-acquisition rate of 15 Hz. Young's moduli values (slope of stress-strain curve) between 1% and 10% strain (between 0.18- and 1.8-mm elongation) were calculated for each sample.

Proliferation assay. MC3T3-E1 cells were seeded onto the gels and polystyrene substrates in 12-well plates at a density of 25,000 cells/cm2, and their proliferation rate was assessed by quantifying cell numbers. Briefly, at 12, 24, 48, and 72 h after being seeded, cells were harvested from triplicate samples after being incubated in the presence of trypsin-EDTA for a prolonged length of time (>10 min) to ensure that all cells were removed from the substrates. Cells were then diluted in an isotonic buffer and subsequently counted using a cell counter (model ZM; Beckman Coulter, Fullerton, CA). Fresh growth medium was added to the remaining wells every 24 h throughout the proliferation assay.

Migration assays using time-lapse video microscopy. For cell migration assays, polyacrylamide gels were fabricated and functionalized in custom cell culture chambers as described previously (2). Briefly, a rectangular coverglass (no. 1, 45 x 50 mm; Fisher Scientific, Pittsburgh, PA) was flamed in a Bunsen burner, soaked in 0.1 N NaOH, and then air dried. After being dried, a small aliquot of 3-aminopropyltriethoxysilane (Sigma) was spread evenly onto the glass surface and allowed to sit for 5 min. Coverslips were then washed thoroughly using distilled water, incubated in a solution of 0.5% glutaraldehyde (Sigma) in PBS for 30 min, and then washed extensively with distilled water. These activated coverslips were then attached with vacuum grease to a 70 x 50 x 5-mm Plexiglas plate with a 35-mm-diameter annulus bored through the center. Gels were fabricated in the annulus by dispensing 200 µl of the acrylamide-bis-acrylamide solution of known concentrations directly onto the coverslip. The solution was covered with a small circular coverglass and allowed to polymerize. After polymerization, the round coverglass was removed carefully and the resulting hydrogel was functionalized as described earlier. Gels fabricated in this fashion were ~3.8 cm2 in area (similar to a 12-well dish) and ~0.7 mm thick.

Cells were then seeded at subconfluence (~4,000 cells/cm2) in these custom chambers in standard growth medium and allowed to attach and spread for 12 h. The serum concentration of the medium was reduced to 2% FBS just before video microscopy. Time-lapse microscopy was conducted using an inverted Nikon TE300 microscope equipped with an environmental chamber (to maintain temperature, humidity, and CO2 levels), an automated stage controller (Prior Scientifics, Temecula, CA), a digital camera (Photometrics CoolSnap fx; Roper Scientific, Tucson, AZ), and MetaMorph software (Universal Imaging, Downingtown, PA). Individual cells (i.e., those exhibiting no intercellular contact during the entire viewing period) were chosen at random in each of five different fields of view, and their centroids were tracked for 10- to 12-h periods at 5-min time intervals. The position of each cell centroid at the end of each interval was used to calculate cell velocity by dividing the displacement (denoted by the square root of {Delta}x2 + {Delta}y2, where x and y are position coordinates) by the time interval (5 min). Overall mean cell migration speed was then determined by averaging the speeds calculated at each interval during the entire 10- to 12-h period. Averaged data from each cell were weighted equally to those of all other cells and samples to calculate mean ± SD speed.

Immunofluorescence microscopy. Quantitative differences in collagen density coupled to the hydrogel surfaces were assessed using fluorescence microscopy staining with a mouse anti-collagen type I MAb (1:500 dilution in PBS; Chemicon International, Temecula, CA), followed by TRITC-conjugated donkey anti-mouse IgG (1:200 dilution in PBS; Jackson ImmunoResearch, West Grove, PA). Images were quantified using ImageJ software (National Institutes of Health, Bethesda, MD) by evaluating the red channel intensities from multiple images under the same conditions. These fluorescence intensity values were then normalized to the values obtained from the low-collagen-density images coupled to the 11.78-kPa hydrogels (which was arbitrarily assigned a value of 1). Focal adhesion and actin stress fibers in adherent MC3T3-E1 cells were visualized using standard fluorescence microscopy. Cells were seeded at a density of 7,000–8,000 cells/cm2 on the compliant hydrogels in 24-well tissue culture plates or on Lab-Tek chamber slides for 12 h in standard growth medium. The medium was changed to one containing 2% FBS for another 12 h to mirror the conditions used in the migration analysis. At the end of this 24-h period, cells were fixed using 4% formaldehyde (Sigma) in PBS at 4°C for 20 min. Mouse anti-vinculin IgG MAb (1:250 dilution in Abdil; Sigma), followed by TRITC-conjugated donkey anti-mouse IgG, was used to localize punctate focal adhesion structures at the cell-ECM interface. Filamentous actin (F-actin) stress fibers were visualized using Oregon Green 488 phalloidin stain (1:40 dilution in Abdil; Molecular Probes, Eugene, OR). Cell nuclei were stained with 4',6-diamidino-2-phenylindole dihydrochloride (1 µg/ml; Sigma) in PBS for 5 min.

Immunoblot analysis and immunoprecipitation. To quantify vinculin levels in focal adhesions, Triton X-100-insoluble and total cell lysates were analyzed as previously described (34). Briefly, total lysates were generated from cells cultured for 24 h (12 h in medium containing 10% FBS, followed by 12 h in medium containing 2% FBS ± nocodazole) on various substrates by washing them twice with cold PBS, followed by 5-min incubation with a lysis buffer [25 mM Tris, pH 7.4, 0.4 M NaCl, 0.5% SDS, and protease inhibitors (10 µg/ml aprotinin, 10 µg/ml leupeptin, 5 µg/ml pepstatin A, 1 mM PMSF, 1 mM NaF, and 1 mM sodium orthovanadate), all of which were obtained from Sigma]. To assay for focal adhesion-associated vinculin, cells were permeabilized for 10 min using permeabilizing buffer (10 mM HEPES, pH 6.9, 50 mM NaCl, 3 mM MgCl2, 0.5% Triton X-100, 300 mM sucrose, 1 mM EGTA, and protease inhibitors) before lysis. Equal protein lysates were then subjected to electrophoresis and Western blot analysis using a mouse anti-vinculin MAb (1:1,000 dilution; Sigma), followed by a 1:5,000 dilution of horseradish peroxidase (HRP)-conjugated goat anti-mouse IgG antibody (Santa Cruz Biotechnology, Santa Cruz, CA) and ECL substrate. The ratio of Triton X-100-insoluble vinculin to total vinculin was determined by performing quantitative densitometry of the developed films.

To assay FAK activity, cultured cells were washed with cold PBS twice and then lysed for 5 min with RIPA lysis buffer (10 mM Tris, pH 7.2, 158 mM NaCl, 1 mM EDTA, 0.1% SDS, 1% Triton X-100, 1% sodium deoxycholate, and protease inhibitors). FAK was immunoprecipitated by incubating equal protein amounts (600 µg) with protein G beads (35 µl) and 5 µg of mouse anti-FAK (clone 4.47) IgG MAb (Upstate Biotechnology, Lake Placid, NY) overnight at 4°C. The bead-antibody-protein complexes were collected by centrifugation at 13,000 rpm, washed three times with 1% BSA in RIPA buffer, and then subjected to electrophoresis and Western blot analysis. Blots were probed with antibodies to either active FAK [mouse anti-FAK (phosphorylated Y397, pY397) IgG; BD Transduction Laboratories, San Jose, CA] or total FAK [mouse anti-FAK MAb IgG (clone 4.47); Upstate Biotechnology, Lake Placid, NY], followed by the HRP-conjugated secondary antibody and ECL substrate. The ratio of Y397-FAK to total FAK for each condition was determined by performing quantitative densitometry of developed films.

Von Kossa staining. To assess the osteoblastic differentiation of MC3T3-E1 cells, the cells were seeded onto gel and polystyrene substrates in 12-well plates at a density of 100,000 cells/cm2 (well above confluence) and incubated for 4 days in differentiation medium ({alpha}-MEM culture medium containing 10% FBS supplemented with 10 mM beta-glycerophosphate, 50 µg/ml ascorbic acid, and 100 nM dexamethasone). Cells were stained using the Von Kossa method with a commercial kit (American MasterTech Scientific, Lodi, CA) according to the manufacturer's instructions.

Statistical analysis. All statistical analyses were performed using InStat 2.01 software for Macintosh. Data are means ± SD unless otherwise noted. In cases in which statistical comparisons were made between three or more groups of data, we performed one-way ANOVA, followed by the Student-Newman-Keuls posttest to compare two data sets at a time. In cases where only two sets of data were compared, Student's unpaired t-test was performed. P < 0.05 denotes statistical significance.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Characterization of substrate mechanical properties and ECM functionalization. To address the influence of ECM compliance on the phenotype of the pre-osteoblastic MC3T3-E1 cell line, we utilized polyacrylamide model substrates based on the fact that their mechanical properties can be manipulated easily by varying the relative amounts of acrylamide and bis-acrylamide as previously described (2). We systematically quantified the bulk macroscopic mechanical properties of the polyacrylamide substrates using an MTS Synergie 100 mechanical test frame capable of applying small deformations at defined rates as previously described (32). Tensile tests were performed on polyacrylamide substrates containing 8% acrylamide plus either 0.1%, 0.2%, or 0.3% bisacrylamide. Hydrogels were deformed at a rate of 1 mm/min, and the Young's moduli determined from the slope of the stress-strain curve between 1–10% strain. The resulting elastic (i.e., Young's) moduli (E) values for each of the substrates used in this study were as follows: 11.78 ± 2.78 kPa for 8% acrylamide, 0.1% bisacrylamide; 21.6 ± 6.17 kPa for 8% acrylamide, 0.2% bisacrylamide; and 38.98 ± 3.78 kPa for 8% acrylamide, 0.3% bisacrylamide (Fig. 1A). Each of these values is statistically significant compared with the others (P < 0.01), yielding model substrates spanning a threefold difference in stiffness levels between the softest and stiffest hydrogels. From here on, we refer to these three substrates as E = 11.78 kPa, E = 21.6 kPa, and E = 38.98 kPa.


Figure 1
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  		Fig. 1. Characterization of the mechanical properties and collagen functionality of polyacrylamide substrates. A: Young's moduli (E) of acrylamide hydrogels were determined via standard tensile testing. Acrylamide (linear chain, component A) was maintained at 8%, whereas N,N'-methylene-bis-acrylamide (cross-linker, component B) was varied between 0.1%, 0.2%, and 0.3%. Each column is significantly different from the others. Error bars, means ± SD. n > 3; *P < 0.01. B: representative immunofluorescent photomicrographs of polyacrylamide hydrogels (21.6 kPa, left, and 38.98 kPa, right) coupled with the 2 different theoretical collagen concentrations used in this study (top, 10 µg/ml, which translates to 5 µg/cm2; bottom: 100 µg/ml, which translates to 50 µg/cm2). Scale bar, 100 µm. C: quantification of images reveals a uniform fluorescence intensity for a given concentration, regardless of substrate stiffness, with a 2-fold increase in fluorescence intensity when theoretical collagen concentration was increased from 10 µg/ml (black bars) to 100 µg/ml (gray bars). ***P < 0.001.

 
To confirm the presence of varying densities of type I collagen on the surface of the acrylamide hydrogels, we used immunofluorescent staining to visually assess the differences between the gels functionalized with either 10 µg/ml (which translates to a theoretical density of 5 µg/cm2 on our gels) or 100 µg/ml (which translates to a theoretical density of 50 µg/cm2) collagen solutions. A distinct, visible increase in collagen density was evident using immunofluorescent staining in the absence of cells at two different substrate stiffness levels (E = 21.6 kPa and E = 38.98 kPa) (Fig. 1B). There was no qualitative difference in collagen immunofluorescence between substrates of different stiffness levels functionalized using solutions with the same starting collagen concentration. Quantification of these types of images confirmed this qualitative assessment (Fig. 1C), revealing no significant differences in collagen content across substrates of different cross-linking density when the same starting amount of collagen was used. This quantification also revealed that a theoretical 10-fold increase in the starting ligand density translates in actuality to an approximately 2-fold increase in collagen density.

Influence of substrate stiffness on MC3T3-E1 migration depends on ECM ligand density. Using substrates of three different values of E and polystyrene as a control, the influence of ECM compliance on the speed of random motility in the MC3T3-E1 cells was assessed. Cells were seeded onto the collagen-modified polyacrylamide hydrogels and polystyrene controls at a subconfluent density to minimize intercellular interaction. Time-lapse video microscopy was used to visualize cell migration over a period of 10–12 h. On substrates functionalized with a solution of low-concentration collagen (10 µg/ml or 5 µg/cm2), cell migration speed increased as ECM stiffness increased (Fig. 2A, black bars), with the maximum speed of 0.42 ± 0.04 µm/min observed on the control polystyrene surface and the minimum speed of 0.25 ± 0.01 µm/min observed on an 11.78-kPa substrate. The difference between these two speeds was statistically significant (P < 0.01), but the migration speed of cells cultured on the stiffest hydrogel was not statistically different from those cultured on polystyrene. However, when cells were seeded onto hydrogels and polystyrene controls modified with the higher concentration of collagen (100 µg/ml or 50 µg/cm2), migration speeds exhibited biphasic dependence on the substrate compliance (Fig. 2A, gray bars). Specifically, the maximum speed of 0.34 ± 0.02 µm/min was observed on the 21.6-kPa hydrogel, whereas the minimum speed of 0.24 ± 0.03 µm/min was observed on the 11.78-kPa hydrogel as well as on the control polystyrene substrate.


Figure 2
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  		Fig. 2. Cell motility and proliferation rates are influenced by ECM rigidity. A: mean cell speeds of MC3T3-E1 cells were determined as a function of substrate compliance for 2 different collagen densities by tracking individual cells. Data represent means cell speed (µm/min) ± SE from at least 3 separate experiments (n = 3). At low collagen densities (black bars), the migration speed on the 11.78-kPa gel was significantly different from that on every other surface (**P < 0.01), whereas at high collagen densities (gray bars), the migration speed on the 11.78-kPa gel was significantly different from only that of the 21.6-kPa gel. **P < 0.01. At both collagen densities, the mean cell speed on the 21.6-kPa gels was significantly different from that under all other conditions. *P < 0.05. Differences in mean cell speeds on the 38.98-kPa gels and polystyrene were not significant at both collagen densities. On both the 38.98-kPa gels and polystyrene, migration speeds at low collagen density were statistically different (#P < 0.01) from those at high collagen densities (compare black bars with gray bars). B: MC3T3-E1 cells cultured on hydrogels with E = 11.78 kPa ({blacklozenge}), E = 21.6 kPa ({blacksquare}), E = 38.98 kPa ({blacktriangleup}), and the control polystyrene surface (bullet) were counted at 12, 24, 48, and 72 h postseeding. Approximately 70% of cells (originally seeded at 25,000 cells/cm2) were attached at 12-h time point across all conditions, indicating insignificant differences in plating efficiency. Statistically significant (P < 0.001) increases in cell number were observed on all substrates from one time point to the next. In addition, differences in cell numbers across all substrates were statistically significant at 24, 48, and 72 h postseeding. Error bars, means ± SD; n = 3. *P < 0.01.

 
The proliferative capacity of the pre-osteoblastic MC3T3-E1 cell line was also assessed as a function of compliance on substrates functionalized with the lower collagen density (Fig. 2B). All substrates were initially seeded at a density of 25,000 cells/cm2. Approximately 70% of the cells were attached after 12 h on all substrates, indicating equivalent plating efficiencies. Twenty-four hours after seeding, cell counts on the softer substrates (E = 11.78 kPa and E = 21.6 kPa) were significantly less compared with the stiffer substrates (E = 38.98 kPa and polystyrene). Similar trends were observed in 48- and 72-h cultures, with more than twice as many cells present on polystyrene as on the softest hydrogel substrate (E = 11.78 kPa) at the end of the 3-day proliferation assay.

Substrate stiffness regulates the organization and steady-state assembly of actin stress fibers and focal adhesions independently of cell spreading. To probe the mechanism by which ECM compliance modulates the proliferation and migration of MC3T3-E1 cells, actin-mediated contractility was assessed qualitatively by visualizing the organization and assembly of actin stress fibers and focal adhesions and quantitatively by measuring the steady-state recruitment of vinculin to focal adhesions. Immunofluorescence microscopy revealed a clear increase in the assembly of actin stress fibers and focal adhesions as the stiffness of the underlying ECM was increased for both collagen densities tested (Fig. 3A). For the low ligand densities (Fig. 3A, top), cells cultured on the most compliant substrate (E = 11.78 kPa) exhibited an F-actin network that was disorganized and poorly defined, along with small, ill-defined focal adhesions. In contrast, cells cultured on substrates that were three times as stiff (E = 38.98 kPa) assembled the actin cytoskeleton into a robust stress fiber network that terminated in well-defined vinculin-containing focal adhesions comparable to those observed in cells cultured on control glass surfaces. For the high ligand density (Fig. 3A, bottom), similar trends were observed regarding the assembly of the F-actin network and focal adhesions. It is also noteworthy that the assembly of these structures was enhanced on substrates functionalized with higher collagen densities (Fig. 3A, compare top row with bottom row) and that the effect of ligand density was most pronounced on softer gels (E = 11.78 kPa and E = 21.6 kPa). Quantitative Western blot analysis of Triton X-100-insoluble vinculin levels (Fig. 3, B and C) corroborated these qualitative assessments. Increasing the rigidity of the underlying ECM shifted the steady-state levels of vinculin in focal adhesions from 43 ± 5.4% of total vinculin on the softest substrate (E = 11.78 kPa) to 84 ± 3.7% on polystyrene when cells were cultured at low ligand density (Fig. 3, black bars). Likewise, when cells were cultured on substrates presenting a high ligand density, the ratio of insoluble to total vinculin increased from 61 ± 1.1% on E = 11.78 kPa substrates to 84.2 ± 5.3% on polystyrene (Fig. 3, gray bars). These results reveal that the expected increases in the ratio of insoluble to total vinculin induced by increased ECM ligand density were more pronounced on compliant surfaces. Furthermore, we report herein that the spreading of MC3T3-E1 cells remained statistically unchanged across the range of compliance and the ECM ligand density values investigated (Fig. 3D).


Figure 3
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  		Fig. 3. Organization and steady-state assembly of actin stress fibers and focal adhesions are sensitive to ECM rigidity independent of changes in cell spreading. A: MC3T3-E1 cells cultured on compliant substrates functionalized with solutions containing either low (top row, 10 µg/ml) or high (bottom row, 100 µg/ml) collagen concentrations were fixed 24 h post seeding and stained for filamentous actin (F-actin; green) and vinculin (red). Representative x100 magnification immunofluorescent photomicrographs as a function of increasing ECM rigidity (left to right) and ligand density (top to bottom). Scale bar, 20 µm. B: representative Western blot analysis of Triton X-100-insoluble and total vinculin levels are shown as a function of ECM compliance for each collagen concentration tested (10 µg/ml and 100 µg/ml). T, total cellular vinculin; I, Triton X-insoluble vinculin; 10 and 100, initial collagen concentrations (in µg/ml) used to functionalize the substrates. C: quantification of Western blots was performed using scanning densitometry of levels of vinculin associated with focal adhesions normalized to total vinculin content for each condition. Black bars show lower collagen density (10 µg/ml, or 5 µg/cm2), whereas gray bars express high-collagen-density data (100 µg/ml, or 50 µg/cm2). Values are means ± SD; n ≥ 3. ***P < 0.001. **P < 0.01. *P < 0.05. #P < 0.001, low vs. high collagen density regarding any one compliance. D: spread cell area of MC3T3-E1 cells cultured for 12–24 h quantified as a function of both ECM compliance and ligand density (black bars, low density; gray bars, high density). Data are not significantly different. Values expressed in µm2 are means ± SE of data obtained from at least 3 different data sets (n = 3).

 
Microtubule depolymerization increases the levels of focal adhesion vinculin and reduces migration speed. Cell migration is a highly coordinated biophysical process (21) involving the extension of a progressing lamellipodium, followed by attachment of integrins to form nascent focal complexes (7, 22). The subsequent generation of contractile forces through actin-myosin interactions induces the maturation of nascent focal complexes to focal adhesions (11), which then must be remodeled for forward displacement of the cell body to occur (41). An emerging hypothesis based on our prior work and the work of many other researchers is that cell movement is maximized when actin-mediated tractional stresses are optimally balanced, both by the ECM itself and by the MT cytoskeleton. On the basis of this argument, we took a rather blunt approach to disrupting this balance of forces by depolymerizing MTs with nocodazole. As expected, in MC3T3-E1 cells cultured on gels functionalized with low-density collagen, fluorescent staining of the cytoskeleton and focal adhesions confirmed that MT depolymerization using nocodazole enhanced actin-mediated contractility, which manifested as increases in stress fibers and focal adhesions (Fig. 4A).


Figure 4
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  		Fig. 4. Microtubule (MT) depolymerization using nocodazole {methyl [5-(2-thienylcarbonyl)-1H-benzimidazol-2-yl]carbamate}increases actin contractility on all substrates and negates the influence of ECM rigidity on cell migration speed. A: MC3T3-E1 cells were cultured on E = 11.78 kPa, E = 21.6 kPa, and E = 38.98 kPa gels and on rigid substrates (glass, in this case) functionalized with low collagen concentration (10 µg/ml) in the presence of nocodazole (10 µg/ml) for 12 h post seeding. After an additional 12 h of culture in the presence of nocodazole, cells were fixed and stained for F-actin (green) and vinculin (red). Representative x100 magnification immunofluorescent photomicrographs are shown. Scale bar, 20 µm. B: spread cell area of MC3T3-E1 cells cultured for 24 h in absence (black bars) or presence (gray bars) of nocodazole was quantified. Data are not significantly different. Values are means ± SE (in µm2) of data obtained in at least 3 different sets of experiments; n = 3. C: representative Western blot of Triton X-100-insoluble and total vinculin levels as a function of ECM rigidity in cells ± nocodazole. Absence of nocodazole in medium is indicated by –, whereas presence of 10 µg/ml nocodazole in the culture medium is indicated by +. D: quantification of Western blot analysis was performed using scanning densitometry, with levels of vinculin associated with focal adhesions normalized to total vinculin content for each condition. Black bars, absence of nocodazole; gray bars, presence of nocodazole. Values are means ± SD; n = 5. ***P < 0.001. *P < 0.05. #P < 0.05 medium with vs. without nocodazole regarding any one compliance (gray vs. black bars). E: MC3T3-E1 cells treated with nocodazole 12 h after seeding were subjected to time-lapse video microscopy 2 h after nocodazole was introduced, and monitored for an additional 10–12 h to quantify mean cell speed. Values regarding all substrates were significantly less than values observed for cells that migrated in absence of nocodazole. *P < 0.05. ***P < 0.001. No significant differences were observed across all conditions in presence of nocodazole. Values are means ± SE of cell speeds (µm/min) on basis of values obtained in 3 separate experiments; n = 3.

 
The qualitative interpretations of the fluorescent images were confirmed by quantifying the relative ratio of vinculin in the Triton X-insoluble lysates to total cellular vinculin using Western blot analysis (Fig. 4, C and D). As noted earlier, steady-state levels of focal adhesion-associated vinculin steadily increased with ECM stiffness (Fig. 4D, black bars), whereas the total cellular vinculin levels remained unchanged across all conditions. Depolymerizing MTs with nocodazole increased the levels slightly (Fig. 4D, gray bars), with the ratio of Triton X-insoluble vinculin to total vinculin increasing from 43 ± 5.4% in the absence of drug to 49 ± 0.4% in the presence of drug on the most compliant substrate (E = 11.78 kPa), indicating an increase in actin-mediated contractility. On polystyrene, the ratio in the absence of drug (84 ± 3.7%) was not statistically significant from that in the presence of drug (86 ± 2.4%). Functionally, MT depolymerization reduced the migration speeds of cells cultured on the most rigid substrate (polystyrene) to values comparable to those on the softest substrate (E = 11.78 kPa) (Fig. 4E). The reduction in the migration speeds was statistically significant (P ≤ 0.05) compared with the values observed in the absence of the drug for all substrate conditions.

Tuning ECM compliance regulates phosphorylation of FAK on Y397. Next, we addressed the hypothesis that ECM compliance regulates the activation of FAK, an important mediator of integrin-mediated signaling, in the MC3T3-E1 pre-osteoblastic cell line. MC3T3-E1 cells were cultured on the compliant substrates and on control polystyrene substrates for 24 h, and the phosphorylation of tyrosine residue 397 was monitored using immunoprecipitation and Western blot analysis (Fig. 5). Qualitatively, increasing the stiffness of the underlying substrate resulted in increasing levels of pY397-FAK, but the levels of total FAK remained largely unchanged (Fig. 5A). Quantitative densitometric analysis of the ratio of pY397-FAK to total FAK normalized to the value of the control polystyrene substrate (which was arbitrarily chosen to be 1) mirrored this qualitative trend. Specifically, the Y397-FAK/FAK ratio on the softest gel (E = 11.78 kPa) was determined to be 32% of the value for cells cultured on polystyrene. This ratio increased to 49% on the intermediate stiffness gels (E = 21.6 kPa) and to 68% on the stiffest gel (E = 38.98 kPa) (Fig. 5B, black bars). The trend clearly shows that in the absence of any other cues, FAK phosphorylation at Y397 is regulated by the mechanical properties of the underlying matrix. However, when cells were cultured in the presence of nocodazole, there was no statistically significant difference between the pY397-FAK/FAK ratio in the presence (Fig. 5B, gray bars) or absence of the drug.


Figure 5
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  		Fig. 5. Levels of Y397-phosphorylated (pY397) FAK increase with ECM rigidity. A: levels of pY397-FAK and total FAK as a function of substrate stiffness were detected using standard immunoprecipitation and immunoblot analysis techniques as shown in these representative blots. B: following scanning densitometry, levels of pY397-FAK were normalized to levels of total FAK, and the ratios were then compared with levels on the control polystyrene substrate. Relative levels of pY397-FAK significantly increased with increasing stiffness of underlying substrate in the absence of nocodazole (black bars). **P < 0.01. ***P < 0.001. The same dependence of FAK activity on ECM compliance was observed when MTs were depolymerized using nocodazole (gray bars). *P < 0.05. ***P < 0.001. Marginal increases in normalized levels of pY397-FAK to total FAK that occur in the presence of nocodazole on all substrates compared with untreated controls were not statistically significant. Data are means ± SD; n ≥ 3.

 
Osteoblastic differentiation of MC3T3-E1 cells is influenced by ECM rigidity. The final element of this study was to determine whether ECM compliance influences osteoblastic differentiation of MC3T3-E1 cells. To eliminate proliferative effects from differentiation effects, cells were seeded at confluent densities on collagen-modified hydrogel substrates at two different stiffness levels (E = 21.6 kPa and E = 38.98 kPa) and on polystyrene and were then cultured in the presence of osteogenic supplements (beta-glycerophosphate, ascorbic acid, and dexamethasone) for 4 days. Using the von Kossa staining method to determine the extent of ECM mineralization (an indicator of a mature osteoblast phenotype), we found that MC3T3-E1 cells cultured on polystyrene deposited mineral to the greatest extent, characterized by many focal mineral deposits (Fig. 6). Cells on the most compliant surface tested (E = 21.6 kPa) mineralized the ECM to a lesser degree, but there was clear evidence that cells on a slightly stiffer substrate (E = 38.98 kPa) exhibited an intermediate degree of differentiation.


Figure 6
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  		Fig. 6. ECM rigidity influences mineralization of the ECM by MC3T3-E1. MC3T3-E1 cells were seeded at confluence and cultured in presence of osteogenic supplements on gels and polystyrene for 4 days. Mineral deposits were detected using the Von Kossa staining method and appear as focal black spots in these representative photomicrographs. Scale bar, 100 µm.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The importance of the ECM in normal and pathological tissue development is widely recognized and well documented (1, 3). However, since the discovery that integrin cell adhesion receptors trigger intracellular biochemical signaling cascades (14), the majority of ECM research has focused on how ligand identity regulates cell signaling and behavior, often overlooking the fact that the ECM was first hypothesized to play a purely structural role. Given the interest of many investigators in the development of injectable hydrogel carriers for bone tissue-engineering applications (23, 33), the present study was designed to determine whether manipulating substrate mechanical properties influences the behavior and function of pre-osteoblastic cells. This objective stemmed from the hypothesis that ECM chemistry and mechanics coordinately regulate cell fate and tissue development in general, an idea supported by a number of recent reports suggesting that ECM compliance regulates cell function (9, 20) and morphogenesis (15, 29). Using collagen-modified polyacrylamide model substrates with tunable mechanical properties, we report herein that ECM compliance modulated by ligand density can influence the migration, proliferation, and differentiation of the MC3T3-E1 pre-osteoblast cell line. Although the precise mechanisms of these phenomena remain to be determined, our structural (i.e., vinculin and F-actin) and signaling (i.e., FAK) data suggest that ECM chemistry and mechanics trigger coordinated biophysical and biochemical changes in the cell, which may ultimately manifest a change in function.

Numerous previous studies defined and characterized a new form of cell migration known as durotaxis (or mechanotaxis), in which the speed and direction of cell migration are dictated solely by ECM compliance (8, 26, 30, 43). We recently reported a novel biphasic dependence of smooth muscle cell migration speed on ECM stiffness in which the value of the optimal substrate stiffness at which migration speed is maximized depends on the density of immobilized ECM ligand (32). Herein we report that the migration of MC3T3-E1 cells exhibits a similar biphasic dependence on ECM compliance on substrates functionalized with high ligand density. However, MC3T3-E1 migration speeds increase as rigidity increases on substrates modified with low collagen density. We postulate a generalized interpretation that the dependence of migration on ECM compliance is in fact biphasic, but that the optimal stiffness shifts to lower values when more ECM ligand is present and shifts to higher values when less ECM ligand is present. In the case of low collagen density described herein, an attenuated migratory capacity was not observed on polystyrene, because the available collagen density was too low. Furthermore, our present results reveal that the optimal stiffness for bone cell migration is notably different from that observed in smooth muscle cells (32). Future studies will expand the range of ligand densities and focus on the molecular mechanisms by which ECM compliance differentially regulates the migratory properties of smooth muscle cells and osteoblasts.

Our results also show that both MC3T3-E1 proliferation and osteogenic differentiation are sensitive to small perturbations in ECM compliance. Of particular interest is the finding that both are maximized on rigid substrates, in opposition to the concept that proliferation and differentiation are mutually exclusive genetic programs that can be switched by ECM and other factors (17, 28). This concept was recently supported by evidence that soft substrates support osteoblastic differentiation better than rigid substrates (20), which is in disagreement with the results reported herein. However, there are several distinctions between our findings and those described in the previous report (20). First, in our study, MC3T3-E1 cells were attached to full-length type I collagen as opposed to short Arg-Gly-Asp (RGD)-containing peptide sequences. {alpha}vbeta3 and {alpha}5beta1 integrins are commonly involved in cell adhesion to RGD sequences, whereas {alpha}2beta1 is the predominant integrin used for binding to type I collagen. This difference therefore raises the tantalizing possibility that the engagement of unique subsets of integrins differentially regulate the phenotypic switch between proliferation and differentiation, a possibility supported by recent studies published by other groups (18, 36). Second, our mineralization studies were conducted with fully confluent cells to eliminate the effects of proliferation. The degree of confluence, possibly sensed via cadherin-mediated intercellular adhesion, blocks cell cycle progression, thereby permitting differentiation. In this context, our results indicate that a more rigid surface is ideal for osteoblastic differentiation once proliferation has been eliminated.

Our findings support the argument that the intrinsic mechanical properties of the ECM are an important determinant of cell function. However, an alternative interpretation is based on the argument that changing the cross-linking density of the polyacrylamide gel system can induce changes in the surface chemistry of the gels. Specifically, manipulating cross-linking density may influence the relative surface hydrophilicity of gels possessing different moduli values, which may affect the covalent coupling or physical presentation of the type I collagen on the acrylamide surfaces. However, our immunofluorescence microscopy results indicate that significant differences in the levels of type I collagen were absent when substrates of two different stiffness levels were functionalized with the same ligand density, suggesting that changing the cross-linking density did not significantly alter the covalent coupling of the ECM proteins. Furthermore, polyacrylamide gels have been used as a model substrate by many other investigators (2, 8, 9, 26, 30–32, 43) and are now widely accepted as an effective means by which to manipulate ECM mechanics. Nevertheless, because an extensive surface characterization (e.g., contact angle measurements) was not performed in this study, we cannot rule out the possibility that the effects we observed were due solely to changes in surface chemistry (18, 24).

In addition, because gels that possess different mechanical properties have been reported to support cell spreading to differing degrees (8, 32), one could argue that the effects of ECM compliance are actually due to a change in cell spreading. In fact, many papers have shown that cell function is directly linked to cell spreading (4, 17, 27, 28). One recent study documented that constraining cell shape by using patterned substrates controlled the commitment of mesenchymal stem cells to either adipogenic or osteoblastic fates by regulating RhoA-mediated tension in the actin cytoskeleton (27) and contributed significantly to the emerging paradigm that differentiation and morphogenesis in general can be regulated by tension in the actin cytoskeleton (29). Herein we have described how the spreading of MC3T3-E1 cells occurs independently of ECM compliance and ligand density, at least for the range of compliance values and the two ligand densities tested. However, cells cultured on stiff substrates are characterized by an increased assembly of actin stress fibers and by the recruitment of vinculin from cytoplasmic pools to the sites of focal adhesion, both of which indicate increased RhoA activity and a higher degree of contractility. Thus our interpretation is that ECM rigidity, perhaps directly in this cell type or indirectly by controlling the spreading of other cell types, regulates cytoskeletal tension, which in turn is a critical determinant of cell fate.

To probe this hypothesis a bit further, we used nocodazole to depolymerize MTs and disrupt a presumptive cytoskeletal force balance. Admittedly, this method is a rather brute force approach confounded by the fact that MT depolymerization enhances the activity of RhoA (25). Regardless, MT depolymerization disrupted cell motility while triggering only small, insignificant increases in the area of cell spreading (Fig. 4B). Likewise, the absence of MTs induced no significant differences in the influence of compliance on FAK activity. By contrast, we observed a perceptible yet significant increase in F-actin and focal adhesion-associated vinculin after MT depolymerization in MC3T3-E1 cells cultured on the more compliant surfaces, suggesting that MTs bear more of the load generated by the contractile actin network when the ECM is unable to do so. Upsetting this presumptive force balance by depolymerizing MTs in turn disrupts the influence of ECM compliance on MC3T3-E1 motility.

In summary, the results presented herein demonstrate that the intrinsic mechanical properties of the ECM affect the migration, proliferation, and differentiation of the preosteoblastic MC3T3-E1 cell line and that ECM ligand density modulates these effects. We speculate that modifying these ECM properties might modulate the degree of contractility (i.e., tension) in the actin cytoskeleton. Given recent evidence that tension in the actin cytoskeleton acts as a master switch to control the differentiation of bone marrow-derived mesenchymal stem cells into committed osteoblasts (27), our findings may be particularly relevant in support of efforts to develop synthetic ECM analogs for tissue engineering and regenerative medicine applications, suggesting that simply tuning material compliance may enable predictable control over differentiation.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was partially supported by National Institute of Dental and Craniofacial Research Grant 1R03-DE-016117-01A1 (to A. J. Putnam) and internal setup funds provided by the University of California, Irvine. S. R. Peyton was partially supported by an Achievement Rewards for College Scientists Foundation fellowship.


   ACKNOWLEDGMENTS
 
We are grateful to C. Ghajar, A. Kundu, and N. Malavia for stimulating discussions.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. J. Putnam, Dept. of Chemical Engineering and Materials Science, Univ. of California, Irvine, 916 Engineering Tower, Irvine, CA 92697-2575 (e-mail: aputnam{at}uci.edu)

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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