1 Department of Physiology, University of Massachusetts Medical School, Worcester 01605; and 2 Department of Biomedical Engineering, Boston University, Boston, Massachusetts 02215
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ABSTRACT |
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One of the hallmarks of oncogenic transformation is anchorage-independent growth (27). Here we demonstrate that responses to substrate rigidity play a major role in distinguishing the growth behavior of normal cells from that of transformed cells. We cultured normal or H-ras-transformed NIH 3T3 cells on flexible collagen-coated polyacrylamide substrates with similar chemical properties but different rigidity. Compared with cells cultured on stiff substrates, nontransformed cells on flexible substrates showed a decrease in the rate of DNA synthesis and an increase in the rate of apoptosis. These responses on flexible substrates are coupled to decreases in cell spreading area and traction forces. In contrast, transformed cells maintained their growth and apoptotic characteristics regardless of substrate flexibility. The responses in cell spreading area and traction forces to substrate flexibility were similarly diminished. Our results suggest that normal cells are capable of probing substrate rigidity and that proper mechanical feedback is required for regulating cell shape, cell growth, and survival. The loss of this response can explain the unregulated growth of transformed cells.
mechanical signaling; cell cycle; cell shape; traction force; cancer
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INTRODUCTION |
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ALTHOUGH IT HAS BEEN RECOGNIZED for many years that adhesions with substrates or other cells regulate the growth of normal but not transformed cells, little is known about the nature of such anchorage-dependent regulation. To date most emphasis has been placed on characterizing the integrin receptor complexes (14), the extracellular matrix (ECM)-integrin binding interactions (19), and the downstream signaling events (10). However, physical forces at adhesion sites may play an equally important role, as evidenced by the responses of cell morphology, growth, apoptosis, and gene expression to mechanical forces such as fluid shear stress (6) or substrate stretching (1, 11, 20). It is possible that the main function of receptor binding is to establish a physical linkage between the cytoplasm and the outside environment, whereas subsequent physical interactions through such linkages elicit the actual enzymatic reactions on the cytoplasmic side of adhesion sites.
Our recent observations further indicate that normal cells not only respond to mechanical forces but actively probe substrate flexibility, most likely by applying contractile forces to the substrate and responding to the feedback of counterforces (21, 23). When plated on flexible substrates, normal cells show enhanced motility and reduced tyrosine phosphorylation (21). Thus an attractive hypothesis is that normal cells may regulate their motility, growth, and apoptosis according to mechanical properties of the substrate, whereas transformed cells may be defective in either detecting or responding to such mechanical signals.
Although previous observations have provided strong evidence that cell-substrate adhesions play a key role in regulating cell growth (14), it has not been possible to identify specifically the role of physical parameters because of the use of substrates that differ in both chemical and physical properties. To address this problem, we recently developed ECM-coated polyacrylamide substrates, which, through minor changes in the concentration of cross-linkers, allowed the regulation of flexibility over a wide range without altering their chemical properties. In this study, we cultured normal or H-ras-transformed 3T3 cells on substrates of different flexibility and compared their rates of growth and apoptosis, as well as cell spreading area and traction forces exerted on the substrate. We found that normal cells are much more sensitive to substrate flexibility than transformed cells. Our data can explain the growth advantage of transformed cells in vivo, where they are able to survive and grow independent of mechanical input from the surrounding tissues.
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METHODS |
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Preparation of polyacrylamide substrate. Polyacrylamide substrates coated with collagen I were prepared essentially as described previously (23, 28). The flexibility of the substrate was manipulated by maintaining the total acrylamide concentration while varying the bis-acrylamide concentration. Experiments with growth and apoptosis were performed with substrates of 5% total acrylamide and bis-acrylamide between 0.012 and 0.06%. Measurements of cell spreading and traction forces were performed with substrates of 8% total acrylamide and either 0.03 or 0.06% bis-acrylamide. This change was necessary for generating high-quality substrate deformation vectors appropriate for computer analysis (too soft a substrate causes surface fluorescent beads to lie on different planes of focus as cells exert forces). Before cells were plated, the acrylamide gels were soaked for 30 min in DMEM at 37°C.
Cell culture and microscopy. NIH 3T3 cells (ATCC, Rockville, MD) and H-ras-transformed NIH 3T3 cells (PAP2) were kindly supplied by Dr. Ann Chambers (London Regional Cancer Center, London, Ontario, Canada; Refs. 2, 3), and were cultured in DMEM (Sigma, St. Louis, MO), supplemented with 10% donor calf serum (JRH Biosciences, Lenexa, KS), 2 mM L-glutamine, 50 µg/ml streptomycin, 50 U/ml penicillin, and 250 µg/ml amphotercin B (GIBCO-BRL, Gaithersburg, MD). Images were recorded with a Zeiss ×40, NA 0.75 Plan-Neofluar objective on a Zeiss Axiovert 10 microscope, using a cooled charge-coupled device camera (Series 200, Photometrics, Tucson, AZ).
Assay of DNA synthesis. Twenty-four or forty-eight hours after plating, NIH 3T3 and PAP2 cells were incubated with 100 µM 5-bromodeoxyuridine (BrdU; Sigma) in DMEM for 1.5 h at 37°C. Cells were then processed for immunofluorescence staining for BrdU as follows: cellular DNA was digested with DNase I (1 mg/ml in 10 mM Tris · HCl, 20 mM MgCl2, pH 7.5) for 20 min at 37°C before incubation with primary monoclonal antibodies against BrdU (1:200; Clone BU-33, Sigma). The secondary antibody was Alexa 546-labeled goat anti-mouse IgG (H+L) antibody (1:200; Molecular Probes, Eugene, OR). Nuclei were counterstained with Hoechst 33258 (10 µg/ml, Sigma) for 10 min at room temperature before observation with fluorescence microscopy. The percentage of BrdU(+) cells was calculated by counting 200 cells in multiple fields in each experiment. The results are shown as means ± SE from five independent experiments.
Assay of apoptosis. Quantitation of apoptotic cells by terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) was performed using the In Situ Cell Death Detection Kit (Boehringer Mannheim, Indianapolis, IN) at 24 or 48 h after cells were plated on collagen-coated polyacrylamide substrates. Briefly, cells were fixed with 4% formaldehyde in PBS, pH 7.4, for 30 min at room temperature, then permeabilized with 0.1% Triton X-100, 0.1% sodium citrate for 2 min on ice. After incubation with the TUNEL reaction mixture for 60 min at 37°C, cells were counterstained with Hoechst 33258 (10 µg/ml, Sigma) for 10 min at room temperature before observation with fluorescence microscopy. The percentage of apoptotic cells was calculated by counting 200 cells in multiple fields in each experiment. The results are shown as means ± SE from five independent experiments.
Calculation of traction forces and cell spreading area. Traction forces generated by the cell were determined as described previously (7, 17). The deformation of the substrate due to cell-generated stresses was detected based on the displacement of embedded fluorescent beads near the substrate surface. Young's modulus of the substrate was estimated based on the depression created by the weight of a 0.65-mm steel ball, using an equation derived from the Hertz theory as was applied previously in atomic force microscopy (17, 25). Calculation of traction was carried out on a supercomputer, using the displacement vectors, the cell boundary, the Young's modulus, and the Poisson's ratio as the input. Average traction was calculated within regions that generated significant tractions. Briefly, the uncertainty of calculation at each pixel (7) was compared with the magnitude of traction. Pixels with a corresponding traction below the estimated uncertainty were excluded from averaging. This procedure minimizes the impact of cell spreading on average traction. Otherwise, because tractions were usually confined in limited peripheral areas while most regions near the nucleus generated no significant force (7), global averaging would cause peripheral exerted tractions to be diluted according to the degree of cell spreading. Cell spreading area was measured by counting the number of image pixels within the cell boundary using custom software.
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RESULTS |
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To test the hypothesis that normal cells respond not only to the occupancy of the integrins by anchored ECM molecules but also to the rigidity of the substrate, we cultured 3T3 cells on flexible collagen I-coated polyacrylamide sheets. Substrates of similar chemical property but different rigidity were prepared by maintaining a constant total concentration of acrylamide while varying the concentration of bis-acrylamide (23, 28). The substrates used for the measurement of cell growth and apoptosis rates have a Young's modulus varying between 4.7 and 14 kN/m2. In the following discussion, these substrates will be referred to as flexible and stiff substrates, respectively.
To study the effects of transformation on cellular responses to substrate flexibility, we compared normal NIH 3T3 cells with PAP2 cells, a line of H-ras-transformed NIH 3T3 cells selected for their ability to metastasize in chick embryos (2, 3). Unlike some highly transformed cells, PAP2 cells require some ECM interactions for their growth, as they did not multiply or survive on bare polyacrylamide substrate without collagen I coating. Thus the following results on growth and apoptosis reflect differential downstream responses to integrins bound to collagen-coated surfaces of different flexibility.
Effects of substrate flexibility on cell growth and apoptosis.
We first studied the effects of substrate flexibility on cell growth by
measuring the percentage of cells incorporating BrdU (Fig.
1). Cells were plated at a density of
1,000/cm2 onto either stiff or soft substrates. For normal
cells, an approximately twofold difference in the rate of BrdU
incorporation was observed 24 h after plating on flexible versus
stiff substrates (Fig. 1B). This difference became even more
pronounced at 48 h (~4-fold). For H-ras-transformed
cells, the rate of growth was not affected significantly by the
substrate flexibility. It is also worth noting that, on stiff
substrates at a limited cell density, H-ras-transformed 3T3
cells showed no growth advantage over nontransformed cells. However, on
flexible substrates, transformed cells grew at a rate approximately
twofold higher than that for nontransformed cells after 48 h of
plating (Fig. 1B).
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Effects of substrate flexibility on cell spreading and traction
forces.
The effects of substrate flexibility are likely linked to changes in
cell morphology, spreading, and/or mechanical interactions with the
substrate. Previous studies have correlated the rate of cell growth
with the degree of cell spreading on stiff substrates of different
chemical properties (8). We found that when cultured on
polyacrylamide substrates, normal cells showed a significant decrease
in spreading area when the Young's modulus decreased from 33 to 14 kN/m2 (Table 1).
H-ras-transformed cells showed no apparent decrease in
spreading area over this range of flexibility (Table 1) and became less
spread only upon further decrease in Young's modulus to <10
kN/m2.
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DISCUSSION |
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To address specifically the impact of mechanical input on cell growth, we cultured cells on flexible polyacrylamide substrates that have similar chemical properties but differ dramatically in their flexibility. The growth and motile behaviors of cells on stiff substrates are generally similar to those on glass or plastic surfaces. However, striking differences are observed as the flexibility increases, indicating that normal cells respond to not only the occupancy of the ECM receptors but mechanical signals transmitted through these receptors.
It has long been recognized that the degree of cell spreading is correlated with the rate of cell growth. Folkman and Moscona (8) first demonstrated that DNA synthesis in nontransformed cells increases as the cell becomes increasingly spread on more adhesive substrates, whereas transformed cells grew independently of cell shape or substrate adhesiveness. This is supported by a subsequent study indicating a progressive loss of shape-responsive metabolic control in transformed cells (29). Recent experiments by Ingber and coworkers (4, 13, 15, 26) further demonstrated changes in growth and apoptosis rates of normal cells under equivalent integrin occupancy but different geometry. However, although most previous studies have emphasized the importance of cell shape (4, 8, 12, 15), little is known about the physical/chemical parameters linking cell spreading/cell shape with the progression of cell cycle.
From both previous and the present studies (21), we propose that it is the mechanical input associated with shape change that regulates the cell cycle. To detect chemically identical but physically different surfaces, cells must rely on an active probing mechanism using contractile forces. Mechanical input from the substrate, transmitted through adhesion receptors, then activates downstream signals that regulate both the degree of cell spreading and the rate of growth. On stiff substrates, resistance to mechanical probing may lead to protein conformational changes and activation of signaling enzymes at the adhesion sites (5, 22). The response in turn causes an increase in traction forces and in cell spreading (Table 1). Activation of downstream chemical events may follow as a result of the direct activation of regulatory pathways or as a consequence of changes in cell shape or surface-to-volume ratio (4), leading to increased DNA synthesis and decreased apoptosis (Figs. 1 and 2). In contrast to normal cells, H-ras-transformed cells appear to be locked in a nonresponsive state in terms of substrate flexibility. They exert a similar magnitude of traction forces irrespective of the mechanical input from the substrate, which may then lead to a sustained mechanical stimulation of cell growth. Therefore, although additional defects in signal transduction are likely, the lack of response of traction forces to substrate flexibility in PAP2 cells may be sufficient to cause defects in the regulation of cell growth. For other types of transformed cells, the regulatory mechanism may breakdown at other levels, for example, through degradation of the ECM (3), deactivation of the force-sensing mechanism, or changes in the expression of integrin receptors (23).
The defects in mechanical response of H-ras-transformed cells may lead to significant growth advantages in the body, which consists largely of flexible tissues. At low cell densities on flexible substrates, normal cells suffered from both increased apoptotic death and reduced growth, whereas transformed cells maintained their rates of growth and apoptosis as on stiff substrates. At high densities, cell-cell contacts and aggregation became dominant. It protects normal cells from apoptosis but also inhibits their growth, whereas transformed cells grow independently of aggregate formation (Fig. 3). Thus transformed cells show a clear growth advantage over normal cells on flexible substrates at both low and high cell densities.
Cellular responses to substrate flexibility, stretching, and fluid shear may share a common mechanism involving active probing the environment and responses to mechanical forces. For nontransformed cells, this mechanism likely plays an important role during embryonic development and wound healing, allowing cellular proliferation or apoptosis to be regulated in response to changes in physical properties of the environment. Conversely, defects in mechanical signals may allow transformed cells to survive through the blood stream and to proliferate in diverse tissue environment.
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ACKNOWLEDGEMENTS |
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We gratefully acknowledge Dr. Ann F. Chambers (Department of Microbiology and Immunology, University of Western Ontario, London, Ontario, Canada) for H-ras-transformed NIH 3T3 (PAP2) cells.
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FOOTNOTES |
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This work was supported by grants from National Aeronautics and Space Administration and National Institutes of Health.
Address for reprint requests and other correspondence: Y. L. Wang, Univ. of Massachusetts Medical School, 377 Plantation St., Rm. 327, Worcester, MA 01605 (E-mail: yuli.wang{at}umassmed.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.
Received 30 November 1999; accepted in final form 8 June 2000.
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