© 1997 American Heart Association, Inc.
Articles |
Mechanical Strain Tightly Controls Fibroblast Growth Factor-2 Release From Cultured Human Vascular Smooth Muscle Cells
the Cardiovascular Division, Department of Medicine (G.C.C., W.H.B., D.S.G., P.L., R.T.L.), Brigham and Women's Hospital, Harvard Medical School, Boston Mass, and the Division of Health Sciences and Technology (G.C.C., A.J.G., M.L.G., R.T.L.) and the Department of Mechanical Engineering (A.J.G., R.T.L.), Massachusetts Institute of Technology, Cambridge, Mass.
Correspondence to Richard T. Lee, MD, Cardiovascular Division, Brigham and Women's Hospital, 75 Francis St, Boston, MA 02115. E-mail RTLEE@BICS.BWH.HARVARD.EDU
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Abstract |
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Although fibroblast growth factor-2 (FGF-2) participates in the response to vascular injury, the role of cellular deformation in FGF-2 release is incompletely understood. To test the hypothesis that mechanical strain tightly controls FGF-2 release, a novel device was used to impose homogeneous and uniform biaxial strain to human vascular smooth muscle cells. Release of FGF-2 increased with the number of cycles of strain (14%, 1 Hz); 1, 9, and 90 cycles of strain, respectively, released 0.55±0.06%, 2.9±0.3%, and 5.5±1.3% of the total cellular FGF-2 (versus 0.00±0.40% for control, P<.05), but release was not further increased for strain of 90 to 90 000 cycles. Mechanical release of FGF-2 depended on both the frequency and amplitude of deformation. For example, strain (90 cycles, 1 Hz) at 4% amplitude released only 0.1±0.1% of the total FGF-2, but strain at 14% and 33% amplitudes, respectively, released 5.7±0.5% and 19.0±3.0% of the FGF-2 cellular pool (P<.05), suggesting a strain amplitude threshold for FGF-2 release. Injury to a subpopulation of cells increased with the frequency and amplitude of strain, but cells were not injured by strains below 10% amplitude. Strain following pretreatment with heparin released 12.6±1.6% of the total FGF-2 (versus 15.8±0.9% for strain alone, P<.05), indicating that most FGF-2 was liberated from the nuclear or cytoplasmic pools and not from low-affinity extracellular receptors. Conversely, strain in the presence of heparin released 25.2±3.5% of the total FGF-2 (versus 15.6±2.6% for strain alone, P<.05). Thus, cellular strain closely modulates the release of intracellular FGF-2 from human vascular smooth muscle cells, but FGF-2 release is negligible in response to the smaller strains that occur in the normal artery. In addition, larger mechanical strains lead to transfer of intracellular FGF-2 to the extracellular low-affinity receptors, where FGF-2 may be displaced by heparin. These observations provide insight into the mechanisms by which deforming vascular injury, such as that produced by arterial interventions, may elicit a proliferative response.
Key Words: atherosclerosis • biomechanics • vascular smooth muscle • fibroblast growth factor
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Introduction |
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Mechanical forces may regulate cellular function in physiological and pathological states. Vascular cells are subjected to a dynamic mechanical environment modulated by pulsatile pressure and oscillatory shear forces. The accompanying stresses may regulate normal vascular tone1 and contribute to atherogenesis,2 the vascular hypertrophy associated with hypertension,3 and the acute rupture of atherosclerotic lesions.4 5 The mechanical forces associated with certain therapeutic interventions may also induce important biological responses. Coronary balloon angioplasty, a common treatment for occlusive vascular disease, imposes extreme vessel wall strains, which may play a major role in restenosis.6 7
Fibroblast growth factor-2 (FGF-2) participates in the early smooth muscle cell migratory and proliferative responses following arterial injury.8 FGF-2 may exert autocrine or paracrine mitogenic effects on endothelial cells and vascular smooth muscle cells, because these two cell types as well as a variety of other cells both produce and respond to this mediator.9 10 11 12 FGF-2 occurs in multiple forms13 14 distributed differentially in the cytoplasm and nucleus.15 16 17 18 FGF-2 may bind to receptors on a broad range of target cells to regulate growth, differentiation, tissue regeneration, and angiogenesis. However, despite its apparent extracellular mode of action, FGF-2 is not found in vesicles9 and lacks a classical hydrophobic leader sequence for secretion by the exocytotic pathway.19
FGF-2 has been shown to be released from cells in vitro by cytosolic leakage through injured cell membranes.20 21 22 23 Biochemical stimuli24 25 and also mechanical trauma, such as scraping21 and crushing,26 have been shown to induce FGF-2 release from cultured cells. Wounding by needle puncture causes skeletal myofibers to release a variable amount of FGF-2 that correlates with the frequency of membrane disruptions.27 However, healthy cells under little or no mechanical stimulus also may release FGF-2. In cultured endothelial cells, neutralizing antibodies to FGF-2 inhibit basal levels of protease production and DNA synthesis. Physiological activities, such as contraction of cardiac myocytes28 and migration of endothelial cells,29 induce FGF-2 release, suggesting that its elaboration requires neither cell death nor extreme injury.
Once released, FGF-2 may bind not only to specific cell surface receptors but also (with lower affinity) to heparan sulfate proteoglycans and other pericellular matrix components.30 31 These low-affinity nonsignaling receptors may enhance growth factor activity32 by raising local growth factor concentrations or promoting receptor binding33 or by other mechanisms. The matrix may also serve as an FGF-2 reservoir, which may be released rapidly under certain conditions.34 35 36 37
Although FGF-2 can modulate a variety of cellular functions in vitro, the function of this growth factor in physiological and pathological states is unclear in part because the regulation of its release is incompletely understood. The ability of a broad range of mechanical stimuli to induce release of FGF-2 in vitro suggests that FGF-2 release and function in vivo may vary with local mechanical conditions. Many cells that synthesize FGF-2 also experience mechanical deformation, or strain, in vivo; depending on the type of cell and anatomic location, these strains may be cyclical or intermittent, short or long in duration, and rapid or slow in onset. Thus, understanding the specific role of mechanical strain in FGF-2 release may help elucidate the function of this growth factor.38 Additionally, the relative roles of the matrix and intracellular reservoirs of FGF-2 have not been characterized for a mechanical stimulus. We have recently characterized FGF-2–mediated DNA synthesis in response to a transient mechanical compression of a three-dimensional smooth muscle cell–collagen matrix culture.39 Although the DNA synthetic response correlated with the magnitude of the compressive strain, the dependence of FGF-2 release on cellular deformation, the extent of cellular injury, and the role of extracellular FGF-2 binding could not be readily assessed.
In the present study, we used a device that applies homogeneous and uniform biaxial mechanical strain to cultured cells, allowing precise control of strain amplitude, to address the following hypotheses: (1) brief oscillatory mechanical strain induces rapid release of FGF-2 from human vascular smooth muscle cells; (2) release of FGF-2 and cellular injury depend on the amplitude, frequency, and the number of cycles of the mechanical strain; (3) FGF-2 is released primarily from the intracellular pool; and (4) once released, FGF-2 may be sequestered by the extracellular low-affinity receptors. The results described below indicate that FGF-2 release is tightly controlled by mechanical strain and is particularly increased at amplitudes of strain that would occur during arterial interventions, such as balloon angioplasty.
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Materials and Methods |
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Mechanical Strain Device
Mechanical tensile deformation was applied to a thin transparent membrane on which cells were cultured, an approach that produces controlled cellular strain and visualization of cells. Modified from Schaffer et al,40 the device used in this study differs from several other membrane stretch devices in that it provides a nearly homogenous and uniform biaxial strain profile, ie, strains that are equal at all locations on the membrane and in all directions. Cells on up to four individual dish membranes may be deformed by controlled displacement of a platen assembly (Fig 1
). Each culture dish consists of a plastic (Kynar or polysulfone) cylinder (height, 32 mm; inner diameter, 76.2 mm; and outer diameter, 88.9 mm) and a circular silicone elastomeric membrane (Silastic, 0.005-inch thickness, Specialty Mfg), which serves as the culture surface. In a diaphragmatic configuration, the membrane is secured with a tightly fitting rubber O-ring (Somerville Rubber) to a groove in the bottom rim of the dish. The dish is autoclavable, covered by a sterile plastic Petri dish lid, and individually mounted to the top cassette with a locking collar.
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The membrane is deformed from below by a slightly smaller, circular platen (aluminum or polytetrafluoroethylene-impregnated Delrin) that is free to move through a hole in the cassette. Friction between the platen and membrane is minimized by a chemically inert, silicone-impermeant grease (Braycote 804, Castrol); a low-friction interface is important for achieving spatially uniform strains.40
The platens are mounted on a rigid aluminum plate secured to a center steel shaft and two lateral shafts, which are guided by glass-filled polytetrafluoroethylene bearings. The center shaft rides on an eccentric circular cam shaft that is driven by a direct-current feedback-controlled gear motor; at the lowest position on the cam, the membranes are in slight contact with the platens but remain in their initial undeformed state. Thus, the membranes undergo cyclical tensile deformation as the platen assembly moves sinusoidally with a frequency and amplitude determined by the motor speed and the cam size, respectively. The motor speed can be varied within a continuous range of 0 to 1 Hz, and the cams produce displacement amplitudes of 3.1, 5.4, 7.3, 9.4, and 17 mm, corresponding here to nominal peak strains of 2%, 5%, 10%, 15%, and 30%.
The plane strains in a thin circular membrane deformed by a circular platen have been shown previously to be homogeneous and biaxially uniform.40 41 Membrane strains for the current device were measured to determine the strain amplitude for each cam and verify strain uniformity. Thirty-five landmarks were attached to the surface of the membrane. Under oscillatory membrane deformation (1 Hz), the positions of the landmarks relative to a rectangular coordinate system in the plane of the membrane were measured with a video imaging system (Qualisys Inc), which tracks the movement of landmarks at an acquisition frequency of 60 Hz and a spatial resolution of 1:50 000 along the diagonal of the field of view. From the undeformed and maximally deformed membrane configurations, the associated membrane strains in the vicinity of each landmark were determined by computing the strain tensor for each group of three landmarks, as previously described.42 The strain components were transformed with respect to a cylindrical coordinate system at the center of the circular membrane. The radial (Fig 2A) and circumferential (Fig 2B) strains were essentially equal (ie, biaxially uniform) for a given cam displacement and independent of radial position on the membrane (ie, homogeneous).
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Cell Culture
DMEM and other cell culture reagents were obtained from GIBCO BRL. Human vascular smooth muscle cells were derived from explants of discarded portions of saphenous veins obtained during coronary bypass surgery performed at Brigham and Women's Hospital. Smooth muscle cells were cultured in DMEM with 10% FCS at 37°C and 5% CO2. These conditions are selective for growth of smooth muscle cells over endothelial cells.43 The explant and culture methods were identical to those used in previous studies of cultured vascular smooth muscle cells.44 Cells were cultured through passages 3 through 5 before transfer to membrane dishes for use in mechanical strain experiments. Sterilized membrane dishes were coated with human plasma fibronectin overnight at 4°C, washed with PBS, and plated with cells at a density of 5x103 cells/cm2. Cells were incubated overnight in DMEM supplemented with 10% FCS to permit adhesion and spreading. For serum-free conditions, cells were washed twice with DMEM and cultured for 24 hours in defined serum-free IT medium (equal volumes of DMEM and Ham's F-12 supplemented with 1 µmol/L insulin and 5 µg/mL transferrin). Before the experiment, serum-free medium was again changed to minimize residual growth factors.
Application of Mechanical Strain
For serum-free conditions, cells were washed twice with DMEM and cultured for 24 hours in defined serum-free medium. Before the experiment, serum-free medium was again changed to minimize residual growth factors. Mechanical strain was applied at a specified constant frequency and amplitude, and control parallel dishes received no mechanical strain but were otherwise treated identically.
To verify that membrane strain was transferred to cells, control experiments were performed under static membrane deformation. Cells were deformed by manually lowering the dish onto a platen, identical in dimensions to that in the strain device, mounted on a microscope stage. Fluorescent microspheres (Fluoresbrite, 1-µm diameter, Polysciences) were suspended in culture media and allowed to settle onto the cell surface for 30 minutes. Excess microspheres were eluted to leave a random array of remaining microspheres on the cell surfaces. For these measurements, microspheres from neighboring cells were used, so that these measurements did not determine intracellular strains. The microsphere displacements were derived from fluorescence photomicrographs taken in the undeformed and deformed configurations; ink dot markings on the membrane were used as landmarks so that the same aggregates of microspheres were photographed. Under these conditions, there was no significant difference between the cell-to-cell and membrane strains.
Measurement of FGF-2
To evaluate the time course of FGF-2 release, media aliquots were collected at various times after beginning strain. At the end of the experiment, the combined intracellular and receptor-bound FGF-2 was harvested with 2 mol/L NaCl, pH 7.5, and three cycles of freeze/thaw. In some experiments, the cell layer was eluted with 2 mol/L NaCl, pH 7.5, and 2 mol/L NaCl, pH 4.0, to harvest FGF-2 bound to the low- and high-affinity receptors, respectively.45 46 47 Alternatively, FGF-2 bound to the low-affinity receptors was harvested by treatment with heparan sulfate, heparitinase,30 or heparin.48 Media, elutions, and cell lysates were assayed for FGF-2 with a quantitative enzyme immunoassay (Quantikine human FGF basic immunoassay, R&D Systems).
Measurement of Cellular Injury
Fluorescent labels were obtained from Sigma Chemical Co. To evaluate the acute detachment of cells from the culture substrate, cell adhesion was measured before and immediately after strain. Cellular nuclei were stained with acridine orange (2 µg/mL) and photographed under fluorescence microscopy at random locations marked on the underside of the membrane. Cells were subjected to mechanical strain and rephotographed at the identical locations.
To assess plasma membrane integrity, cells were subjected to mechanical strain and immediately stained with propidium iodide and fluorescein diacetate (10 µg/mL) for several minutes. Propidium iodide and fluorescein diacetate stains have been used previously to identify injured cells and are more sensitive markers than lactate dehydrogenase release.49 50 Alternatively, plasma membrane disruption was detected by fluoresceinated dextran uptake, which has been used previously to assess sublethal cell injury in the context of FGF-2 release.21 22 27 Before mechanical strain, cells were changed to serum-free media with fluoresceinated dextran (molecular weight, 10 000; 2.5 mg/mL). Cells were subjected to mechanical strain, incubated for 10 minutes, and washed twice with PBS. After labeling with either method, cells were photographed under fluorescence microscopy at several random membrane locations.
Statistics
Data are presented as mean±1 SD. Comparison among groups of continuous variables was performed using ANOVA. For comparison between specific groups of continuous variables, the two-sample Student's t test was used. A value of P<.05 was considered statistically significant. The findings reported below are representative of results of at least three independent experiments from at least two different cell donors.
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Results |
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Release of FGF-2 During Dynamic Strain
Continuous oscillatory mechanical strain (1 Hz, 14%) of human vascular smooth muscle cells induced the rapid release of FGF-2 (Fig 3
); after 10 minutes of strain, media contained 3.2±0.2% of the total initial cellular FGF-2 (versus 0.3±0.2% for unstrained control, P<.005); after 30 minutes, media contained 3.5±1.2% of the total FGF-2 (versus 0.4±0.2% for control, P<.005); and after 60 minutes, media contained 3.8±1.1% of the total FGF-2 (versus 0.3±0.3% for control, P<.001). The total initial cellular FGF-2 in each dish was estimated as the combined media and cell-associated (intracellular and receptor-bound) FGF-2 at the end of the experiment. Additional experiments were performed to assess the dependence of release on characteristics of the mechanical strain.
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) of human vascular smooth muscle cells, relative to control (
) (n=3 to 7 for each measurement; error bars denote 1 SD).
Dependence of FGF-2 Release on Characteristics of Strain
Release of FGF-2 increased with the number of cycles of mechanical strain (Fig 4); 60 minutes after one cycle of strain (1 Hz, 14% amplitude), media contained 2.9±1.5% of the total initial cellular FGF-2 (versus 0.9±0.4% for unstrained control, P=NS); 60 minutes after 9 cycles of strain, media contained 4.8±0.4% of the total FGF-2 (P<.05 versus control); and after 90 cycles, media contained 7.8±0.7% of the total FGF-2 (P<.005 versus one cycle). More than 90 cycles released no additional FGF-2; after 900 cycles and 3600 cycles of strain, respectively, media contained 7.2±1.2% and 6.2±0.9% (P=NS versus 90 cycles) of the initial total cellular FGF-2, suggesting a limited releasable FGF-2 pool for a given combination of strain frequency and amplitude. In a separate experiment, cells were exposed to strain (1 Hz, 14% amplitude) of either 90 cycles or 
90 000 cycles with a media change at 150 minutes after beginning strain; cumulative FGF-2 in the media increased comparably for both strain protocols before the media change but decreased to baseline control thereafter (data not shown). Thus, FGF-2 was rapidly released in response to either brief (1-minute) or prolonged (24-hour) oscillatory strain, but release did not continue 150 minutes after strain was begun. Further characterization of FGF-2 release was performed primarily for 90 cycles of strain.
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), and 90 cycles (90 seconds) (
); however, FGF-2 was not further increased for 900 cycles (15 minutes) (
) (n=3 for each measurement; error bars denote 1 SD).
Cyclic strain at 14% amplitude induced FGF-2 release in a frequency-dependent fashion (Fig 5); 60 minutes after beginning strain (90 cycles, 0.25 Hz), media contained 4.8±0.8% of the total initial cellular FGF-2 (versus 0.4±0.6% for control, P<.01 by ANOVA); for strain at 0.5 Hz, media contained 10.4±0.8% of the total FGF-2 (P<.001 versus 0.25 Hz); and for strain at 1 Hz, media contained 13.8±1.2% of the total FGF-2 (P<.05 versus 0.5 Hz). Mechanical release of FGF-2 was also dependent on the amplitude of strain (1 Hz) (Fig 6); 60 minutes after beginning strain (90 cycles, 4%) amplitude, media contained 0.1±0.1% (versus 0.0±0.1% for control, P=NS) of the total initial cellular FGF-2; for strain at 14% amplitude, media contained 5.7±0.5% (P<.05 versus 4% amplitude); and for strain of 33% amplitude, media contained 18.9±2.7% (P<.001 versus 14% amplitude). In another experiment, 90 cycles (1 Hz) at 14% amplitude strain induced rapid release of FGF-2, whereas 90 000 cycles at 4% amplitude induced little release throughout the 24-hour period (data not shown). Although the percentage of FGF-2 released by a given mechanical stimulus varied between cell preparations, the dependence on frequency and amplitude was always observed, and the amplitude threshold between 4% and 14% was a stringent requirement for FGF-2 release.
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Cellular Injury
To evaluate the possibility that the increase in FGF-2 in the media was caused simply by acute detachment of cells from the membrane during the mechanical strain, photographs of identical regions of membranes were compared both before and after strain. No significant detachment occurred acutely after mechanical strains used in the present study. In addition, total cell number both before and after mechanical strain was compared with the percentage of total cellular FGF-2 released by image analysis of random regions of membranes. These studies demonstrated negligible decreases in cell numbers under conditions of significant FGF-2 release. For example, after mechanical strain (90 cycles, 1 Hz, 14%), the change in number of cells was -0.7±1.1% compared with the fraction of cellular FGF-2 released (11.5±0.6%, P<.001). Thus, although we cannot exclude the possibility that cellular detachment and subsequent lysis contributed to a small component of FGF-2 found in the media, the vast majority was derived from the intact cell layer.
Cellular injury by mechanical strain was detected by dual staining with fluorescein diacetate and propidium iodide. Propidium iodide stains the nuclei of injured cells by diffusing through disrupted plasma membranes, whereas fluorescein diacetate stains healthy cells with intact plasma membranes; together, these markers have been shown to be more reliable than either lactate dehydrogenase or trypan blue exclusion in the assessment of cell injury.50 51 Mechanical strain led to increased propidium iodide staining of a subpopulation of cells distributed diffusely on the culture membrane. The percentage of propidium iodide–stained cells increased with the amplitude of strain (90 cycles, 1 Hz) (Fig 7A). This amplitude dependence of cellular injury was also detected by fluoresceinated dextran uptake (Fig 7B); because dextran is retained only by cells whose membranes have resealed,21 27 52 these observations suggest transient sublethal injury to cellular strain.
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Role of Low-Affinity Receptors in FGF-2 Release
Heparin-like molecules and certain matrix degradative enzymes have been shown to either release FGF-2 prebound to the low-affinity receptors or compete with these receptors for binding of free FGF-2.30 In control experiments, treatment of quiescent smooth muscle cells with heparin, heparitinase, or heparan sulfate caused comparable release of FGF-2 (1% of the total) into the media; elution with 2 mol/L NaCl released a negligible amount of FGF-2 (data not shown). To evaluate the relative contributions by the intracellular and the low-affinity receptor FGF-2 pools to the mechanical release of FGF-2, cells were pretreated with heparin (5 µg/mL) before mechanical strain. Heparin treatment released <1% of the total cellular FGF-2 (Fig 8). Mechanical strain (90 cycles, 33%, 1 Hz) after heparin treatment and removal of heparin released 12.6±1.6% of the total initial cellular FGF-2 (versus 15.8±0.9% for strain alone, P<.05). Thus, most FGF-2 was released into the media from the nuclear or cytoplasmic pools and not from the low-affinity receptors.
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To assess the amount of FGF-2 that is sequestered by the low-affinity receptors after mechanical release, cells were exposed to strain (90 cycles, 33%, 1 Hz) and incubated with heparin (5 µg/mL) to block receptor binding of the FGF-2 as it was being released. After 60 minutes, media with and without heparin, respectively, contained 25.2±3.5% and 15.6±2.6% (P<.05) of the total initial cellular FGF-2 (Fig 9). Conversely, parallel cells were exposed to strain, incubated for 1 hour, and treated with fresh heparin-supplemented media to liberate any FGF-2 that was potentially transferred by mechanical strain to the low-affinity receptors. Media with and without heparin, respectively, contained 11.0±0.7% and 4.2±0.1% (P<.001) of the total initial cellular FGF-2. Thus, approximately one third of the mechanically released FGF-2 was subsequently bound to the low-affinity receptors.
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Discussion |
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Although FGF-2 is a ubiquitous cytokine that can mediate a variety of cellular responses in vitro, the regulation of its release is incompletely understood. FGF-2 has been referred to as a "wound hormone" by McNeil and colleagues,27 28 53 who made the important observation that mechanical injury to cells releases FGF-2. Both overt trauma and mild mechanical stimuli may induce release of FGF-2, suggesting that FGF-2 may serve important but potentially different roles in physiological and pathological states.
The present data demonstrate tight control of FGF-2 release by cellular strain and are summarized as follows. Release of FGF-2 was regulated by a strain-amplitude threshold. At amplitudes below 10%, prolonged (105 cycles at 1 Hz) oscillatory strain did not cause release of FGF-2; however, brief (102 cycles) oscillatory strain above 10% amplitude induced rapid release of the growth factor, even after the stimulus was discontinued. When strain exceeded the amplitude threshold, FGF-2 release increased with the amplitude and frequency of strain. In addition, release of FGF-2 increased with the number of cycles of strain but was maximal for a relatively brief stimulus (102 cycles, 1 Hz). FGF-2 release began within 5 minutes of the onset of strain and was complete within 150 minutes, even for prolonged (105 cycles) oscillatory strain. Taken together, these data suggest that oscillatory cellular strain above an amplitude threshold induces an acute FGF-2 release response determined primarily by the frequency and amplitude of the early (102 cycles) mechanical stimulus. In addition, FGF-2 release does not appear to require an ongoing injury if the acute stimulus is sufficient.
The rapid and limited nature of FGF-2 release in response to cellular strain is consistent with a cell injury–mediated mechanism. Cellular injury occurred in parallel with FGF-2 release; a subpopulation of cells was injured for strain above an amplitude threshold of 10%, and injury increased with the amplitude and frequency of strain. One interpretation of these observations is that cellular injury is determined independently by the strain amplitude and frequency. Alternatively, cellular injury may be determined by a single parameter, the strain rate; for sinusoidal strain, e=Asin(
t), the strain rate (de/dt) is a function of both the frequency () and the amplitude (A) of strain: de/dt=Acos(t). Thus, whether cellular injury and FGF-2 release increased with the amplitude of strain independent of the increase in strain rate was not distinguished in these experiments. Amplitude dependence could be tested by performing experiments in which the amplitude is increased but the frequency is correspondingly adjusted to maintain a constant strain rate. Strain rate has been shown to regulate injury to astrocytes caused by a single strain impulse of constant magnitude.49
Cells subjected to external mechanical manipulation, such as scraping22 53 or puncture,27 suffer transient resealable cell membrane disruptions through which FGF-2 may be released. Our experiments demonstrate that cellular perturbation via deformation of the underlying substrate induces a transient increase in cell membrane permeability.27 54 Although plasma membrane disruptions were not localized, one possible scenario is that mechanical failure occurs near integrins at focal adhesions as they transmit forces from the underlying matrix.55 56 57 The process may be similar to the disruption and detachment of focal adhesions observed at the trailing edge of migrating cells,58 since migration is associated with release of intracellular FGF-2, possibly through permeabilized cell membranes.29
In addition to the intracellular FGF-2 pool, the extracellular receptor-bound FGF-2 is also a potential source of mechanically releasable growth factor. FGF-2 binds to both high-affinity signaling receptors and (with lower affinity) to heparan sulfate and other cell surface proteoglycans.31 32 These low-affinity receptors may serve as a high-capacity reservoir for FGF-2, which may be released enzymatically.34 37 Heparin and heparin-like molecules may competitively inhibit binding of the low-affinity receptors to free FGF-2. In these experiments, treatment with heparin, heparan sulfate, or heparitinase demonstrated release of FGF-2 primarily from the intracellular compartment, with only a small fraction arising from the low-affinity receptor binding. It is important to note that the relative baseline intracellular and extracellular distributions of FGF-2 may be altered by cell density or other culture conditions.48 In addition, the monolayer cell culture may not adequately simulate FGF-2 binding to and potential mechanical release from a three-dimensional matrix in vivo.
The low-affinity receptors are required for some FGF-induced biological responses and may help stabilize,59 present, and promote binding of the growth factor to the high-affinity signaling receptors.33 Subsequent dimerization of the high-affinity receptors may also depend on interactions with these proteoglycans.60 Another function of the low-affinity receptors may be to minimize FGF-2 diffusion away from the site of release.32 Previous studies suggest the potential for the level of FGF-2 in the culture medium to underestimate total FGF release.29 39 In this study, mechanical strain induced the transfer of FGF-2 from the intracellular cytoplasmic and/or nuclear compartments to the extracellular receptors. Thus, the low-affinity receptors can increase the effective concentration of the growth factor near the site of release, presumably enhancing the probability of interactions with the high-affinity receptors. The role of cell surface proteoglycans in concentrating FGF-2 near the site of injury may be particularly important for targeting appropriate angiogenic responses.61
Interestingly, neighboring cells exposed to nearly identical membrane substrate strain were nonuniformly injured, reflecting possible differences in intracellular strains. Vascular smooth muscle exposed to biaxial deformation experiences higher local strains in the pseudopods than in the cell body.42 Because the strain profile in a given cell under these experimental conditions may depend on its particular cytoskeletal arrangement and focal adhesion distribution, sensitivity to injury may be variable within a population of cells. Under baseline conditions, FGF-2 was localized by immunofluorescence to the nucleus and cytoplasm with no perceptible differences between adjacent cells; staining intensity decreased modestly after strain but remained evenly distributed between cells (data not shown). These observations raise the possibility that unlike cellular injury, FGF-2 is released uniformly from the population of strained cells. Alternatively, these observations may reflect different sensitivities in the methods used to detect injury and to localize FGF-2. In addition, the existence of two discrete intracellular FGF-2 compartments may complicate the estimation of total FGF-2 by immunostaining.
Responses to mechanical loading have been studied in vascular smooth muscle and other types of cells using a variety of devices. Mediated by various signal transduction mechanisms,1 62 63 64 the observed responses include early gene induction,65 66 proliferation,67 68 69 morphological changes,70 71 matrix metabolism,72 73 and release of growth factors and other cytokines.26 Despite the diversity of these mechanoresponses, the present study imposing a homogeneous and uniform biaxial strain to all cells within a culture underscores the potential regulation by specific components of the applied strain. Although the frequency of oscillation is controlled, certain studies that impose spatially nonhomogeneous and biaxially nonuniform strains to cells may be difficult to interpret. With a spatially varying strain profile, neither the strain amplitude nor the strain rate is consistently delivered to all cells within the culture well. Under these conditions, the observed response might not be controlled directly by mechanical strain but may be modulated in a paracrine fashion by an underlying strain amplitude–dependent or strain rate–dependent response, such as FGF-2 release. With a biaxially nonuniform strain profile, cells within the culture dish may undergo variable compressive, tensile, and shear strains depending on their orientation on the membrane; such components of strain may differentially regulate some cellular responses. Thus, the experimental application of homogeneous and biaxially uniform mechanical stimulation may be critical for understanding the regulation of certain mechanoresponses. It should also be noted that in vivo arterial strains are not symmetric and that extrapolation of these experiments to cellular strains in the arterial wall requires caution. However, it is intriguing that normal arterial strains are 10% (depending on the age of the patient and the artery type),74 the approximate threshold of strain for FGF-2 release in these experiments.
The dependence of FGF-2 release on mechanical strain suggests a potential model for the regulation of FGF-2 release in pathophysiological states. Under long-term oscillatory deformation, vascular smooth muscle cells may synthesize and maintain an intracellular FGF-2 pool of which little is released. This reservoir remains intact and is depleted only when cellular strain exceeds an amplitude threshold. Under conditions of elevated heart rate or of mechanical stress, such as hypertension, or at focal regions of atherosclerotic plaque,4 75 76 the strain rate may be sufficient to injure a subset of cells, which then release FGF-2. Under conditions of extremely high mechanical strain, such as during balloon angioplasty, a majority of cells may be injured, rapidly releasing a large amount of FGF-2, particularly to extracellular low-affinity receptors.
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Acknowledgments |
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This study was supported in part by a Grant-in-Aid from the American Heart Association, Massachusetts Affiliate, a grant from the Goldsmith Foundation, and grants from the National Institutes of Health (HL-47840, HL-48743, HL-54759, and AR-41352).
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Footnotes |
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This manuscript was sent to Howard E. Morgan, Consulting Editor, for review by expert referees, editorial decision, and final disposition.
Received August 27, 1996; accepted October 11, 1996.
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D. J. Tschumperlin and S. S. Margulies Equibiaxial deformation-induced injury of alveolar epithelial cells in vitro Am J Physiol Lung Cell Mol Physiol, December 1, 1998; 275(6): L1173 - L1183. [Abstract] [Full Text] [PDF]
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D. W. Courtman, A. Cho, L. Langille, and G. J. Wilson Eliminating Arterial Pulsatile Strain by External Banding Induces Medial but Not Neointimal Atrophy and Apoptosis in the Rabbit Am. J. Pathol., December 1, 1998; 153(6): 1723 - 1729. [Abstract] [Full Text] [PDF]
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K. Yamamoto, Q. N. Dang, R. A. Kelly, and R. T. Lee Mechanical Strain Suppresses Inducible Nitric-oxide Synthase in Cardiac Myocytes J. Biol. Chem., May 8, 1998; 273(19): 11862 - 11866. [Abstract] [Full Text] [PDF]
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J.-H. Yang, W. H. Briggs, P. Libby, and R. T. Lee Small Mechanical Strains Selectively Suppress Matrix Metalloproteinase-1 Expression by Human Vascular Smooth Muscle Cells J. Biol. Chem., March 13, 1998; 273(11): 6550 - 6555. [Abstract] [Full Text] [PDF]
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K. Tamura, Y. E. Chen, M. Lopez-Ilasaca, L. Daviet, N. Tamura, T. Ishigami, M. Akishita, I. Takasaki, Y. Tokita, R. E. Pratt, et al. Molecular Mechanism of Fibronectin Gene Activation by Cyclic Stretch in Vascular Smooth Muscle Cells J. Biol. Chem., October 27, 2000; 275(44): 34619 - 34627. [Abstract] [Full Text] [PDF]
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R. T. Lee, C. Yamamoto, Y. Feng, S. Potter-Perigo, W. H. Briggs, K. T. Landschulz, T. G. Turi, J. F. Thompson, P. Libby, and T. N. Wight Mechanical Strain Induces Specific Changes in the Synthesis and Organization of Proteoglycans by Vascular Smooth Muscle Cells J. Biol. Chem., April 20, 2001; 276(17): 13847 - 13851. [Abstract] [Full Text] [PDF]
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T. P. Quinn, M. Schlueter, S. J. Soifer, and J. A. Gutierrez Mechanotransduction in the Lung: Cyclic mechanical stretch induces VEGF and FGF-2 expression in pulmonary vascular smooth muscle cells Am J Physiol Lung Cell Mol Physiol, May 1, 2002; 282(5): L897 - L903. [Abstract] [Full Text] [PDF]
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R. Dono, J. Faulhaber, A. Galli, A. Zuniga, T. Volk, G. Texido, R. Zeller, and H. Ehmke FGF2 Signaling Is Required for the Development of Neuronal Circuits Regulating Blood Pressure Circ. Res., January 11, 2002; 90 (1): e5 - e10. [Abstract] [Full Text] [PDF]
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G. W. De Keulenaer, Y. Wang, Y. Feng, S. Muangman, K. Yamamoto, J. F. Thompson, T. G. Turi, K. Landschutz, and R. T. Lee Identification of IEX-1 as a Biomechanically Controlled Nuclear Factor-{kappa}B Target Gene That Inhibits Cardiomyocyte Hypertrophy Circ. Res., April 5, 2002; 90(6): 690 - 696. [Abstract] [Full Text] [PDF]
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