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J Appl Physiol 98: 242-249, 2005. First published August 20, 2004; doi:10.1152/japplphysiol.01006.2003
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Vascular smooth muscle cell glycocalyx influences shear stress-mediated contractile response

Kristy M. Ainslie,1 Jeffrey S. Garanich,2 Randal O. Dull,3 and John M. Tarbell4

Biomolecular Transport Dynamics Laboratory, Departments of 1Chemical Engineering and 2Bioengineering, The Pennsylvania State University, University Park, Pennsylvania; 3Department of Anesthesiology, The University of Utah, Salt Lake City, Utah; and 4Department of Biomedical Engineering, The City College of New York, New York, New York

Submitted 15 September 2003 ; accepted in final form 2 August 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
This study addressed the influence of the rate of shear stress application on aortic smooth muscle cell (SMC) contraction and the role of specific glycosaminoglycans in this mechanotransduction. Rat aortic SMCs were exposed to either a step increase in shear stress (0 to 25 dyn/cm2) or a ramp increase in shear stress (0 to 25 dyn/cm2 over 5 min) in a parallel plate flow chamber, and cell contraction was characterized by cell area reduction. SMCs contracted at levels similar to those reported previously and equally in response to both a step and ramp increase in shear stress. When the cells were pretreated with heparinase III or chondroitinase ABC to remove the glycosaminoglycans heparan sulfate and chondroitin sulfate, respectively, from the glycocalyx, the contraction response to increases in shear stress was significantly inhibited. These studies indicate that specific components of the SMC glycocalyx play an important role in the mechanotransduction of shear stress into a contractile response and that the rate of application of shear stress does not affect the SMC contraction.

glycosaminoglycans; myogenic response

SMOOTH MUSCLE CELLS (SMC) normally reside in the tunica media of the arterial wall and are not exposed directly to fluid flow shear stress associated with blood flow. SMCs can be exposed directly to blood flow when the intima and internal elastic lamina (IEL) are damaged in procedures such as angioplasty or in the anastomotic region of vascular grafts (27, 41). Experiments in animal models of atherosclerosis and intimal hyperplasia have shown that luminal SMCs may be present from days to months after a vascular procedure (10, 19). These exposed SMCs will experience vascular fluid shear stresses on the order of 10–20 dyn/cm2 (time-averaged value). SMCs present in intact vessels are shielded from blood flow but are, however, exposed to fluid shear stress due to transmural interstitial flow. Wang and Tarbell (50) predicted wall shear stresses associated with transmural interstitial flow on the order of 1–3 dyn/cm2. Furthermore, it has been predicted that the most superficial layer of SMCs, lying directly beneath the IEL, may be exposed to higher levels of shear stress due to the funneling of flow through the fenestral pores in the IEL, on the order of 10–50 dyn/cm2 (51).

Recent in vitro work by Civelek et al. (8) has shown that a step increase in shear stress elicits contraction in rat aortic SMCs. Cells starved of serum, which alters the cell phenotype into the contractile state, exhibited a contractile response at 11 dyn/cm2 of shear stress (8). Most recently, an in vivo study has shown that transmural flow plays a significant role in the myogenic response (24). This group perfused arterioles with an osmotic supplement (BSA-Ficoll) to reduce transmural flow at fixed pressure and observed a significant attenuation of myogenic contraction in response to a step increase in pressure. Ficoll is a nonionic synthetic polymer of sucrose, whose concentration can be varied to alter the osmotic pressure of the luminal media. These in vivo results support the model proposed by Wang and Tarbell (50) and the in vitro work reported by Civelek et al. (8), indicating dependence of the myogenic response on transmural flow and associated shear stress.

Bayliss initially described the myogenic response as the contraction of a vessel (reduction of diameter) after an increase in pressure, and it is now known to be a critical mechanism in the control of blood flow (26, 28). It is widely believed that the myogenic response is driven by the initial stretch of SMC that occurs right after a step increase in pressure (12, 18). However, recent studies have shown that, when pressure is increased in a ramp fashion without an accompanying stretch, a myogenic response still develops that is comparable to the step response in the steady state (22). A hypothesis of the present study is that the increase in pressure that drives the myogenic response produces an increase in transmural interstitial flow and associated shear stress by a classic Starling mechanism (transmural flow is proportional to the transmural pressure differential) and that it is the fluid shear stress on SMC that drives the steady-state myogenic response. Civelek et al. (8) showed that a step increase in fluid shear stress induced SMC contraction in vitro. In the present study, we ask whether a ramp increase in fluid shear stress can induce SMC contraction comparable to a step increase in the steady state.

In addition to that of Civelek et al. (8), there have been a number of other studies that have shown that SMCs are responsive to fluid shear stress (20, 33, 43, 47, 51). The mechanism by which SMCs sense and transduce changes in shear stress into a cellular response has not been determined. It has been suggested in the literature that the glycocalyx is responsible for the sensing of flow on the endothelial cell (EC) surface (42, 44). The glycocalyx of cardiovascular cells (ECs, SMCs, and fibroblasts) is a surface layer consisting primarily of proteoglycans and glycoproteins that are incorporated into the cell membrane. Proteoglycans contain a protein backbone with extended polysaccharide branches composed of glycosaminoglycans (GAGs). The most common protein backbones found in the intima and media are syndecans and glypicans. Syndecans span the cell membrane and possess continuous extracellular and intracellular domains (5, 36). They have been linked to several cellular functions, including cell proliferation and cell-matrix and cell-cell adhesion (6). In contrast, glypicans terminate in the cell membrane (36). Heparan sulfate and chondroitin sulfate are the most frequently found GAGs in the intima and media of an artery (36, 38). On the SMC surface ~50–60% of the GAGs are chondroitin sulfate, and the remainder are predominantly heparan sulfate (30). Glycoproteins, composed of a protein backbone with much smaller sugar residues, such as integrins and selectins, are also present on the EC and SMC surface (36).

Suarez and Rubio (48) determined that, in guinea pig hearts, coronary flow stimulates glycolytic flux through shearing forces acting on specific components of the glycoclayx of the ECs lining the heart. Recent work by Florian et al. (16) demonstrated that the heparan sulfate GAGs on EC surfaces are integral to the shear-mediated production of nitric oxide (NO). Depletion of the heparan sulfate GAGs with heparinase III treatment significantly reduced shear-mediated NO production by ECs (16). These studies have led us to hypothesize that the glycocalyx is a mechanosensor on SMCs that mediates the contraction response of fluid shear stress observed by Civelek et al. (8).

To assess SMC contraction in response to shear stress, we employed a parallel-plate flow chamber and programmable syringe pump. The contraction responses to a 25 dyn/cm2 step and 5-min ramp up to 25 dyn/cm2 were determined and compared with each other and a no-flow control. Our results indicate that the step and ramp induce indistinguishable steady-state contraction responses. To determine the role of the glycocalyx in shear-induced SMC contraction, the enzymes heparinase III and chondroitinase ABC were applied to selectively cleave heparan sulfate and chondroitin sulfate GAGs, respectively, from the glycocalyx. It was observed that enzymatic pretreatment significantly inhibited shear-induced SMC contraction.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Chemicals were purchased from Sigma (St. Louis, MO), unless otherwise noted.

SMC culture.   Rat aortic SMCs were enzymatically isolated from the thoracic aortas of adult male Sprague-Dawley rats (6–7 wk old, ~150 g), as described elsewhere (2). The cell isolation protocol was reviewed and approved by The Institutional Animal Care and Use Committee at The Pennsylvania State University. Cells were grown to confluency in DMEM-F12 supplemented with 100 U/ml penicillin and 100 µg/ml streptomycin (1% P/S) and 10% FBS in culture flasks. SMCs were positively identified by their characteristic "hill-and-valley" morphology.

The contractile phenotype was induced by starving the cells of FBS in culture flasks containing DMEM-F12 + 1% P/S for 2–5 days before experiments. The cells were then detached with trypsin-EDTA (0.07%) and plated on 35 x 75-mm, 1.7-mm-thick quartz slides (Fredrich & Dimmock, Millville, NJ) with DMEM-F12 + 1% P/S at a density of 150,000 cells/slide. Once plated, the cells were starved for 2 additional days in petri dishes. Serum-starved SMCs were characterized by their expression of two contractile proteins, smooth muscle myosin heavy chain (SM-MHC) and smooth muscle {alpha}-actin (described in detail below). All experiments were conducted 2 days postplating, and only passages 3–8 were used.

SM-MHC and {alpha}-actin expression.   Glass slides, 38 x 75 mm, 1-mm thick (Corning Glass Works, Corning, NY), were used in place of quartz slides for these experiments. An immunofluorescence protocol described elsewhere (3) was used to determine SM-MHC expression. A monoclonal IgG1 mouse anti-rat MHC primary antibody (Santa Cruz Biotechnology, Santa Cruz, CA) specific for SM-MHC and a fluorescently labeled Alexa Fluor 488 goat anti-mouse IgG secondary antibody (Molecular Probes, Eugene, OR) were used.

To determine smooth muscle {alpha}-actin expression, starved SMC were fixed via exposure to an ethanol (Fisher Scientific, Fair Lawn, NJ) gradient (70% for 4 min followed by 95% for 20 min). Slides were placed in a –20°C freezer overnight and rehydrated in phosphate buffer solution without calcium and magnesium (PBS; Cellgro, Herndon, VA) for 5 min. Cells were then incubated at 37°C with a monoclonal anti-{alpha}-smooth muscle actin clone 1A4 alkaline phosphatase conjugate for 1 h. Slides were rinsed with PBS for 5 min and incubated with an alkaline phosphatase substrate (SIGMAFAST Fast Red TR/Naphthol AS-MX tablet set) for 15 min at room temperature.

Following all incubations, a drop of Fluoromount-G mounting media (Southern Biotechnology Associates, Birmingham, AL) was added to slides used in both MHC and {alpha}-actin expression experiments. Slides were observed with a x20 objective on a Nikon Eclipse TE2000-E inverted microscope. Negative controls (no primary antibody added) were conducted in both sets of experiments. Cells were counted as "positive" for MHC or {alpha}-actin expression if they visually exhibited significantly more staining than their respective negative control. For each contractile protein, four to five fields of 2–12 cells were counted on each of three to five separate slides.

Shear apparatus.   The parallel plate flow chamber was a modification of the design of Frangos et al. (17). The quartz slide with attached preconfluent cells formed the bottom plate of the flow chamber, and a polycarbonate plate formed the top. A Silastic gasket (SF Medical, Hudson, MA) was used to maintain a uniform gap between the two parallel plates. All three components were held together by an applied vacuum. The chamber was inverted and placed onto the stage of a microscope (Olympus IMT-2). A field of view with ample isolated cells was chosen to be recorded for the time course of the experiment. The image of the cells was recorded for 30 s before the flow was started and then during 30 min of flow. Wall shear stress was calculated assuming fully developed laminar flow between infinite parallel plates, by using the following equation: {tau} = 6 µ/bh2, where {tau} is the wall shear stress, µ is the viscosity, is the fluid flow rate, b is the width of the flow channel, and h is the height of the flow channel (17).

For the ramp and high-viscosity step experiments, the flow chamber was connected to two disposable plastic syringes (BD Scientific) mounted in a KD Scientific dual-loaded syringe pump (KDS210C; KD Scientific, New Hope, PA). The programmable feature of the pump allowed for ramp or step operation. The circulating media was DMEM-F12 + 1% P/S with dextran (average molecular weight: 75,000) at room temperature. Dextran was added to create a high-viscosity DMEM-F12 + 1% P/S solution, measured with a cone and plate viscometer to have a viscosity of 7.03 cP. For low-viscosity step experiments and glycocalyx flow experiments, the chamber was placed in a closed continuous-flow loop circulating DMEM-F12 + 1% P/S at room temperature (viscosity was 0.72 cP). For both flow systems, the same ultimate shear stress level was imposed (25 dyn/cm2).

The microscope was interfaced to a charged-coupled device camera, which was connected to a videocassette recorder and television to record all experiments. Image processing software, Image-Pro Express (Media Cybernetics, Des Moines, IA), was used to gather data from videotapes and calculate cell areas. To do this, the outlines of cells had to be manually traced by using the computer’s mouse, then cell areas at each time point were determined by the software, and reduction in cell area over time was used as the criterion for contraction. Cell area reduction has been used previously as a measure of contraction (8, 15). Individual cell areas at each time point were normalized with respect to their area at 0 min to account for differences in cell size. By this method, each cell had a normalized area of 1 at 0 min; thus the percent area reduction was calculated at subsequent time points.

Enzymatic treatment of glycocalyx.   Heparinase III and chondroitinase ABC were used to enzymatically cleave specific components of the SMC’s glycocalyx. The enzymes were used individually at a concentration of 0.2 U/ml in DMEM + 1% P/S. The preconfluent monolayers of SMC on quartz slides were treated with one of the two enzyme solutions for 30 min preshear in a petri dish placed inside of a 95% air and 5% CO2, 37°C incubator. The slides were washed with fresh DMEM + 1% P/S at 37°C before mounting in the flow chamber.

Protease activity.   The protease activity of the heparinase III and chondroitinase ABC was assayed by using a RediPlate 96-EnzCheck Protease Assay kit (Molecular Probes). A broad spectrum of proteases ranging from metallo-serine, acid, and sulfhydryl proteases can be detected with this kit, and it has been used previously to indicate protease activity of GAG-depleting enzymes (16). The kit was used in accordance with manufacturer’s instructions. The protease activity of 15 mU/ml heparinase III and chondroitinase ABC was compared against known concentrations of chymotrypsin and pronase at 1, 3, and 24 h. All samples of chymotrypsin and pronase were run in triplicate.

Fluorescent labeling of the glycocalyx.   Digital fluorescence imaging was implemented to assess the degree of enzymatic removal of the glycocalyx. For the heparan sulfate GAG imaging, a primary antibody was used that binds specifically to heparan sulfate (HepSS-1; US Biological, Swampscott, MA) (25). The secondary antibody, which bound specifically to HepSS-1 and contained a fluorescent attachment, was Alexa-Fluor 488 goat anti-mouse IgM (Molecular Probes). A biotinylated lectin (Bandeiraea simplicifolia) that binds specifically to the N-acetyl-D-galactosamine residue present on chondroitin sulfate was also employed. A fluorescent-labeled avidin conjugate (NeutriAvidin Alexa-Fluor 350; Molecular Probes) was used to label the lectin.

For GAG visualization, SMCs were grown to confluency on 25-mm glass coverslips in culture-treated six-well plates. The primary antibody or lectin was diluted in PBS to a concentration of 2 µl HepSS-1/ml PBS or 0.005 mg lectin/ml PBS. The primary solutions were incubated with the confluent coverslips in a 95% air and 5% CO2, 37°C incubator for 15 min. The coverslips were washed with fresh PBS at 37°C. The secondary treatments were diluted in PBS to concentrations equal to those of their respective primary treatment. The secondary solutions were incubated with their respective primary treated coverslips for 15 min in an incubator. The coverslips were washed with fresh PBS and inverted on a glass slide.

Labeled GAGs were imaged by using a wide-field fluorescence objective at x20 on an Olympus BX60 digital microscope. Overall fluorescence intensity was calculated for a series of image fields by using Image Pro Plus (Media Cybernetics, Des Moines, IA). Each series of images was a 4 x 4 grid of unique neighboring fields of view. Each slide was imaged in three locations, and three slides were imaged for each treatment case (n = 9). Control cases were performed in which coverslips were imaged in the absence of enzymatic treatment. Other coverslips were treated with enzyme solution, at concentrations used in experimental protocols, for 30 min before primary antibody or lectin incubation. The enzymatic-treated coverslips were washed with fresh PBS at 37°C before antibody or lectin incubation.

Statistics.   Values are means ± SE. Significant differences between groups were analyzed by the general linear model ANOVA by using statistical analysis software (Minitab, State College, PA). The model performs univariate ANOVA, analysis of covariance, and regression for each variable. Comparisons were made by using the Tukey-Kramer method for pairwise comparison at each time point. As part of the general linear model ANOVA results, t-test P values, based on regression analysis, were presented for each comparison. A P value < 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
SM-MHC and {alpha}-actin expression.   Eighty-seven of 122 cells counted (71%) stained "positive" for SM-MHC, whereas 51 of 70 cells (73%) stained "positive" for smooth muscle {alpha}-actin. These results are consistent with those of others who have reported SM-MHC and smooth muscle {alpha}-actin expression in growth-arrested rat aortic SMC (32, 39) and verify the presence of the contractile phenotype in a majority of starved cells.

Shear rate vs. shear stress.   Figure 1 presents the contraction response (area reduction) of SMCs to a step increase in fluid flow with media of different viscosities in which the flow has been adjusted to maintain constant shear stress. The responses to 7.03- and 0.72-cP media were not significantly different at any of the measured time points. The normalized areas at the 30-min time points were 55.1 ± 3.5% (7.03 cP) and 61.4 ± 3.3% (0.72 cP). Control experiments were also performed in which preconfluent SMC seeded quartz slides were incubated in high-viscosity media without flow and observed for 30 min at room temperature. After 30 min of exposure, the normalized area was 99.6 ± 3.8% (n = 13). The normalized area response of cells in the high-viscosity media was not significantly different from the control experiments at any of the measured time points.



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  		Fig. 1. Normalized area response of rat aortic smooth muscle cells (RASMC) to a step in fluid flow with media of different viscosities. Shear stress for both cases was 25 dyn/cm2. High-viscosity media was prepared with dextran added to low-viscosity media. No statistical significance is observed between the 2 groups at any of the measured time points. Values are means ± SE; n, total no. of cells observed. The normalized areas at the 30-min time points are 55.1 ± 3.5% (7.03 cP) and 61.4 ± 3.3% (0.72 cP).

 
Ramp vs. step in shear stress.   SMCs were exposed to 25 dyn/cm2 shear stress induced with a step in flow (Step) and a ramp in flow over 5 min with steady flow for the remaining 25 min (Ramp). These responses were compared with a no-flow control (Fig. 2). The step and ramp area reductions were significantly greater than the control response at the 5-min (P < 0.05) and greater (P < 0.01) time points. The ramp and step responses were not significantly different from each other at any of the measured time points. The normalized areas at the 30-min time points were 91.2 ± 3.5% (Control), 66.3 ± 4.8% (Ramp), and 55.1 ± 3.5% (Step).



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  		Fig. 2. The normalized area response of RASMC to step and ramp fluid flow introduction. Ramp fluid flow introduction was from 0 to 25 dyn/cm2 over 5 min. The step fluid flow introduction was 0 to 25 dyn/cm2 at time 0. Values are means ± SE; n, total no. of cells observed. *Significantly different from control (no flow), P < 0.05. The ramp and step responses were not significantly different from each other at any of the time points measured. The normalized areas at the 30-min time points were 91.2 ± 3.5% (control), 66.3 ± 4.8% (ramp), and 55.1 ± 3.5% (step).

 
Glycocalyx component enzymatic digestion.   Figure 3 presents the normalized area responses to 25 dyn/cm2 shear stress with and without enzymatic digestion of the glycocalyx GAGs, heparan sulfate and chondroitin sulfate. Control experiments were performed without enzymatic treatment, with (25 dyn/cm2) and without shear stress (Control). At 10 min and beyond, the normalized response for the chondroitin sulfate treatment was significantly different from the response to 25 dyn/cm2 without enzymatic treatment but was not significantly different from the no-flow control response. In addition, at 20 min and beyond, both the chondroitin and heparin sulfate treatments showed this behavior. The normalized area responses at the 30-min time point were 89.3 ± 3.8% (Control), 87.1 ± 6.8% (CS Shear), 81.1 ± 3.0% (HP Shear), and 61.4 ± 3.3% (25 dyn).



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  		Fig. 3. The change in normalized cell area over 30 min in response to 25 dyn/cm2 (step) with and without enzymatic digestion of glycoclayx components. Data are shown for the area response in the absence of enzymatic treatment and shear stress (control), without enzymatic treatment but with exposure to shear (25 dyn), enzymatic treatment with chondroitinase ABC and exposure to shear stress (CS shear), and enzymatic treatment with heparinase III and exposure to shear stress (HP shear). Values are means ± SE; n, total no. of cells observed. *Significantly different from 25 dyn/cm2, P < 0.05. The normalized area responses at the 30-min time point are 89.3 ± 3.8% (control), 61.4 ± 3.3% (25 dyn), 87.1 ± 6.8% (CS shear), and 81.1 ± 3.0% (HP shear).

 
Control experiments were performed in which preconfluent SMC seeded quartz slides were bathed in 0.2 U/ml chondroitinase ABC or 0.2 U/ml heparinase III and observed for 30 min at room temperature. After 30 min of exposure, the normalized areas were 94.5 ± 11.2, 86.2 ± 7.3, and 81.2 ± 10.0% for the control, chondroitinase ABC (n = 10), and heparinase III (n = 14), respectively. These normalized area responses of unsheared cells pretreated with enzymes were not significantly different from the control experiments (no enzyme or shear) at any time point.

To demonstrate that cells were still capable of contracting to a known agonist after enzyme treatment, preconfluent monolayers were treated with 51 mM KCl after 30 min of enzymatic treatment. The results of this experiment are presented in Fig. 4. These cells displayed a normalized area response that was not significantly different from the response to KCl treatment without enzymatic treatment at all time points beyond 5 min. At the 30-min time point, the normalized areas were 57.2 ± 4.3, 51.2 ± 6.0, and 57.8 ± 4.5% for the 51 mM KCl without enzymatic treatment (n = 21), with 0.2 U/ml chondroitinase treatment (n = 12), and with 0.2 U/ml heparinase treatment (n = 15), respectively.



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  		Fig. 4. Change in normalized cell area over 30 min in response to 51 mM KCl with and without enzymatic digestion of glycoclayx components. Data are shown for the area response to enzymatic treatment with chondroitinase ABC and exposure to KCl (CS and 51 mM KCl), enzymatic treatment with heparinase III and exposure to KCl (HP and 51 mM KCl), and absence of enzymatic treatment but with exposure to KCl (51 mM KCl). Values are means ± SE; n, total no. of cells observed. *Significantly different from CS and 51 mM KCl, P < 0.05. The normalized area responses at the 30-min time point are 51.2 ± 6.0% (CS and 51 mM KCl), 57.8 ± 4.5% (HP and 51 mM KCl), and 57.2 ± 4.3% (51 mM KCl).

 
Verification of glycocalyx component enzymatic digestion.   The results of our protease activity assay indicate that neither heparinase nor chondroitinase displays protease activity. After an incubation of 3 h, the protease activity of chymotrypsin and pronase varied from 214 to 420 and 135 to 520 fluorescence units, as the concentration increased from 0.16 to 1.25 µM, respectively. The results for these two proteases indicate the upper (520 fluorescence units) and lower (135 fluorescence units) limits of the assay. The protease activity of the heparinase produced readings between 104 and 121 fluorescence units, and the activity of the chondroitinase was between 103 and 105 fluorescence units. These results indicate that the protease activity of these two GAG-depleting enzymes is below the sensitivity of the assay. The lower detection limit of this assay is very low indeed, being nearly 40 times less than the protease activity required to show any detectable influence on the glycocalyx of ECs (1, 13).

Previously, Chen and Wight (7) reported that chondroitinase ABC treatments were capable of removing a large fraction of chondroitin sulfate proteoglycans on the SMC surface. A study by Simionescu et al. (46) concluded that heparinase treatments degrade substantial portions of heparan sulfate GAGs on the surface of SMCs. To verify the selective removal of GAG components, confluent SMCs were labeled with an antibody specific to heparan sulfate or a lectin that binds to chondroitin sulfate and were visualized with and without enzymatic treatment. Representative images demonstrating the selective action of the enzymes are displayed in Figs. 5 and 6. Each image is a composite of 16 different fields of view from a single coverslip. The fluorescence intensity was normalized with respect to control and presented as normalized means ± SE in Table 1. Heparinase III significantly reduced the intensity of the heparan sulfate fluorescence, but not the intensity of chondroitin sulfate fluorescence. Chondroitinase ABC significantly reduced the intensity of chondroitin sulfate fluorescence but not heparan sulfate fluorescence. It should be noted in Fig. 5 that, although the intensity of the heparan sulfate image has not been altered by the chondroitinase treatment, the appearance (organization) of heparan sulfate has been altered somewhat by the removal of chondroitin sulfate, as this has altered molecular interactions in the glycocalyx.



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  		Fig. 5. Glycocalyx imaging of heparan sulfate with and without enzymatic treatment. Control represents heparan sulfate without enzyme treatments. Enzymatic treatments were 0.2 U/ml heparinase III (HP) and 0.2 U/ml Chondroitinase ABC (CS).

 


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  		Fig. 6. Glycocalyx imaging of chondroitin sulfate with and without enzymatic treatment. Control represents chondroitin sulfate imaging without enzyme treatment. Enzymatic treatments are as described in Fig. 5 legend.

 

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  		Table 1. Normalized fluorescence intensity for control and enzyme-treated preconfluent smooth muscle cells

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
This study addressed several issues related to the contraction of SMC in response to fluid flow-induced shear stress that was recently demonstrated by Civelek et al. (8). It described the first experiments that begin to characterize the mechanotransduction mechanism for fluid shear stress on SMCs. In addition, it provided more support for the hypothesis that fluid shear stress is a mechanical signal that mediates the myogenic response.

Figure 1 demonstrates that SMCs respond in a similar manner to an equivalent step increase in shear stress by using media of different viscosities. Shear stress is the product of viscosity and shear rate for Newtonian fluids, such as the experimental media employed in this study. The area reduction in response to a step in shear stress with high-viscosity media (7.03 cP) is not significantly different from the response for normal viscosity media (0.78 cP). This indicates that the contraction response is dependent on the level of shear stress, not shear rate.

Figure 2 shows that the rate of change of shear stress onset does not significantly alter the ultimate contraction response. Hill et al. (22) and Mellander and colleagues (20, 45), using intact resistance vessels in vivo and ex vivo, reported myogenic responses to a 5-min ramp increase in pressure that reached steady-state contraction levels comparable in magnitude to the step responses. Our observations are in good agreement with these studies. When comparing the results presented in Figs. 1 and 2 with those of Hill et al. (22), it is observed that the time course of the myogenic response is somewhat slower in the in vitro model compared with the ex vivo model. The difference in time course may be reconciled by realizing that, in the in vitro model, contraction requires detachment of SMC focal adhesions from the rigid substrate that resist the contraction, whereas contraction in the ex vivo model does not require cellular detachment from the deformable extracellular matrix.

Evidence that the mechanism by which rat aortic SMCs sense fluid flow-induced shear stress is linked to the glycocalyx of the cell is presented in Fig. 3. Depletion of heparan sulfate and chondroitin sulfate GAG chains on the surface by highly specific enzymes significantly reduced the contraction response of the arterial myocytes. The activity and selectivity of the heparinase III and chondroitinase ABC utilized to deplete GAGs were demonstrated by immunostaining, as seen in Figs. 5 and 6 and quantified in Table 1.

It has been demonstrated previously that the presence of heparinase alters not only the thickness (13, 35) but also the barrier function of the glycocalyx (29). The concentration of heparinase used in the present study and its effect on SMC heparan sulfate removal are supported by numerous references. Dull et al. (14) reported that, at a concentration of 15 mU/ml of heparinase III, a 67% removal of labeled sulfate (35SO4) was observed in bovine lung microvascular ECs. Haldenby et al. (21) noted a 27–51% reduction in fluorescence intensity with heparinase treatments using wheat germ agglutinin in vitro. Work by Florian et al. (16) reports a 45% reduction in fluorescence with heparinase treatment on bovine aortic ECs that completely blocks shear-induced NO production. Aortic ECs typically possess 50–90% heparin sulfate GAGs on their surface (23), compared with SMCs that have 40–60% heparin sulfate GAGs (34). Our results indicate a reduction of 38% with heparinase treatment, consistent with previous studies (36). Bourin et al. (4) labeled the N-acetyl-D-galactosamine component of chondroitin sulfate on rabbit heart ECs and reported a 45% reduction in the concentration of label after 30-min incubation with chondroitinase ABC. Consistent with that study, we report a 35% reduction in fluorescence with chondroitinase ABC treatment.

Three major signaling mechanisms for the myogenic response to mechanical stimuli have been suggested in the literature: membrane depolarization (11, 40), cytoskeleton rearrangement (11), and G-protein coupling (9, 52). Each of these mechanisms could be activated through linkages with the glycocalyx; however, most studies of the linkage of the glycocalyx to cell signaling have been performed in vascular ECs.

Membrane depolarization.   Siegel and colleagues (44) have developed a model for EC sensing of flow wherein the heparan sulfate proteoglycans, which are coiled due to hydrogen bonding at no-flow conditions, become unfurled in response to increased flow. The unfurled proteoglycan exposes additional binding sites for Na ions. The additional binding of Na ions begins a signal transduction chain mediated through voltage-dependent ion channels, mainly Na2+ channels, that depolarize the membrane (44). Na ion voltage-operated channels have been shown to be present in vascular SMCs, and Na2+ channels on the SMC surface have been observed to be similar to those found on the EC surface (49).

Cytoskeleton rearrangement.   On the SMC surface, chondroitin sulfate and heparan sulfate GAGs can be present on syndecans-1 and -4 (5, 37, 46) and on glypicans (38). The intracellular domain of syndecan-1 has been shown to associate with the actin cytoskeleton (5, 6, 37). The coupling of syndecans to the actin cytoskeleton could result in mechanical stress being transmitted from extracellular to intracellular domains.

G-protein coupling.   A previous study by Civelek et al. (8) concluded that rat aortic SMCs respond to fluid shear stress via PKC and RhoA pathways that are calcium independent. Oh et al. (31) have reported that syndecan-4, present on the SMC surface, binds to PKC and regulates PKC’s activity and distribution. The sensing of flow by the GAGs and syndecan body could activate the PKC pathway, leading to calcium-independent cellular contraction. Thus the contraction response of rat aortic SMCs to fluid shear stress could be sensed through the glycocalyx by one or more of these plausible pathways. Further studies will be required to determine the specific pathways that are involved.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grant HL-35549 and National Aeronautics and Space Administration Grant NAG3.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 ACKNOWLEDGMENTS
REFERENCES
 
The authors thank Elaine Kunze and coworkers at the Center for Quantitative Cell Analysis and Aaron Mulivor for their expertise.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. M. Tarbell, Dept. of Biomedical Engineering, Steinman Hall, 2nd Floor, The City College of New York/CUNY, Convent Ave. at 140th St, New York, NY 10031 (E-mail: tarbell{at}ccny.cuny.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.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 ACKNOWLEDGMENTS
 REFERENCES
 

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