Heparin Modulates the Composition of the Extracellular ... - NCBI - NIH

AmericanJournal ofPathology, Vol. 137, No. 2, August 1990 Copyright © American Association ofPathologists

Heparin Modulates the Composition of the Extracellular Matrix Domain Surrounding Arterial Smooth Muscle Cells Alan D. Snow,* Robert P. Bolender,t Thomas N. Wight,* and Alexander W. Clowest From the Departments ofPathology,* Biological Structure, and Surgery,J University of Washington, Seattle, Washington

Heparin and related molecules influence vascular wall structure by their ability to inhibit smooth muscle cell (smc) proliferation and migration. However, little is known as to whether heparin has an effect on the extracellular matrix. In the present study, the effect of heparin on the content and regional distribution ofelastin, collagen, andproteoglycans (PGs) in blood vessels following experimental injury was determined. Two groups of rats were subjected to left common carotid balloon injury and were infused with either 0.9% saline or heparin in a saline solution, for 2 weeks. Using a new morphometric method ofanalysis, the authors determined changes in volumes of elastin, collagen, and PGs contained within an 'extracellular matrix domain (ECM domain), 'the average envelope of connective tissue surrounding each smc. Heparin treatment inhibited intimal thickening and decreased the elastin content in the ECM domain in the upper and lower arterial intima. Collagen also was found to be significantly decreased 5. 0-fold and 7.6-fold in the ECM domains of upper and lower intima, respectively, of heparin-treated animals. The decrease in both elastin and collagen was balanced by a significant increase in amorphous and filamentous electron-dense material. Heparin also caused a significant 1.8-fold and 1.9fold increase in the PG content in the ECM domain in the upper and lower intima, respectively. Immunohistochemical analysis, using antibodies to elastin and PG subclasses, supported the morphometric observations. This study has shown that heparin administered in vivo can alter the accumulation and distribution of each of the major vascular ECM components in a specific and differential manner. (Am JPathol 1990; 13 7:313-330)

The extracellular matrix (ECM) in all tissues influences a variety of cellular events which form the basis for both development and disease.1 The ECM is in a constant state of flux and changes as cells modulate their behavior. Such ECM changes in turn may have a profound effect on the physiologic properties of the tissue, as well as on the metabolic properties of the cells themselves. The major ECM components surrounding smooth muscle cells in the arterial wall are collagens, elastic fibers, and proteoglycans (PGs).2 These macromolecules are distributed in the intimal, medial, and adventitial layers of the vascular wall in such a manner as to confer both structural integrity and viscoelasticity. Blood vessels undergo a marked intimal thickening in response to mechanical injury,3'4 and this process is believed to reflect one of the early stages in the development of the atherosclerotic lesion. A number of studies have shown that part of this thickening is a consequence of the migration and proliferation of arterial smooth muscle cells3 and part is due to the accumulation of components of the ECM4 such as elastin,5 collagens,6 and PGs.2 Additional studies have demonstrated that intimal thickening after arterial injury can be reduced by heparin infusion, which appears to act by inhibiting the migration and proliferation of arterial smooth muscle cells.7-10 However, the effect of heparin on the ECM surrounding these cells is not known. The objective of the present study is to define the regional composition and distribution of the major ECM components (elastin, collagen, and PGs) in blood vessels after experimental injury and to determine the nature of the ECM changes when the injury response is interrupted by heparin. To study the extracellular matrix as an experimental variable of smooth muscle cells, we devised a new method of analysis by defining the extracellular matrix as a specific compartment. This compartment, the ECM domain, provided a means for detecting changes in the concentration and composition of individual ECM compoSupported by the NIH grants HL 18645 and HL 27769. Accepted for publication March 9, 1990. Address reprnt requests to Dr. Alan D. Snow, Department of Pathology SM-30, University of Washington, Seattle, WA 98195.

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nents in tissues in various physiologic states. Combining this approach with immunohistochemical data has allowed us to define specifically the nature of the ECM changes after arterial injury in the presence or absence of heparin.

Materials and Methods Balloon Injury Model A total of eight 5-month-old male Sprague-Dawley rats (Tyler Laboratories, Seattle, WA) each were subjected to left common carotid balloon injury.34 The right carotid artery was left untraumatized and served as controls for each animal. Immediately after injury, a silastic catheter was inserted into the left jugular vein and connected to a 2-week, 2-ml miniosmotic infusion pump (Alzet, Alza Corporation, Palo Alto, CA), which was placed in the subcutaneous tissue over the back of the animal. The osmotic pumps contained either heparin (Sigma type 11; 0.30 mg/ kg of body weight/hour) in a 0.9% saline solution, or a saline solution only. Previous studies have shown that the rate of delivery for the 2 ml solution in the 2-week osmotic pump is 5 ,ul/hour, with active pumping occurring approximately 4 hours after pump placement.9 The solutions were sterilized, using a 0.22-,i filter, before loading into the pumps.

Tissue Sampling, Fixation, and Processing At 2 weeks, animals were anesthetized and then killed by cardiac puncture. To determine whether heparin was properly delivered during the 2-week experiment, blood was taken for demonstration of prolongation of the whole blood clotting time.34 The aortic arch and both the left and right common carotids were removed and washed extensively in buffer and divided into small pieces by cross-sectioning. Length measurements of the entire carotid were made with a centimeter ruler under the dissecting microscope, and vessel segments for fixation were taken from the middle portions of each vessel, using random systematic sampling.1' These areas were devoid of

endothelium.12"3 Small pieces of tissue from both the left (injured) and right (uninjured) carotid arteries of each animal were taken, starting from the middle portion of each vessel, and alternating tissue segments were immediately placed in fixatives for either light or electron microscopy. For histochemistry and immunocytochemistry, some tissue segments were fixed in a solution of 90% ethanol and 10% formaldehyde for 24 hours at -20°C, processed routinely

into paraffin, and serial sectioned at 6 ,u. For electron microscopy and ultrastructural morphometry, pieces of tissue were fixed in 0.025 mol/l (molar) sodium acetate buffer (pH 5.6) containing 2.5% glutaraldehyde and 0.2% Cuprolinic blue (BDH Chemicals, Vancouver, BC) with 0.3 mol/l and 0.7 mol/l magnesium chloride."4 After fixation overnight at room temperature, the tissues were washed 3 times (10 minutes each wash) in the same solution, but without Cuprolinic blue. The tissues were further washed 3 times in 1% aqueous sodium tungstate and then dehydrated through graded alcohols with the 30% and 50% ethanol concentrations containing 1 % sodium tungstate. The tissues then were infiltrated with propylene oxideMedcast (Ted Pella, CA) and eventually embedded in Medcast. Thin sections were mounted on copper grids and stained with uranyl acetate and lead citrate. Sections were viewed and photographed on a JOEL 1 OB electron microscope (Japan Optics Electron Laboratory, Tokyo, Japan) at 60 kV.

Histochemistry Adjacent serial sections of carotid arteries from both saline- and heparin-treated animals were stained with the following stains: 1) hematoxylin and eosin (H&E), 2) Verhoeff's hematoxylin for the detection of elastin,15 3) van Gieson's stain for the detection of collagen,15 and 4) alcian blue (pH 5.7) with 0.3 mol/l and 0.7 mol/l magnesium chloride, according to the method of Scott and Dorling,16 for the detection of sulfated (0.3 mol/l magnesium chloride) and highly sulfated (0.7 mol/l magnesium chloride) glycosaminoglycans.

Immunocytochemistry Adjacent serial sections also were used for immunocytochemical studies. After pilot experiments to ensure that the antibodies were used at dilutions that provided minimal background staining, the following primary antibodies were used: 1) a polyclonal antibody (1 :100 dilution) known as alpha-2, which recognizes insoluble elastin17"18 (gift of Dr. Robert Mecham, Washington University, St. Louis, MO); 2) a polyclonal antibody (1:50 dilution) that recognizes the protein core of the heparan sulfate proteoglycan'9 (gift of Dr. John Hassell, University of Pittsburgh, Pittsburgh, PA); 3) a polyclonal antibody known as TAP-1 (1:10 dilution), which recognizes the glycosaminoglycan chains of chondroitin sulfate proteoglycans' (gift of Steve Carlson, University of Washington, Seattle, WA); and 4) a polyclonal antibody (1:10 dilution) that recognizes the protein core of the dermatan sulfate proteoglycan21 (gift

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of Hans Kresse, University of Munster, West Germany). This latter antibody was additionally used after sections were incubated in 0.1 mol/l TRIS-HCI buffer (pH 8.0) containing chondroitinase ABC (Sigma Chemical, St. Louis, MO) at an enzyme concentration of 2.0 units/ml at 370C for 3 hours.22 The specificity of all antibodies were determined by preabsorption of the antibodies with the appropriate antigens before immunostaining. 19-21,23 Immunostaining of tissue sections was accomplished using the peroxidase-antiperoxidase (PAP) method.24 To rule out nonspecific binding of antibodies, sections also were stained according to the PAP technique using TRISbuffered saline (TBS) instead of the primary antibody.

Measurement of Arterial Intimal Area The cross-sectional areas of arterial lumen and intima were obtained by making tracings of individual sections by means of a camera lucida and a Hewlett-Packard digitizing pad. Measurements of intimal area were made using the X6.3 objective of a Leitz Dialux 20 light microscope (Ernst Leitz, Wetzlar Corp., Wetzlar, Germany). Comparisons were made between intimal cross-sectional areas of saline vs. heparin-treated animals.

Magnification Calibration A carbon grating replica containing 21,575 lines per centimeter (E. F. Fullam Inc., Schenectady, NY) was used to calculate the final magnification of the electron micrographs being used for morphometry.

Volume Density of Smooth Muscle Ce/l and Matrix Compartments To determine the volume densities of smooth muscle cells and extracellular matrix compartments in the control and experimental tissues, a series of electron micrographs were taken at a final magnification of X8000. A pilot experiment indicated that three to four vessel segments from each of three to four animals, using 10 electron micrographs per region of the intima, gave a standard error less than 10% of the mean. A random systematic sampling11 was used to collect electron micrographs from both the upper (luminal side) and lower portions of the intima. Volume densities were estimated using a 108-point square lattice test grid (Pgr,id) with the number of linear probes per point probe (k) equal to 2, and the length (D) of the linear probe in the test grid equal to 2.0 cm. When the test grid was placed over the electron micrographs, points that fell

on either smooth muscle cells, 'other cells,' or matrix were counted. Relatively few 'other cells' were found within the intimal wall and these were essentially all identified as lymphocytes based on morphologic criteria. Nuclear profiles per unit area (NA) were counted for smooth muscle cell (smc) and 'other cells' and used for numerical density (Nv) estimations. For each animal, three vessel segments were analyzed, giving typically a total of 30 electron micrographs from the upper intima and 30 electron micrographs from the lower intima per animal. With four animals per group, a total of 120 electron micrographs per intimal region (upper and lower) were analyzed. Volume densities were calculated for smooth muscle cells as: Vv (smc/intima) = Psmc/(Psmc + Poc + Pmx) and for the matrix as: Vv (mx/intima) = Pmx,(Psmc + Poc + Pmx). The intima was used as the reference volume and Pmx, Psmc, and Poc referred to points on the matrix, smooth muscle cells, and 'other cells,' respectively. To determine volume densities for each of the components of the intimal ECM, electron micrographs were taken at a final magnification of X 60,000. Electron micrographs were taken from each of two regions within the ECM of the intima, including the upper intima (the 1/3 of the intimal wall closest to the lumen) and lower intima (the 1/3 of the intimal wall closest to the internal elastic lamina). Three to four vessel segments from each of four animals were analyzed using 10 electron micrographs per region (ie, upper and lower intima). For ECM analysis, five components were defined and counted. These included: 1) elastin, which appeared on electron micrographs as pale, amorphous islands surrounded by a darker border; 2) co/lagen fibrils, which were seen in either longitudinal or cross section and were characterized by their distinctive banding pattern (64 nm periodicity on longitudinal section); 3) proteoglycans, which with Cuprolinic blue appeared as tapered filaments of variable length; 4) 'space,' which represented areas unoccupied by staining components; and 5) a compartment referred to as 'other,' which represented amorphous, unidentified filamentous material. The volume densities of elastin, collagen, 'space,' and 'other' were estimated using a 108-point square lattice test grid with Pg§d = 108, k = 2, and D = 2.0 cm. A separate test grid was used for PG analysis. This latter grid had a Pgrd = 945 points, k = 2, and D = 0.7 cm. For each animal, three vessel segments were analyzed, giving a total of about 30 electron micrographs per intimal region. With three animals per group, a total of 60 electron micrographs per intimal region (upper and lower) were analyzed.

Extracellular Matrix Domains of Smooth Muscle Cells The 'extracellular matrix domain' is defined as the volume of extracellular matrix on average surrounding a smooth

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muscle cell. An ECM domain was calculated by dividing the volume density (VJ) of a matrix compartment (ie, elastin, collagen, PGs, etc.) by the numerical density (NV) of smooth muscle cells: V = VV/NV. Smooth muscle cells were counted according to Gittes and Bolender,25 using the expression NV = w2/4 X NA/B X 1 /KS, where NA is the number of nuclear profiles counted in a known area of section, B is the mean nuclear boundary, and K. is a correction factor for nuclear shape. PCS System II software running under MS-DOS26 was used to collect stereologic data and make Vv, NV, and ECM domain estimates. Because the correction factor for shape was only about 5% for both saline- and heparin-treated animals, it was not included in the NV estimates.

Mean Surface Area of Nuclei and Smooth Muscle Cell Volume mean surface area (S) of smc nuclei and smc volumes (V) were calculated, respectively as S = 1 6/13 X B2

The

X Ks and V = 1 /Nv, according to Gittes and Bolender.25

Statistical Analysis For statistical purposes, all results compared the two groups (saline-treated and heparin-treated) using the Student's t-test27 and the Mann-Whitney U-test.28 Statistical significance was set at the 99% (P < 0.01) and 95% (P < 0.05) levels.

Results

Reduction of Intimal Thickening Following Heparin Treatment All heparin-treated animals demonstrated prolonged whole blood clotting time (>6 minutes), indicating that heparin was properly delivered during the 2-week experimental period. Marked intimal thickening was apparent in the saline-treated animals 2 weeks after carotid injury (Figure 1 A). In comparison with saline-treated animals (0.13 mm2 + 0.01), measurements of intimal cross-sectional areas were 2.7-fold less (P < 0.01) in the heparintreated animals (0.05 mm2 0.01) (Figure 1 B). These observations are in agreement with previous studies describing the heparin effect.9 10'12

Histochemistry Histochemical staining of de-endothelialized carotid arteries for elastin, collagen, and PGs demonstrated distinct differences in heparin-treated compared with salinetreated animals, particularly evident within the intimal portions of the blood vessels. In the carotid arteries of the injured saline-treated animals (Figure 1 C), elastin, as demonstrated by intense staining with the Verhoeff's hematoxylin stain, was evident in both the thickened intima and media of the artery. The heparin-treated animals, however, displayed a marked decrease in elastin staining in the intima, whereas elastin staining in the media was still apparent (Figure 1 D). Collagen was observed throughout the thickened intima in the injured carotid arteries of saline-treated animals using the van Gieson's stain (Figure 1 E). In comparison with saline-treated animals, the heparin-treated animals exhibited a decrease in collagen staining in the intima, whereas staining in the media and adventitia was unaffected (Figure 1 F). Strong alcian blue staining in the presence of 0.3 mol/ magnesium chloride was found in the intimas of both saline- and heparin-treated groups (not shown), suggesting that sulfated glycosaminoglycans (GAGs) were present in the intimal region of both groups regardless of treatment. Heparin-treated animals showed a slight increased intensity of alcian blue staining in the presence of 0.7 mol/ magnesium chloride in the intima (Figure 1 H), in comparison with saline-treated animals (Figure 1 G), suggesting the presence of highly sulfated GAGs.16'29-30

Immunocytochemistry The decrease of intimal elastin in the carotid arteries of heparin-treated animals observed histochemically was confirmed using an elastin antibody. A polyclonal antibody that recognizes insoluble elastin18 demonstrated positive immunostaining in both the intima and media of saline-treated animals (Figure 2A). The heparin-treated group, however, exhibited essentially no intimal elastin immunostaining, whereas immunostaining was evident in the media (Figure 2B). No staining in the vessel walls was observed in either of the two groups when TBS was used instead of the primary antibody. Carotid arteries from saline-treated animals demonstrated moderate heparan sulfate proteoglycan (HSPG) immunostaining in the ECM surrounding smooth muscle cells in the intima and media of the vessel wall (Figure 2C). Heparin-treated animals demonstrated an increase in immunostaining with the HSPG antibody in the intima in comparison with saline-treated animals (Figure 2D). Be-

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PGs Figure 1. Histochemnical staininig of de-enidothelialized carotid arteries (2 weeks afaer initimal injury) from saline-treated (Figs. A, C, E, G} anid heparini-treated (Pigs. B, D, F, H) animals. A and B: I-IE C and D: VVG without collageni counterstain; E and F: VVG without elastini staini; G and H: a/cian b/tie (PHI 5. 7) with U. 7 miol MgC12 added. (A) Marked initimal thickening is observed in a carotid artery from a salinie-treated animiial intima (i), mnedia (nO' liar: SOOpti (B) lIn comparisoii with Figure A, heparin-treated aninials demonstrate a decrease in initintial thickeninig. iiitiina (i) mLedia (in) Bar, 300pu (C) Carotid artery from saline-treated animial deinonst rating e/astini staininig in both initima (i anid mtedia (in). Note stioiig staiiiiiig of the elastic lamellae (arrowheads). Bar, 100 pA. (D) In coinparison uwith Figurie C, heparin -treated animials demionstrate a marked decrease in elastint staining in the iiititina (i, uhereas staining]o)r elastini in the media (vi) is still presenitt liar, 100 p. (E) Carotid artery from saliiie-treated animal demioiistratinig collagen staining (pink deposits) (arrouheads) in ivitimnal matrix between smooth muscle cells. intima (i0; media (in) liar, /00 p. (F) In coniparisoui uwith Figuire F, heparini-treated amiimna/s showed a decrease ini col/ageii staining in the intima (i). Note stroiig stainiiig of collageni iii adi'eiititia (a). Bar, 100 pA. (G) Carotid artery from salimie-treated aiiimal demonstrating slight stainiiingfor highly, su/;7ted GAGs iii iiitiiiia (arrouhead). Note stroiig staininig of mast cells (duie topreseiice ofheparin) in adventitia (arrows). liar, 300 pA (H) In comiparison with Figure G, hep)arimi -treated anzimals shooed stronig staiiiiiig for highly, sulfated GAGs in initimta (arrowhead) liar, 300 p.

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DSPG Figure 2. Immunohistochemical staining of de-endothelialized carotid arteries (2 weeks after intimal injury) from saline-treated (Figs. A, C, E, G) and heparin-treated (Figs. B, D, F, H) animals. A and B: elastin C and D: HSPG antibody; E and F: CSPG antibody; G and H: DSPG antibody; (A) Positite elastin immunostaining in inztima antibodj; (i) and media (m) in carotid arteryfrom salinetreated animal. Note positive staining of elastic lamellae in media (arrowhead). Bar, 150 p. (B) In contrast to Figure A, heparintreated animals displayed a marked decrease in elastin immunostaining in the intima (i), whereas elastin immunostaining in media (m) was still evident. Bar, 150 pA. (C) Moderate HSPG immunostaining in the intima (i) and media (m) of a animal. Bar, 150 A. (D) In comparison with Fig. C, heparin-treated animals displayed strong HSPG immunostainingsaline-treated in the intima (i). Note also strong HSPG immunostaining of vessels in adventitia (arrowheads). Bar, 150 p. (E) Weak CSPG immunostaining in the intima (i) of saline-treated animal. Bar, 150. (F) In comparison with Figure E, heparin-treated animals demonstrated strong CSPG immunostaining in the intima (i). Bar, 150 p. (G, H) Both saline-treated (Figure G) and heparin-treated (Figure H) animals displayed uweak DSPG immunostaining in the intima (i). Strong immunostainingfor DSPG waspresenlt in the adventitia (a) in both groups, a location where DSPGs are known to be present. Bar, 8Opl.


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cause the HSPG antibody recognizes the protein core," the increased immunostaining observed in the intima of heparin-treated animals was due to the presence of HSPGs, and rules out the possibility that the positive immunostaining was due to infused heparin trapped within the de-endothelialized vessel wall. A polyclonal antibody to chondroitin sulfate proteoglycans (CSPGs) revealed moderate immunostaining in the intimal regions in the carotid arteries of saline-treated animals (Figure 2E). This immunostaining was markedly increased in the intima of heparin-treated animals (Figure 2F). Negative controls, using either preabsorbed antibody (not shown), a different antibody of the same gamma G immunoglobulin (IgG) class (Figure 2G, H), or TBS, instead of the primary anti-CSPG, exhibited no staining in these areas. An affinity-purified antibody recognizing the protein core of the small dermatan sulfate proteoglycan (known as DSPG-1l) produced only slight immunostaining in the intima in both saline-treated (Figure 2G) and heparintreated (Figure 2H) animals. Strong staining in both groups was apparent in the adventitia in association with bundles of collagen fibrils (Figure 2G, H), as previously described.31'32 Chondroitinase treatment of the sections did not alter the degree of immunostaining with the DSPG

antibody.

Electron Microscopy Electron microscopy was used to confirm observations made at the light microscopic level. The saline-treated animals demonstrated an abundance of elastin in the intimal ECM, in many instances coalescing to form elastic lamellae (Figure 3A). In addition, the saline-treated animals showed bundles of collagen fibrils throughout the entire intima, appearing more prevalent in the lower intima (Figure 3A). Conversely, carotid arteries from the heparintreated animals demonstrated a markedly reduced amount of both elastin and collagen in the intimal ECM (Figure 3B). The elastin in the intimas of heparin-treated animals appeared as small islands (Figure 3B). The reduction in both collagen and elastin contrasted markedly with an increase in the content of PGs in the intimas of heparin-treated animals. Proteoglycans were preserved and visualized by the use of the cationic dye Cuprolinic blue (Figure 4). Cuprolinic blue staining at 0.3 mol/l magnesium chloride preserved all three classes of PGs known to be present in the arterial wall (ie, HSPGs, CSPGs, and DSPGs)33.34 (Figure 4). Cuprolinic blue-positive filaments, ranging in length from 40 to 80 nm, were present in the ECM of both saline-treated and heparintreated animals. High-magnification electron micrographs

demonstrated primarily PGs in the intimal ECM of heparintreated animals (Figure 5B). The saline-treated animals exhibited an intimal ECM containing elastin, collagen, and PGs (Figure 5A).

Ultrastructural MorphometryVolume Densities The volume density of smooth muscle cells in the intima did not differ between saline- and heparin-treated animals (Table 1). Smooth muscle cells occupied 29.2% and 27.8% of the upper intima in saline and heparin-treated animals, respectively, whereas a significant difference was found in the lower intima in comparison with the upper intima (Table 1). The majority of the intima was occupied by ECM, with no difference found between salinetreated and heparin-treated animals (Table 1). The 'other' type of cells observed occasionally in the intima were rounded cells with a large nuclear-to-cytoplasmic ratio, typical of lymphocytes. In both groups, lymphocytes occupied less than 1% of the total volume density of the intimal wall (Table 1). Neutrophils or other cell types were not observed in the intimal layer in either group. Volume densities of each of the ECM components constituting the intima also were determined (Table 2). The results indicated that heparin treatment caused a 4.4fold and 2.2-fold difference in the volume occupied by elastin in the upper and lower intima, respectively. Heparin-treated animals also demonstrated a significant 4.5fold and 8.6-fold difference in the volume occupied by collagen in the upper and lower intima, respectively. Conversely, the volume occupied by PGs was found to differ 2-fold in both the upper and lower intima in heparintreated animals. Although the volume occupied by space did not differ between the two groups, heparin treatment caused a 1.8-fold and 2.9-fold difference in the volume occupied by amorphous material (defined as 'other') in the upper and lower intima, respectively.

Smooth Muscle Cell Volumes We determined the average volume of each primary component in the ECM surrounding each smooth muscle cell. No significant differences were found between each parameter in saline- vs. heparin-treated animals (Table 3). Nuclear numerical density also was used to calculate smc volumes for the two groups. No significant difference was found in the volume per smooth muscle cell between saline- and heparin-treated animals in either the upper (saline group 811 ± 36 U3; heparin group 711 ± 57 s3) or lower intima (saline group 657 ± 33 83; heparin group 598

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proteoglycans within the intima of the inijured carotid artery. The volume ofproteoglycans (PGs) in the ECM domain (in the

experimentally induced atherosclerosis. A recent study using cDNA clones for collagen and elastin has demonstrated that increases in these molecules after arterial injury correlate with an increase in mRNA levels coding for elastin and type and Ill procollagen.5 Increased synthesis and deposition of ECM components is a general response of a variety of cells to various forms of injury and during wound repair.40'41 Injury is often accompanied by cell proliferation, and at present it is unclear whether these ECM changes are directly related to the stimulation of cell division. A number of reports indicate that the synthesis of ECM components is increased when arterial smooth muscle cells are stimulated to divide.3-4247 However, it is also clear that ECM molecules also may be increased by other factors released at wound sites, such as transforming-growth factor (TGF) beta, which does not cause cell proliferation to occur.48 Thus, the regulation of ECM deposition after arterial injury appears to be exceedingly complex.

upper and lower initima) of saline- and heparin-treated animals wasfound to differ significantly, 2 weeks after iniitial inti-

mal injury. A significant (P < 0.01) 1.8-fold and 1.9-fold inin the volume of PGs in the ECM domainuwas found in the upper and lower intima in heparin-treated aniimals, recrease

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was found surrounding the smooth muscle cells in the upper intima, whereas smooth muscle cells in the lower intima were surrounded by abundant collagen fibrils.

Such results suggest that smooth muscle cells in different parts of the same tissue process fibrillar collagen differently. Evidence that arterial smooth muscle cells throughout intimal fibromuscular thickenings differ is derived from studies that demonstrate that, after carotid injury, cells nearest the lumen (upper intima) are actively proliferating, whereas cells in the lower intima are quiescent.3-12 Therefore, differences in the collagen content in these two regions may be a reflection of the growth state of the cells. Additionally, in situ hybridization has been used to demonstrate that certain populations of smooth muscle cells in the outer media within hypertensive pulmonary arteries exhibit increased expression of tropoelastin mRNA compared with smooth muscle cells in the inner media.38 Such studies further support the concept that smooth muscle cells within the vascular wall differ phenotypically with regard to the ECM that they produce.

Heparin and Related Molecules Modulate Smooth Muscle Cell Phenotype Heparin has been shown to dramatically alter the response of blood vessels to experimental injury. Clowes and Karnovsky7 were the first to demonstrate that infusion of heparin into animals subject to arterial balloon injury caused a significant reduction in the intimal hyperplastic response. Subsequent studies have shown that heparin is effective in inhibiting the proliferation and migration of arterial smooth muscle cells in response to mechanical injury. 34910,12,49 Treatment of vascular smooth muscle cells in vitro with heparin or heparan sulfate derived from vascular cells causes inhibition of growth,5'056inhibition of migration away from experimental wounds57 and modulation to a growth factor-responsive phenotype.' Therefore, heparin appears to be altering the proliferative response by having an effect on the behavior of the smooth muscle cells themselves. We now know from the results of the present study that, in addition to effects on cellular behavior, heparin has a dramatic effect on the types and quantities of ECM molecules that surround arterial smooth muscle cells.

Changes in ECM Domains During Injury and Repair

Heparin Differentially Affects Collagen, Elastin, and Proteoglycans in the Extracellular Matrix

It is well documented that components of the ECM such collagen,6 elastin,5 and PGs3-39 increase in blood vessels after mechanical injury, during hypertension, and in

The response is not uniform in that the major fibrous proteins, collagen and elastin, are decreased in the presence of heparin, whereas the more viscous PGs are modestly

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Figure 8. Heparin treatment increases the volume occupied by amorphous andfilamentous materialfollowing carotid injury. A significant (P < 0.01) increase in the volume occupied by amorphous andfllamentous material (defined as other) in the ECM domain wasfound in the upper and lower intima in heparin-treated animals. This increase is believed primarily due to improperly formed or degraded elastin and collagen that occurred in the presence of heparin.

increased. Indeed, it is unlikely that there is one common mechanism to explain the effect of heparin on these different components of the ECM. Various possibilities for each set of these molecules being affected will be discussed. One possibility is that heparin causes a decrease in the synthesis of collagen and elastin. Previously, heparin has been shown to decrease the synthesis of collagen by chondrocytes59 and intestinal smooth muscle cells in vitro.Y-61 In the chondrocyte study,59 there was an accumulation of collagen precursors, found primarily in the cell layer compartment, that appeared to be the results of heparin's inhibition of the activity of the NH2-terminal protease. Such results suggest that heparin's affect on decreasing collagen deposition may involve interference with collagen processing. Although Majack and Bornstein62 demonstrated no overall difference in collagen production when rat arterial smooth muscle cells were treated with heparin, there was a significant decrease in the amounts of type IlIl relative to type procollagen and an increase in the synthesis of a nonfibrillar short chain collagen. In a more recent study, Tan et al63 demonstrated that collagen production, as measured by the incorporation of [3H]proline into [3H]hydroxyproline, was inhibited 91% to 95% in human arterial smooth muscle cells in the presence of heparin and ECGF. In addition to having a direct effect on collagen and elastin synthesis, heparin may interfere with processing and the fiber-forming mechanism of these proteins. For example, heparin inhibits the oxidation of lysine in collagen by lysyl oxidase, which is a critical step in the forma-

tion of the collagen fibril.64 It is interesting to note that heparin has little effect on elastin oxidation by Iysyl oxidase,64 suggesting that heparin's affect on regulating collagen and elastin content may be operating through different mechanisms. Heparin may interfere with elastin fibrillogenesis by altering the rate and extent of accretion of tropoelastin units to the nascent elastin fibril. For example, inhibition of lysyl oxidase by inhibitors such as beta-amino propionitrile in growing chick and rat aortas reveals abnormal elastin aggregates permeated by glycosaminoglycans.65 66 These observations suggest that either endogenous or exogenous GAGs such as heparin may interact with positively charged groups in tropoelastin to prevent aggregation. Heparin also appears to influence the structural organization of the major fibrous proteins in the ECM. For example, a number of studies have recently demonstrated that heparin disrupts collagen fibers in a hydrated gel and prevents the contraction of these gels by mesenchymal cells.67Y70 Because heparin is elevated in fibrotic and in scar tissue, the abnormal arrangement of collagen fibrils and decreased tensile strength of scar tissue may in part be the consequence of the discontinuous packing of the collagen fibrils promoted by heparin.71 The decrease in collagen and elastin in arteries infused with heparin also may be due to the increased degradation or turnover of these molecules. For example, heparin has been shown to increase the amount and activity of a bone collagenase,72 and it is interesting to note that osteoporosis develops in patients given large dosages of heparin in the treatment of clotting disorders.73 A number of reports indicate that heparin suppresses wound healing and scarring in burned, wounded, or surgically treated tissues, presumably through its ability to stimulate collagen degradation and inhibit its cross-linking.64 Heparin may be also influencing elastase activity. For example, Jordan et a174 recently demonstrated that heparin promotes the inactivation of anti-thrombin Ill by neutrophil elastase. Although the mechanism by which this occurs is uncertain, evidence indicates that interaction of heparin with elastase, as well as with anti-thrombin 111, is necessary for accelerated degradation. This stimulatory effect of heparin on elastase activity is in contrast to a more commonly observed inhibition of neutrophil and leukocyte elastase in the presence of sulfated GAGs such as heparin.75,76 It may be that elastases synthesized by resident cells of the arterial wall differ in their sensitivity to heparin inactivation or activation. In the present study, accompanying the marked decrease in elastin and collagen estimated by ultrastructural morphometry there was a significant increase in the amorphous, unidentifiable fibrillar components as defined by the 'other' category. Such components may represent


327


Saline-treated

Figure 9. Extracellular matrix domains in the intima of heparin-treated and salinetreated animals. The concept of the extracelluilar matrix domain is demonstrated in thisfigure. Each smooth muscle cell (smc) in the intima is surrounded by a envelope of ECM (- ). As determined from the present study, comparisons of the quantities of elastin, collagen, and proteoglycans within the ECM domain of heparin-treated vs. saline-treated animals is demonstrated. Note that the ECM domain of heparin-treated animals contain lesser quantities of elastin and collagen, and a greater quantity of proteoglycans in comparison with saline-treated animals.

\ , L,

i

Heparin-treated

f'-

-

, e

,\

------,,-

- - -----

elastin

proteoglycans

at

collagen

A

"other"

SMC smooth muscle cell N

nucleus

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accumulation of precursor or breakdown products of collagen and elastin. The changes in the PG content of the arterial lesions in the presence or absence of heparin was determined by preserving the PGs in the tissue as discrete structures using Cuprolinic blue.14'32 This procedure has been used effectively in the past to map the content and composition of arterial PG34 and has been shown to provide a reliable estimate of the involvement of PG in various tissues.3277 Using this approach, we have demonstrated that, in contrast to the decrease in collagen and elastin content, there is an increase in the PG content in arteries after injury in animals treated with heparin. A few studies have demonstrated that heparin can cause an increase in the incorporation of radiolabeled sulfate by cultured cells such as fibroblasts78 and arterial smooth muscle cells,47'79 suggesting that heparin may have a direct effect on PG synthesis and/or turnover. Heparin has been shown to effectively inhibit the activity of heparanase, an enzyme that degrades heparan sulfate chains.8082 In addition, Au et al83 also have recently demonstrated that heparin decreased the synthesis of tissue plasminogen activator by cultured arterial smooth muscle cells, which in turn influence the degradation activity of plasmin. Because plasmin degrades a wide variety of macromolecules, including PGs,4 heparin's affect on PG turnover may be partially mediated by this response. In summary, an analysis of the matrix in the extracellular domain surrounding smooth muscle cells in heparintreated arteries demonstrates a marked decrease in interstitial elastin and collagen and an increase in proteoglycan (Figure 9). These results provide support for the hypothesis that heparin and related endogenous molecules may not only regulate vascular wall structure by inhibiting smooth muscle cell proliferation, but also by altering the

composition of the ECM associated with the smooth muscle cells.

References 1. Hay E: Cell Biology of the Extracellular Matrix. New York, Plenum Press, 1981 2. Wight TN: Matrice extracellulaire et atherosclerose., Les Malaides De La Paroi Arterielle. Edited by J Camilleri, CL Berry, J Fiessinger, J Bariety. Paris, Medicine-Sciences/Flammarion, 1987, pp 163-173 3. Clowes AW, Reidy MA, Clowes MM: Kinetics of cellular proliferation after arterial injury: I. Smooth muscle growth in the absence of endothelium. Lab Invest 1983, 49:327-333 4. Clowes AW, Reidy MA, Clowes MM: Mechanisms of stenosis after arterial injury. Lab Invest 1983, 49:208-215 5. Boyd CD, Kniep AC, Pierce RA, Deak SB, Karboski C, Miller DC, Parker Ml, Mackenzie JW, Rosenbloom J, Scott GE: Increased elastin mRNA levels associated with surgically induced intimal injury. Connect Tissue Res 1988,18:65-78 6. Barnes MJ: Collagens in atherosclerosis. Coll Relat Res

1985, 5:65-97 7. Clowes AW, Karnovsky MJ: Suppression by heparin of smooth muscle cell proliferation in injured arteries. Nature

1977, 265:625-626 8. Guyton JR, Rosenberg RD, Clowes AW, Karnovsky MJ: Inhibition of rat arterial smooth muscle cell proliferation by heparin. In vivo studies with anticoagulant and non-anticoagulant heparin. Circ Res 1980, 46:625-634 9. Clowes AW, Clowes MM: Kinetics of cellular proliferation after arterial injury: II. Inhibition of smooth muscle growth by heparin. Lab Invest 1985, 52:611-616 10. Clowes AW, Clowes MM: Kinetics of cellular proliferation after arterial injury: IV. Heparin inhibits rat smooth muscle mitogenesis and migration. Circ Res 1986, 58:839-845

328

Snow et al


11. Wiebel ER: Stereological methods, Practical Methods for Biological Morphometry. London, Academic Press, 1981, pp 82-91 12. Clowes AW, Clowes MM, Reidy MA: Kinetics of cellular proliferation after arterial injury: III. Endothelial and smooth muscle growth in chronically denuded vessels. Lab Invest 1986,

54:295-303 13. Reidy MA, Clowes AW, Schwartz SM: Endothelial regeneration: V. Inhibition of endothelial regrowth in arteries of rat and rabbit. Lab Invest 1983, 49:569-575 14. Scott JE: Collagen-proteoglycan interactions. Localization of proteoglycans in tendon by electron microscopy. Biochem J 1980,187:887-891 15. Mallory FB: Pathological Technique. New York, Hafner Publishing Co, 1961 16. Scott JE, Dorling J: Differential staining of acid glycosaminoglycans (mucopolysaccharides) by Alcian blue in salt solutions. Histochemie 1965, 5:221-233 17. Mecham RP, Whitehouse LA, Wrenn DS, Parks WC, Griffen GL, Senior RM, Crouch EC, Stenmark KR, Voelkel NF: Smooth muscle-mediated connective tissue remodeling in pulmonary hypertension. Science 1987, 237:423-426 18. Wrenn DS, Mecham RP: Immunology of elastin. Methods Enzymol 1987,144:246-259 19. Hassell JR, Robey PG, Barrach H, Wilczek J, Rennard Si, Martin GR: Isolation of a heparan sulfate-containing proteoglycan from basement membrane. Proc Natl Acad Sci USA 1980, 77:4494-4498 20. Carlson SS, Wight TN: Nerve terminal anchorage protein 1 (TAP-1) is a chondroitin sulfate proteoglycan: Biochemical and electron microscopic characterization. J Cell Biol 1987, 105:3075-3086 21. GlossI J, Beck M, Kresse H: Biosynthesis of proteodermatan sulfate in cultured human fibroblasts. J Biol Chem 1984,259: 14144-14150 22. Saito H, Yamagata T, Suzuki S: Enzymatic methods for the determination of small quantities of isomeric chondroitin sulfates. J Biol Chem 1968, 243:1536-1542 23. Snow AD, Mar H, Nochlin D, Kimata K, Kato M, Suzuki S, Hassell J, Wight TN: The presence of heparan sulfate proteoglycans in the neuritic plaques and congophilic angiopathy in Alzheimer's disease. Am J Pathol 1988,133:456-463 24. Sternberger LA: The unlabeled antibody peroxidase-antiperoxidase (PAP) method, Immunocytochemistry. Edited by HM Zimmerman, New York, Grune and Stratton, 1973, pp 1-26 25. Gittes F, Bolender RP: Counting cell nuclei with random sections: The effect of shape and size. Micron Microsc Acta 1987,18:59-67 26. Bolender RP, Pentcheff ND: Users manual: PCS system 11. Health Science Center for Educational Resources SB-56, Seattle, University of Washington, 1985 27. Snedecor GW, Cochran WG: Statistical methods. Ames, IA, The Iowa State University Press, 1967 28. Zar JH: Biostatistical Analysis. New Jersey, Prentice Hall, 1974

29. Dorling J: 'Critical electrolyte concentration' method in histochemistry. J Med Lab Tech 1969, 26:124-130 30. Scott JE: Histochemistry of alcian blue: Ill. The molecular basis of staining by alcian blue 8GX and analogous phthalocyanins. Histochemie 1972, 32:191 -212 31. Scott JE, Haigh M: Proteoglycan-collagen interactions in intervertebral disc. A chondroitin sulphate proteoglycan associated with collagen fibrils in rabbit annulus fibrosus at the d-e bands. Biosc Rep 1986, 6:879-888 32. Van Kuppevelt THMSM, Rutten TLM, Kuyper CMA: Ultrastructural localization of the proteoglycans in tissue using Cuprolinic blue according to the critical electrolyte concentration method: Comparison with biochemical data from the literature. Histochem J 1987,19:520-526 33. Volker W, Schmidt A, Buddecke E: Compartmentation and characterization of different proteoglycans in bovine arterial wall. J Histochem Cytochem 1986,34:1293-1299 34. Volker W, Schmidt A, Buddecke E: Mapping of proteoglycans in human arterial tissue. Eur J Cell Biol 1987, 45:72-79 35. Bolender RP: Correlation of morphometry and stereology with biochemical analysis of cell fractions. Int Rev Cytol 1978, 55:247-289 36. Bertram JF, Sampson PD, Bolender RP: Influence of tissue composition on the final volume of rat liver blocks prepared for electron microscopy. J Electron Microsc Tech 1986, 4: 303-314 37. Bolender RP: Methods for decreasing the statistical variance of stereological estimates. Anat Rec 1983, 207:89-108 38. Prosser IW, Stenmark KR, Suthar M, Crouch EC, Mecham RP, Parks WC: Regional heterogeneity of elastin and collagen gene expression in intralobar arteries in response to hypoxic pulmonary hypertension as demonstrated by in situ hybridization. Am J Pathol 1989,135:1073-1078 39. Wight TN, Lark MW, Kinsella MG: Blood vessel proteoglycans, Biology of Extracellular Matrix; Biology of Proteoglycans. Edited by TN Wight, RP Mecham. Orlando, FL, Academic Press, 1987, pp 267-300 40. Dvorak HF: Tumors: Wounds that do not heal. Similarities between tumor stroma generation and wound healing. N Engl J Med 1986, 315:1650-1659 41. Couchman JR, Hook M: Proteoglycans and wound repair, Molecular and Cellular Biology of Wound Repair. Edited by RAF Clark, PM Henson. New York, Plenum Press, 1988, pp 437-470 42. Holderbaum D, Ehrhart LA: Modulation of types and Ill procollagen synthesis at various stages of arterial smooth muscle cell growth in vitro. Exp Cell Res 1984,153:16-24 43. Hollman J, Thiel J, Schmidt A, Buddecke E: Increased activity of chondroitin sulfate synthesizing enzymes during proliferation of arterial smooth muscle cells. Exp Cell Res 1986, 167:484-494 44. Majack RA, Coates-Cook S, Bornstein P: Platelet-derived growth factor and heparin-like glycosaminoglycans regulate thrombospondin synthesis and deposition in the matrix by smooth muscle cells. J Cell Biol 1985,101:1059-1070 45. Breton M, Berrou E, Brahimi-Horn MC, Deudon E, Picard J: Synthesis of sulfated proteoglycans throughout the cell cy-


329


cle in smooth muscle cells from pig aorta. Exp Cell Res

46.

47.

48.

49. 50.

51.

52.

53.

54.

55.

56.

57.

58. 59.

60.

61.

62.

1986,166:416-426 Schmidt A, Bunte A, Buddecke E: Proliferation dependent changes of proteoglycan metabolism in arterial smooth muscle cells. Biol Chem 1987, 368:277-284 Wight TN, Potter-Perigo S, Aulinskas T: Proteoglycans and cell proliferation. Am J Respir Dis 1989,140:1132-1135 Chen JK, Hoshi H, McKeehan WL: Transforming growth factor beta specifically stimulates synthesis of proteoglycan in human arterial smooth muscle cells relative to endothelial cells. Proc Natl Acad Sci USA 1987, 84:5287-5291 Clowes AW, Rosenberg RD, Clowes MM: Regulation of arterial smooth muscle cell proliferation by heparin in vivo. Surg Forum 1983, 34:357-360 Castellot JJ Jr, Addonizio ML, Rosenberg RD, Karnovsky MJ: Cultured endothelial cells produce a heparin-like inhibitor of smooth muscle cell growth. J Cell Biol 1981, 90:372379 Castellot JJ Jr, Favreau LV, Karnovsky MJ, Rosenberg RD: Inhibition of vascular smooth muscle cell growth by endothelial cell-derived heparin. Possible role of a platelet endoglycosidase. J Biol Chem 1982, 57:11256-11260 Hoover RL, Rosenberg RD, Haering W, Karnovsky MJ: Inhibition of rat arterial smooth muscle cell proliferation by heparin: II. In vitro studies. Circ Res 1980, 47:578-583 Fritze LMS, Reilly CP, Rosenberg RD: An antiproliferative heparan sulfate species produced by post confluent smooth muscle cells. J Cell Biol 1985,100:1041-1049 Benitz WE, Lessler DS, Coulson JD, Bernfield M: Heparin inhibits proliferation of fetal vascular smooth muscle cells in the absence of platelet derived growth factor. J Cell Physiol 1986,127:1-7 Wright TC, Castellot JJ, Petitou M, Lormeau TC, Choay J, Karnovsky MJ: Structural determinants of heparin's growth inhibitory activity. Interdependence of oligosaccharide size and charge. J Biol Chem 1989, 264:1534-1542 Scoft-Burden T, Buhler FR: Regulation of smooth muscle proliferative phenotype by heparinoid-matrix interactions. Trends in Pharm Sci 1988, 9:94-98 Majack RA, Clowes AW: Inhibition of vascular smooth muscle cell migration by heparin-like glycosaminoglycans. J Cell Physiol 1984,118:253-256 Chamley-Campbell JH, Campbell GR: What controls smooth muscle phenotype? Atherosclerosis 1981, 40:347-357 Brown CC, Balian G: Effect of heparin on synthesis of short chain collagen by chondrocytes and smooth muscle cells. J Cell Biol 1987,105:1007-1012 Graham MF, Drucker DEM, Perr HA, Diehelmann RF, Ehrlich HP: Heparin modulates human intestinal smooth muscle cell proliferation, protein synthesis, and lattice contraction. Gastroenterology 1987, 93:801-809 Graham MF, Drucker DEM, Diegelmann RF, Elson CO: Collagen synthesis by human intestinal smooth muscle cells in culture. Gastroenterology 1987, 92:400-405 Majack RA, Bornstein P: Heparin regulates the collagen phenotype of vascular smooth muscle cells: Induced synthesis of an Mr 60,000 collagen. J Cell Biol 1985,100:613-619

63. Tan EML, Levine E, Sorger P, Unger GA, Hacobian N, Planck B, lozzo RV: Heparin and endothelial cell growth factor modulate collagen and proteoglycan production in human smooth muscle cells. Biochem Biophys Res Comm 1989, 63:84-92 64. Gavriel P, Kagan HM: Inhibition by heparin of the oxidation of lysine in collagen by lysyl oxidase. Biochemistry 1988, 27: 2811-2815 65. Fornieri C, Baccarani-Contri M, Quaglino D Jr, Pasquali-Ronchetti I: Lysyl oxidase activity and elastin/glycosaminoglycan interactions in growing chick and rat aortas. J Cell Biol 1977,105:1463-1469 66. Pasquali-Ronchetti I, Bressan GM, Fornieri C, BaccaraniContri M, Volpin D: Elastin fibers associated glycosaminoglycans in beta-aminopropionitrile induced lathyrism. Exp Mol Pathol 1984, 40:235-245 67. Ehrlich HP, Griswold TR, Rajarathanum JBM: Studies on vascular smooth muscle cells and dermal fibroblasts in collagen matrices. Effects of heparin. Exp Cell Res 1986,164:154-162 68. Guidry C, Grinnell F: Heparin modulates the organization of hydrated collagen gels and inhibits gel contraction by fibroblasts. J Cell Biol 1987,104:1097-1103 69. McPherson JM, Ledger PW, Ksander G, Sawamura SJ, Conti A, Kincaid S, Michaeli D, Clark RAF: The influence of heparin on the wound healing response to collagen implants in vivo. Coll Relat Res 1988,1:83-100 70. McPherson JM, Swamura SJ, Condell RA, Rhee W, Wallace DG: The effects of heparin on the physicochemical properties of reconstituted collagen. Coll Relat Res 1988,1:65-82 71. Bienenstock JA, Befus AD, Denburg JA: Mast cell heterogeneity: Basic questions and clinical implications, Mast Cell Differentiation and Heterogeneity. Edited by J Bienenstock, AD Befus, JA Denburg. New York, Raven Press, 1986, pp 391-402 72. Sabamot S, Goldhaber P, Glicher MJ: Mouse bone collagenase. The effect of heparin on the amount of enzyme released in tissue culture and on the activity of the enzyme. Calcif Tissue Res 1973,12:247-258 73. Jaffe MD, Willis PW: Multiple fractures associated with longterm sodium heparin therapy. J Am Med Assoc 1965, 193: 152-154 74. Jordan RE, Kilpatrick J, Nelson RM: Heparin promotes the inactivation of anti-thrombin by neutrophil elastase. Science 1987, 237:777-780 75. Redini F, Tixier J, Petitou M, Choay J, Robert L, Hornebeck W: Inhibition of leucocyte elastase by heparin and its derivatives. Biochem J 1988, 252:515-519 76. Redini F, Lafuma C, Hornebeck W, Choay J, Robert L: Influence of heparin fragments on the biological activities of elastase(s) and protease inhibitor. Biochem Pharmacol 1988, 37:4257-4261 77. lozzo RV, Wight TN, Bolender RP: Proteoglycan changes in the intercellular matrix of human colon carcinoma: An integrated biochemical and stereological analysis. Lab Invest 1982, 47:124-138 78. Wever J, Schachtschabel 0, Sluke G, Wever G: Effect of short or long term treatment with erogenous glycosamino-

330

Snow et al

4/P Auigust 1990, Vol. 137, No. 2

79.

80.

81.

82.

glycans on growth and glycosaminoglycan synthesis of human fibroblasts (WI38) in culture. Mech Ageing Dev 1980, 14:89-99 Harris SA, Gajdusek C, Schwartz SM, Wight TN: The stimulatory effect of heparin on proteoglycan biosynthesis by arterial smooth muscle cells in vitro (abstr). Fed Proc 1981, 40:623a Irimura T, Nakajima M, Nicolson GL: Chemically modified heparins as inhibitors of heparan sulfate specific endoglucuronidase (heparanase) of metastatic melanoma cells. Biochemistry 1986, 25:5322-5328 Bar-Ner M, Kramer MD, Schirrmacher V, lshai-Michaeli R, Fuks Z, Vlodavsky I: Sequential degradation of heparan sulfate in the subendothelial extracellular matrix by highly metastatic lymphoma cells. Int J Cancer 1985, 35:483-491 Bar-Ner M, Eldor A, Wasserman L, Matzner Y, Cohen IR, Fuks Z, Vlodavsky I: Inhibition of heparanase-mediated deg-

radation of extracellular matrix heparan sulfate by non-anticoagulant heparin species. Blood 1987, 70:551-557 83. Au YPT, Kenagy RD, Clowes AW: Heparin inhibits tissue type plasminogen activator gene expression in cultured baboon vascular smooth muscle cells. J Cell Biol 1989, 107: 504(Abstr) 84. Richardson M, Hatton MWC, Moore S: The plasma proteases, thrombin and plasmin, degrade the proteoglycan of rabbit aorta segments in vitro: An integrated ultrastructural and biochemical study. Clin Invest Med 1988,11:139-150

Acknowledgments The authors thank Dr. Jurgen Fingerle for surgical assistance. In addition, the technical assistance of Stephanie Lara and Selena Certeza was greatly appreciated.