endothelial cell growth factor - Europe PMC

Proc. Natl. Acad. Sci. USA Vol. 84, pp. 7124-7128, October 1987

Cell Biology

Human vascular smooth muscle cells both express and respond to heparin-binding growth factor I (endothelial cell growth factor) (acidic fibroblast growth factor/atherosclerosis/paracrine/autocrine)

JEFFREY A. WINKLES*t, ROBERT FRIESEL*t, WILSON H. BURGESS*t, RICHARD HOWK*, TEVIE MEHLMAN*t, ROBERT WEINSTEINt, AND THOMAS MACIAG*t§ *Biotechnology Research Center, Meloy Laboratories, Inc., 4 Research Court, Rockville, MD 20850; and tDepartment of Biomedical Research, St. Elizabeth's Hospital of Boston, 736 Cambridge Street, Boston, MA 02135

Communicated by Earl P. Benditt, June 16, 1987

binding growth factor (basic fibroblast growth factor) (13). The major serum mitogen for connective tissue cells, including vascular smooth muscle cells (14, 15) is platelet-derived growth factor (PDGF). However, endothelial cells do not possess cell-surface receptors for PDGF and, therefore, are not responsive to PDGF (16). PDGF purified from platelets exists as a dimer of partially homologous A and B chains (17). The PDGF B chain is encoded by the SIS gene, the cellular homolog to the transforming gene v-sis of simian sarcoma virus (17, 18). The control of vascular cell proliferation in vivo presumably requires the precise temporal and spatial regulation of growth factor delivery. Although the delivery of PDGF to smooth muscle cells located at sites of vessel wall injury was initially thought to involve only the platelet and macrophage, it is now known that human endothelial cells express mRNA for the A and B chains of PDGF (19-21) and bovine endothelial cells synthesize PDGF-like proteins (22, 23). These data indicate a potential endothelial cell-derived paracrine mechanism for the control of smooth muscle cell growth within the vessel wall. An autocrine mechanism involving PDGF may also exist; cultured rat arterial smooth muscle cells express PDGF mRNA (24, 25) and synthesize PDGF-like proteins (26). In contrast, the control of endothelial cell proliferation in the vessel wall is not well established. Although polypeptide mitogens for endothelial cells are well characterized from tissue sources (27), the absence of these polypeptides in plasma (28) makes the circulatory system an unlikely route of delivery to sites of endothelial cell injury. Reports have indicated that one endothelial cell mitogen, class II heparin-binding growth factor, is expressed by bovine endothelial cells themselves (29, 30). We report that cultured human aortic and umbilical vein smooth muscle cells, but not cultured human umbilical vein endothelial cells (HUVEC), express HBGF-I mRNA. Human aortic smooth muscle cells (HASMC) also express an HBGF-I-like polypeptide, possess HBGF-I receptors on their cell surface, and respond to HBGF-I as a mitogen. These results suggest that vascular cell proliferation in vivo may be regulated by both paracrine and autocrine mechanisms.

ABSTRACT The control of vascular endothelial and smooth muscle cell proliferation is important in such processes as tumor angiogenesis, wound healing, and the pathogenesis of atherosclerosis. Class I heparin-binding growth factor (HBGFI) is a potent mitogen and chemoattractant for human endothelial cells in vitro and will induce angiogenesis in vivo. RNA gel blot hybridization experiments demonstrate that cultured human vascular smooth muscle cells, but not human umbilical vein endothelial cells, express HBGF-I mRNA. Smooth muscle cells also synthesize an HBGF-I-like polypeptide since (i) extract prepared from smooth muscle cells will compete with III-labeled HBGF-I for binding to the HBGF-I cell surface receptor, and (it) the competing ligand is eluted from heparinSepharose affinity resin at a NaCl concentration similar to that required by purified bovine brain HBGF-I and stimulates endothelial cell proliferation in vitro. Furthermore, like endothelial cells, smooth muscle cells possess cell-surface-associated HBGF-I receptors and respond to HBGF-I as a mitogen. These results indicate the potential for an additional autocrine component of vascular smooth muscle cell growth control and establish a vessel wall source of HBGF-I for endothelial cell division in vivo.

The two major cellular components of the vasculature are the endothelial and smooth muscle cells. The endothelium constitutes a monolayer of cells lining the inner surface of all blood vessels and functions as a nonthrombogenic interface between blood and tissue. Smooth muscle cells are the predominant cellular constituent of the medial layer of the arterial wall. Both of these cell types normally exist in vivo in a quiescent state (1-3) but will proliferate and migrate in response to vessel wall injury (3, 4). In addition, endothelial cells proliferate during the neovascularization associated with tumor growth and a variety of diseases (4, 5), and intimal smooth muscle cell hyperplasia is a major characteristic of the developing atherosclerotic lesion (6-8). Thus, factors that regulate vascular cell growth in vivo are of interest. Vascular endothelial and smooth muscle cells will proliferate in vitro in response to purified polypeptide mitogens. Both acidic and basic angiogenic heparin-binding endothelial cell growth factors have been described (9). The polypeptides within the acidic class, including those termed endothelial cell growth factor (10), acidic fibroblast growth factor (11), and a retina-derived growth factor (12), represent various molecular weight forms of the same mitogen. To simplify the terminology, the acidic form has been designated class I heparin-binding growth factor (HBGF-I) by Lobb et al. (9), and we intend to conform to their nomenclature. HBGF-I has ==50% amino acid sequence identity with class II heparin-

Abbreviations: HBGF-1, class I heparin-binding growth factor; PDGF, platelet-derived growth factor; HASMC, human aortic smooth muscle cells; HUVEC, human umbilical vein endothelial cells; LE-I1, murine lung capillary endothelial cells. tPresent address: Laboratories of Molecular Biology, Jerome H. Holland Biomedical Research Laboratory, American Red Cross, 15601 Crabbs Branch Way, Rockville, MD 20855. §To whom reprint requests should be addressed at: Laboratories of Molecular Biology, Jerome H. Holland Biomedical Research Laboratory, American Red Cross, 15601 Crabbs Branch Way, Rockville, MD 20855.

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

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MATERIALS AND METHODS Cell Culture. HUVEC were a generous gift from Michael Gimbrone (Harvard Medical School). HUVEC were grown in medium 199, 10% (vol/vol) fetal bovine serum (HyClone, Logan, UT), crude HBGF-I at 100 ,ug/ml, and heparin at 5 units/ml as described (31, 32). HUVEC were used between passage 4 and 15 for all experiments. Murine lung capillary endothelial cells (LE-II) were a generous gift from A. B. Schreiber (Meloy Laboratories, Rockville, MD) and were grown in Dulbecco's modified Eagle's medium (DMEM), 5% (vol/vol) fetal bovine serum, and 5% (vol/vol) charcoaltreated fetal bovine serum as described (33). Human smooth muscle cells were grown on fibronectin-coated (10 ug/cm2) tissue culture dishes in medium 199, 10% (vol/vol) fetal bovine serum. The two strains used in this work were initially isolated from the abdominal aorta of a 19-year-old male (HASMC) and from umbilical vein (HUVSMC). The HUVSMC were a generous gift from Charles Selden (Johns Hopkins University, Baltimore). Human foreskin fibroblasts (Meloy Laboratories) were grown in DMEM, 10% (vol/vol) fetal bovine serum, 1% sodium pyruvate, and 1% nonessential amino acids. Preparation of RNA and RNA Blot Hybridization. Total RNA from cultured cells was prepared as described by Sargent et al. (34) except the LiCl precipitation step was omitted. Fifteen micrograms of total RNA was denatured in 2.2 M formaldehyde/50% (vol/vol) formamide and subjected to electrophoresis in a 1.25% agarose gel containing 2.2 M formaldehyde. The gels were strained with ethidium bromide at 0.5 ,ug/ml and photographed to verify that each lane contained an equal amount of undegraded RNA. The RNA was transferred to nylon filters (Zetabind, Cuno, Meriden, CT) by electroblotting; the filters were then air dried for 30 min, irradiated with short-wave UV light for 5 min, and baked in a vacuum oven at 80°C for 2 hr. Hybridization to 32p_ labeled nick-translated DNA probes and subsequent washing of the filters was carried out essentially as described by Church and Gilbert (35) except denatured salmon sperm DNA at 100 ,g/ml was included in the hybridization solution, and a final high-stringency wash was performed at 65°C in 36 mM NaCl/2 mM NaH2PO4, pH 7.2/0.2 mM EDTA. The DNA fragments used as probes were the 2.2-kilobase (kb) EcoRI fragment of human HBGF-I cDNA clone 1 (36) and the 0.9-kb Pst I-Xba I fragment of plasmid pVsis (37) obtained from K. Robbins (National Institutes of Health, Bethesda, MD). Preparation of Cell Extracts. HASMC or HUVEC were grown to confluence and then harvested by washing twice with cold isotonic phosphate-buffered saline (PBS) and gently scraping the dishes with a rubber policeman. Cells were pelleted at 800 x g, resuspended in 1.0 ml of 50 mM Tris HCl/10 mM EDTA, pH 7.4 (extraction buffer, EB), and disrupted by sonication (Microprobe Heat Systems, Farmingdale, NY). Insoluble material was pelleted by centrifugation (12,000 x g, 10 min), and the resulting supernatant was either used immediately or frozen at -70°C. Protein concentrations of cell extracts were determined by the method of Lowry et al. (38). Heparin-Sepharose Chromatography of HASMC Extract. HASMC were washed, scraped, and pelleted as described above. Cells (-2 x 108) were resuspended in 30 ml of EB and disrupted with a Polytron homogenizer. The homogenate was centrifuged at 15,000 x g for 20 min, and the resulting supernatant was then incubated with 5 ml of heparinSepharose (Pharmacia) for 1 hr at 40C with end-over-end mixing. The heparin-Sepharose was packed into a column and then sequentially washed with 50 ml of EB, with 50 ml of EB containing 0.65 M NaCl, with 30 ml of 1.2 M NaCI, and

Proc. Natl. Acad. Sci. USA 84 (1987)

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with 30 ml of 2.0 M NaCl. Fractions (2.5 ml) were collected and stored at 40C until assayed. HBGF-I Receptor Binding Assay. HBGF-I was purified from bovine brain as described by Burgess et al. (39); the polypeptide used for these studies corresponds to the a form of endothelial cell growth factor. HBGF-I was iodinated as described (40); receptor competition assays were also performed as described (41) with some modifications. Briefly, LE-I1 cells were grown to confluence in 24-well cluster plates (Costar, Cambridge, MA). Cells were washed once in binding buffer (BB; DMEM, 25 mM Hepes, bovine serum albumin at 5 mg/ml, pH 7.4) and then received various concentrations of HUVEC extract, HASMC extract, or purified bovine HBGFI in the presence of 125I-labeled HBGF-I at 10 ng/ml (specific activity, 105 cpm/ng). The assay was also performed using an equal volume of the HASMC extract heparin-Sepharose fractions. After incubation at 40C for 90 min, cultures were washed three times with cold BB, and the cell-associated radioactivity was determined after solubilizing the cells with 1.0 ml of 0.1 M NaOH. Chemical Cross-Linking of 1251-Labeled HBGF-I to Cells. Chemical cross-linking of 125I-labeled HBGF-I to cells was performed as described (40). Briefly, HUVEC and HASMC were grown to confluence in 100-mm tissue culture dishes. Cells were washed once with BB and then incubated in 3.5 ml of BB containing 125I-labeled HBGF-I at 10 ng/ml with or without unlabeled HBGF-I at 1.0 ,g/ml for 90 min at 40C. Binding was terminated by washing three times with BB, and cross-linking was initiated by the addition of disuccinimidyl suberate to a final concentration of 0.3 mM. Samples were processed for electrophoresis as described (40) and separated on 7.5% polyacrylamide gels. Gels were stained, destained, dried, and exposed to Kodak X-Omat AR film at -70°C with an intensifying screen. For some experiments, LE-II cells were grown to confluence in 35-mm dishes, and HUVEC or HASMC cell extracts were tested for their ability to compete with 125I-labeled HBGF-I in the cross-linking assay. LE-II cells were washed once with BB and incubated with 1251_ labeled HBGF-I at 10 ng/ml with or without 150 ,ug of either HUVEC or HASMC extract for 2 hr at 4°C. Cells were then washed, exposed to disuccinimidyl suberate, and processed as described above. [3H]Thymidine Incorporation Assays. Fractions eluted from heparin-Sepharose chromatography of the HASMC extract were assayed for mitogenic activity on LE-II cells by [3H]thymidine incorporation as described (33). The mitogenic responsiveness of HASMC to HBGF-I was determined by growing HASMC to confluence in 24-well cluster plates and then incubating them for 2 days in 2% (vol/vol) calf serum. Various concentrations of purified bovine HBGF-I were then added, and the cells were incubated at 37°C for 18 hr. HASMC were then pulsed with [3H]thymidine at 0.5 ,tCi/ml (New England Nuclear, 6.7 Ci/mmol; 1 Ci = 37 GBq) and further incubated at 37°C for 6 hr. Cells were subsequently washed once with PBS, and the trichloroacetic acid-precipitable radioactivity was determined as described (33).

RESULTS Smooth Muscle Cells Express HBGF-I mRNA. Human HBGF-I cDNA clones have been obtained by oligonucleotide screening of a human brain stem cDNA library (36). We identified cell types expressing HBGF-l mRNA by hybridizing a nick-translated HBGF-I probe to RNA gel blots containing equivalent amounts of RNA prepared from various cultured cells. As shown in Fig. 1, lanes 1-4, human foreskin fibroblasts and two different human vascular smooth muscle cell strains, but not HUVEC, express a major HBGF-I transcript of 4.4 kb. A similarly sized poly(A)+ RNA has also been detected in human brain stem tissue (36). We

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Proc. Natl. Acad. Sci. USA 84 (1987)

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FIG. 1. RNA gel blot analysis of HBGF-I (ECGF, endothelial cell growth factor) and PDGF-B mRNA expression. Fifteen micrograms of total RNA prepared from human foreskin fibroblasts (lanes 1 and 5), abdominal aorta smooth muscle cells (lanes 2 and 6), umbilical vein smooth muscle cells (lanes 3 and 7), and umbilical vein endothelial cells (lanes 4 and 8) were electrophoresed on a 1.25% agarose/formaldehyde gel and transferred to a nylon membrane. The blot containing lanes 1-4 was hybridized to nick-translated HBGF-I cDNA, and the blot containing lanes 5-8 was hybridized to nicktranslated v-sis DNA. The migration of 28S and 18S rRNA as determined by ethidium bromide staining is shown on the left.

also detect two minor transcripts of 3.3 and 2.8 kb. The two human smooth muscle cell strains analyzed here were derived from abdominal aorta (lane 2) and umbilical vein (lane 3). Three additional smooth muscle cell strains (two derived from abdominal aorta and one from internal mammary artery) also express HBGF-I mRNA (data not shown). Similar RNA gel blots were hybridized to a v-sis probe to analyze PDGF-B chain mRNA expression (lanes 4-8). We did not detect PDGF-B mRNA expression in human foreskin fibroblasts or human smooth muscle cells. However, as reported, HUVEC express a 3.7-kb PDGF-B mRNA (19-21, 25). Since the same HUVEC RNA preparation was used for lanes 4 and 8, the PDGF-B mRNA hybridization demonstrates that the RNA used for lane 4 is undegraded. Smooth Muscle Cells Express an HBGF-I-Like Polypeptide. The presence of HBGF-I mRNA in human vascular smooth muscle cells suggests that HBGF-I is synthesized by these cells. We determined whether an HBGF-I-like polypeptide was present in HASMC by asking (i) whether a HASMC extract could compete with 1251-labeled HBGF-I for binding to the HBGF-I cell surface receptor, and (ii) whether any such competing ligand in the HASMC extract had structural and biological properties similar to purified bovine brain A

HBGF-I. We have shown that 125I-labeled HBGF-I will bind specifically to a membrane receptor on several cell types (28) and can be cross-linked covalently to an apparent Mr 150,000 cell surface protein present on LE-II cells (40). Various other polypeptide growth factors will not compete for 1251-labeled HBGF-I binding in either the direct binding or cross-linking assay (28, 40). The degree and specificity of cell extract competition for 1251I-labeled HBGF-I binding to the HBGF-I receptor were measured by preparing extract from cells that express (HASMC) and do not express (HUVEC) the HBGF-I mRNA transcript. A constant amount of 125I-labeled HBGF-I and various amounts of each extract were incubated with LE-II cells, and the cell-associated radioactivity was quantitated. Extract from HASMC but not HUVEC competed with 1251-labeled HBGF-I for binding to LE-II cells (Fig. 2A). The competing activity in the HASMC extract is both heatand trypsin-sensitive and, therefore, is polypeptide in nature. We confirmed that the competing ligand was binding specifically to the HBGF-I receptor by testing the same extracts for competition in a covalent cross-linking assay. As shown in Fig. 2B, although this qualitative assay revealed some competition using 150 ,ug of the HUVEC extract protein (lane 3), the same amount of HASMC extract protein competed significantly more with 125I-labeled HBGF-I for receptor occupancy (lane 4). Since additional HUVEC extracts did not compete in the cross-linking assay, the competition observed in lane 3 may be due to nonspecific cross-linking inhibitors in this particular extract preparation. The HASMC extract was then chromatographed on heparin-Sepharose to determine if the HBGF-I-like protein in HASMC could in fact bind heparin. The column breakthrough and fractions eluting at four different NaCl concentrations were collected and first tested for their ability to compete with 125I-labeled HBGF-I for binding to LE-II HBGF-I cell surface receptors. There was minimal competing activity in the breakthrough; as shown in Fig. 3A, all of the heparin-binding competing activity could be eluted with either 1.2 or 2.0 M NaCl. The activity eluting with 1.2 M NaCl has an affinity for heparin similar to that of HBGF-I (33, 39), whereas the activity eluting with 2.0 M NaCl may represent class II heparin-binding growth factor, which has a higher affinity for heparin (9) and has been reported to interact with the HBGF-I receptor (42). The breakthrough and heparinSepharose binding fractions were then tested for their endothelial cell mitogenic activity, as assayed by the stimulation of [3H]thymidine incorporation in LE-IT cells (Fig. 3B).

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FIG. 2. (A) Analysis of HBGF-I (ECGF, endothelial cell growth factor) receptor competing activity in HASMC and HUVEC cell extracts. Cell extracts were prepared from HASMC (A) or HUVEC (o) and used in a HBGF-I receptor competition assay. A standard curve showing the competing activity of unlabeled bovine HBGF-I (O) is also shown. (B) Cross-linking competition assay of HASMC and HUVEC cell extracts. LE-I1 cells were incubated in the presence of 'l25-labeled HBGF-I (251I-ECGF) at 10 ng/ml (lanes 1-4) and no additions (lane 1), bovine HBGF-I at 1.0 ,g/ml (lane 2), HUVEC extract at 150 ,ug/ml (lane 3), or HASMC extract at 150 ,ug/ml (lane 4) for 90 min. Cells were then washed, cross-linked with disuccinimidyl suberate, and processed for electrophoresis. An autoradiograph of the polyacrylamide gel is shown. The migration of high molecular weight protein size markers (Bio-Rad) is shown on the left.

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FIG. 4. Cross-linking of 1251_ labeled HBGF-I to HASMC and HUVEC. 125I-labeled HBGF-1 was incubated with HASMC (lanes 1 and 2) or HUVEC (lanes 3 and 4) in the presence (lanes 2 and 4) or absence (lanes 1 and 3) of excess unlabeled HBGF-1, and cross-linking was performed. An autoradiograph of the polyacrylamide gel is shown.

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density in vitro (data not shown). These results demonstrate that HASMC, like HUVEC, contain HBGF-I receptors and respond to HBGF-I as a mitogen.

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FIG. 3. (A) HBGF-I receptor competing activity of fractions from HASMC cell extract purified over heparin-Sepharose. Aliquots (10 ,ul) of each fraction from a HASMC extract purified over heparinSepharose were tested for their ability to compete with 125I-labeled HBGF-I for binding to LE-Il cells. The data are plotted as the % inhibition of binding of 1251-labeled HBGF-I (1251-ECGF, where ECGF is endothelial cell growth factor) at 10 ng/ml for each fraction. The concentration of NaCI used to elute the various fractions is shown along the top. (B) Mitogenic activity of fractions from a HASMC extract purified over heparin-Sepharose. The same heparinSepharose fractions were also tested for their mitogenic activity in the LE-II cell [3H]thymidine incorporation assay. Aliquots were incubated at a 1:100 dilution with LE-II cells for 18 hr at 37°C and then pulsed with [3H]thymidine. a

The majority of the HASMC-derived endothelial cell mitogenic activity binds to heparin-Sepharose; the active fractions are present in the 0.65, 1.2, and 2.0 M NaCl elution pools. The results of these two assays indicate that HASMC express numerous heparin-binding endothelial cell mitogens with different structural properties, of which HBGF-I-like molecules are one class. Smooth Muscle Cells Contain Biologically Active HBGF-I Cell Surface Receptors. We next determined whether HASMC have HBGF-I receptors and respond to HBGF-I as a mitogen. The presence of HBGF-I receptors on HASMC was tested using the 1251I-labeled HBGF-I cross-linking assay. As shown in Fig. 4, an apparent Mr 150,000 cross-linked 1251-labeled HBGF-I-HBGF-I receptor complex was detected in both HASMC and HUVEC (lanes 1 and 3); in addition, complex formation was inhibited by an excess of unlabeled HBGF-I (lanes 2 and 4). The effect of HBGF-I on HASMC proliferation in vitro was investigated using both [3H]thymidine incorporation and cell growth assays. HASMC incorporate [3H]thymidine into DNA in response to HBGF-I in a dose-dependent manner (Fig. 5). Half-maximal stimulation of radiolabel incorporation occurs at -1.0 ng of HBGF-I per ml. HASMC also divide in response to exogenous HBGF-I when seeded at low cell

We have demonstrated by RNA gel blot hybridization analysis and HBGF-I receptor binding competition experiments that cultured human smooth muscle cells, but not endothelial cells, express HBGF-I mRNA and HBGF-I-like protein. This result is consistent with a preliminary report indicating cultured smooth muscle cells derived from normal porcine aorta and human carotid plaques express an endothelial cell mitogen (43). In addition to being an endothelial cell mitogen (33), HBGF-I is also an endothelial cell chemoattractant (44) and can induce angiogenesis in vivo (11). HBGF-I could, therefore, modulate neovascularization by the vasa vasorum, a process which occurs during atherosclerosis (45). The mechanism by which smooth muscle cell-derived HBGF-I could interact with neighboring endothelial cells is at present unknown; HBGF-I does not contain a recognizable signal peptide sequence (36) and, therefore, may not be a secreted polypeptide in the classical sense. HBGF-I could be transported out of the smooth muscle cell through its association with binding proteins or heparin-related glycosaminoglycans; alternatively, it may only be released after cell damage. Bovine brain HBGF-I has been described (13) as a mitogen for bovine vascular smooth muscle cells. Similarly, we find 400

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FIG. 5. The effect of HBGF-I (ECGF, endothelial cell growth factor) on [3HIthymidine incorporation by HASMC. HASMC were plated in 24-well plates, grown to confluence in 10% (vol/vol) fetal bovine serum and then starved for 2 days in 2% (vol/vol) calf serum. After 2 days, various concentrations of purified bovine HBGF-1 were added, and [3H]thymidine incorporation was measured. The response to 20% (vol/vol) fetal calf serum (CS) is also shown.

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that human smooth muscle cells contain HBGF-I receptors and will proliferate in vitro in response to added HBGF-I. Although it is tempting to speculate that HBGF-I may act as an autocrine regulator of smooth muscle cell growth in vivo, we presently have no data to directly support this possibility. In addition to HBGF-I, human smooth muscle cells also proliferate in response to PDGF. Monocytes/macrophages, endothelial cells, and smooth muscle cells have been identified as potential nonplatelet sources of PDGF for vascular smooth muscle cells in vivo. Although not permanent residents of the vessel wall, monocytes infiltrate the wall in response to injury and during the pathogenesis of atherosclerosis (7). In culture, activated human monocytes express PDGF-B mRNA and secrete PDGF-like proteins (46). Human endothelial cells also express PDGF mRNA (19-21, 25) and bovine endothelial cells synthesize PDGF-like proteins (22, 23). Various studies have demonstrated that PDGF gene expression by endothelial cells can be modulated. For example, the expression of PDGF-B mRNA decreases during human endothelial cell differentiation in vitro (47) and increases in response to thrombin (48). In addition, secretion of PDGF-like proteins decreases when bovine endothelial cells are treated with modified low density lipoproteins (49) and increases in response to chemical injury (23), thrombin (50), and factor Xa (51). Under certain conditions rat smooth muscle cells can also express PDGF. Primary cultures of adult rat arterial smooth muscle cells express PDGF-A mRNA (24) and PDGF-like proteins (26), and pup rat aortic smooth muscle cells passaged serially express PDGF-I mRNA (25) and secrete PDGF-like proteins (52). The regulation of human smooth muscle cell proliferation in vivo may, therefore, be quite complex; involving both a paracrine mode, as exemplified by the delivery of macrophage- or endothelium-derived PDGF and an autocrine mode, as suggested by the synthesis of HBGF-I or PDGF-A by vascular smooth muscle cells themselves. We thank Linda Peterson for expert secretarial assistance. R.F. performed this work in partial fulfillment of the requirements for the degree of Doctor of Philosophy from the Department of Biochemistry, George Washington University, School of Medicine, Washington, DC 20037. This work was supported in part by Grants HL 01734 and AG 00599 (to R.W.) and HL 32348 and 35627 (to T.M.) from the National Institutes of Health. 1. Gimbrone, M. A., Jr., Cotran, R. S. & Folkman, J. (1974) J. Cell Biol. 60, 673-681. 2. Schwartz, S. M. & Benditt, E. P. (1977) Circ. Res. 41, 248-255. 3. Clowes, A. W., Reidy, M. A. & Clowes, M. M. (1983) Lab. Invest. 49, 327-332. 4. Maciag, T. (1984) Prog. Hemostasis Thromb. 9, 33-58. 5. Folkman, J. (1985) Adv. Cancer Res. 43, 175-203. 6. Schwartz, S. M., Campbell, G. R. & Campbell, J. H. (1986) Circ. Res. 58, 427-444. 7. Ross, R. (1986) N. Engl. J. Med. 314, 488-500. 8. Benditt, E. P. & Benditt, J. M. (1973) Proc. Natl. Acad. Sci. USA 70, 1753-1756. 9. Lobb, R. R., Harper, J. W. & Fett, J. W. (1986) Anal. Biochem. 154, 1-14. 10. Burgess, W. H., Mehiman, T., Marshak, D. R., Fraser, B. A. & Maciag, T. (1986) Proc. Natl. Acad. Sci. USA 83, 7216-7220. 11. Thomas, K. A., Rios-Candelore, M., Gimenez-Gallego, G., DiSalvo, J., Bennett, C., Rodkey, J. & Fitzpatrick, S. (1985) Proc. Natl. Acad. Sci. USA 82, 6409-6413. 12. Baird, A., Esch, F., Gospodarowicz, D. & Guillemin, R. (1985) Biochemistry 24, 7855-7860. 13. Esch, F., Baird, A., Ling, N., Ueno, N., Hill, F., Denoroy, L., Klepper, R., Gospodarowicz, D., Bohlen, P. & Guillemin, R. (1985) Proc. Natl. Acad. Sci. USA 82, 6507-6511.

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