Chronic administration of growth hormone-releasing factor increases wound strength and collagen maturation in granulation tissue

JOURNAL OFSURGICAL RESEARCH 51,297-302 (1991)

Chronic Administration of Growth Hormone-Releasing Factor Increases Wound Strength and Collagen Maturation in Granulation Tissue DOMINIQUE R. GARREL, M.D.,**’ PIERF~ETTEGAUDREAU, PH.D.,? LIMINZHANG, ISABELLEREEVES, B.Sc.,? ANDPAULBRAZEAU, PH.D.?

M.&L,*

*Burn Center, Hotel-Dieu Hospital, 3840 St-Urbain, Montreal, Quebec, Canada H2 W 1 T8; ana’ tNotre-Dame Research Center, 1560 Shmbrooke East, Montreal, Quebec, Canada H2W IT8 Submitted

for publication

April 4, 1990

this effect is responsible for the enhanced cutaneous wound strength seen at 10 and 14 days postwounding in GRF-treated rats. Ol99lAcademicPre~~,Inc.

The effects of chronic administration of growth hormone-releasing factor (GRF) on wound healing were studied in rats. Cutaneous wound strength was measured by tensometry at 5, 10, and 14 days postwounding in rats implanted with a slow-release pellet which contained a compressed mixture of a fatty acid and [desamino Tyr’, D-Ala’, Ala’“]hGRF(l-29)NH, or the fatty acid alone. There was a significant increase in wound tensile strength in GRF-treated rats compared to controls at each measurement: Day 5,130 k 12 vs 9’7 + 14 g; Day 10,402 + 18 vs 280 + 11 g; Day 14,830 f 17 vs 614 + 14 g (P < 0.01 for each value). Granulation tissue obtained from subcutaneously implanted polyvinyl alcohol sponges encased in silicone tubing was also studied. The amount of collagen deposited in the granulation tissue was estimated by measuring the hydroxyproline (Hyp) content of sponges retrieved 5, 10, and I4 days postinsertion from GRF-treated and control rats. Hyp content (nmole/mg sponge) was similar in both treated and control animals at each measurement: Day 5, 1.7 f 0.2 vs 2.2 + 0.2; Day 10,31.9 +- 4.1 vs 26.7 + 0.4; and Day 14, 41.6 k 7.3 vs 38.6 + 4.4. Hyp/proline, Hyp/glycine, and glycine/total amino acid ratios, evaluated after 10 days, were also similar in both groups. Collagen from the granulation tissue of sponges retrieved after 14 days from treated and control rats was studied by electron microscopy (magnifications, 7,100 and 22,720). The intercellular spaces in the sponges of the control rats were sparsely covered by collagen fibrils, in contrast to the intercellular spaces in the sponges of the treated rats, which were completely filled with collagen fibrils. These results demonstrate that chronic administration of GRF for 14 days increases wound strength during the early stage of healing in rats. In addition, GRF does not stimulate collagen deposition in granulation tissue, but accelerates collagen maturation instead. We postulate, therefore, that

1 To whom correspondence

Hospital,

INTRODUCTION The effects of growth hormone (GH) on the process of wound healing are not well understood. Whether GH plays a physiologic role in tissue repair has been debated, because wounds in GH-deficient subjects heal normally. Until recently, it was not possible to study the effects of GH administration on wound healing because naturally occurring GH was very expensive and its availability was scarce. However, with the advent of recombinant DNA human GH (hGH), these effects can now be explored. In recent studies, hGH increased cutaneous wound strength in rats when administered around the time of injury [I] and increased the tensile strength of subcutaneously implanted sponges [2]. However, these studies used large amounts of GH; moreover, daily GH injections do not reproduce its physiologic pulsatile secretion pattern. This pattern may be important since periodic GH administration promotes growth and IGF-I secretion better than continuous infusion [3,4]. GH secretion is controlled by two hypothalamic hormones: somatostatin, which is inhibitory, and growth hormone-releasing factor (GRF) which is stimulatory [5]. The first of these two peptides plays a major role in determining GH pulsatility [6]. Since its isolation and characterization, GRF biological activity has been extensively explored in both animals and humans [7]. Because of its short halflife [8, 91, GRF must be administered continuously to produce a prolonged increase in both basal and stimulated GH secretion. The effects of continuous stimulation with no down-regulation are clearly shown in reported cases of GRF-secreting tumors that cause acromegaly in man, where both basal and pulsatile GH secretions increased [lo]. In healthy subjects, continu-

should be addressed. 297

0022-4804/91$1.50

Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

298

JOURNAL

OF SURGICAL

RESEARCH:

ous, 24-hr GRF infusion stimulates both the basal and the magnitude of pulsatile GH secretion [ 111. In male rats, where bursts of GH secretion occur every 3.12 hr [6], continuous GRF administration increases both basal and stimulated GH secretion [ 121. Accordingly, we implanted rats with a newly designed pellet that slowly released a trisubstituted analog of human GRF (l29)NH2 for up to 14 days to study the effects of chronic stimulation of GH secretion on wound healing.

METHODS

Animals and wounds. Male Sprague-Dawley rats (250 g), obtained from Charles River Canada, Inc. (StConstant, Canada), were housed in individual cages and fed ad libitum. Each rat was weighed daily. The rats were randomly assigned to either GRF-treated or control groups. Each study group contained six to nine rats. Full-thickness cutaneous incisions which included the panniculus carnosus were made longitudinally on the dorsal skin of the animals with a No. 22 scalpel blade under pentobarbital anesthesia (50 mg/kg body wt). Care was taken not to damage underlying muscular tissue. The incisions were immediately closed with dermalon 3-O sutures placed 1 cm apart. Daily urinary collection and food intake measurements were taken in rats whose wounds were studied 14 days postwounding. Blood samples were taken by cardiac puncture after 30 min of pentobarbital anesthesia on the day of sacrifice. The study protocol was approved by the animal care committee of our institution. GRF administration. GRF was chronically administered via a slow-release pellet implanted subcutaneously into the abdomen of the animals. The pellet, weighing 50 mg, consisted of a pyrogen-free, compressed mixture of a fatty acid (85%) and the GRF analog (15%) [desamino Tyr’, D-Ala2, Ala15]hGRF(1-29)NH, (patent pending). Control animals were implanted with a pellet containing only the fatty acid. The GRF analog was synthesized by solid-phase methodology [13] and purified by high-pressure liquid chromatography (HPLC) [14]; purity, 98%; peptide content, 83%. After 14 days, the pellets containing the GRF analog were removed and their residual content was assessed by HPLC after extraction in dimethyl sulfoxide. Using an unimplanted GRF pellet as an external standard for quantification, it was determined that 63 + 10% of the GRF contained in the pellets had been released. Wound breaking strength. Wound breaking strength was measured 5, 10, and 14 days postinjury. The dorsal skin of each rat was excised under pentobarbital anesthesia and cut into three strips, 1 cm wide by 6 cm long each, with four razor blades positioned onto a rigid support. Each strip was placed between the clamps of a 1101 Instron tensometer and the amount of force required to break the strip was recorded. Tension was applied at

VOL.

51, NO. 4, OCTOBER

10 mm/min. fashion.

1991

All measurements

were made in a blind

Dermal thickness. Dermal thickness was measured with a micrometer 14 days postwounding. Dorsal skin samples were excised 1 cm from the original incisions, fixed in 5% formalin, embedded in paraffin, and stained with hematoxylin phloxin safran. Studies ofgranulation tissue. Granulation tissue was obtained from implants made of 1 X 1 X 30-mm pieces of polyvinyl alcohol (PVA) sponge threaded into perforated silicone tubes, as described by Cohen and Diegelmann [15]. The tubes were perforated every 1 mm and the ends were sealed with silastic glue after insertion of the sponge. The implants were thoroughly washed and autoclaved and then inserted dorsally, 1 cm from and parallel to the midline, with a No. 12 needle placed subcutaneously through a stab wound. Wound strength and granulation tissue were studied separately, in different groups of rats. Implants were recovered 5, 10, and 14 days postinsertion for biochemical analysis. Ultrastructural studies. The distal extremity (3 mm) of each sponge retrieved 14 days after insertion was fixed in 3% glutaraldehyde solution and later embedded in resin. Sponges from both treated and control groups were studied. Four sections of each sponge were stained with uranyl acetate and lead and examined under electron microscopy (Phillips Model 400) at magnifications of 7,100 and 22,720. The collagen fibril diameters of 20 fibrils from each sponge were measured on electron micrographs, using a caliper (Mitutoyo No. 500; precision, l/100 per mm). Biochemistry: Amino acids. Total hydroxyproline (Hyp) and other amino acids were analyzed by cationexchange HPLC (40 X 250-mm column, 60“(Z), using postcolumn ortho-phthalaldehyde derivatization and inline conversion with sodium hypochlorite after hydrolysis of sponges in 6 N HCl (llO°C for 18 hr) and quantified using appropriate external amino acid standards [16]. The sensitivity limit of the assay was 250 pmole. Inter- and intra-assay variations were 8 and 9%, respectively. Ratios of Hyp to proline, Hyp to glycine, and glytine to total amino acid content were also evaluated. Biochemistry: Nitrogen. Urinary nitrogen concentrations were measured by chemoluminescence with an Antek analyzer (Antek Instruments, Houston, TX). Biochemistry: Growth hormone. Serum growth hormone concentrations were measured with an anti-murine GH serum, generously given by Dr. Sinha (Whittier Institute for Diabetes and Endocrinology, La Jolla, CA [17]). The reference preparation RP-1, used as standard, and NIADDK-GH-I-4, used as iodination standard, were kindly provided by Dr. Raiti (National Institutes of Arthritis, Diabetes, Digestive and Kidney Diseases, Bethesda, MD). The intra- and inter-assay variations were 5 and 8%.I rewectivelv. ” I

GARREL

‘Ooo

ET AL.:

GROWTH

HORMONE-RELEASING p c 0.01

1 EI

Control 1 Treated

T

n = 6 to 9 rats per group

P-=

p < 0.01 -

5

10 DAYS POST WOUNDING

14

FIG. 1. Wound breaking strength of control and GRF-treated animals was measured on three skin strips per rat with a 1011 Instron tensometer. Speed of tension was 10 mm/min. Values represent the mean f SEM.

Mean values for each parameter Statistical analysis. studied were compared by analysis of variance (SPSS). A treatment effect was tested analyzing the variance of all data pooled together. Statistical significance was established at P < 0.05. RESULTS

Weight gain and food intake were Animal growth. not significantly different in treated and control rats throughout the study. Daily urinary nitrogen excretion was virtually identical in both treated and control rats at 14 days postwounding, as was dermal thickness (1.47 + 0.20 and 1.47 f 0.30 mm, respectively). As shown in Fig. 1, wound Wound breaking strength. breaking strength was significantly higher in treated rats (P < 0.01) at all times. The differences in the force required to break the wounds between treated and control groups were 36, 47, and 35% at 5, 10, and 14 days, respectively. Hemorrhagic wounds, which represented less than 3% of the total wounds, were discarded. No infection was detected. Percentages of variations for the wound breaking strength of the three skin samples measured per rat ranged from 3 to 15%. Granulation tissue. The hydroxyproline content of the sponges is shown in Table 1. No difference was seen between treated and control groups at any time. In addition, the ratios of Hyp/proline, Hyp/glycine, and glytine/total amino acids at 10 days postwounding did not differ significantly between treated and control rats, and were 28 f 6.9% vs 29.3 +- 7.4%, 27.1 + 6.7% vs 22.6 f 5.5%, and 10.7 f 0.2% vs 11.3 f 0.6%, respectively. The typical characteristics of sponges retrieved after 14 days from treated and control rats are shown in Figs. 2A and 2B. The entire extracellular matrix of sponges from treated rats was filled with collagen fibrils. In contrast, only small areas of the extracellular matrix of all sections of sponges from control rats were filled with colla-

FACTOR

AND

WOUND

299

HEALING

gen fibrils. The mean diameter of the fibrils was 70 nm for both treated and control groups. No microfibrillar structures were apparent in the intercellular spaces at 7,100 magnification; however, at 22,720 magnification, some microfibrillar structures were seen (see Fig. 2 insets). At the end of Serum growth hormone concentrations. each treatment period, serum GH concentrations (ng/ ml) were significantly higher in GRF-treated animals than in controls: Day 5,239 + 110 vs 110 f 34 (P < 0.01); Day 10,167 k 18 vs 72 +- 27 (P < 0.05); Day 14,347 rf: 72 vs 103 +- 31 (P < 0.01). DISCUSSION

In this study, GRF-treated rats showed increased wound strength without increased collagen deposition in granulation tissue. Wound healing is a complex cascade of molecular and cellular events which lead to the closure and repair of injured tissue. Collagen deposition and maturation from fibroblasts that migrate to the wound constitute the main biological events that promote wound strength. In incisional skin wounds, collagen may start to accumulate as early as 24 hr after injury. This accumulation increases for 1 to 2 weeks and then plateaus [18,19]. The amount of collagen deposited in the wound cannot be measured directly because of the collagen in normal skin surrounding the wound. Several experimental models have been designed for the study of wound collagen deposition. Among these are subcutaneously implanted chambers or sponges, wherein fibroblasts migrate and collagen is deposited [20]. During the first weeks following injury, wound strength is directly related to the amount of collagen deposited in granulation tissue [21]. Our results are in agreement with this phenomenon, as both cutaneous wound strength and collagen deposition in granulation tissue increased during the first 2 weeks after injury. Wound breaking strength and collagen deposition in subcutaneously implanted materials have also been the main determinants of the efficacy of wound healing enhancing agents.

TABLE Total

Hydroxyproline

1

Content (nmole/mg)

of the Sponges

Days postwounding

A Control Treated

2.2

*

:94

:99 0.2

1.7 * 0.2*

26.7

f

0.4

31.9 f 4.1*

41.6 38.5

2 +

7.0 4.0*

Note. Values represent the mean f SEM. The number in parentheses indicates the sample size. * P > 0.05, compared to controls.

300

JOURNAL

OF SURGICAL

RESEARCH:

VOL.

51, NO. 4, OCTOBER

1991

FIG. 2. Electron micrographs (magnification, 7,100) of sponges retrieved after 14 days from control and GRF-treated rats. Four sections of each sponge sample were examined after staining with uranyl acetate and lead. In sponges from control rats (A), very few collagen fibrils were seen and a nonfibrillar substance occupied most of the area between cells. In contrast, in sponges from treated animals (B), all sections showed collagen fibrils filling the intercellular spaces. Fibrils from treated sponges measured 70 nm in diameter. Insets (top right) are X22,720 magnification.

Among such agents, vitamin A, anabolic steroids, and bone cartilage powder have been shown to increase cutaneous wound breaking strength and wound collagen deposition in rats [22]. More recently, growth factors, which are present in wounds and which may play an important role in cell migration and activity during the entire wound healing process, have been successfully used to increase cutaneous wound strength [23]. As in earlier studies, these reports have been associated with increased wound infiltration by fibroblasts [25-281 and increased collagen production in wound chambers. Our finding that wound breaking strength increases with no concomitant increase in sponge Hyp content differs from previous studies on wound healing enhancing agents. Hyp is specific to collagen molecules (with the exception of elastin, which is present in very small

amounts in granulation tissue) and can be used as a reliable index of collagen deposition. Since proline and glytine make up a third of the amino acids in collagen, the ratios of Hyp/proline and Hyp/glycine would have changed if proline hydroxylation had been altered during collagen synthesis [29]. However, no differences were found in any of these parameters in the sponges retrieved from GRF-treated rats. This suggests that GRF administration did not stimulate collagen deposition and did not change proline hydroxylation. In addition, the ratio of glycine to total amino acids did not differ in treated rats, suggesting that the proportion of collagen molecules to other extracellular matrix proteins was unmodified by GRF administration. Wound strength depends not only on the amount also but on the chemical stability of the collagen in the

GARREL

ET AL.:

GROWTH

HORMONE-RELEASING

wound. Type I collagen molecules, which are the most abundant molecules in wounds, aggregate into fibrils in the extracellular space and form covalent cross-links to strengthen the healing tissue [21]. Such aggregation is a complex phenomenon [29]. Studies in the chick embryo have shown that intracellular packaging of procollagen molecules, as well as complex interactions between the cells and the extracellular space, occurs during the deposition of newly formed collagen fibrils [30]. Our ultrastructural findings suggest that GRF dramatically increases collagen stability. This increased stability may be responsible, at least in part, for the increased wound strength of GRF-treated rats. The predominant intercellular substance in the sponges implanted for 14 days in control rats was unidentifiable, as it lacked the characteristic features of collagen fibrils. Indeed, there was only indirect evidence to suggest that the substance was in fact collagen. The microfibrillar aspect of the substance was evident at the higher (22,720) magnification only. Furthermore, Hyp data did not differ between sponges of control and treated animals, suggesting that collagen had been deposited equally in the sponges of both groups. It could be hypothesized that this substance is collagen that has been adulterated by the fixation and staining procedures. This would suggest that the collagen was more stable in sponges from treated rats. Further investigation is needed to identify the extracellular matrix components of the granulation tissue that was harvested in the sponges. Our data do not give information about the site or mechanism of the accelerated collagen maturation observed with GRF administration. It is possible that different types of collagen and other components of the extracellular matrix, such as fibronectin and proteoglycans, could play a role in this maturation, and that GRF administration could have other effects during the first days after wounding. Indeed, wounds contain very little collagen at this time, when wound strength depends on epithelialization rather than on collagen deposition [31]. The mechanisms responsible for our observations regarding wound strength are probably related to increased GH secretion. The stimulation of cellular events inside the wound only, rather than a general GH-induced improvement in all anabolic processes, is suggested by the lack of change in body weight, nitrogen retention, and dermal thickness in GRF-treated rats. The improvement in wound healing could have been due to a direct effect of GH on fibroblasts [32] or to a stimulation of IGF-I, which is present in wound fluid [33]. Whether enhanced collagen maturation is responsible for the increased wound strength in GRF-treated rats remains to be established by direct analysis of wound collagen. However, such a possibility is substantiated by a recent report on the effect of GH on collagen deposition and tensile strength in subcutaneously implanted PVA sponges [2]. In this study, the increased tensile

FACTOR

AND

WOUND

HEALING

301

strength in the sponges from GH-treated rats was not paralleled by a proportional increase in collagen deposition. Therefore, it is possible that GH increases wound strength by enhancing the quality of wound collagen. Our results not only add to the understanding of wound healing physiology; they also have potential clinical applications. To our knowledge, increased cutaneous wound strength without increased collagen deposition has not been reported before and GRF may be the first agent found to improve wound strength by enhancing collagen maturation. Further investigation should determine whether GRF administration has the potential to affect collagen maturation in wound tissue itself. ACKNOWLEDGMENTS The authors are grateful to Ms. Nora Sheppard and Guy Charette of the Electron Microscopy Laboratory, Shriner’s Hospital for Crippled Children, Montreal, for the electron microscopy studies, and to Michel Van Der Rest for his advice and helpful participation in our discussions. This work was supported by a grant from Hotel-Dieu Hospital Foundation and the Medical Research Council of Canada.

REFERENCES 1.

2.

3.

Hollander, D. M., Devereux, D. F., Marafino, B. J., and Hope, H. Increased wound breaking strength in rats following treatment with synthetic human growth hormone. Surg. Forum 2: 612, 1988. Jorgensen, P. H., and Andreassen, T. T. The influence of biosynthetic human growth hormone on biochemical properties and collagen formation in granulation tissue. Horn. Metub. Res. 20: 490,1988. Clark, R. G., Jansson, J. O., Issakson, O., and Robinson, I. C. A. F. Intravenous growth hormone: Growth responses to patterned infusions in hypophysectomised rats. J. Endocrinol.

104:53,1985. 4. Maiter, D., Underwood,

5.

L., Maes, E. M., Davenport, M. L., and Ketelslegers, M. Different effects of intermittent and continuous growth hormone (GH) administration on serum somatomedin C/Insulin-like growth factor I and liver GH receptors in hypophysectomised rats. Endocrinology 123: 1053, 1988. Wehrenberg, W. B., Ling, N., Bohlen, P., Esch, F., Brazeau, P., and Guillemin, R. Physiological roles of somatocrinin and somatostatin in the regulation of growth hormone secretion. Biothem. Biophys. Res. Commun. 109: 562,1982.

6. Tannenbaum,

G. S., Painson, J. S., Gurd, W., Reeves, I., and Brazeau, P. Growth hormone secretory dynamics in response to a sustained-release preparation of a growth hormone-releasing factor analog: Evidence for an ultradian rhythm of somatostatin secretion. In Proceedings, 72nd Annual Meeting of the Endocrine Society, Atlanta, 1990. [Abstract 1061. I. Gelato, M. C., and Merriam, G. R. Growth hormone-releasing hormone. Annu. Rev. Physiol. 48: 569, 1986. 8. Frohman, L. A., Downs, T. R., Wilhams, T. C., Heimer, E. P., and Felix, A. M. Rapid enzymatic degradation of growth hormone-releasing hormone by plasma in vitro and in vivo to a biologically inactive product cleaved at the NH2 terminus. J. Clin. Invest. 78: 906, 1986. 9. Mowles, T. F., Stricker, P., Eisenbeis, H., Heimer, E., Felix, A. M., and Brazeau, P. Effects of novel GRF analogs on GH secretion in vitro and in vivo. International Symposium on GRF, GH and somatomedins. Endocrinol. Jpn. 34 (Suppl.): 148, 1987.

302 10.

11.

12.

13. 14.

15.

16. 17.

18. 19.

20.

21.

JOURNAL

OF SURGICAL

RESEARCH:

VOL.

Chang, J. L. C., Christofides, N. D., Kraezlon, M. E., Keshavarzian, A., Burrin, J. M., Woolf, I. L., Hodgson, H. J. F., and Bloom, S. R. Growth hormone secretion dynamics in a patient with ectopic growth hormone releasing factor production. Am. J. Med. 79: 135,1985. Vance, M. L., Kaiser, D. L., Evans, W. S., Furlanetto, R., Vale, W., Rivier, J., and Thorner, M. 0. Pulsatile growth hormone secretion in normal man during continuous infusion of human growth hormone-releasing factor (l-40). J. Clin. Znuest. 75: 1584,1985. Sugihara, H. I., Wakabayashi, Minami, S., Takahashi, F., Shibasaki, T., and Ling, N. Effects of a-methyl-p-tyrosine and an antiserum to rat growth hormone (GH) releasing factor (GRF) on plasma GH secretory profile during continuous infusion of human GRF in rats. Brain Res. 475: 128, 1988. Merrifield, R. B. Solid phase peptide synthesis. 1. The synthesis of a tetrapeptide. J. Am. Chem. Sac. 85: 2149, 1963. Gaudreau, P., Baillet, M., Lepine-Frenette, C., Gorys, V., and Brazeau, P. Preparative high performance liquid chromatography purification of mammalian somatocrinin. In Proceedings, 5th International Symposium of HPLC of Proteins, Peptides and Polynucleotides. Toronto, Canada, 1985. [Abstract 7091. Diegelman, R. F., Lindblad, W. J., and Cohen, I. K. A subcutaneous implant for wound healing studies in humans. J. Surg. Res. 40: 229,1986. Lee, K. S., and Dresher, D. G. Fluorometric amino acid analysis with ophthaldialdehyde (OPA). Znt. J. Biochem. 9: 457, 1978. Sinha, Y. N., Selby, F. W., Lewis, U. J., and Vanderlaan, W. P. Studies of GH secretion in mice by a homologous radioimmunoassay for mouse GH. Endocrinology 91: 784, 1972.

22.

Ross, R., and Bendit, E. P. Wound healing and collagen formation. J. Biophys. Biochem. Cytol. 11: 671, 1961. Madden, J. W., and Peacock, E. E. Studies on the biology of collagen during wound healing. Rate of collagen synthesis and deposition in cutaneous wound of the rat. Surgery 64: 288,1968. Goodson, W. H., III, and Hunt, T. K. J. Development of a new miniature method for the study of wound healing in human subjects. J. Surg. Res. 33: 394, 1982. Harris, D. R. Healing of the surgical wound-I-Basic considerations. J. Am. Acad. Dermatol. 1: 197, 1979.

31.

23.

24.

25.

26.

27.

28.

29. 30.

32.

33.

51, NO. 4, OCTOBER

1991

Herrman, J. B., and Woodward, S. C. An experimental study of wound healing accelerators. Am. Surg. 1: 26, 1972. Mustoe, T. A., Pierce, G. F., Thomason, A., Gramates, P., Sporn, M. B., and Deuel, T. F. Accelerated healing in incisional wounds in rats induced by transforming growth factor beta. Science 2 19: 1333,1987. McGee, G. S., Davidson, J. M., Buckley, Sommer, A., Woodward, S. C., Aquino, A., Barbour, R., and Demetrion, A. A. Recombinant basic fibroblast growth factor accelerates wound healing. J. Surg. Res. 45: 145, 1988. Brown, G. L., Curtsinger, L. J., Brigthwell, D., Ackerman, D. M., Tobin, G. R., Polk, H. C., Nascimento, C. G., Venezuela, P., and Shultz, G. Enhancement of epidermal regeneration by biosynthetic epidermal growth factor. J. Exp. Med. 163: 1319, 1986. Laato, M., Niinikoski, J., Gerdin, B., and Lebel, L. Stimulation of wound healing by epidermal growth factor. Ann. Surg. 203: 379,1985. Sporn, M. B., Roberts, A. B., Shull, H. H., Smith, J. M., Ward, J. M., and Sodey, J. Polypeptide transforming growth factors isolated from bovine sources and used for wound healing in vivo. Science 219: 1329,1983. Sprugel, K. H., McPherson, J. M., Clowes, A. W., and Ross, R. In A. Barbul, E. Pines, M. Caldwell, and T. K. Hunt (Eds.), Growth Factors and Other Aspects of Wound Healing. New York: A. R. Liss, 1988. p. 77. Van Der Rest, M. In R. L. Cruess (Ed.), The MuscuZaskeZetal System. New York: Churchill Livingston, 1979. p. 65. Trelstad, R. L., and Birk, D. E. In R. L. Trelstad (Ed.), The role of the Extracellular Matrix in Development. New York: A. R. Liss, 1984. p. 513. Gilman, T., and Penn, J. Studies of the repair of cutaneous wounds. Med. Proc. 2: (Suppl.) 121, 1956. Cook, J. J., Haynes, K. M., and Werther, G. A. Mitogenic effects of growth hormone in cultured human fibroblasts: Evidence for action via local, insulin-like growth factor I production. J. Clin. Znuest. 81: 206, 1988. Spencer, E. M., Skover, G., and Hunt, T. K. In A. Barbul, E. Pines, M. Caldwell, and T. K. Hunt (Eds.), Growth Factors and Other Aspects of Wound Healing. New York: A. R. Liss, 1988. p. 103.