The FASEB Journal express article 10.1096/fj.00-0490fje. Published online October 6, 2000.
The angiogenesis inhibitor endostatin impairs blood vessel maturation during wound healing Wilhelm Bloch,* Katharina Huggel,† Takako Sasaki,‡ Richard Grose,† Philippe Bugnon,† Klaus Addicks,* Rupert Timpl,‡ and Sabine Werner† *Institute of Anatomy, University of Cologne, Köln, Germany; †Institute of Cell Biology, Swiss Federal Institute of Technology, Zürich, Switzerland; and ‡Max-Planck-Institute of Biochemistry, Martinsried, Germany Corresponding author: Dr. Sabine Werner, Institute of Cell Biology, Swiss Federal Institute of Technology, ETH Hönggerberg, CH-8093 Zürich, Switzerland. E-mail:
[email protected] ABSTRACT Endostatin is a cleavage product of collagen XVIII that strongly inhibits tumor angiogenesis. To determine if endostatin affects other angiogenic processes, we generated full-thickness excisional wounds on the back of mice that were systemically treated with recombinant murine endostatin. No macroscopic abnormalities of the wound healing process were observed. Histological analysis revealed normal wound contraction and re-epithelialization, but a slight reduction in granulation tissue formation and reduced matrix deposition at the wound edge. The blood vessel density in the wounds of endostatin-treated mice was not affected. However, ultrastructural analysis demonstrated severe abnormalities in blood vessel maturation. The wound vessels in the endostatin-treated mice were narrowed or closed with an irregular luminal surface, resulting in a severe reduction in the number of functional vessels and extravasation of erythrocytes. Endostatin treatment did not affect the expression level and localization of collagen XVIII mRNA and protein. Furthermore, the angiogenesis regulators vascular endothelial growth factor, angiopoietin-1, and angiopoietin-2 were normally expressed in the wounds of endostatin-treated mice. However, expression of the major wound matrix proteins fibronectin and collagens I and III was significantly reduced. This reduction is likely to explain the reduced density of the wound matrix. Our results demonstrate that endostatin treatment reduces the number of functional blood vessels and the matrix density in the granulation tissue, but does not significantly affect the overall wound healing process. Key words: angiogenesis • angiopoietin • collagen XVIII • skin • VEGF
I
njury to adult tissues initiates a series of events including inflammation, new tissue formation, and tissue remodelling (1, 2). The formation of new tissue is initiated by migration and proliferation of the keratinocytes at the wound edge within 1 day after injury. New stroma, often called granulation tissue, begins to form after a delay of 2 or 3 days. This tissue consists of inflammatory cells, fibroblasts, loose connective tissue, and numerous
capillaries that endow the neostroma with its granular appearance. The new capillaries are generated by sprouting from pre-existing vessels in a process called angiogenesis. The latter involves migration and proliferation of endothelial cells, lumen formation, and stabilization of the new vessels by recruitment of associated supporting cells (3, 4). Wound angiogenesis is essential to support the regenerating tissue with oxygen and nutrition (1), but the extent of angiogenesis required for normal repair has not been examined. Angiogenesis does not only occur during wound healing but also during embryonic development, during the female reproductive cycle, and after cardiac ischemia (3–5). Under these conditions, the formation of new blood vessels is beneficial. However, undesirable angiogenesis occurs in various diseases such as diabetic retinopathy. Most importantly, it is a prerequiste for the growth of solid tumors (6). Consequently, the development of drugs that inhibit tumor angiogenesis is one of the most important goals of modern cancer research (4, 5). One of the most promising molecules for the inhibition of tumor angiogenesis is endostatin. The latter is a 180-amino acid polypeptide that is part of the larger carboxyl-terminal domain NC1 of collagen XVIII. It was originally isolated from conditioned medium of hemangioendothelioma cells as an inhibitor of endothelial cell proliferation (7, 8). It is important to note that systemic administration of purified endostatin strongly inhibited angiogenesis and growth of Lewis lung carcinoma metastases in mice (7). The antitumor effect was subsequently confirmed in other tumor models using either purified endostatin or endostatin gene therapy (9–14); however, it has remained unclear whether wound angiogenesis is also affected by endostatin treatment. We have recently prepared recombinant endostatin in highly soluble form and determined its crystal structure (15, 16). This material was shown to inhibit cytokine-induced angiogenesis and to affect various parameters of endothelial cell growth and migration (12, 17, 18). The data also showed that proteolytically released endostatin is a distinct component of the extracellular matrix of various normal tissues (15, 19). In this study, we analyzed the effect of our recombinant murine endostatin on the healing process of full-thickness excisional wounds in mice. We demonstrate that endostatin treatment does not significantly change the number of newly formed blood vessels in the wound but results in narrowed or closed blood vessels with an irregular luminal surface. In spite of this abnormal vessel architecture, the overall wound healing process was only modestly affected. MATERIALS AND METHODS Animals Balb/c mice and B2D2F2 mice were obtained from the animal care facility of the Max-PlanckInstitute of Biochemistry, or from RCC (Füllinsdorf, Switzerland). They were housed and fed according to federal guidelines, and all procedures were approved by the local authorities. Endostatin treatment of mice
Female Balb/c mice or B2D2F2 mice (12 wks of age) were injected with 0.3 mg/kg s.c. murine endostatin in 100 µl 0.9% saline/0.02M ammonium acetate pH 6.8. Control animals were injected with solvent alone. Injection was performed daily at 11:00 a.m. for 2 days before wounding, immediately after injury, and for 5–7 consecutive days. Mice were killed at days 5 and 7 after wounding. In one experiment, endostatin was injected daily for 7 days after wounding, and the mice were then killed 7 days after the last endostatin injection (14 days after injury). Endostatin and immunological assays The purification of recombinant mouse endostatin from transfected human kidney cells and a radioimmunoinhibition assay for its quantitation have been described (15). The extraction of tissues at 4°C with neutral buffer pH 7.4 containing 10 mM EDTA followed by the same buffer containing detergents was performed as recently described (19). Wounding and preparation of wound tissue Mice were anesthetized with a single intraperitoneal injection of ketamine/xylazine. The hair on the animals’ back was cut and the skin was wiped with 70% ethanol. Four full-thickness excisional wounds (6 mm diameter, 3–4 mm apart, two wounds on each side of the spinal cord) were generated on the back of each animal by excising skin and panniculus carnosus as described by Munz et al. (20). The wounds were allowed to dry to form a scab. Animals were killed at varioius time points after injury. For RNA isolation or preparation of tissue lysate, the wounds on the left side including 2 mm of the epithelial margins were isolated and immediately frozen in liquid nitrogen. The back skin that was excised upon generation of the wound, as well as belly skin that was removed after the mice were killed, were used as controls. For histology, electron microscopy, and immunohistochemistry, the complete wounds on the right side were isolated, bisected, fixed, and embedded as described below. Tissue fixation, embedding, histology, and electron microscopy For routine histology, wounds were fixed overnight in 4% paraformaldehyde in PBS and subsequently embedded in paraffin. Six-micrometer sections were stained with hematoxylin/eosin or Masson trichrome stain. For immunohistochemistry, wounds were fixed overnight in 95% ethanol/1% acetic acid and subsequently embedded in paraffin. For the preparation of thin and ultrathin sections, wounds were fixed overnight in 2% paraformaldehyde/2% glutaraldehyde in 0.1M cacodylate buffer pH 7.2 and subsequently rinsed in cacodylate buffer (0.1M) three times, followed by 1% uranyl acetate. The sections were then incubated in 70% ethanol for 8 h. Thereafter, dehydration was performed in a series of graded ethanol, and specimens were then embedded in araldite. Thin sections of plastic-embedded tissue were cut with a glass knife on a Reichert ultramicrotome (Reichert, Bensheim, Germany) and stained with methylene blue. Ultrathin sections (30–60 nm) for electron microscopic observation were processed on the same microtome with a diamond knife and placed on copper grids. The transmission electron microscopy was performed with a Zeiss 902A electron microscope (Zeiss, Oberkochem, Germany).
Immunofluorescence and immunohistochemistry For immunofluorescence ethanol/acetic acid-fixed paraffin sections were dewaxed and washed in PBS. The FITC-conjugated PECAM antibody (Pharmingen, San Diego, Calif.) was diluted 1:250 in PBS containing 12% BSA and 0.02% NaN3 and the slides were incubated for 1 h at room temperature with this antibody. After three 10-min washes with PBS, sections were washed once with 10 mM Tris/HCl (pH 8.8) and coverslipped with mounting medium. Immunohistochemical staining of ethanol/acidic acid fixed paraffin sections with a rabbit polyclonal collagen I antiserum (1:300 diluted; Chemicon, Temecula, Calif.) was performed with the avidin-biotin-peroxidase complex system (Vector, Burlingame, Calif.) as described by the manufacturer. Antibody-binding cells were visualized using 3-amino-9-ethylcarbazole (AEC) as a substrate for the peroxidase. After development the slides were rinsed with water, counterstained with hematoxylin, and mounted. Collagen XVIII immunohistochemistry was performed on 4% paraformaldehyde-fixed paraffin sections. After deparaffination, the sections were placed in a solution of 3% hydrogen peroxide and 60% methanol in PBS for 60 min to block endogenous peroxidase activity. They were subsequently permeabilized with 0.25% Triton X-100 in 0.1M PBS. Incubation with a 1:500 dilution (5 µg/ml) of an affinity-purified rabbit antibody against endostatin (15) in PBS/0.8% bovine serum albumin was performed overnight at 4°C. After rinsing with PBS, sections were incubated for 1 h in a 1:400 dilution of a biotinylated anti-rabbit antibody (Dako, Glostrup, Denmark). A streptavidin-horseradish-peroxidase conjugate (Amersham, Buckinhamshire, England) (1:100 diluted) was used as a detection system. After a 60-min incubation with this reagent, sections were stained for 30 min with 3,3`-diaminobenzidine-tetrahydrochloride in 0.05M Tris/HCl/0.1% hydrogen peroxide. Staining without the primary antibody was used as a negative control in all experiments. RNA isolation and RNase protection assay RNA isolation was carried out as described by Chomczynski and Sacchi (21). RNase protection assays were performed as described by Werner et al. (22). As a loading control, 1 µg of all RNA samples was loaded on a 1% agarose gel before hybridization and stained with ethidium bromide. Alternatively, a radiolabelled GAPDH riboprobe was included in the hybridization mix. All protection assays were carried out at least in duplicate. As templates, we used a 200-bp fragment corresponding to nt 1248–1447 of the murine angiopoietin-1 cDNA (23); a 200-bp fragment corresponding to nt 1146–1345 of the murine angiopoietin-2 cDNA (24); a 316-bp fragment corresponding to nt 139–455 of the murine VEGF-A120 cDNA (25); a 150-bp murine collagen α1(I) chain cDNA (kindly provided by Dr. Reinhard Fässler); a 220-bp fragment corresponding nt 15394–15613 of the murine collagen α1(III) chain cDNA (accession number X52046); a 162-bp fragment from the 3’-end of the mouse fibronectin cDNA (kindly provided by Dr. P. Ekblom); a 176-bp murine IL-1β cDNA fragment (26); a 307-bp murine TNF-α cDNA fragment (26); a 392-bp fragment corresponding to nt 3476–3867 of the murine collagen XVIII cDNA (27); and a 120-bp fragment of the murine GAPDH cDNA (kindly provided by Dr. J. Regenbogen).
In situ hybridization In situ hybridization was performed with a 550-nt digoxigenin-dUTP labelled riboprobe corresponding to the carboxyl-terminal part of collagen XVIII. Labelling was performed with the digoxigenin DNA labelling kit (Roche, Mannheim, Germany) as described by the manufacturer. In situ hybridization was carried out as described by Wevers et al. (28). The hybridized probe was detected with an alkaline phosphatase-conjugated digoxigenin antibody (Roche; 1:100 diluted). Development was performed with 5-bromo-4-chloro-3-indolylphosphate (BCIP) and nitroblue tetrazolium chloride (NBT) (Roche). RESULTS To determine the effect of endostatin on wound healing, recombinant mouse endostatin was purified to homogeneity as described (15). Three different preparations were used that were all shown to inhibit angiogenesis or tumor growth in animal models (17) (T. Sasaki and R. Timpl, unpublished data). Five independent wound-healing experiments were performed, each with five control animals and five endostatin-treated animals. The wounded tissue was analyzed at day 5 (three experiments), day 7 (1 experiment), and day 14 (1 experiment) after wounding. A dose of 0.3mg/kg/day was injected subcutaneously in the belly. This dose was shown to suppress the growth of subcutaneous tumors in mice (Sorensen et al., personal communication). Treatment was started 2 days before wounding and continued during the wound-healing process. The wounds were harvested 24 h after the last endostatin injection. At this time point, no elevated levels of endostatin were observed in the serum and in the wound tissue of the treated mice as determined by radioimmunoinhibition assay (Table 1), demonstrating a rapid turnover of the injected protein. No obvious abnormalities in wound closure or wound appearance were observed in the endostatin-treated mice. The lack of severe wound-healing abnormalities was confirmed by histological staining of sections from the middle of the wound. As shown in Figure 1, reepithelialization, contraction, and overall wound closure appeared normal after endostatin treatment. Furthermore, no obvious differences in the inflammatory response were observed. The latter finding was confirmed by the identical expression of the proinflammatory cytokines interleukin-1β, and tumor necrosis factor-α in buffer- and endostatin-injected animals (data not shown). However, in the endostatin-treated mice, the extent of granulation tissue was reduced and the connective tissue appeared less dense. Furthermore, Masson trichrome staining revealed reduced collagen deposition at the wound edge. These features were particularly obvious at day 7 after wounding (Fig. 1). Most importantly, significant hemorrhage was observed at day 5 (Fig. 2B, D, E) and day 7 (not shown) in the wound tissue of the endostatin-treated mice as demonstrated by the large numbers of extravasated erythrocytes. By contrast, erythrocytes were almost exclusively found within blood vessels in the control animals (Fig. 2A, C). These endostatin-dependent abnormalities were also observed in another mouse strain (B2D2F2) (data not shown). To determine the reason for the hemorrhage, we first prepared semi-thin sections of the wounds and stained them with methylene blue. As shown in Figure 3, no significant difference in vessel density was observed between endostatin-treated mice and control mice, indicating that
endostatin does not affect the extent of blood vessel formation in the wound (Fig. 3A, B). However, significant differences in the quality of the vessels within the wound were observed. In the buffer-injected animals, the vessels were round, open, and filled with blood cells (Fig. 3A). By contrast, a large percentage of the vessels in the endostatin-treated mice were closed and irregular in shape (Fig. 3B). This phenomenon was further confirmed by electron microscopy (Fig. 4). In the controls, a normal lumen was seen, vessels were filled with blood cells (Fig. 4A), and even the sprouting vessels contained erythrocytes (Fig. 4B). In the endostatin-injected mice, however, the vessels were narrowed (Fig. 4C) or closed (Fig. 4D), and they had an irregular luminal surface. The endothelial cells of such vessels were enlarged, and they revealed a large number of vacuoles and short cytoplasmic protrusions (Fig. 4E). Furthermore, the contrast of the endothelial cells appeared different, indicating severe cell damage. In contrast to the abnormalities seen in the newly formed wound vessels, the pre-existing vessels in the dermis adjacent to the wound appeared normal (Fig. 4F). To determine the effect of endostatin treatment on the quality of the healed wound, we performed an additional wound-healing experiment where the endostatin treatment was stopped at day 7 after injury. One week later the mice were killed and the wounds were analyzed histologically. As shown by Masson trichrome staining (Fig. 5, top panel), both endostatintreated and control wounds were fully healed at this time point. Consistent with the reduced density of the wound matrix seen at day 5 and 7 after injury (see above), the extent of the remaining scar was slightly reduced in the endostatin-treated mice, and the deposited collagen appeared less dense (Fig. 5, top and middle panel), demonstrating that endostatin treatment even improves the quality of the healed wound. Similarly, as in early wounds, no significant differences in vessel density were observed between both treatment groups as demonstrated by immunostaining with the endothelial cell marker PECAM (Fig. 5, lower panel). It is interesting that most vessels appeared normal at this time point as determined by histological analysis of ultra-thin sections (Fig. 3D) and electron microscopy (data not shown), but a few vessels that revealed the characteristic abnormalities (see above) remained (data not shown). As a next step we determined whether endostatin treatment affects expression and/or localization of the endogenous collagen XVIII. Because expression of this gene has as yet not been analyzed in wounded skin, we first performed RNase protection assays with RNAs from normal skin and from wounds at varioius stages after injury of non-treated Balb/c mice. As shown in Figure 6, the mRNA expression levels of collagen XVIII did not significantly change after skin injury. A slight reduction in the expression of this gene was observed at late stages (14d) after wounding. In situ hybridization revealed the presence of collagen XVIII mRNA in epidermal keratinocytes and hair follicle keratinocytes of nonwounded skin but not in blood vessels (Fig. 7A). In the wounded tissue, expression of collagen XVIII was detected throughout the hyperproliferative epithelium and in endothelial cells of microvessels in control mice (Fig. 7B, C) and endostatintreated mice (Fig. 7D and data not shown). Neither expression levels nor distribution of collagen XVIII transcripts were affected by the endostatin treatment (Figs. 6B and 7, and data not shown). The expression of collagen XVIII mRNA correlated with the deposition of immunoreactive protein in the epidermis of normal and wounded skin as well as around vessels and in the extracellular matrix within the granulation tissue (Fig. 8). Again, no differences between control and endostatin-treated mice could be detected. Furthermore, a radioimmunoassay of wound extracts (Table 1) did not reveal any differences in the amounts of extractable collagen XVIII-
derived polypeptides between control and endostatin-treated mice. Yet the total amount of extractable endostatin from wound tissue (40–50 µg/g wet weight) was about fourfold higher compared with normal skin (19), indicating increased collagen XVIII production and/or endostatin release during wound repair. To gain insight into the mechanisms that might underlie the phenotype observed in the endostatin-treated wounds, we analyzed the expression of VEGF, angiopoietin-1, and angiopoietin-2. These factors have been shown to be major players involved in the angiogenic response (29). As previously demonstrated in our laboratory (30), expression of VEGF increased in wounded skin during the period when angiogenesis occurs (days 2–7). This time course of expression was not affected by endostatin treatment (data not shown). Furthermore, no differences in the expression of the corresponding protein were detected by immunostaining with a VEGF-specific antiserum (data not shown). To determine expression of the angiopoietins, we first cloned cDNA probes corresponding to murine angiopoietin-1 and -2, and we analyzed the time course of expression of these factors during the normal healing process. As demonstrated in Figure 9A, a slight and transient decline in angiopoietin-1 expression was observed after wounding, and a second decline followed at day 14 after injury. Angiopoietin-2 expression increased after skin injury with maximal levels being found between days 3 and 7 after injury. At day 14 after wounding, angiopoietin-2 mRNA levels had declined to basal levels (Fig. 9A). This time course of angiopoietin-2 expression correlated well with the one observed for VEGF, demonstrating that both factors are up-regulated during the period when wound angiogenesis occurs. No differences in angiopoietin-1 and -2 expression were observed in the endostatintreated mice during the period when angiogenesis occurs (Fig. 9B), indicating that neither a difference in VEGF nor in angiopoietin expression is responsible for the blood vessel abnormalities seen in the granulation tissue of endostatin-treated animals. However, angiopoietin-2 expression was reduced in 14d wounds of endostatin-treated mice compared with control mice (Fig. 9B). To determine the reason for the less dense appearance of the connective tissue in the wounds of the endostatin-treated mice, we analyzed the expression of major extracellular matrix molecules by RNase protection assay. As shown in Figure 10, the mRNA levels of collagen α1 (I) chain and fibronectin were significantly reduced in the wounds of the endostatin-treated animals at all time points. Furthermore, reduced expression of collagen III was observed at day 14 after injury. These results demonstrate that endostatin treatment leads to suppression of the expression of major wound matrix proteins. This finding is likely to provide an explanation for the less dense connective tissue seen in the wounds of the endostatin-treated mice. DISCUSSION Inhibition of angiogenesis is one of the most promising novel approaches for the treatment of cancer. Thus, several antiangiogenic substances have been shown to inhibit the growth of primary tumors and also of metastases in animals models, and some of these drugs are already being tested in patients. In contrast to conventional chemotherapy, development of resistance has as yet not been observed with antiangiogenic therapy (8), indicating that long-term treatment with these drugs should be effective. In addition to this advantage, the almost complete lack of angiogenesis in the adult organism should minimize the side effects of antiangiogenic therapy.
An important exception, however, is the wound-healing process, which strongly depends on the growth of new blood vessels in the injured tissue. Because tumor patients often undergo extensive surgery, inhibition of the wound-healing response might create a serious problem in patients treated with antiangiogenic molecules. We therefore determined the effect of endostatin on the healing process of full-thickness excisional wounds in mice. Endostatin was chosen because it is one of the most promising antiangiogenic molecules currently being tested in clinical trials (31). Furthermore, the mechanisms of endostatin action have only partially been elucidated, so the detailed analysis of the wounds in endostatin-treated mice might provide further insights into the effects of endostatin at the cellular level. It is interesting that we did not observe any macroscopic differences in wound closure or appearance between control mice and endostatin-treated animals. Furthermore, histological analyses did not reveal significant differences in re-epithelialization and contraction, demonstrating that wounds can heal under endostatin treatment. These results are consistent with a previous report where the effect of human endostatin on the healing process in athymic nude mice was analyzed. These wounds revealed a similar tensile strength compared with control wounds, although a histological analysis was not performed (32). In this study, however, a 100fold higher concentration of endostatin was used. Therefore, these results are not directly comparable with our data. The most interesting finding of our study was the severe blood vessel abnormalities observed in the endostatin-treated animals. A large number of extravasated erythrocytes were observed, indicating impaired blood vessel integrity. This hypothesis was confirmed by ultrastructural analysis. A large proportion of the newly formed vessels in the granulation tissue were narrowed or closed, thereby inhibiting the normal blood flow. Thus, a significant reduction in the number of functional vessels was observed in the wounds of endostatin-treated mice. The endothelial cells of these vessels were enlarged and vacuolized. Their shape was strikingly changed, and multiple small protrusions were observed. These morphological changes are a typical sign of cell damage that might eventually lead to cell death. This finding is consistent with the effect of endostatin on endothelial cell apoptosis in vitro (18, 33). Interestingly, the observed blood vessel abnormalities were at least partially reversible, because only few abnormal vessels were observed within 7 days after endostatin withdrawal. In contrast to the abnormal blood vessel morphology, no obvious reduction in the density of newly formed vessels was observed in the endostatin-treated mice. This demonstrates that endothelial cell migration, proliferation, and angiogenesis can occur in the wounds of endostatintreated mice. Most importantly, the strong mitogenic activity of wound endothelial cells seems to compensate for the endothelial cell loss caused by endostatin as demonstrated by the normal vessel density. By contrast, the balance between endothelial cell proliferation and endostatininduced cell death might be different in tumors, possibly because of their lower angiogenic activity. This hypothesis is supported by results of a recent study, demonstrating that even in highly angiogenic tumors the angiogenesis is 4–20 times less intense as compared with the physiologcal angiogenesis seen in the growing ovarian corpus rubrum (34). Similar differences might exist between wound angiogenesis and tumor angiogenesis. Thus, the apoptotic effect of
endostatin on endothelial cells might be sufficient to inhibit tumor angiogenesis but not the much stronger angiogenesis occurring under physiological conditions. To further prove this hypothesis it will be important to determine the effect of endostatin and other angiogenesis inhibitors on ovarian angiogenesis. To understand the mechanisms of endostatin action during wound healing, we analyzed its effect on endogenous endostatin. Because no data have as yet been available regarding the expression and processing of collagen XVIII in skin wounds, we first determined the expression of this type of collagen in normal and wounded skin. Using a cDNA and an antibody specific for the endostatin part of collagen XVIII, we demonstrated expression of both the mRNA and the protein in keratinocytes in normal skin and in wounds. By contrast, only the newly formed blood vessels in the wound bed, but not the pre-existing vessels in the adjacent dermis, expressed this gene, indicating a specific role of endogenous collagen XVIII and/or its cleavage products in wound angiogenesis. However, no differences in collagen XVIII expression were observed between control and endostatin-treated mice, suggesting that exogenous endostatin does not affect expression of the endogenous gene. A series of previous studies have documented the important role of VEGF and of angiopoietins in angiogenesis. Whereas VEGF is required for various processes involved in angiogenesis including endothelial cell proliferation (35), angiopoietin-1 has been identified as an important factor for the stabilization of blood vessels (36). It is thought to play an essential role in mediating interactions between endothelial cells and surrounding support cells (29, 37). This stabilizing effect of angiopoietin-1 is antagonized by angiopoietin-2, a naturally occurring antagonist that binds to the same receptor (24). Consistent with the destabilizing function of angiopoietin-2, it is expressed at sites of vascular remodelling in the adult, such as in the female reproductive tract (24) and in certain areas of tumors where angiogenesis occurs (38). Interestingly, expression of angiopoietin-2 together with VEGF seems to induce the angiogenic response, whereas the presence of angiopoietin-2 in the absence of VEGF leads to vessel regression (39). The results described in our study provide the first indication for a similar role of angiopoietin-2 and VEGF in wound angiogenesis, because expression of both factors was induced with a similar kinetics after skin injury. However, this expression pattern was not modified by endostatin during the period when wound angiogenesis occurs, indicating that abnormalities in VEGF or angiopoietin expression are not involved in the phenotypic abnormalities seen in the endostatin-treated mouse wounds. However, it seems possible that endostatin inhibits the biological activity of at least some of these factors in the wound, because it has been shown to inhibit VEGF-induced endothelial cell migration in vitro (12) and basic fibroblast growth factor-induced angiogenesis in the chorioallantoic membrane assay (17). Some of these latter activities are lost by mutation of the heparin-binding site of collagen XVIIIderived endostatin, which is associated with a restricted basic surface area (16–18). These and other endostatin mutants could now become useful reagents to examine the mechanism of the distinct, although reversible, endostatin effect on vessel stability. In spite of the normal wound closure seen in the endostatin-treated mice, we reproducibly observed a reduced connective tissue density in early and late wounds of these animals, indicating that endostatin treatment even improves the quality of the healed wound. This effect on the matrix is at least partially regulated at the mRNA level, because we found a significantly
reduced mRNA expression of the major wound matrix molecules collagen types I and III and fibronectin. Although we cannot exclude the possibility that endostatin directly affects the expression of these genes, an indirect effect based on the reduced number of functional blood vessels seems more likely. Taken together, we demonstrated striking and unexpected effects of endostatin on blood vessel maturation during wound healing. Our results suggest that the large number of newly formed functional vessels in the wound is not required for healing under normal circumstances. Thus, inhibition of angiogenesis to a certain extent seems to be tolerable for the wound-healing process, an observation that is likely to be important when use of endostatin is considered in tumor patients who undergo surgery. ACKNOWLEDGMENTS We thank Dr. Ingrid Renner-Müller and Petra Renner for help with the endostatin injections, and Christiane Born and Andreas Stanzel for excellent technical assistance. This work was partially supported by grants from the German Ministry for Research and Development (S. W.), and by EC grant BI04-CT96-0537 (R. T.). REFERENCES 1. Clark, R. A. F. (1996) Wound repair: overview and general considerations. In The Molecular Biology of Wound Repair (Clark, R. A. F., ed). pp. 195–248, Plenum, New York 2. Martin, P. (1997) Wound healing: aiming for perfect skin regeneration. Science 276, 75–81 3. Risau, W. (1997) Mechanisms of angiogenesis. Nature 386, 671–674 4. Augustin, H. G. (1998) Antiangiogenic tumour therapy: will it work? Trends Pharmacol. Sci. 19, 216–222 5. Ferrara, N., and Alitalo, K. (1999) Clinical applications of angiogenic growth factors and their inhibitors. Nat. Med. 5, 1359–1364 6. Folkman, J. (1995) Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat. Med. 1, 27–31 7. O`Reilly, M. S., Boehm, T., Shing, Y., Fukai, N., Vasios, G., Lane, W. S., Flynn, E., Birkhead, J. R., Olsen, B. R., and Folkman, J. (1997) Endostatin: an endogenous inhibitor of angiogenesis and tumor growth. Cell 88, 277–285 8. Boehm, T., Folkman, J., Browder, T., and O`Reilly, M. S. (1997) Antiangiogenic therapy of experimental cancer does not induce acquired drug resistance. Nature 27, 404–407
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Table 1
Fig. 1
Figure 1. Reduced granulation tissue formation in the endostatin-treated mice. Full-thickness excisional wounds were made on the back of female control mice (Ctr; upper panel) and endostatin-treated mice (ES; lower panel). Mice were killed at day 7 after injury. Masson trichrome stains of paraformaldehyde-fixed paraffin sections (6 µm) from the middle of the wounds are shown. G: Granulation tissue, HE: hyperproliferative epithelium, Es: Eschar. Magnification 25x.
Fig. 2
Figure 2. Hemorrhage in the granulation tissue of endostatin-treated mice. Full-thickness excisional wounds were made on the back of female control mice (Ctr) and endostatin -treated mice (ES). Mice were killed at day 5 after injury. Paraformaldehyde-fixed paraffin sections (6 µm) from the middle of the wounds were stained with hematoxylin/eosin. (A) and (C): Wounds of control mice; (B), (D), and (E): wounds of endostatin-treated mice. G: Granulation tissue, HE: hyperproliferative epithelium, E: Eschar. Arrows indicate normal vessels in the granulation tissue of control mice and extravasated erythrocytes in the granulation tissue of endostatin-treated mice. Magnification 200x (A, B), and 400x (C, D, E).
Fig. 3
Figure 3. Normal vessel-density in endostatin-treated mice. Methylene-blue stained, semi-thin sections from the middle of 7d (A, B) and 14d (C, D) wounds of control mice (A, C) and endostatin-treated mice (B, D) are shown. Note the similar vessel density in the wounds of control mice and endostatin-treated mice, the round, open, and blood cell-filled vessels in the control mice, and the closed and irregularly-shaped vessels in 7d wounds of the endostatin-treated animals. Vessels are indicated with arrows. Scale Bar: 25 µm.
Fig. 4
Figure 4. Blood vessel abnormalities in the granulation tissue of endostatin -treated mice. Electron micrographs of newly formed blood vessels in the granulation tissue of 7d wounds from control mice (A, B) and endostatin-treted mice (C, D, E), as well as vessels in the normal dermis of endostatin -treated mice (F) are shown. Note the open and erythrocyte-filled resting (A) and sprouting (B) vessels in the control mice, and the narrowed (C) or closed (D) vessels with irregular luminal su rface in the endostatin -treated mice. Endothelial cells in the granulation tissue of endostatin treated mice have a large number of vacuoles and short cytoplasmic processes (E). By contrast, endothelial cells in the normal dermis adjacent to the wound appear normal (F). Scale bar: 4.5 µm (A, B, D, F); 3 µm (C); 2 µm (E).
Fig. 5
Figure 5. Reduced connective tissue but normal vessel density in healed wounds of endostatin-treated mice. Fullthickness excisional wounds were made on the back of female control mice (Ctr) and endostatin-treated mice (ES). Mice were killed at day 14 after injury. Masson trichrome stains of sections (6 µm) from the middle of the wounds are shown in the top panel. Immunohistochemistry with a collagen I antibody is shown in the middle panel. Immunofluorescence with an FITC-conjugated antibody against PECAM is shown in the lower panel. Sc: scar tissue, E: epidermis. Magnification 25x (top and middle panel); 100x (lower panel).
Fig. 6
Figure 6. Expression of collagen XVIII mRNA in normal and wounded skin of control mice and endostatintreated mice. Balb/c mice were wounded as described in Materials and Methods, and sacrificed at different time points after injury (1d, 3d, 5d, 7d, and 14d after injury) as indicated. Skin represents normal, non-wounded back skin. (A) 20 µg total cellular RNA from normal and wounded skin were analyzed by RNase protection assay for the expression of collagen XVIII mRNA. 1 µg of each RNA sample was loaded on a 1% agarose gel and stained with ethidium bromide. A picture of the RNA gel is shown below the RNase protection assays. (B) RNA was isolated from wounded skin of control mice and endostatin-treated mice at days 5 and 14 after injury and analyzed by RNase protection assay for the presence of collagen XVIII and GAPDH mRNAs. 20 µg tRNA was used as a negative control in both experiments. 1000 cpm of the hybridization probes were loaded in the lanes labeled “probe” and used as size markers.
Fig. 7
Figure 7. In situ hybridization demonstrating expression of collagen XVIII mRNA in normal and wounded skin. Sections from non-wounded skin (A) and from 5d wounds (B, C, D) of control mice (A,B,C) and endostatin-treated mice (D) were analyzed by non-radioactive in situ hybridization for the presence of collagen XVIII mRNA. Note the strong signals in hair follicle and epidermal keratinocytes of non-wounded skin (A) and in the hyperproliferative wound epithelium (B, C). Strong expression was also observed in the mesenchyme at the wound edge (B), particularly in blood vessels (C, D, indicated by arrows). E: Epidermis, D: Dermis, PC: Panniculus Carnosus, H: Hair follicle, G: Granulation tissue, HE: Hyperproliferative epithelium. Scale bar: 40 µm (A, D); 50 µm (C); 250 µm (B).
Fig. 8
Figure 8. Immunohistochemical detection of collagen XVIII in non-wounded and wounded skin. Paraformaldehyde-fixed frozen sections from non-wounded back skin and from 5d wounds were stained with an affinitypurified antiserum directed against the endostatin fragment of collagen XVIII. Strong signals were found in the epidermis but not in the dermis of non-wounded skin (A). At the wound edge, a strong deposition of the protein was found around vessels (indicated by arrows) and in the extracellular matrix (B). E. Epidermis, D: Dermis, G: Granulation tissue. Scale bar: 30 µm.
Fig. 9
Figure 9. Expression of angiopoietin-1 and –2 mRNAs in non-wounded and wounded skin of control mice and endostatin-treated mice. Balb/c mice were wounded as described in Materials and Methods, and sacrificed at different time points after injury. (A) 20 µg total cellular RNA from normal and wounded skin were analyzed by RNase protection assay for the expression of angiopoietin- 1 and -2 mRNAs as indicated. 1 µg of each RNA sample was loaded on a 1% agarose gel and stained with ethidium bromide. A picture of the RNA gel is shown below the RNase protection assays. (B) RNA was isolated from wounded skin of control mice and endostatin-treated mice at days 5 and 14 after injury and analyzed by RNase protection assay for the presence of angiopoietin-1, angiopoietin-2, and GAPDH mRNAs. The same batch of RNAs was used for the angiopoietin-1 and –2 protection assays shown in (A) and (B), respectively. 20 µg tRNA was used as a negative control in all experiments. 1000 cpm of the hybridization probes were loaded in the lanes labeled “probe” and used as size markers.
Fig. 10
Figure 10. Expression of extracellular matrix molecules in the wounds of control mice and endostatin-treated mice. 20 µg total cellular RNA from 5d wounds and 14d wounds of control mice and endostatin-treated mice were analyzed by RNase protection assay for the presence of collagen I, collagen III, fibronectin, and GAPDH mRNAs. The same batch of RNAs was used for all protection assays. 20 µg tRNA was used as a negative control in all experiments. 1000 cpm of the hybridization probes were loaded in the lanes labeled “probe” and used as size markers.
