Leptin is an autocrine/paracrine regulator of wound healing

The FASEB Journal express article 10.1096/fj.03-0068fje. Published online August 15, 2003.

Leptin is an autocrine/paracrine regulator of wound healing Ali Murad,*,§ Anjali K. Nath,†,§ Sung-Tae Cha,* Erhan Demir,* Jaime Flores-Riveros,‡ and M. Rocío Sierra-Honigmann* *Division of Plastic Surgery, Department of Surgery, Cedars-Sinai Research Institute and University of Southern California, Los Angeles, California; †Department of Biology, Yale University School of Medicine, New Haven, Connecticut; and ‡Hollis-Eden Pharmaceuticals, San Diego, California. §Both authors contributed equally.

Corresponding author: M. Rocío Sierra-Honigmann, Cedars-Sinai Research Institute 8700 Beverly Blvd., Los Angeles, CA 90048. E-mail: [email protected]

ABSTRACT

Leptin, a 16 kDa pleiotropic cytokine primarily expressed in adipose tissue, has been shown to cause multiple systemic biological actions. Recently, leptin has also been documented as an important component of the wound healing process and its receptor appears to be expressed in wound tissue. We have previously demonstrated that leptin is a potent angiogenic factor exerting direct effects on endothelial cells and that transcription of its encoding gene is regulated by hypoxia. Here, we hypothesize that leptin expression is acutely up-regulated in the ischemic tissue of experimental wounds. Using a combination of in situ hybridization and quantitative RT-PCR experiments, we show that leptin expression is rapidly and steadily up-regulated in skin tissue from incisional and excisional wounds. By immunohistochemistry, we demonstrate increased and sustained leptin protein levels in basal keratinocytes, blood vessel walls, and fibroblasts. To determine whether leptin is required for normal healing, excisional wounds were treated with neutralizing anti-leptin antibodies. This treatment markedly hampered healing progression and prevented wound closure and contraction. Finally, a transient rise in circulating blood leptin levels was detected within the first 24 h after inflicting the injury; we present evidence suggesting that this elevation is due to increased leptin production at the ischemic wound site. We conclude that leptin is acutely up-regulated in the injured skin and propose that this local production of leptin serves a critical functional role as an autocrine/paracrine regulator of normal wound healing.

Key words: hypoxia • ischemia • tissue injury

he process of wound healing is a highly regulated dynamic cascade of cellular and biochemical events, which normally result in the successful repair of injured tissues. Following the rupture of blood vessels caused by traumatic injury to the skin, a fibrin-rich

clot forms, which achieves temporary mechanical closure of damaged tissue, including vascular structures. The dermal microvasculature plays an essential role in tissue repair, whereby severed blood vessels lead to hypoperfusion and local ischemia of the wound site and trigger neovascularization through local synthesis and secretion of angiogenic factors, including VEGFs, FGF-2, TGF-β, and PDGFs (1–3). Endothelial cells (EC) respond to these angiogenic factors by rapidly forming a complex capillary network (4). This process occurs as EC dissolve their cell–

T

cell attachments, migrate outside the vessel into the extracellular matrix, undergo mitosis, and finally re-associate to form new vessels.

Leptin is a hypoxia-inducible pleiotropic cytokine known to participate in multiple cellular and physiological processes. In mice, the genetic defects in leptin production or leptin receptor expression result in a complex phenotype with multiple metabolic and reproductive alterations that include development of morbid obesity, metabolic syndrome, and infertility. The ob/ob (leptin null), and db/db (leptin receptor null), mouse strains are characterized by severe impairment in their ability to repair cutaneous wounds and have been extensively used as models of pathological wound healing (5–7). Because these mice carry naturally occurring mutations in the ob or db loci, they exhibit distinct alterations in food intake behavior and energy metabolism (8, 9). These animals typically develop morbid obesity and insulin resistance and sometimes frank diabetes. It is commonly thought that their defective wound healing is merely secondary to underlying metabolic alterations and diabetic microvascular disease (7). However, in view of several recent reports documenting wound healing-promotion by leptin treatment in normal animals, an alternative explanation might be that such defect results not only from metabolic alterations, but also from a primary impairment in leptin-dependent actions required for normal wound healing (10, 11). In this regard, we (and others) have recently demonstrated that leptin gene transcription is rapidly up-regulated in skin dermal fibroblasts upon exposure to hypoxia (12, 13). This response occurs through a mechanism involving activation of hypoxia inducible factor-1 (HIF-1), which binds to hypoxia response elements (HRE) contained within the leptin gene promoter (14). These observations invite the hypothesis that the hypoxic environment in the wound induces acute expression of leptin, which in turn acts as an early hypoxia-regulated endogenous mediator of wound healing. One important element in the evaluation of this hypothesis is the demonstration that leptin is actively produced in the wound bed and that its functional integrity must be intact. Therefore, to test this hypothesis, we used a model of full-thickness wounds inflicted on the skin of normal adult mice. We present data demonstrating the presence and acute in situ production of leptin in these experimental wounds. Functional neutralization of leptin with an anti-leptin antibody severely impairs wound healing. Finally, we demonstrate a transient increase in circulating leptin after injury; such elevation in leptin levels appears to derive from the wound site itself. Our findings provide compelling evidence that leptin plays a fundamental role in normal wound healing.

MATERIALS AND METHODS

Animals

Adult C57BL/6J mice (Jackson Laboratory, Bar Harbor, ME) were used at 8 weeks of age. Severe combined immunodeficient (SCID) beige mouse (Taconic Farms, Germantown, NY) were 5- to 8-weeks old. The mice were singly housed in microisolated cages and fed ad libitum with autoclaved mouse chow and water. All protocols used were previously approved by appropriate regulatory committees from the Cedars Sinai Research Institute and the Yale University Animal Care and Use.

Human skin-SCID chimeras

Surgical specimens of discarded normal adult human breast skin were obtained through the Yale University Department of Pathology under a protocol approved by the ethics review board. The superficial portions of the skin were harvested in 500- to 700-µm-thick sheets by using a surgical dermatome and were cut in ~7 × 7-mm pieces, which were kept in lactated Ringers at 4°C until transplantation (no longer than 6 h). SCID mice were anesthetized by inhalation of methoxyflurane (Pitman-Moore, Mundelein, IL). The two fragments measuring 5 × 5-mm skin portions were excised from the back of each mouse and replaced by the human skin grafts, which were fixed with surgical staples (3M, St. Paul, MN). Grafts were allowed to heal for 4 to 6 weeks before each experiment was initiated.

Wounding procedure and topical treatments

Mice were anesthetized before surgery by subcutaneous administration of 0.05 ml of a ketamine/xylazine mixture (55 mg/ml and 6.25 mg/ml, respectively; Fort Dodge Animal Health, Fort Dodge, IA). Both the left and right sides on the back of each mouse were shaved and disinfected with 70% ethanol. Using a surgical skin marker (Precision Dynamics Corp., San Fernando, CA), reference lines 8-mm in length were first drawn bilaterally along the paramedial dorsal region, 1.5 cm from the spine, and full-thickness incisional wounds were then performed with fine surgical scissors. For excisional wounds, anesthetized mice were placed on a surgical table resting on one of their flanks while retracting the skin from the body wall with forceps to avoid internal injury. Two full-thickness, circular excisions were then made simultaneously by punching through using a disposable 3 mm biopsy punch (Biopunch, Fray Corp., Amherst, NY). Wounds were collected post-euthanasia by excising a tissue flap containing the wound in bloc. Then a circular area containing the wound in the center was cut from the flap using a 6 mm biopsy punch. The tissue flaps were full-thickness and included the skin, hypodermis, and muscle layers. For antibody treatments, the excisional wounds received 5 µg of neutralizing polyclonal goat anti-mouse leptin antibody (R&D Systems, Minneapolis, MN) or 5 µg goat IgG (controls) in a volume of 10 µl. Antibody treatments were repeated daily for the times indicated.

Wound tissue processing and immunohistochemistry

The wound areas and surrounding tissue were excised and bisected mid-transversally, fixed overnight in buffered formalin (Sigma, St. Louis, MO), and then embedded in paraffin. Hematoxylin and eosin (H&E) staining was performed by using 4 µm sections. After removal of paraffin and inactivation of endogenous peroxidase, sections were treated for antigen retrieval with 10 mM sodium citrate (pH 6.0) for 15 min at 95°C. Staining for leptin was performed immunohistochemically by using a polyclonal goat anti-leptin antibody (R&D Systems, Minneapolis, MN), followed by a biotin-conjugated donkey anti-goat antibody and developed by the streptavidin/HRP system by using an ABC kit (Vector, Burlingame, CA). Anti-leptin antibody was pre-adsorbed with an excess of leptin (10 µg/ml) for 1 h at room temperature and served as the negative control.

In situ hybridization

RNA sense and antisense probes for leptin were labeled with digoxigenin (DIG; Boehringer, Indianapolis, IN) by in vitro transcription from a full-length mouse leptin cDNA subcloned between the XhoI and BamHI sites of pBluescript II/KS+ (Stratagene, La Jolla, CA). Frozen sections of 10 µm were cut and fixed in 2% paraformaldehyde and then digested for 2 min at 4°C in a mixture of 100 mM sodium citrate/0.1% NP-40. The slides were hybridized with the DIG-labeled probes at 37°C for 12 h, washed, and then incubated with anti-digoxigenin-AP Fab’ fragment conjugates followed by color development with NCIP/BTP (Boheringer, Indianapolis, IN).

RNA extraction and quantitative RT-PCR

Total RNA was isolated from mouse skin samples by using two consecutive extractions and Trizol® (Invitrogen, Carsbald, CA) to ensure RNA purity. Before cDNA synthesis, the samples were digested with DNase 1 to eliminate any residual genomic DNA contamination (Ambion, Austin, TX). cDNA synthesis was performed using SuperScript™ II (Invitrogen, Carlsbad, CA). Quantitative PCR (qPCR) was performed by using a Biorad iCycler iQ real-time PCR machine and HotStarTaq™ DNA polymerase (Qiagen, Valencia CA). Taqman® probe-based chemistry with its dual-labeled fluorogenic probe (prb), labeled with 5′-FAM/3′-BHQ (Biosearch Technologies, Novato, CA), flanked by forward (fwd) and reverse (rev) primers, was chosen for qPCR. The sequences used were as follows (all sequences are 5′-3′):

Leptin: prb - cgcagtcggtatccgccaagcag, fwd - aaccctcatcaagaccattgtc, rev - tggtccatcttggacaaactc. CD-31: prb - ccaagctgggatcctgtccg, fwd - caggaccacgtgttagtgtt, rev - actcctgatgggttctgact. Endoglin: prb - ccagctgcggcatgaaagtg, fwd - gacactgacgaccatcttgt, rev - ggaaactgatgatcacctca. VEGFR1: prb - tgagaccaagcccaaggcctcc, fwd -aactccacctccatgtttga, rev - tcgctattctcaagtctatcttca. VEGFR2: prb - tcctgggactgtggcgaagatg, fwd - gaagattgtaaaccgggatg, rev - ttggtcactcttggtcacac. Collagen I (α1): prb -ctcacccttgtcaccacggg, fwd - agactggtcctgctggt, rev - atgcctctgtcaccttgtt. Collagen III (α1): prb- tggtgccctggaagaatggagaac, fwd - tcctggtcctaaaggaaatg, rev - attctttccagcagacctga. Collagen IV(α 1): prb - tgaacctggatttccgggagtac, fwd - acaggcacaagttaaggaaa, rev - aggttcacctttctctccat. Tenascin C: prb - ccctcccaaagaccttattgtgacag, fwd -tctgcaatgaaggttacaca, rev - tgccagatttacagtctcct .

The internal gene control was β2-microglobulin. Amplification reactions containing no RT or cDNA template were used as negative controls.

Serum leptin measurement

A separate group of C57BL/6J mice was used to study blood leptin levels after wounding. Two incisional wounds were made for each animal in this group. Serum samples were obtained at the indicated times after wounding by cardiac puncture followed by euthanasia. All specimens were collected at the same time of day to account for circadian variations in leptin levels. Serum leptin levels were measured by using a species-specific RIA kit (Linco Research, St. Charles, MO).

RESULTS

Local expression of the leptin gene at the wound site

To determine whether leptin is a physiological effector involved in the wound healing process, we made two small contralateral incisional full-thickness wounds on the paramedial dorsal region of adult C57BL/6J mice. The wounds were examined to ensure that the paniculus carnosum was severed. The borders of the incision were well separated, clearly exposing the underlying muscle fascia. After various periods of time, the mice were killed and tissue blocks containing the wounds and a portion of normal surrounding tissue were excised for in situ hybridization and immunohistological examination (see Materials and Methods). To provide a comprehensive microscopic assessment of the wound, we prepared sections by encompassing the keratinocyte epidermal layer and dermis, subdermal tissue, and muscle fascia. Sections were performed transversally across the edges of the wound, such that they contain the advancing keratinocyte epidermal wedge underneath the wound crust or scab.

To determine whether leptin is synthesized within the wound, in situ hybridization experiments were performed on frozen sections of control and wounded skin collected 6 h after the incision. The skin sections included the epidermis and dermis without the hypodermis. Leptin mRNA was detected by using digoxigenin-labeled anti-sense leptin RNA oligonucleotides. Sections of normal mouse skin hybridized to the antisense leptin probe showed leptin mRNA signal in some cells of the basal keratinocyte layer and few isolated cells in the dermis (Fig. 1a, c). In contrast, numerous clusters exhibiting a strong hybridization signal were detected in sections from the wound edge prepared 6 h (Fig. 1b and d) and 24 h (Fig. 1e) after skin incision. In the wound samples, the hybridization signal is scattered throughout the dermis, a pattern that is also distinct in the basal keratinocyte layer of the epidermis. In the dermis, many of the cells exhibiting positive hybridization signal display elongated morphology, probably corresponding to fibroblasts (right arrows, Fig. 1d). The control sense oligonucleotide leptin probe showed no reactivity when either normal (not shown) or wounded skin sections were used (Fig. 1f). To measure the level of leptin gene expression, we performed real-time PCR experiments by using RNA prepared from wounds at various times. Typically, wounds for RNA collection were created by using a biopsy punch. Two bilateral wounds were made on each animal, and four mice were used for each time point. Generally, the tissue from eight wounds was pooled to prepare RNA; pooling of multiple wounds was adopted in order to minimize individual variations. Consistent with the in situ hybridization results shown above, leptin mRNA expression reached a maximum as early as 6 h post-wounding. Of note, this increase in leptin gene expression was sustained throughout the healing process. On Day 5, leptin mRNA levels remained elevated and were found to diminish only after 10 days, at which time repair is complete and scar remodeling is actively taking place (Fig. 1g). Thus, it is apparent that leptin is actively produced in the wound environment throughout the inflammatory and proliferative phases of tissue repair. However, leptin up-regulation appears to begin immediately after inflicting the wound. The observed time frame is consistent with the kinetics of leptin gene expression (in dermal fibroblasts), which we have recently shown to be subject to induction by hypoxia via HIF-1 complexes binding to specific HRE consensus sequences within the leptin promoter (12).

Progressive increase of leptin expression and content in wounds

To evaluate the presence of leptin protein in wound tissue, wounds collected at various times after incision were examined. Immunohistochemical staining was performed by using a polyclonal antibody directed against mouse leptin; normal non-wounded skin was used as control (Fig. 2). Specific staining indicative of cell-associated leptin in intact, unwounded skin was observed in areas containing adipose tissue. However, mild immunoreactivity was also present in non-adipose tissue cell types, such as keratinocytes in the epidermal basal layer, skeletal muscle, and cellular elements of the dermis (Fig. 2a). Following the injury, a robust immunohistochemical staining signal was detected throughout the wound and wound edges. The elevation in leptin was observed to develop progressively, thereby underscoring the continued expression of leptin in the course of the wound repair process. As early as 6 h after skin incision, the wounds already exhibit distinctive immunostaining for leptin (Fig. 2b, c), which became markedly stronger after 24 h (Fig. 2d, e) and remained conspicuous even at 48(Fig. 2f, g) and 72 h (Fig. 2h). At a higher magnification, it is evident that abundant wound-resident cells with myofibroblast-like appearance were intensely immunoreactive to the leptin antibodies (Fig. 2k). Although leptin is known to be expressed constitutively in adipose tissue, our findings suggest that after wounding, adipocytes may also increase their level of leptin production (Fig. 2l). Notably, not all cells displayed immunoreactivity to the leptin antibodies (Fig. 2k, l). In particular, inflammatory cells recruited within the first 48 h after the injury do not display leptin immunoreactivity (Fig. 2f). For control, serial sections were routinely exposed to anti-leptin antibodies in the presence of excess soluble recombinant murine leptin to compete with binding of the antibody to the tissue in the histological section. The absence of immunostaining in control sections demonstrates that positive immunoreactivity detected on the tested sections is specific for leptin (Fig. 2i).

Arrest of wound healing by neutralizing anti-leptin antibodies

To establish whether there might be a functional requirement for leptin during wound healing, 5 µg of a neutralizing polyclonal antibody raised against recombinant mouse leptin was applied to an excisional skin wound every day for 3 days. In a control group of animals, wounds were treated with the same amount of purified nonimmune goat IgG. On Day 3, the wounds were collected and processed for histological evaluation. The degree of reepithelialization was measured by computer-assisted morphometry as the transversal distance between the wound margins at the center of the spindle-shaped incision defect. This distance was, on average, ~1.6 times wider in wounds treated with the anti-leptin antibody when compared with control wounds treated with nonimmune IgG (Fig. 3c). In addition, the thickness of the granulation tissue present in the wound was significantly reduced in the mice treated with anti-leptin antibodies (Fig. 3b). To further evaluate the effects of antibody on healing progression, experiments were conducted to measure the expression of various mRNA transcripts known to be elevated in wounds (Fig. 3d). The neutralization of soluble leptin in the wound environment induced a significant down-regulation in the transcriptional expression level of several extra cellular matrix molecules, such Tenascin C and collagen III. These changes may help explain the visibly reduced matrix density found in antibody-treated wounds (Fig. 3b). In addition, mRNA expression for endothelial markers, such as CD-31, endoglin and VEGFR-2 (FLK-1) was also diminished. These results strongly suggest that wound-associated leptin production plays an important functional role in

epidermal reepithelialization, neovascularization, and development of granulation tissue, which are key events in the wound repair.

Increased serum leptin levels after wounding

To evaluate whether the increase of leptin detected in the injured tissue had an impact on circulating leptin, we measured blood leptin levels in serum after wounding. Incisional wounds were created in adult C57BL/6J mice, and serum leptin was measured at various times thereafter. Circulating leptin levels increased rapidly, reaching a peak by 12 h and returning to baseline after 24 h. Leptin serum concentration increased by an average of sixfold, from 2 ng/ml to 12 ng/ml (Fig. 4a). To determine whether the peak of circulating leptin originates from the wound site, we inflicted wounds on human skin grafted on human-SCID mouse chimeras. We found that human leptin was not detectable in unwounded human skin-SCID mouse chimeras. However, detectable human leptin appeared in the mouse circulation after wounding; in contrast, mouse leptin levels did not change significantly (Fig. 4a, b). Moreover, circulating murine leptin in wounded human skin-SCID chimeras was comparable with the levels of circulating leptin in unwounded C57BL/6J mice (Fig. 4a, b). These results suggest that increased production of leptin in wounds contributes to the sharp increase in circulating leptin after the injury. The spillover of wound leptin into the circulation may likely occur due to a combination of factors, such as increased local permeability of the blood vessels in the periphery of the wound, clearance of wound fluid, and direct up-regulation of leptin synthesis in the wound as documented in this report.

DISCUSSION

In this report, we demonstrate that wound-resident cells rapidly engage in leptin synthesis shortly following an incision injury of the skin. Leptin production appears to continue throughout the entire process of healing at the wound site. Several previous observations suggest that leptin is an essential component of tissue repair. First, naturally occurring mutations of the leptin or leptin receptor gene (ob/ob and db/db) in mice result in a severe wound healing impairment phenotype (7, 15, 16). Of note, the healing defect appears to be independent from the diabetic phenotype because, although ob/ob mice only occasionally develop full-scale diabetes, they regularly manifest a wound-healing deficit (17, 18). Second, macroscopic assessment of cutaneous wounds following topical or systemic treatment with leptin restores healing in leptin-deficient ob/ob mice, but not in leptin receptor-deficient db/db mice (11, 7). Third, we have recently shown that leptin is a hypoxia-inducible gene and it is therefore plausible that its expression might be up-regulated in ischemic tissues, such as the wound bed (12). In this regard, leptin is a known pro-angiogenic factor and it is likely to be an active participant in the neovascularization process that accompanies wound healing (19, 20). Taken together, these observations coupled with the results reported herein suggest that normal wound healing requires acute local production of leptin and a functionally intact leptin signaling system at the wound site.

In the classical context, leptin is regarded as a circulating hormone produced primarily by adipose tissue, fulfilling a predominant biological role to regulate feeding and energy homeostasis through CNS efferent pathways (8). However, a growing number of reports depict a rather broad spectrum of physiological actions for leptin, some of which may reflect direct, peripheral effects. For example, leptin has been shown to exert direct effects on T cells and monocytes, causing the release of cytokines that include GM-CSF, G-CSF, TNF-α, and IL-6

(21–23). Our own studies have documented that leptin directly causes angiogenesis on endothelial cells, which express the Ob-Rb long form (bearing the entire cytoplasmic signaling domain) of the leptin receptor. Also, leptin induces neovascularization in corneas from normal rats but not in corneas from fa/fa Zucker rats, which lack functional receptors (19). Additionally, a number of previous reports have documented the production of leptin in a variety of non-adipose cells, including placental trophoblasts, mammary epithelial cells, gastric fundic mucosa, and ovarian follicle cells (24–30). In this study, we show that cells residing within the wound environment created by an incisional dermal insult engage in a dramatic burst of leptin production shortly thereafter. When considered collectively, the above observations are consistent with a pleiotropic character for leptin, a feature shared by other multifunctional cytokines to which leptin is structurally related, including IL-6, oncostatin M (OSM), ciliary neurotrophic factor (CNTF), leukemia inhibitory factor (LIF), interleukin 11 (IL-11), and cardiotropin 1 (CT-1) (31–33). Thus, our results give strength to the concept of leptin as a functionally pleiotropic cytokine that shares overlapping biological effects with other IL-6 cytokine family members.

Normal progression of healing in wounds involves formation of granulation tissue, which requires neovascularization. A variety of soluble factors, such as FGF-2, TGF-α and -β, TNF-α, and VEGF are actively produced in wounds and stimulate angiogenesis either directly or through chemoattracted macrophages that actively secrete angiogenic molecules (34). The previously demonstrated role of leptin as a potent angiogenic factor, coupled with our own findings showing its presence in wounds, suggests that the healing augmentation effect of leptin may be the result of its ability to induce neovascularization during tissue repair. However, it is conceivable that leptin may also modulate other important aspects of the wound repair process, in addition to its pro-angiogenic activity. For example, the recent report that leptin promotes platelet aggregation, coupled with its known stimulating effects on macrophages and lymphocytes, suggests a more comprehensive, multifunctional role during wound healing, possibly including subsequent remodeling of the scar tissue as well (21–23, 35). In particular, the study of the relationship between leptin induction and other cytokines actively produced at the wound site should provide insight into the potential role that leptin might play as an upstream regulator of the cytokine activation cascade that accompanies tissue repair.

ACKNOWLEDGMENTS

We thank the staff of Yale Dermatopathology laboratory, Yale Animal Services, and Cedars-Sinai Research Institute Animal Services facilities for their technical assistance. We are also grateful to Jordan Pober and Parvez Sultan for the SCID mice.

REFERENCES

1. Dvonch, V. M., Murphey, R. J., Matsuoka, J., and Grotendorst, G. R. (1992) Changes in growth factor levels in human wound fluid. Surgery 112, 18–23

2. Howdieshell, T. R., Riegner, C., Gupta, V., Callaway, D., Grembowicz, K., Sathyanarayana, and McNeil, P. L. (1998) Normoxic wound fluid contains high levels of vascular endothelial growth factor. Ann. Surg. 228, 707–715

3. Nissen, N. N., Polverini, P. J., Koch, A. E., Volin, M. V., Gamelli, R. L., and DiPietro, L. A. (1998) Vascular endothelial growth factor mediates angiogenic activity during the proliferative phase of wound healing. Am. J. Pathol. 152, 1445–1452

4. Christofidou-Solomidou, M., Bridges, M., Murphy, G. F., Albelda, S. M., and DeLisser, H. M. (1997) Expression and function of endothelial cell alpha v integrin receptors in wound-induced human angiogenesis in human skin/SCID mice chimeras. Am. J. Pathol. 151, 975–983

5. Greenhalgh, D. G., Sprugel, K. H., Murray, M. J., and Ross, R. (1990) PDGF and FGF stimulate wound healing in the genetically diabetic mouse. Am. J. Pathol. 136, 1235–1246

6. Albertson, S., Hummel, R. P., 3rd, Breeden, M., and Greenhalgh, D. G. (1993) PDGF and FGF reverse the healing impairment in protein-malnourished diabetic mice. Surgery 114, 368–372; discussion 372–363

7. Tsuboi, R., Shi, C. M., Rifkin, D. B., and Ogawa, H. (1992) A wound healing model using healing-impaired diabetic mice. J. Dermatol. 19, 673–675

8. Friedman, J. M., and Halaas, J. L. (1998) Leptin and the regulation of body weight in mammals. Nature 395, 763–770

9. Lee, G. H., Proenca, R., Montez, J. M., Carroll, K. M., Darvishzadeh, J. G., Lee, J. I., and Friedman, J. M. (1996) Abnormal splicing of the leptin receptor in diabetic mice. Nature 379, 632–635

10. Frank, S., Stallmeyer, B., Kampfer, H., Kolb, N., and Pfeilschifter, J. (2000) Leptin enhances wound re-epithelialization and constitutes a direct function of leptin in skin repair. J. Clin. Invest. 106, 501–509

11. Ring, B. D., Scully, S., Davis, C. R., Baker, M. B., Cullen, M. J., Pelleymounter, M. A., and Danilenko, D. M. (2000) Systemically and topically administered leptin both accelerate wound healing in diabetic ob/ob mice. Endocrinology 141, 446–449

12. Ambrosini, G., Nath, A. K., Sierra-Honigmann, M. R., and Flores-Riveros, J. (2002) Transcriptional activation of the human leptin gene in response to hypoxia. Involvement of hypoxia-inducible factor 1. J. Biol. Chem. 277, 34601–34609

13. Grosfeld, A., Andre, J., Hauguel-De Mouzon, S., Berra, E., Pouyssegur, J., and Guerre-Millo, M. (2002) Hypoxia-inducible factor 1 (HIF-1) transactivates the human leptin gene promoter. J. Biol. Chem.

14. Semenza, G. L., and Wang, G. L. (1992) A nuclear factor induced by hypoxia via de novo protein synthesis binds to the human erythropoietin gene enhancer at a site required for transcriptional activation. Mol. Cell. Biol. 12, 5447–5454

15. Goodson, W. H., III, and Hunt, T. K. (1986) Wound collagen accumulation in obese hyperglycemic mice. Diabetes 35, 491–495

16. Tsuboi, R., and Rifkin, D. B. (1990) Recombinant basic fibroblast growth factor stimulates wound healing in healing-impaired db/db mice. J. Exp. Med. 172, 245–251

17. Hummel, K. P., Coleman, D. L., and Lane, P. W. (1972) The influence of genetic background on expression of mutations at the diabetes locus in the mouse. I. C57BL-KsJ and C57BL-6J strains. Biochem. Genet. 7, 1–13

18. Coleman, D. L., and Hummel, K. P. (1973) The influence of genetic background on the expression of the obese (Ob) gene in the mouse. Diabetologia 9, 287–293

19. Sierra-Honigmann, M. R., Nath, A. K., Murakami, C., Garcia-Cardena, G., Papapetropoulos, A., Sessa, W. C., Madge, L. A., Schechner, J. S., Schwabb, M. B., Polverini, P. J., et al. (1998) Biological action of leptin as an angiogenic factor. Science 281, 1683–1686

20. Cao, R., Brakenhielm, E., Wahlestedt, C., Thyberg, J., and Cao, Y. (2001) Leptin induces vascular permeability and synergistically stimulates angiogenesis with FGF-2 and VEGF. Proc. Natl. Acad. Sci. USA 98, 6390–6395

21. Lord, G. M., Matarese, G., Howard, J. K., Baker, R. J., Bloom, S. R., and Lechler, R. I. (1998) Leptin modulates the T-cell immune response and reverses starvation-induced immunosuppression. Nature 394, 897–901

22. Santos-Alvarez, J., Goberna, R., and Sanchez-Margalet, V. (1999) Human leptin stimulates proliferation and activation of human circulating monocytes. Cell. Immunol. 194, 6–11

23. Gainsford, T., Willson, T. A., Metcalf, D., Handman, E., McFarlane, C., Ng, A., Nicola, N. A., Alexander, W. S., and Hilton, D. J. (1996) Leptin can induce proliferation, differentiation, and functional activation of hemopoietic cells. Proc. Natl. Acad. Sci. USA 93, 14564–14568

24. Hoggard, N., Hunter, L., Duncan, J. S., Williams, L. M., Trayhurn, P., and Mercer, J. G. (1997) Leptin and leptin receptor mRNA and protein expression in the murine fetus and placenta. Proc. Natl. Acad. Sci. USA 94, 11073–11078

25. Senaris, R., Garcia-Caballero, T., Casabiell, X., Gallego, R., Castro, R., Considine, R. V., Dieguez, C., and Casanueva, F. F. (1997) Synthesis of leptin in human placenta. Endocrinology 138, 4501–4504

26. Masuzaki, H., Ogawa, Y., Sagawa, N., Hosoda, K., Matsumoto, T., Mise, H., Nishimura, H., Yoshimasa, Y., Tanaka, I., Mori, T., et al. (1997) Nonadipose tissue production of leptin: leptin as a novel placenta-derived hormone in humans. Nat. Med. 3, 1029–1033

27. Henson, M. C., Castracane, V. D., O'Neil, J. S., Gimpel, T., Swan, K. F., Green, A. E., and Shi, W. (1999) Serum leptin concentrations and expression of leptin transcripts in placental trophoblast with advancing baboon pregnancy. J. Clin. Endocrinol. Metab. 84, 2543–2549

28. Bado, A., Levasseur, S., Attoub, S., Kermorgant, S., Laigneau, J. P., Bortoluzzi, M. N., Moizo, L., Lehy, T., Guerre-Millo, M., Le Marchand-Brustel, Y., et al. (1998) The stomach is a source of leptin. Nature 394, 790–793

29. Smith-Kirwin, S. M., O'Connor, D. M., De Johnston, J., Lancey, E. D., Hassink, S. G., and Funanage, V. L. (1998) Leptin expression in human mammary epithelial cells and breast milk. J. Clin. Endocrinol. Metab. 83, 1810–1813

30. Antczak, M., Van Blerkom, J., and Clark, A. (1997) A novel mechanism of vascular endothelial growth factor, leptin and transforming growth factor-beta2 sequestration in a subpopulation of human ovarian follicle cells. Hum. Reprod. 12, 2226–2234

31. Lambert, P. D., Anderson, K. D., Sleeman, M. W., Wong, V., Tan, J., Hijarunguru, A., Corcoran, T. L., Murray, J. D., Thabet, K. E., Yancopoulos, G. D., et al. (2001) Ciliary neurotrophic factor activates leptin-like pathways and reduces body fat, without cachexia or rebound weight gain, even in leptin-resistant obesity. Proc. Natl. Acad. Sci. USA 98, 4652–4657

32. Akira, S. (1997) IL-6-regulated transcription factors. Int. J. Biochem. Cell Biol. 29, 1401–1418

33. Touw, I. P., De Koning, J. P., Ward, A. C., and Hermans, M. H. (2000) Signaling mechanisms of cytokine receptors and their perturbances in disease. Mol. Cell. Endocrinol. 160, 1–9

34. Tonnesen, M. G., Feng, X., and Clark, R. A. (2000) Angiogenesis in wound healing. J. Investig. Dermatol. Symp. Proc. 5, 40–46

35. Nakata, M., Yada, T., Soejima, N., and Maruyama, I. (1999) Leptin promotes aggregation of human platelets via the long form of its receptor. Diabetes 48, 426–429

Received March 4, 2003; accepted June 25, 2003.

Fig. 1

Figure 1. Increased expression of leptin mRNA in wounds demonstrated by in situ hybridization and real-time RT-PCR. Leptin RNA probes were hybridized to frozen sections prepared from normal mouse skin (a, c) or from the margins of wounds collected 6 h (b, d) and 24 h (e, f) post-incision. Specific mRNA hybridization signal (arrows) appears to be in the keratinocyte epidermal layer and dermis. Specific antisense (e) and negative control sense (f) are shown. E = epithelium. wb = wound bed. D = dermis. (Scale bars: a and b = 125 µm; c and d = 10 µm; e and f = 20 µm). Quantitative (real-time) RT-PCR (g) was used to measure leptin mRNA expression in excisional wounds after 6, 24, and 48 h, 5 and 10 days. Values are adjusted to volume of skin/wound tissue collected. A total of eight wounds were pooled for each time point. Values are shown as average ± SEM (n=6).

Fig. 2

Figure 2. Immunohistochemical detection of leptin in full-thickness incisional wounds. Sections were prepared from normal mouse skin (a) or from wounds collected at 6 (b, c); 24 (d, e); 48 (f, g); and 72 (h, i) hours and 10 (j) days post-incision. Immunostaining was conducted by using a rabbit polyclonal antibody to mouse leptin. A negative control (i) to demonstrate specificity of the immunostaining was prepared by using a serial section from the 48 h wound tissue block and was incubated with the anti-leptin antibody in the presence of an excess of soluble recombinant mouse leptin (10 µg/ml). Higher magnification photomicrograph of the 48 h section showing details of the wound bed microvasculature (g, white arrows) and the dermis around the wound with myofibroblast-like cells displaying intense immunoreactivity (k, black arrows). Subcutaneous adipose tissue at 48 h showing increased immunoreactivity containing some interstitial cells with no immunoreactivity (l, black arrows). Scale bars are 50 µm for (a, b, d, f, h, i, j; 20 µm for c, e, g; and 10 µm for k and l).

Fig. 3

Figure 3. Neutralizing anti-leptin antibodies impair normal healing. Microscopic appearance of sections from excisional dermal wounds on Day 3 after receiving daily treatments of 5 µg of goat IgG (a) or 5 µg neutralizing anti-leptin antibodies (b). Sections were stained with H&E. A representative photomicrograph of both treatment groups depicting the progress of healing is shown. The saline group exhibits a greater degree of contraction and re-epithelialization of the wound. Arrows indicate location of the dermis on the borders of the wounds (D). The anti-leptin antibody-treated wound has failed to contract and has little detectable granulation tissue, and the matrix density is notably reduced. Morphometric analysis of wound size from IgG controls and anti-leptin treated wounds in wild-type mice by measurement of the linear distance between dermis borders(c) (average ± SEM; n=6; P=0.005). Quantitative (real-time) RT-PCR results from 3-day excisional wounds treated daily with 5 µg neutralizing anti-leptin antibody compared with control wounds treated with 5 µg goat IgG (d). Values are adjusted to volume of skin/wound tissue collected and are relative to unwounded skin. A total of eight wounds were pooled for each time point. * Values of Collagens I and III shown are equal to 1/10 of the actual values to fit the scale. Data shown is from an experiment done in triplicate. VEGFR1 = VEGF Receptor-1 or FLT-1; VEGFR2 = VEGF Receptor-2 or FLK-1; COL-1 = Collagen I (α1), COL III = Collagen III (α1); COL-IV = Collagen IV(α 1); TENASCIN = Tenascin C. Values are shown as average ± SEM (n=6).

Fig. 4

Figure 4. Increased production of leptin in the wound milieu causes a transient increase in serum leptin. C57/BLK mice (a) received bilateral full thickness incisional wounds (8 mm). Serum was obtained at indicated times by cardiac puncture (n=3). All serum samples were collected at same time of day. To determine whether the 10–12 h peak of circulating leptin originates from the wound site, SCID beige mice-human chimeras were wounded on the human skin graft and sera were collected after 8 h (n=3). Mouse (b) and human (c) leptin concentrations were measured by two species-specific RIA.