Abstract
Leptin, a 16-kD circulating hormone secreted mainly by white adipose tissue, is a product of the obese (ob) gene. Leptin acts on human marrow stromal cells to enhance differentiation into osteoblasts and inhibit differentiation into adipocytes. Leptin also inhibits bone formation through a hypothalamic relay. To obtain a better understanding of the potential role of leptin in bone formation, the localization of leptin in endochondral ossification was examined immunohistochemically. High expression of leptin was identified in hypertrophic chondrocytes in the vicinity of capillary blood vessels invading hypertrophic cartilage and in a number of osteoblasts of the primary spongiosa beneath the growth plate. The hypertrophic chondrocytes far from the blood vessels were negative for leptin. Moreover, we detected the production and secretion of leptin by a mouse osteoblast cell line (MC3T3-E1) and a mouse chondrocyte cell line (MCC-5) by RT-PCR, immunocytochemistry, and Western blotting. Leptin enhanced the proliferation, migration, tube formation, and matrix metalloproteinase-2 (MMP-2) activity of human endothelial cells (HUVECs) in vitro. These findings suggest the possibility that leptin exerts its influence on endochondral ossification by regulating angiogenesis.
ENDOCHONDRAL OSSIFICATION and intramembranous ossification are two major processes that control skeletogenesis (bone formation). Bones of the vertebral column, pelvis, and extremities are formed by endochondral ossification. In this type of bone formation, cartilage tissue is first formed and is eventually replaced by bone tissue. The first indication of endochondral bone formation in a long bone is a local enlargement of the chondrocyte (hypertrophy) in the middle of the shaft of the cartilage model (diaphysis). At the same time, blood vessels invade hypertrophic cartilage, branch, and grow forward at either end of the center of ossification (Trueta 1963; Streeten and Brandi 1990; Stanka et al. 1991). Mesenchymal stem cells (including osteoprogenitor cells) and hematopoietic stem cells migrate with perivascular connective tissue of the invading blood vessels into the hypertrophic and calcified cartilage. Osteoprogenitor cells differentiate into osteoblasts and begin to deposit bone matrix on the spicules of calcified cartilage. The same series of events, i.e., hypertrophic conversion of chondrocytes, blood vessel invasion, and bone matrix formation by osteoblasts also occur at the epiphyses (Alini et al. 1996). In both primary and secondary centers of ossification, the invasion of blood vessels into hypertrophic and calcified cartilage with mesenchymal stem cells appears to be a prerequisite for endochondral ossification. From this point of view, angiogenic factors that can regulate the proliferation, migration, and/or function of endothelial cells are considered to play an important role in bone formation.
Leptin, a 16-kD circulating hormone secreted mainly by white adipose tissue, is a product of the obese (ob) gene (Zhang et al. 1994). It influences body weight homeostasis through its effects on food intake and energy expenditure by negative feedback at the hypothalamic nuclei (Friedman and Halaas 1998). Recent studies have demonstrated that leptin is produced by other tissues, such as placenta (Masuzaki et al. 1997), fetal bone/cartilage (Hoggard et al. 1997), stomach (Bado et al. 1998), skeletal muscles (Wang et al. 1998), brain, and pituitary gland (Jin et al. 1999; Morash et al. 1999). In addition to its effects on the central nervous system, leptin has various physiological actions on lipid metabolism (Bryson et al. 1999), hematopoiesis (Gainsford et al. 1996), thermogenesis (Hwa et al. 1996), ovarian function (Spicer and Francisco 1997; Zachow and Magoffin 1997), and angiogenesis (Bouloumie et al. 1998; Sierra-Honigmann et al. 1998). Furthermore, leptin acts on human marrow stromal cells to enhance differentiation into osteoblasts and to inhibit differentiation into adipocytes (Thomas et al. 1999). Other experiments have shown that leptin inhibits bone formation through a hypothalamic relay (Ducy et al. 2000). On the basis of these facts, leptin may also play an important physiological role in skeletogenesis.
To elucidate a possible role of leptin in bone formation, we immunohistochemically investigated the expression of leptin in endochondral ossification in mouse. Gene and protein expression of leptin in MC3T3-E1, a mouse osteoblast cell line, and MCC-5, a mouse chondrocyte cell line (Kitaoka et al. 2001), was also analyzed by reverse transcription-polymerase chain reaction (RT-PCR), immunocytochemistry (ICC), and Western blotting. The effect of leptin on human umbilical vein endothelial cells (HUVECs) was also investigated in vitro. As a result, leptin is suggested to regulate angiogenesis at the ossification center of endochondral bone formation by enhancing proliferation, migration, and differentiated function of endothelial cells.
Materials and Methods
Animals
ICR mice (Clea Japan; Tokyo, Japan) were used. The housing care and experimental protocol were approved by the Animal Care and Use Committee of the University of Tokushima School of Dentistry.
Immunohistochemistry (IHC)
Hindlimbs of 1-week-old and neonatal ICR mice were removed, fixed in Mildform 10 N (Wako Pure Chemical Industries; Osaka, Japan), decalcified with 10% formic acid/10% sodium citrate, and embedded in paraffin. Sections were cut 3 μm thick and transferred onto poly-L-lysine-coated glass slides. After deparaffinization with xylene and rehydration with descending concentrations of ethanol, endogenous peroxidase was blocked by treatment with 3% H2O2 in methanol for 35 min at room temperature (RT). After treatment with 10% normal rabbit serum for 10 min at RT, sections were incubated with goat anti-mouse leptin antibody (Genzyme; Minneapolis, MN) diluted 1:200 in PBS, pH 7.4, containing 1% bovine serum albumin (BSA) overnight at 4C. After washing with PBS, the localization of leptin was visualized by a Histofine SAB-PO (G) Kit (Nichirei; Tokyo, Japan) and a diaminobenzidine (DAB) Substrate Kit (Nichirei). Sections were counterstained with hematoxylin and mounted. The specificity of the immunoreaction was confirmed by (a) incubation with goat IgG (Chemicon; Temecula, CA) instead of the primary antibody or (b) preabsorption of anti-leptin antibody with recombinant mouse leptin (Diaclone Research; Besan, France).
Fifteen-day-old embryos were also fixed in Mildform 10 N and embedded in paraffin. Sagittal sections were cut at 3 μm thickness and transferred onto poly-L-lysine-coated glass slides. After deparaffinization and rehydration, antigenicity of leptin was recovered by immersing sections in boiling distilled water for 10 min. The following IHC observation was performed as mentioned above. All the histological observations were performed using consecutive sections to understand the three-dimensional distribution of leptin-positive cells.
Cell Culture
MC3T3-E1, a mouse osteoblast cell line, was cultured in α-minimum essential medium (α-MEM) containing 10% fetal bovine serum (FBS), 50 μg/ml <sc<L-ascorbic acid phosphate magnesium salt n-hydrate (Wako), 1 × Glutamax (Gibco BRL; Gaithersburg, MD) 100 U/ml penicillin, and 100 μg/ml streptomycin (Gibco) (hereafter termed MC3T3-E1 growth medium).
MCC-5, a mouse chondrocyte cell line derived from costal cartilage of 8-week-old male transgenic mice harboring temperature-sensitive simian virus 40 large T-antigen driven by an SV40 promoter, was established in our laboratory (Kitaoka et al. 2001). This cell line was cultured in α-MEM containing 0.5% FBS, 50 μg/ml <sc<L-ascorbic acid phosphate magnesium salt n-hydrate, 1 × Glutamax, 100 U/ml penicillin, and 100 μg/ml streptomycin (hereafter termed MCC-5 growth medium).
HUVECs (Kurabo Biomedical Business; Osaka, Japan) were cultured in HuMedia-EB2 (Kurabo Biomedical Business) containing 2% FBS, 10 ng/ml human epidermal growth factor (EGF), 1 μg/ml hydrocortisone, 50 μg/ml gentamicin, 50 ng/ml amphotericin, 5 ng/ml human basic fibroblast growth factor (bFGF), and 10 ng/ml heparin (hereafter termed HUVEC growth medium).
Reverse Transcription-Polymerase Chain Reaction (RT-PCR)
Total RNA was extracted from cells at confluence using TRIzol reagent (Gibco), and the cDNA was generated from 1 μg of total RNA by using the Superscript Preamplification System (Gibco) with random primers. PCR was carried out in 50 μl reaction mixture using PCR master (Roche Diagnostics; Mannheim, Germany). The primers for the leptin (GenBank accession no. U18812): forward (337–356), 5′-TGC TGC AGA TAG CCA ATG AC-3′; reverse (478–457), 5′-GAG TAG AGT GAG GCT TCC CAG GA-3′ were as described (Hoggard et al. 1997). The following PCR primers were also used: glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (GenBank accession no. X01677): forward (586–607), 5′-ACC ACA GTC CAT GCC ATC AC-3′; reverse (1037–1017), 5′-TCC ACC ACC CTG TTG CTG TA-3′. The PCR amplification was carried for 40 cycles at 94C for 60 sec, 55C for 60 sec, and 72C for 90 sec (leptin), and for 25 cycles at 94C for 30 sec, 55C for 30 sec, and 72C for 60 sec (GAPDH). PCR products were analyzed by ethidium bromide staining after separation by electrophoresis on a 2% agarose gel.
Immunocytochemistry
MC3T3-E1 and MCC-5 were seeded in two-well glass chamber slides (Nalge Nunc International; Naperville, IL). At semiconfluence, cells were fixed in 95% ethanol for 15 min at RT. Fixed cells were incubated with 1% BSA in PBS for 30 min at RT and then with anti-mouse leptin antibody diluted 1:200 in PBS containing 1% BSA for 1 hr at RT. The cells were washed three times with PBS and leptin expression was detected with FITC-conjugated anti-goat IgG (Wako) under a fluorescence microscope (Nikon; Tokyo, Japan).
Western Blotting for Leptin
MC3T3-E1 and MCC-5 were cultured in growth media until they reached confluency. At confluence, the culture media were changed to serum-free media and the cultures were maintained for 24 hr. Conditioned medium of each cell line was collected and concentrated using Centriprep (Millipore; Bedford, MA). The samples (50 μg protein) were separated by the NuPAGE System (Novex; San Diego, CA) using 4–12% Bis-Tris Gel, and electroblotted onto a nitrocellulose membrane (Hybond ECL; Amersham Pharmacia Biotech, Poole, UK). Membranes were blocked with 5% skim milk for 3 hr at RT and then incubated with goat anti-mouse leptin antibody diluted 1:500 in 25 mM Tris/137 mM NaCl/0.05% Tween-20/0.1% BSA, pH 7.4, for 2 hr. Membranes were washed several times and incubated with horseradish peroxidase-conjugated anti-goat IgG antibody (Dako Denmark; Glostrup, Denmark) diluted 1:2000 in the same buffer for 1 hr. After several washes, leptin was visualized using ECL Western Blotting Detection Reagents (Amersham Pharmacia Biotech) and an ECL mini-camera (Amersham Pharmacia Biotech).
Cell Proliferation Assay
The effect of leptin on the proliferation of HUVECs was analyzed using a crystal violet staining method as described previously (Fedarko et al. 1995). Cells were seeded into 24-well culture plates at a cell density of 3 × 103 cells/well. After 6 hr, medium was changed. HUVECs were cultured in HuMedia-EB2 containing 1% FBS and various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of recombinant human leptin (Diaclone Research) or 10 ng/ml VEGF165, which is the most potent isoform of VEGF. After 4 days of culture, cells were rinsed with PBS and fixed with 1% glutaraldehyde in PBS for 15 min. Cells were stained by incubation with 0.02% crystal violet in deionized water for 30 min at RT. The cells were rinsed twice with deionized water and the crystal violet bound to cells was extracted by overnight incubation with 200 μl/well of 70% ethanol at 4C. Absorbance was measured at 570 nm using a microplate reader (Corona; Ibaraki, Japan).
Cell Migration Assay
The effect of leptin on migration of HUVECs was determined using a 24-well chemotaxicell chamber (Kurabo Biomedical Business) with polycarbonate film (8-μm pore size). HUVECs (200 μl of 2 × 105 cells/ml) in serum-free medium containing leptin (0, 0.1, 1, 10, or 100 ng/ml) were applied in the upper compartment of the chamber. The medium (500 μl) containing leptin (0, 0.1, 1, 10, or 100 ng/ml) was added to the lower compartment of the chamber. The chamber was incubated at 37C for 6 hr in 5% CO2 in air. After incubation the filter was removed and the migrated cells were fixed with 1% glutaraldehyde and stained with Giemsa staining solution. The number of migrated cells was counted in 10 random fields at ×200 magnification under a microscope. A checkerboard design was used to distinguish between chemotaxisis (migration direct up a concentration gradient) and chemokinesis (non-oriented increase in cell motility).
Tube Formation Assay
Two hundred and fifty μl Matrigel Growth Factor Reduced (Becton Dickinson; CBP Product, Boston, MA) was added to a 24-well culture plate and was then allowed to solidify at 37C for 1 hr. HUVECs (1 × 104 cells/well) were seeded on the matrigel and cultured in HuMedia-EB2 containing various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of recombinant human leptin or 10 ng/ml VEGF165 at 37C for 12 hr in a humidified atmosphere of 5% CO2 in air. After incubation, three different fields were randomly observed with a phase-contrast microscope and photographed at ×40 magnification, and the lengths of the tube structures were measured.
Zymogram for Matrix Metalloproteinases
HUVECs were grown in standard growth medium until they reached confluence. At confluence, the medium was changed to Humedia-EB2 containing various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of leptin or 10 ng/ml VEGF165. After 6 hr the sample from each conditioned medium was collected. The samples were mixed with Tris-glycine SDS Sample Buffer (Novex) and separated on a 10% Zymogram Gel (Novex) containing gelatin with 1 × Tris-glycine SDS Running Buffer (Novex). After washing in the Renaturing Buffer (Novex) for 30 min at RT and then in Developing Buffer (Novex) for 30 min at RT, the gels were incubated in fresh Developing Buffer for 4 hr, 7 hr, or 12 hr at 37C. Twelve-hour incubation with Developing Buffer showed the most quantitative results in the present assay. The gel was stained with 0.25% Coomassie Brilliant Blue and destained to visualize the MMP bands. Expression levels were measured using NIH Image software.
Statistics
All data are expressed as mean ± SE. Statistical analysis was performed by one-way ANOVA with the Bonferroni test. Values of p<0.05 were considered statistically significant.
Results
Localization of Leptin in Endochondral Ossification
One-week-old Mice. High expression of leptin was identified in hypertrophic chondrocytes, which were characterized by enlarged cell bodies, swelling of nuclei, less stainability of chromatin, and confluence of their lacunae, adjacent to capillary blood vessels invading secondary ossification centers (Figures 1B, 1E, and 2B). However, the hypertrophic chondrocytes that were not adjacent to blood vessels were completely negative for leptin (Figures 1B and 1E). Moreover, no gradation of staining for leptin was noted. A number of osteoblasts in the primary spongiosa beneath the growth plate (Figures 1H and 2F) and some chondrocytes in the hypertrophic zone of the growth plate were noted to express leptin (Figure 2E). In contrast, no staining was observed in the resting or proliferative cartilage (Figures 2C and 2D).
Immunohistochemical localization of leptin in endochondral ossification of 1-week-old mouse femurs. (
Neonatal Mice. A similar pattern of leptin expression was observed in primary spongiosa of neonatal mouse femurs. At this time the invasion of capillary blood vessels into epiphyses was not yet recognized. Leptin was not detected in chondrocytes in epiphyses (data not shown).
Fifteen-day-old Mouse Embryos. Some blood vessels were observed before and after invasion into hypertrophic cartilage (Figure 3). At this primary ossification center, early hypertrophic chondrocytes were positive for leptin (Figures 3B and 3D). In particular, expression of leptin was observed before invasion of blood vessels (Figure 3B). This expression pattern of leptin was substantially same as that in the secondary ossification center.
Immunolocalization of leptin in the epiphyseal growth plate from 1-week-old mice. (
Immunohistochemical localization of leptin in ribs (
Leptin Production by Mouse Osteoblasts and Chondrocytes
To confirm that mouse osteoblasts and chondrocytes produce and secrete leptin in vitro, the gene and protein expression of leptin was investigated by RT-PCR and Western blotting, respectively. Leptin and the long-form variant of the leptin receptor (OB-Rb), which was the most functional isoform in leptin receptors, were detected in MC3T3-E1 and MCC-5 (Figure 4A). Western blotting analysis of conditioned media of MC3T3-E1 and MCC-5 showed a 16-kD band of leptin (Figure 4B). IHC of MC3T3-E1 and MCC-5 showed leptin protein located in the cytoplasm where FITC labeling was observed (Figure 5). mRNA and protein of Types II and X collagen were also detected in MCC-5 (data not shown).
RT-PCR analysis showed that mRNA for OB-Rb was expressed in MC3T3-E1 and MCC-5. This finding suggested that leptin had an effect on proliferation and/or differentiation of osteoblasts and chondrocytes. In support of this hypothesis, we examined the proliferation and the production of extracellular matrix of MC3T3-E1 and MCC-5 in the presence of various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of leptin. However, statistically significant differences were not detected compared with control (no leptin) at each time point (data not shown).
Effects of Leptin on Human Endothelial Cells
First, we confirmed the expression and functionality of the long-form variant of the leptin receptor (OB-Rb) in HUVECs by RT-PCR and immunoprecipitation/Western blotting using anti-Stat3 antibody, anti-Shp-2 antibody, and anti-phosphotyrosine antibody. RT-PCR showed that HUVECs expressed mRNA for OB-Rb, and immunoprecipitation/Western blotting analysis revealed that Stat3 and Shp-2, two downstream regulators of leptin signaling, were tyrosine-phosphorylated by treatment with leptin (data not shown). These results revealed that OB-Rb in HUVECs was functional.
Leptin significantly promoted the proliferation of HUVECs in a dose-dependent manner compared with control (Figure 6). The maximal effect observed with 100 ng/ml leptin was statistically comparable to that elicited by 10 ng/ml VEGF165.
(
To investigate the effect of leptin on endothelial cell migration (chemotaxis and/or chemokinesis), a checkerboard design was used to test the response of HUVECs to leptin (Table 1). As shown in the first column of the table, increasing concentrations of leptin in lower compartments increased the cell migration rate in a dose-dependent manner. This indicated that leptin had a chemotactic activity to HUVECs. Intermediate responses in other treatment combinations below the main diagonal (positive gradients, i.e., lower > upper) also demonstrated that leptin had a chemotactic effect to HUVECs. In addition, along the main diagonal, increasing the equal concentrations of leptin in both upper and lower compartments (i.e., no concentration difference) slightly increased the cell migration rate. It indicated that leptin also slightly induced chemokinesis of HUVECs.
To elucidate the effect of leptin on angiogenic activity of HUVECs, the tube formation assay was performed. As the concentration of leptin was increased, enhanced tube formation was noted (Figure 7A). Significant promotion was observed at 100 ng/ml leptin (40% increase) which was almost equal to that observed in treatment with 10 ng/ml VEGF165 (41% increase). Addition of 100 ng/ml leptin induced marked change in HUVEC morphology, with structural rearrangements leading to formation of capillary-like networks (Figure 7B).
Immunocytochemical localization of leptin in cultured MC3T3-E1 (
The expression of MMP-2 and MMP-9 in HUVECs was evaluated with gelatin SDS-PAGE zymography. MMP-9 production was slightly increased by treatment with leptin (0.1–100 ng/ml) compared with control (Figures 8A and 8B). However, those changes were less than 10% of control value and not statistically significant. In contrast, treatment with 10 ng/ml VEGF165 significantly increased MMP-9 production (Figures 8A and 8B).
Both active (67 kD) and latent (72 kD) forms of MMP-2 were expressed in HUVECs. Total MMP-2 production (active and latent MMP-2) was significantly increased by treatment with leptin (0.1–100 ng/ml) or 10 ng/ml VEGF165 (Figures 8A and 8C). The amount of latent MMP-2 was not affect by treatment with leptin or VEGF165. In contrast, MMP-2 activation was significantly enhanced by treatment with leptin (0.1–100 ng/ml) in a dose-dependent manner (Figures 8A and 8C).
Effect of leptin on proliferation of HUVECs. Cells were seeded into 24-well culture plates at a cell density of 3 × 103 cells/well. After 6 hr, medium was changed to HuMedia-EB2 containing 1% FBS and various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of recombinant human leptin or 10 ng/ml VEGF165. After 4 days of culture, the effect of leptin on HUVEC proliferation was analyzed using a crystal violet staining method. Treatment with leptin promoted proliferation of HUVECs in a dose-dependent manner. Statistically significant difference was analyzed compared with control (no leptin). ∗ p<0.05; ∗∗ p<0.01; ∗∗∗ p<0.001.
Discussion
Leptin, a 16-kD circulating hormone secreted by various tissues and organs, is known to regulate osteoblast differentiation and bone formation. To better understand the role of leptin in bone formation, we investigated leptin expression in endochondral ossification by IHC. Leptin was expressed in the secondary ossification center of femora of 1-week-old mice as well as the primary ossification center of 15-day-old embryos. Interestingly, high levels of leptin were expressed in hypertrophic chondrocytes adjacent to invading capillary blood vessels in primary and secondary ossification centers. In contrast, hypertrophic chondrocytes that were not adjacent to blood vessels were completely negative for leptin. This coexistent localization of leptin and newly formed blood vessels gave us the impression that leptin might have some effect on angiogenesis in endochondral ossification. Hoggard et al. (1997) also demonstrated that high levels of leptin were expressed in the fetal bone and/or cartilage, especially in fetal structures undergoing ossification in 14.5-day-old mouse embryos. Our findings were consistent with Hoggard's findings.
In endochondral ossification, invasion of blood vessels (angiogenesis) into hypertrophic cartilage precedes cartilage resorption and bone formation by osteoblasts (Trueta 1963). Therefore, it has been assumed that vascular invasion is crucial for endochondral ossification, but little is known about how this vascularization is regulated. Some studies have demonstrated that chondrocytes produce a number of factors that positively or negatively regulate angiogenesis (Alini et al. 1996). For example, vascular endothelial growth factor (VEGF), which is known to promote the proliferation, chemotaxis, and tube formation of endothelial cells, immunolocalizes in growth plate cartilage and is an essential coordinator of angiogenesis and bone formation in endochondral ossification (Gerber et al. 1999; Harper and Klagsbrun 1999; Horner et al. 1999; Garcia-Ramirez et al. 2000). Other studies showed that endothelial cell-stimulating angiogenic factor (ESAF) released by chondrocytes during calcification was an activator of matrix metalloproteinases (MMPs) that degrade extracellular matrix components (Brown and McFarland 1992). Furthermore, mice with the homozygous gelatinase B/MMP-9 null mutation undergo a delayed process of vascularization and ossification, which also suggests the implication of MMPs and their inhibitors in the neovascularization process in bone formation (Vu et al. 1998). Therefore, chondrocytes in hypertrophic and calcifying cartilage are considered to regulate angiogenesis in endochondral ossification by producing and secreting a variety of factors. To investigate the possibility that leptin is one of these factors, we performed RT-PCR analysis, Western blotting, and IHC analysis using cultured MCC-5 and MC3T3-E1 cell lines. Both MCC-5 and MC3T3-E1 were shown to produce and secrete leptin protein in vitro. We also detected the expression of mRNA of OB-Rb, which is the most functional isoform in leptin receptors, in MCC-5 and MC3T3-E1. However, leptin had no significant effect on the proliferation or production of extracellular matrix of these cell lines. These results suggested that leptin produced and secreted by osteoblasts and chondrocytes is not an autocrine or paracrine factor.
Checkerboard design used to distinguish between chemotaxis and chemokinesis a
Values are expressed as migrated cell number/field (×200)/6 hr (mean ± SE).
Next, to elucidate the possible role of leptin in angiogenesis in endochondral bone formation, the effects of leptin on the proliferation and function of HUVECs were investigated. These experiments revealed that leptin enhanced the proliferation, chemotaxis, and tube formation of HUVECs. These data were consistent with those of previous reports (Bouloumie et al. 1998; Sierra-Honigmann et al. 1998). Interestingly, the total amount of MMP-2 secreted by endothelial cells was elevated by treatment with leptin compared with control. Moreover, the activation of MMP-2 was also enhanced by leptin in a dose-dependent manner. MMP-2 was reported to be involved in migration, morphogenesis, and tube formation of vascular cells (Schnapeer et al. 1993; Haas et al. 1998; Zhu et al. 2000). From these facts, it is strongly suggested that leptin may stimulate angiogenesis in endochondral bone formation. The mechanisms by which leptin enhances the secretion and activation of MMP-2 are still unknown. However, judging from reports that MT1-MMP, one of the membrane-type MMPs, activates MMP-2 and that endochondral ossification and angiogenesis were impaired in mice deficient in MT1-MMP, MT1-MMP may therefore be involved in the activation of MMP-2 induced by leptin (Holmbeck et al. 1999; Zhou et al. 2000). Very recently, it was reported that leptin increased MMP-2 in cultured cytotrophoblastic cells (Castellucci et al. 2000). These facts suggest that leptin may increase and activate MMP-2 in various types of cells. However, from the fact that highest dose of leptin used in this study was about 50–100-fold as much as that in the peripheral blood, the actual leptin levels produced by chondrocytes in vitro and in vivo should be investigated carefully in the future.
Mature mice deficient in leptin (ob/ob) or its receptor (db/db) were found to have a two- to threefold higher bone mass than age-matched wild type mice (Ducy et al. 2000). This study suggested that leptin behaved as general hormone and inhibited bone formation through a hypothalamic relay. In contrast, another experiment using 4-week-old mice showed that femoral length in ob/ob mice was shorter than that in wild-type mice and was increased by peripherally administered leptin (Steppan et al. 2000). On the basis of these facts, the mechanism by which leptin regulates the bone formation appears to be very complex and age-dependent, and it remains to be elucidated more in detail. In this study we focused on the possible local action of leptin in bone formation. Our data showed that leptin was highly expressed in the hypertrophic chondrocytes adjacent to capillary blood vessels invading the primary and secondary ossification centers and that leptin promoted proliferation, migration, tube formation, and MMP2 activity of HUVECs in vitro. These findings suggested that leptin regulated the bone formation not only as a systemic hormone but also as a local factor by regulating angiogenesis in endochondral ossification. To confirm this, further IHC experiments using ob/ob or db/db mice would be very useful and should be performed in the future.
Effect of leptin on the tube formation of HUVECs on matrigel. (
Effect of leptin on MMP activity of HUVECs. HUVECs were grown in HuMedia-EG2 until they reached confluence. At confluence, the medium was changed to HuMedia-EB2 containing various concentrations (0, 0.1, 1, 10, or 100 ng/ml) of leptin or 10 ng/ml VEGF165. After 6 hr a sample from each conditioned medium was collected and analyzed using the Novex Zymogram System. Expression levels of MMPs were analyzed using NIH Image software. (
Footnotes
Acknowledgements
Supported in part by a grant-in-aid for Scientific Research (no. 12671945) from the Ministry of Education, Science, Sports and Culture of Japan.