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© 1998 American Heart Association, Inc.
Rapid Communication |
Leptin, the Product of Ob Gene, Promotes Angiogenesis
From Institut für Kardiovaskuläre Physiologie, Klinikum der J.W. Goethe-Universität (A.B., R.B.), Frankfurt/Main, Germany; Max-Planck-Institut für physiologische und klinische Forschung (H.C.A.D.), Bad Nauheim, Germany; and INSERM U317, Institut Louis Bugnard, CHU Rangueil (M.L.), Toulouse, France.
Correspondence to Dr Anne Bouloumié, Institut für Kardiovaskuläre Physiologie, Klinikum der J.W. Goethe-Universität, Theodor-Stern-Kai 7, 60596 Frankfurt/Main, Germany. E-mail bouloumie{at}em.uni-frankfurt.de
Abstract
Abstract—The adipocyte-derived cytokine leptin is thought to play a key role in the control of satiety and energy expenditure. Because adipogenesis and angiogenesis are tightly correlated during the fat mass development, we tested the hypothesis that leptin is able to modulate the growth of the vasculature. Experiments were performed using cultured human umbilical venous endothelial cells (HUVECs) and porcine aortic endothelial cells. The presence of 170-kDa endothelial leptin receptor (Ob-R) was assessed in HUVECs by Western blot analysis. Reverse transcriptase–polymerase chain reaction analysis using specific oligonucleotides for the short and long Ob-R forms further revealed the expression of both Ob-R transcripts in endothelial cells. Moreover, leptin evoked a time-dependent tyrosine phosphorylation of a number of endothelial proteins, the most prominent of which were the mitogen-activated protein kinases Erk1/2. Treatment of HUVECs with leptin led to a concentration-dependent increase in cell number that was maximal at 10 ng/mL leptin and equivalent to that elicited by vascular endothelial growth factor. This effect was associated with an enhanced formation of capillary-like tubes in an in vitro angiogenesis assay and neovascularization in an in vivo model of angiogenesis. These results indicate that leptin, via activation of the endothelial Ob-R, generates a growth signal involving a tyrosine kinase-dependent intracellular pathway and promotes angiogenic processes. We speculate that this leptin-mediated stimulation of angiogenesis might represent not only a key event in the settlement of obesity but also may contribute to the modulation of growth under physiological and pathophysiological conditions in other tissues.
Key Words: obesity • adipocyte • growth • cytokine
Angiogenesis, the formation of new blood vessels by capillary sprouting from preexisting vessels, is a major physiological event that occurs for example in the female reproductive system throughout the menstrual cycle and in pregnancy, as well as during wound healing.1 The angiogenic process is under the control of proangiogenic factors, including vascular permeability factor/vascular endothelial growth factor (VEGF), fibroblast growth factor, and antiangiogenic factors such as endostatin and angiostatin.2
Obesity is characterized by an excess of fat mass as a
consequence of adipocyte hypertrophy and
hyperplasia.3 The excessive growth of adipose tissue
requires the formation of new capillaries for proper function. Because
the development of the vascular bed in adipose tissue is tightly
connected to both number and size of adipocytes and adipose tissue
serves as an important conduit for growing blood vessels, it is
conceivable that adipocytes may modulate the growth of the vasculature
in a paracrine manner. Indeed, adipose tissue has regularly been used
in clinical applications to facilitate
revascularization and healing of compromised or
ischemic organs and tissues.4 Moreover,
immortalized preadipocyte cell lines promote the formation of highly
vascularized fat pads after injection into nude mice,5
suggesting that adipocytes per se possess proangiogenic activity.
Several recent reports have shown that adipocytes are not only sites of
energy storage but also are important sources of cytokines such
as tumor necrosis factor-
, growth factors including
VEGF,6 and lipid derivatives such as 1-butyryl
glycerol7 ; some of these are potentially involved in the
regulation of angiogenesis. Recently, leptin, the product of the ob
gene, was identified as an adipocyte-secreted
protein.8 9 10 Leptin appears to play a key role in the
regulation of body weight and more specifically on the control of food
consumption, sympathetic nervous system activation, and
thermogenesis.11 Additional roles have been proposed in
the control of the reproduction,12 13
hematopoiesis,14 and proinflammatory immune
responses.15 The expression and the plasma concentration
of leptin were found to be markedly increased in human obesity and
positively correlated to body fat mass.11 Because leptin
is secreted into the plasma, endothelial cells could be
exposed to much higher concentrations of this cytokine than
other cell types. We therefore assessed whether leptin interacts with
endothelial cells via specific receptors and thereby
modulates cell growth and angiogenic processes.
Materials and Methods
Materials
Chemicals were obtained from either Sigma (Deisenhofen, Germany)
or Merck (Darmstadt, Germany). Human recombinant leptin was provided by
Biomol (Hamburg, Germany) and human recombinant
VEGF165 was provided by Chiron Corp (Emeryville,
Calif). The murine monoclonal phosphotyrosine antibody was purchased
from Upstate Biotechnology (Lake Placid, NY), the murine monoclonal
activated mitogen-activated protein kinase (clone 12D4)
from nanoTools (Biomol, Hamburg, Germany), the goat polyclonal leptin
receptor (Ob-R) C-20 and Ob-R N-20 from Santa Cruz Biotechnology, the
agarose-conjugated antiphosphotyrosine antibody from Oncogene Science
(P-Tyr [Ab-1]-A), and the prestained protein marker from New England
Biolabs. The [P32]dCTP was purchased from
Hartmann Analytic (Braunschweig, Germany).
Cell Culture
HUVECs and porcine aortic endothelial cells
(PAECs), isolated as previously described,16 were seeded
at a density of 45 000 cells/cm2 in culture
dishes containing M-199 medium (Life Technologies) and 10% FCS
(Biochrom, Berlin, Germany) supplemented with penicillin (50 U/mL) and
streptomycin (50 µg/mL). All experiments were performed on quiescent
cells from the first passage.
Western Blot Analysis and In-Gel Protein Kinase
Assays
Cells were lysed with 1% Triton X-100 and 0.1% SDS in 20
mmol/L Tris-HCl, pH 7.4, containing (in mmol/L) NaCl 50, NaF 50,
EDTA 5, sodium pyrophosphate 20,
Na2VO3 1, and protease
inhibitors (100 µg/mL phenylmethylsulfonyl, 1 µg/mL
aprotinin, and 1 µg/mL leupeptin). The protein content of the crude
preparations was measured by the method of Bradford, using serum
albumin as standard. Thirty-microgram proteins were subjected
to SDS/PAGE gel, either directly or after immunoprecipitation using
agarose-conjugated phosphotyrosine antibody, and transferred to
nitrocellulose membranes (Schleicher and Schuel), as previously
described.17 Ponceau staining was performed to verify the
quality of the transfer and the equal amount of protein in each lane.
Proteins were detected using specific antibodies as described (see
Results) and were visualized by enhanced chemiluminescence using a
commercially available kit (Amersham). The autoradiographs were
analyzed by scanning densitometry using as software Image
master 1D (Pharmacia). For in-gel protein kinase assays, 30-µg
proteins were separated by SDS-PAGE. Myelin basic protein (0.4 mg/mL)
was used as substrate that was polymerized in the gel. After protein
renaturation, the kinase reactions were performed in the gel as
previously described.18
Analysis of the Expression of Ob-R by Reverse
Transcriptase–Polymerase Chain Reaction (RT-PCR) Analysis
Total RNA was extracted according to the method of Chomczynski
and Sacchi.19 For RT analysis, 2 µg total RNA
was incubated with 200 U reverse transcriptase (Gibco), dNTP (125
µmol/L), oligo(dT) (200 ng), and reaction buffer in a final volume of
20 µL at 37°C for 60 minutes. In some reaction mixtures, reverse
transcriptase or total RNA was omitted to determine the amplification
of contaminating genomic DNA or cDNA. After a final denaturation at
94°C for 7 minutes, 10 µL cDNA was subjected to PCR consisting of a
denaturation at 94°C for 1 minute, followed by 90 seconds of
annealing at 55°C and 90 seconds of elongation at 72°C for 32
cycles. The last cycle ended with 7 minutes of elongation at 72°C.
The primers used to amplify the long and short forms of the leptin
receptors were chosen as previously described14 : a common
forward primer (5'-GAAGGAGTGGGAAAACCAAAG-3') used in combination with a
specific reverse primer either for the long form
(5'-CATAGGTTACCTCAGTACCCTC-3'), allowing the amplification of 433 bp,
or the short form (5'-CCACCATATGTTAA-CTCTCAG-3'), allowing the
amplification of 365 bp. The PCR contained 0.4 µmol/L of each
primer, dNTP (200 µmol/L), MgCl2 (1
mmol/L) reaction buffer, and 2.5 U Taq polymerase (Promega) in a final
volume of 50 µL. The amplified cDNAs were size-fractionated by
agarose gel electrophoresis, visualized under UV with an ethidium
bromide staining, transferred to a nylon membrane (porablot NY amp,
Macherey-Nagel) and hybridized with
32P–end-labeled oligonucleotides
specific for the long form (5'-CTGTGGTCTCTCTACTTTCAAC-3') or the short
form (5'-CTAATCATGATCACTACAGATG-3'). Autoradiographs were then exposed
for 4 to 8 hours.
Viability Assays
Cell viability was assessed by the use of an MTT
(3-[4,5-dimethylthiazol-2-yl]-2,5-dephenyltetrazolium bromide) assay
allowing the quantification of viable cells. HUVECs were seeded in
96-well plates and incubated the next day with serum-deprived medium
supplemented with 0.1% BSA (Gibco Life Technologies,
Eggenstein-Leopolshafen, Germany) for the next 72 hours, in the
presence of increasing concentrations of leptin or
VEGF165 (0.1 to 100 ng/mL). Twenty microliters of
MTT (5 mg/mL) was then added to 200 µL medium culture and incubated
for 3 hours. At the end of the incubation period, the medium was
removed and the converted dye solubilized with acidic isopropanol.
Absorbance of converted dye was measured at a wavelength of 570 nm with
background subtraction at 630 nm.
Proliferation Assays
The total cell number was determined in HUVECs seeded in 24
wells and incubated the next day with serum-deprived medium
supplemented with 0.1% BSA or with 1% FCS for the next 72 hours in
the presence or absence of leptin. At the time indicated, the cells
were counted after trypsinization using a cell counter (CASY1,
Schärfe System).
Apoptosis Assays
HUVECs were cultured in 12-well plates and incubated the next
day with serum-deprived medium supplemented with 0.1% BSA for 24, 48,
or 72 hours, in the presence of increasing concentrations of leptin (1
to 100 ng/mL). The plates were centrifuged (2000g,
10 minutes). Thereafter, the medium was removed, cells were fixed in
4% formaldehyde, and stained with DAPI (4',6-diamino-2-phenylindole;
0.2 µg/mL in 10 mmol/L Tris-HCl [pH 7], 10 mmol/L EDTA,
and 100 mmol/L NaCl) for 30 minutes. Nuclei were analyzed
by fluorescence microscopy, and apoptotic cells were
counted in a double-blind manner.
In Vitro Angiogenesis Assays
The microcarrier-based fibrin gel angiogenesis assay was
performed as previously described.20 Trypsinized
PAECs were allowed to attach onto cytodex-2 microcarrier beads for 4
hours at 37°C (Sigma) in M-199 supplemented with 10% FCS and
subsequently were grown to confluence for 48 hours. Porcine fibrinogen
(Sigma) was dialyzed overnight against PBS (pH 7.0; 10 mmol/L
phosphate, 140 mmol/L sodium chloride) and then diluted to 1 mg/mL
in PBS, pH 7.0. Packed microcarrier beads (40 µL) were pipetted into
2 mL of fibrinogen solution in 35-mm Petri dishes, and gels were
allowed to polymerize for 30 minutes after addition of 0.625 U/mL
thrombin. The fibrin gels were then equilibrated in M-199 supplemented
with 1 mg/L insulin, 1 mg/mL transferrin, 1 µg/L selenium, 50 U/mL
penicillin, and 50 µg/mL streptomycin for 60 minutes at 37°C, and
fresh medium was subsequently added on top of the gels. The number of
positive beads for angiogenesis was determined under the microscope, by
observers who were blind to the experimental conditions, in 3 randomly
chosen fields by counting the number of capillary-like tubes of a
length >150 µm. For angiogenesis assays with HUVECs,
trypsinized cells were plated directly on the surface of the fibrin
gels and cultured in M-199 supplemented with 0.1% BSA. Where
appropriate, leptin was added to both the fibrinogen solution prior to
polymerization and the medium bathing the cells.
Chick Chorioallantoic Membrane (CAM) Assays
All experiments with chick embryos (n=28) were carried out in
ovo. On 3-day old embryos, a window of 7 to 10 mm in diameter was
cut into the eggshell, resealed with parafilm, and further
incubated until day 9. For the CAM assay, 70 µL of a sterile 1%
methylcellulose solution were mixed with 70 µL of a leptin solution
in sterile water. Seven-microliter aliquots of this mixture, each
containing a total amount of either 200 ng, 1 µg, or 3 µg leptin
were pipetted onto bacteriological-grade Petri dishes, air-dried for 2
hours, and the resulting discs placed onto 9 day-old CAMs for 2 days.
Two to 3 discs were placed onto each CAM about 10 mm apart.
Control discs contained sterile water only. Evaluation of the CAMs was
performed on 11 day-old CAMs. To better visualize the vascular system
of the CAM, 20% Luconyl Blue in PBS (BASF, Ludwigshafen) was injected
into a vitelline vein using glass capillaries. Photographs were taken
using a Zeiss SV6 stereomicroscope with camera adapter. The angiogenic
response was assessed as positive after appearance of a significantly
increased vascularization under or in the vicinity of the disc.
Statistics
Data are expressed as mean±SEM. Statistical analyses
were performed by 1-way ANOVA followed by Bonferroni t test.
Values of P<0.05 were considered statistically
significant.
Results
To determine whether endothelial cells express
leptin receptors, Western blot and RT-PCR analysis were
performed on whole protein and RNA extracts from cultured HUVECs.
Antibodies directed against the carboxyl or the amino terminus of the
human leptin receptor recognized a 170-kDa protein (Figure 1A
), the size of which is
consistent with the human leptin receptor.21 Using
specific primers for Ob-Rb and Ob-Ra, cDNAs of the expected length were
amplified from reverse-transcriptase reaction performed with HUVEC RNA
(Figure 1B). These cDNA fragments were specifically recognized by the
respective Ob-Rb and Ob-Ra radiolabeled
oligonucleotides (Figure 1C). Sequence analysis
confirmed the identities of the cDNA fragments as Ob-Rb and
Ob-Ra.13 These results indicate that cultured HUVECs
express at least 2 forms of the leptin receptor, Ob-Ra and Ob-Rb.
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To further characterize the functionality of Ob-R, Western blot
analysis was performed on whole protein extracts from HUVECs
(30 µg) treated with human recombinant leptin (10 ng/mL) at different
time points (2, 5, 10, or 30 minutes). Leptin enhanced, in a
time-dependent manner, the tyrosine phosphorylation of
endothelial proteins, the most prominent of which
exhibited molecular weights of 180, 170, 130, 91, 86, 76, 60, 56, 52,
44, and 42 kDa (Figure 2A).
Immunoprecipitation using a phosphotyrosine antibody, followed by
Western blot analysis, confirmed this finding (data not shown).
Western blot analysis using antibody directed against the
activated form of extracellular signal-regulated kinases (Erk1
and Erk2) showed that both isoforms are time-dependently
activated in leptin-treated HUVECs (10 ng/mL) (Figure 2B). This
stimulatory effect of leptin on Erk1/2 was confirmed by in-gel kinase
assays using myelin basic protein as substrate (data not shown). These
results demonstrate that endothelial Ob-R is
functionally active and linked to activation of tyrosine
kinase-dependent pathways. Because activation of
mitogen-activated protein kinases, and more specifically, the
Erk1 and Erk2 isoforms, are involved in the control of proliferation
and/or differentiation in various cell types, we studied the effect of
leptin on endothelial cell growth and viability.
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The leptin effect on cell viability was assessed on
endothelial cells cultured with increasing
concentrations of human recombinant leptin (0.1 to 100 ng/mL) for 72
hours in a serum-deprived medium. The number of viable cells per well
in the presence of leptin was compared with control and
VEGF165-treated cells. As shown in Figure 3A, the number of viable cells maintained
in serum-deprived medium for 72 hours decreased slightly compared with
initial cell number, although this decrease did not reach significance.
Leptin dose-dependently increased the number of viable cells. The
maximal effect observed with 10 ng/mL leptin was not significantly
different from that elicited by 10 ng/mL VEGF165
and was associated with a significant increase in the initial cell
number (P<0.05, n=3). This effect was also observed when
cells were cultured in medium supplemented with 1% FCS (Figure 3B). The total number of cells was significantly increased with 10
ng/mL leptin compared with the initial cell number or after the 72-hour
incubation period. To determine whether this survival/proliferative
effect could be linked to an antiapoptotic effect of leptin,
the apoptotic rate was determined by visual analysis of
DAPI-stained cells. The apoptotic rate in untreated cells was
low and not affected by the length of the incubation period in
serum-deprived medium (1.9±0.3% for 24 hours, 2.4±0.7% for 48
hours, and 2.2±0.8% for 72-hour incubation period). Leptin treatment
did not modify the percentage of apoptotic nuclei detected
(2.2±0.3 in 10 ng/mL leptin versus 2.2±0.8 in control cells,
n=3).
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The effect of leptin was studied using in vitro (3D fibrin gels) and in
vivo (CAM) angiogenesis assays. As shown in Figure 4, in HUVECs cultured on fibrin gels,
addition of leptin (100 ng/mL) induced marked changes in HUVEC
morphology (Figure 4B and 4D) compared with control (Figure 4A and 4C),
with structural rearrangements leading to the formation of
capillary-like networks. In a second approach to determine whether the
leptin-promoted angiogenesis was also evident on "adult"
macrovascular endothelial cells, experiments were
performed on microcarrier beads coated with PAECs and included in a
fibrin gel. As shown in Figure 5, 7 days
after fibrin polymerization, addition of leptin (100 ng/mL) in both the
medium and the gel increased the number as well as the length of the
capillary-like structures derived from the cell-coated microcarriers
(Figure 5B and 5E). A similar angiogenic effect was observed with 10
ng/mL leptin (data not shown). A 4- to 5-fold increase in the number of
microcarriers that developed capillary-like tubes of >150 µm in
length was observed with 100 ng/mL leptin.
VEGF165 (100 ng/mL) elicited a comparable
angiogenic effect (Figure 5C).
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Experiments were then performed in vivo on CAMs using methylcellulose
discs containing 0.2, 1, and 3 µg leptin. After 48 hours of contact
with the CAM, the discs containing 1 and 3 µg leptin were associated
with a significant and marked stimulation of neovascularization
(Table). A representative
area of the CAM with a control disc and a disc loaded with 3 µg
leptin is shown in Figure 6.
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Discussion
In the present study, we have demonstrated the presence of functionally active leptin receptors on cultured HUVECs, linked to tyrosine kinase-dependent intracellular pathways. The stimulation of endothelial cells by leptin led to an increase in cell proliferation and/or survival and elicited a marked enhancement of angiogenesis.
Several forms of Ob-R, which differ in their cytoplasmic domains, have
been described on the basis of mRNA analysis. The receptor with
the longest cytoplasmic domain (Ob-Rb) shares sequence similarity with
the corresponding regions of the leukemia inhibitory factor
(LIF) receptor -chain and gp130, whereas the short form
(Ob-Ra) contains the conserved box 1-motif of the hematopoietin
receptor family.21 Ob-Rb receptors, originally described
in the central area, are far less abundant in peripheral
sites than the short Ob-Ra form.21 It was recently
reported that endothelial cells from the human brain
capillaries exhibited specific and saturable binding sites to leptin,
which were thought to play a role in the leptin endocytosis at the
blood-brain barrier.22 The present study extends this
first observation to human macrovascular endothelial
cells and delivers by the use of Western blot and RT-PCR
analysis a characterization of the type of Ob-R expressed on
endothelial cells: Ob-Ra and Ob-Rb. Ob-Rb receptors
have been suggested to play a role in the leptin-mediated control of
cell differentiation, whereas Ob-Ra receptors may be involved in the
leptin endocytosis.21 It is therefore conceivable that
endothelial cells, in addition to their ability to
internalize leptin, probably through Ob-Ra, may also be a target for
the modulating effect of leptin on proliferation and differentiation
through the Ob-Rb form. In particular, HUVECs might be affected by
leptin, because leptin was shown to be produced in the human
placenta23 and found in the venous cord
blood.24
The functionality of endothelial Ob-R was demonstrated by the leptin-induced increase in the tyrosine phosphorylation of several endothelial proteins. The molecular weights from some of them were consistent with those of proteins already described to be activated under leptin treatment in other target cell types, such as members of the Janus family (JAK1 and JAK2), the STAT family (STAT1 and STAT3), and the mitogen-activated protein kinase family Erk1/2.25 The finding of a leptin-induced activation of Erk1/2, as assessed with specific antibody directed against the activated Erk1/2 and in-gel kinase activity, led to the hypothesis that leptin may stimulate endothelial cell growth. Our results clearly indicate that leptin in serum-deprived medium produced a marked increase in the number of viable endothelial cells, whereas the number of cells undergoing apoptosis was unaffected. Moreover, in the presence of serum, a significant increase in total cell number was observed. Thus, leptin treatment of endothelial cells is associated with proliferation and/or survival. The maximal effect was elicited by a physiological plasma concentration of leptin (10 ng/mL) and was equivalent to that evoked by VEGF165 in the same concentration range. Given that VEGF165 is considered to be a major proangiogenic factor, this observation underlines the significance of leptin as an endothelial growth factor. Proliferative effects of leptin have already been reported on hematopoietic or embryonic fibroblast cell lines,14 25 26 but this is the first report on vascular cells. Proliferation of endothelial cells constitutes one key event in the complex angiogenic process.2 Angiogenesis starts by cell-mediated degradation of the basement membrane, followed by the migration and the proliferation of endothelial cells. The morphogenesis of the cells into capillary tubes finishes this process. Using 2 different in vitro models of angiogenesis (ie, endothelial cell–coated microcarrier-induced and monolayer-induced formation of capillary-like tubes in fibrin gels), we demonstrated that the leptin-induced proliferation and/or survival is associated with the enhanced formation of capillary-like tubes in a 3D matrix in a similar manner to that elicited by VEGF165. Moreover, in vivo angiogenic assay using CAMs showed clearly that leptin enhanced the formation of new blood vessels in vivo. This positive angiogenic effect was observed using higher concentrations of leptin than those that were effective in vitro. This is probably a result of leptin diffusion being hindered by methylcellulose. However, our observation is in accordance with a study published during the preparation of this article that describes an angiogenic effect of leptin in the rat corneas.27
Thus, our results demonstrate that leptin could be a potent modulator of the angiogenic process. This peripheral action of leptin, observed in concentrations usually found in plasma or in adipose tissues, opens new perspectives toward links between this adipocyte-secreted cytokine and angiogenesis. It may be argued that this effect of leptin is not of great physiological relevance, because animals with a defect in the leptin system (Ob and Db mice)11 seem to exhibit no manifest alteration in angiogenesis. However, the use of the genetically manipulated mouse as animal model has clearly evidenced that those mice quite often develop compensatory mechanisms to overcome the genetic defect. It is therefore likely that the defect in the leptin system in the Ob or Db mice might be compensated by other angiogenic factors. Experimental models in which angiogenesis is provoked, eg, unilateral ligation of femoral artery, will help to elucidate the modulatory role of leptin in the control of angiogenic processes.
Studies of fetal adipose tissue have indicated a remarkable spatial and temporal relationship between adipogenesis and vasculogenesis.28 In situations of continuing adipose tissue growth (in the adult and during the settlement of obesity), it is documented that angiogenesis is present.28 The factors influencing fat pad development, at both the adipocyte and endothelial cell levels, are at an early stage of characterization. Because obesity in humans is associated with an elevation of leptin in the plasma as well as adipose tissue, it is tempting to speculate that the leptin-mediated cross talk between adipocytes and endothelial cells promotes angiogenesis, which in turn participates in the additional increment of the adipose mass and ultimately to a progressive aggravation of the obese state. The present study opens a promising perspective concerning future investigations of leptin-dependent modulation of angiogenesis more specifically in adipose tissue during the settlement of obesity but also in other tissues under physiological and pathophysiological conditions.
Acknowledgments
This study was supported by the Deutsche Forschungsgemeinschaft (SFB 553, B5). The expert technical assistance of Isabel Winter is gratefully acknowledged. We are grateful to Dr I. Fleming for helpful discussion and advice.
Received June 10, 1998; accepted September 29, 1998.
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G. J. Mick, X. Wang, and K. McCormick White Adipocyte Vascular Endothelial Growth Factor: Regulation by Insulin Endocrinology, March 1, 2002; 143(3): 948 - 953. [Abstract] [Full Text] [PDF]
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P. Vigano, E. Somigliana, R. Matrone, A. Dubini, C. Barron, M. Vignali, and A. M. di Blasio Serum Leptin Concentrations in Endometriosis J. Clin. Endocrinol. Metab., March 1, 2002; 87(3): 1085 - 1087. [Abstract] [Full Text] [PDF]
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 K. Kume, K. Satomura, S. Nishisho, E. Kitaoka, K. Yamanouchi, S. Tobiume, and M. Nagayama Potential Role of Leptin in Endochondral Ossification J. Histochem. Cytochem., February 1, 2002; 50(2): 159 - 170. [Abstract] [Full Text] [PDF]
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 A. Wiecek, F. Kokot, J. Chudek, and M. Adamczak The adipose tissue--a novel endocrine organ of interest to the nephrologist Nephrol. Dial. Transplant., February 1, 2002; 17(2): 191 - 195. [Full Text] [PDF]
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 D. R. Harder, C. Zhang, and D. Gebremedhin Astrocytes Function in Matching Blood Flow to Metabolic Activity Physiology, February 1, 2002; 17(1): 27 - 31. [Abstract] [Full Text] [PDF]
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A. Glasow, W. Kiess, U. Anderegg, A. Berthold, A. Bottner, and J. Kratzsch Expression of Leptin (Ob) and Leptin Receptor (Ob-R) in Human Fibroblasts: Regulation of Leptin Secretion by Insulin J. Clin. Endocrinol. Metab., September 1, 2001; 86(9): 4472 - 4479. [Abstract] [Full Text] [PDF]
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A. Bouloumie, C. Sengenes, G. Portolan, J. Galitzky, and M. Lafontan Adipocyte Produces Matrix Metalloproteinases 2 and 9: Involvement in Adipose Differentiation Diabetes, September 1, 2001; 50(9): 2080 - 2086. [Abstract] [Full Text] [PDF]
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M. Winnicki, B. G. Phillips, V. Accurso, P. van de Borne, A. Shamsuzzaman, K. Patil, K. Narkiewicz, and V. K. Somers Independent Association Between Plasma Leptin Levels and Heart Rate in Heart Transplant Recipients Circulation, July 24, 2001; 104(4): 384 - 386. [Abstract] [Full Text] [PDF]
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H. Yamashita, J. Shao, T. Ishizuka, P. J. Klepcyk, P. Muhlenkamp, L. Qiao, N. Hoggard, and J. E. Friedman Leptin Administration Prevents Spontaneous Gestational Diabetes in Heterozygous Leprdb/+ Mice: Effects on Placental Leptin and Fetal Growth Endocrinology, July 1, 2001; 142(7): 2888 - 2897. [Abstract] [Full Text] [PDF]
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 F. LARCHER, M. DEL RIO, F. SERRANO, J. C. SEGOVIA, A. RAMIREZ, A. MEANA, A. PAGE, J. L. ABAD, M. A. GONZALEZ, J. BUEREN, et al. A cutaneous gene therapy approach to human leptin deficiencies: correction of the murine ob/ob phenotype using leptin-targeted keratinocyte grafts FASEB J, July 1, 2001; 15(9): 1529 - 1538. [Abstract] [Full Text] [PDF]
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 H. Koshiba, J. Kitawaki, H. Ishihara, N. Kado, I. Kusuki, K. Tsukamoto, and H. Honjo Progesterone inhibition of functional leptin receptor mRNA expression in human endometrium Mol. Hum. Reprod., June 1, 2001; 7(6): 567 - 572. [Abstract] [Full Text] [PDF]
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G. De Placido, C. Alviggi, C. Carravetta, M.L. Pisaturo, V. Sanna, M. Wilding, G.M. Lord, and G. Matarese The peritoneal fluid concentration of leptin is increased in women with peritoneal but not ovarian endometriosis Hum. Reprod., June 1, 2001; 16(6): 1251 - 1254. [Abstract] [Full Text] [PDF]
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J. A. Pasco, M. J. Henry, M. A. Kotowicz, G. R. Collier, M. J. Ball, A. M. Ugoni, and G. C. Nicholson Serum Leptin Levels Are Associated with Bone Mass in Nonobese Women J. Clin. Endocrinol. Metab., May 1, 2001; 86(5): 1884 - 1887. [Abstract] [Full Text]
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B. Winters, Z. Mo, E. Brooks-Asplund, S. Kim, A. Shoukas, D. Li, D. Nyhan, and D. E. Berkowitz Reduction of obesity, as induced by leptin, reverses endothelial dysfunction in obese (Lepob) mice J Appl Physiol, December 1, 2000; 89(6): 2382 - 2390. [Abstract] [Full Text] [PDF]
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P. Trayhurn, J. S. Duncan, A. M. Wood, and J. H. Beattie Metallothionein gene expression and secretion in white adipose tissue Am J Physiol Regulatory Integrative Comp Physiol, December 1, 2000; 279(6): R2329 - R2335. [Abstract] [Full Text] [PDF]
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K. Linnemann, A. Malek, R. Sager, W. F. Blum, H. Schneider, and C. Fusch Leptin Production and Release in the Dually in VitroPerfused Human Placenta J. Clin. Endocrinol. Metab., November 1, 2000; 85(11): 4298 - 4301. [Abstract] [Full Text]
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M. C. Henson and V. D. Castracane Leptin in Pregnancy Biol Reprod, November 1, 2000; 63(5): 1219 - 1228. [Abstract] [Full Text]
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 G. Fantuzzi and R. Faggioni Leptin in the regulation of immunity, inflammation, and hematopoiesis J. Leukoc. Biol., October 1, 2000; 68(4): 437 - 446. [Abstract] [Full Text]
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H Mix, A Widjaja, O Jandl, M Cornberg, A Kaul, M Goke, W Beil, M Kuske, G Brabant, M P Manns, et al. Expression of leptin and leptin receptor isoforms in the human stomach Gut, October 1, 2000; 47(4): 481 - 486. [Abstract] [Full Text] [PDF]
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R. F. Gariano, A. K. Nath, D. J. D’Amico, T. Lee, and M. R. Sierra–Honigmann Elevation of Vitreous Leptin in Diabetic Retinopathy and Retinal Detachment Invest. Ophthalmol. Vis. Sci., October 1, 2000; 41(11): 3576 - 3581. [Abstract] [Full Text]
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G. Matarese, C. Alviggi, V. Sanna, J. K. Howard, G. M. Lord, C. Carravetta, S. Fontana, R. I. Lechler, S. R. Bloom, and G. De Placido Increased Leptin Levels in Serum and Peritoneal Fluid of Patients with Pelvic Endometriosis J. Clin. Endocrinol. Metab., July 1, 2000; 85(7): 2483 - 2487. [Abstract] [Full Text]
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J. Alfer, F. Muller-Schottle, I. Classen-Linke, U. von Rango, L. Happel, K. Beier-Hellwig, W. Rath, and H. M. Beier The endometrium as a novel target for leptin: differences in fertility and subfertility Mol. Hum. Reprod., July 1, 2000; 6(7): 595 - 601. [Abstract] [Full Text] [PDF]
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W. A. Banks, C. M. Clever, and C. L. Farrell Partial saturation and regional variation in the blood-to-brain transport of leptin in normal weight mice Am J Physiol Endocrinol Metab, June 1, 2000; 278(6): E1158 - E1165. [Abstract] [Full Text] [PDF]
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A. Z. Zhao, M. M. Shinohara, D. Huang, M. Shimizu, H. Eldar-Finkelman, E. G. Krebs, J. A. Beavo, and K. E. Bornfeldt Leptin Induces Insulin-like Signaling That Antagonizes cAMP Elevation by Glucagon in Hepatocytes J. Biol. Chem., April 6, 2000; 275(15): 11348 - 11354. [Abstract] [Full Text] [PDF]
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G. Uckaya, M. Ozata, A. Sonmez, C. Kinalp, T. Eyileten, N. Bingol, B. Koc, F. Kocabalkan, and I. C. Ozdemir Is Leptin Associated with Hypertensive Retinopathy? J. Clin. Endocrinol. Metab., February 1, 2000; 85(2): 683 - 687. [Abstract] [Full Text]
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P. Sinnaeve, O. Varenne, D. Collen, and S. Janssens Gene therapy in the cardiovascular system: an update Cardiovasc Res, December 1, 1999; 44(3): 498 - 506. [Abstract] [Full Text] [PDF]
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