Molecular Human Reproduction, Vol. 6, No. 9, 811-819,
September 2000
© 2000 European Society of Human Reproduction and Embryology
Uterine physiology |
Adrenomedullin is an autocrine regulator of endothelial growth in human endometrium
1 Nuffield Department of Obstetrics and Gynaecology and 2 Molecular Angiogenesis Laboratory, Imperial Cancer Research Fund, Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Oxford OX3 9DU, UK
Abstract
Human endometrium is a mucosa served by a microvascular blood supply that involves benign angiogenesis under the control of ovarian steroids throughout reproductive life. Adrenomedullin is a multifunctional 52-amino acid peptide involved in numerous physiological and pathological processes, including angiogenesis, growth regulation, differentiation, vasodilation and smooth muscle relaxation. We have previously shown that adrenomedullin is present in the human uterus. To investigate further the role of adrenomedullin in human endometrial angiogenesis, a method for the isolation and culture of non-pregnant endometrial endothelium was developed. Enzymatic dispersion and `Percoll' gradient centrifugation, followed by positive selection using Ulex europaeus agglutinin-coated immunomagnetic beads, yielded pure isolates of endothelium. The cells formed a typical `cobblestone' monolayer within 5–7 days and expressed the classic endothelial markers, CD31 and von Willebrand factor. The presence of adrenomedullin immunoreactivity in endometrial endothelial cells was shown by immunohistochemistry both in vitro and in vivo. Adrenomedullin promotes growth of endothelial cells as measured by [methyl-3H] thymidine uptake. Adrenomedullin also induced cyclic AMP in endometrial endothelial cells. These results demonstrate, for the first time, that adrenomedullin is an autocrine growth factor for human endometrial endothelial cells and is thus involved in endometrial angiogenesis.
adrenomedullin/angiogenesis/endometrium/vascular endothelial growth factor
Introduction
The 52 amino acid peptide, adrenomedullin, has been shown to have a wide range of biological actions, including regulation of cell growth and differentiation (Miller et al., 1996
; Morrish et al., 1996; Isumi et al., 1998a; Kubo et al., 1998), vasodilation (He et al., 1995) and control of hormone secretion (Yamaguchi et al., 1996). We have recently shown that adrenomedullin is expressed in endometrium and is a novel angiogenic factor (Zhao et al., 1998).
Adrenomedullin was initially isolated from the human pheochromocytoma cell line, PC12 (Kitamura et al., 1993a). The adrenomedullin promoter contains several binding sites for activator protein-1 (AP-1), activator protein-2 (AP-2) and a cyclic AMP (cAMP)-regulated enhancer element (Kitamura et al., 1993b; Ishimitsu et al., 1994). Subsequent studies showed a growth promoting effect of adrenomedullin (Martinez et al., 1997; Isumi et al., 1998a) with signalling mediated via cAMP (Withers et al., 1996).
The biological actions of adrenomedullin are mediated via adrenomedullin receptors. As adrenomedullin shares some homology with calcitonin gene-related peptide (CGRP) (Kitamura et al., 1993a), it has been classified as a member of the calcitonin/CGRP/amylin peptide family. These peptides have seven transmembrane domain G-protein-coupled receptors. Binding experiments have shown that the receptor with which adrenomedullin interacts varies in different organ systems. In some, e.g. skin (Eguchi et al., 1994), it binds to its own specific receptor but in others, it interacts with the CGRP receptor (Ishizaka et al., 1994; Withers et al., 1996; Martinez et al., 1997). Molecular characterization has recently shown that the adrenomedullin and CGRP receptors are interrelated. Calcitonin-receptor-like receptor (CRLR), can function as either a CGRP receptor or an adrenomedullin receptor, depending on which receptor-activity-modifying proteins (RAMPs), are expressed. RAMPs are required to transport CRLR to the plasma membrane. RAMP1 presents the receptor at the cell surface as a CGRP receptor; RAMP2- and RAMP3-transported receptors are adrenomedullin receptors (McLatchie et al., 1998; Kamitani et al., 1999).
Although adrenomedullin receptors are present in endothelial cells (Kamitani et al., 1999) and adrenomedullin has been shown to stimulate cAMP production in the large vessel endothelial cells (Isumi et al., 1998b) and to be angiogenic in vivo (Zhao et al., 1998), it has not yet been shown to have an effect on human microvascular endothelium.
Since the discovery that the adrenomedullin gene is more highly expressed in endothelial cells than even in the adrenal medulla (Sugo et al., 1994), the peptide has come to be regarded as a secretory product of vascular endothelium. Immunohistochemistry and in-situ hybridization have shown that, apart from its expression in endothelial cells, adrenomedullin is expressed in diverse cell types from many organs, e.g. cardiomyocytes (Nishimori et al., 1997), zona glomerulosa cells (Kapas et al., 1998), macrophages (Kubo et al., 1998), astrocytes (Kuchinke et al., 1995) and fibroblasts (Isumi et al., 1998a).
Adrenomedullin is also found in cells of the reproductive system, including breast epithelium (Jahnke et al., 1997), placental trophoblast (Marinoni et al., 1998), prostatic epithelium (Libert et al., 1991) and granulosa (Abe et al., 1998) cells. Adrenomedullin mRNA (Upton et al., 1997; Cameron and Fleming, 1998) and binding sites (Upton et al., 1997) have also been found in abundance in the rat uterus. We have previously demonstrated the presence of adrenomedullin in human endometrium (Zhao et al., 1998), and this has been confirmed by other groups (Michishita et al., 1999). The purpose of the present study was to investigate whether adrenomedullin is involved in microvascular endothelial cell growth in human endometrium.
Materials and methods
Source of tissue
All tissues were collected in accordance with the requirements of the Central Oxfordshire Research Ethics Committee, UK (approval number 2519). Uteri were obtained at hysterectomy from non-pregnant women of reproductive age (30–49 years) who had a history of regular menstrual cycles (26–30 days) and had not used either oral or intrauterine contraception and had not received any hormonal treatment for at least 6 months prior to surgery. Uteri were bisected: one half was used for cell isolation and preparation of frozen samples, the other half was processed for histological examination.
Samples of decidua were obtained from first trimester therapeutic abortions of apparently healthy pregnancies. Gestational age was 6–12 weeks. Decidual tissue was separated by morphological examination.
Cell isolation and culture
The method used in the present study was based on that developed for the isolation of decidual endothelium (Grimwood et al., 1995). Incubations were performed at 37°C in a 5% CO2 humidified atmosphere. Washes were carried out using phosphate-buffered saline (PBS) containing 0.1% bovine serum albumin (BSA).
Tissue was collected into McCoy's 5A medium (Gibco, Paisley, UK) containing 10% fetal calf serum (FCS; Autogen Bioclear, Calne, UK) and antibiotics (penicillin 100 IU/ml, streptomycin 100 µg/ml, gentamicin 50 µg/ml, fungizone 1 µg/ml) and stored overnight at 4°C. Then tissue was cut into 1 mm2 pieces and incubated for 2 h, with continuous stirring, in 2 mg/ml collagenase type 1 (Worthington, Lakewood, NJ, USA) in McCoy's medium containing 10% FCS and antibiotics. The resulting cell suspension was filtered through a sterile 100 µm mesh (Nylon Bolting Cloth; Lockertex, Warrington, UK) to remove any undigested material, centrifuged and washed repeatedly with medium.
The endothelial cell population was purified using discontinuous `Percoll' gradient centrifugation (Amersham Pharmacia Biotech, Little Chalfont, UK). Cells were resuspended in 20% Percoll in PBS/BSA and underlayered with 40 and 60% `Percoll' fractions. Separation was achieved by centrifugation at 670 g for 30 min and an enriched endothelial population was recovered from the 40/60% interface. After washing, cells were resuspended in McCoy's medium/10%FCS.
Lectin Ulex europaeus agglutinin-1 (UEA-1; Sigma, Poole, UK) was covalently bound to tosylactivated `Dynabeads' M-450 (Dynal, Oslo, Norway) as previously described (Jackson et al., 1990). Equal volumes of UEA 1 (0.2 mg/ml in 0.5 mol/l borate buffer, pH 9.6) and Dynabeads (2x107 beads/ml) were incubated overnight at 4°C with end-over-end rotation. After coating, the beads were washed three times in PBS using a magnetic particle concentrator (Dynal), resuspended at a final concentration of 2x107 beads/ml in PBS/BSA, and stored at 4°C until use.
Positive selection of endothelial cells using UEA-1 coated beads was performed by incubation for 10 min at 40C using end-over-end rotation. Cells were incubated with beads at a bead:cell ratio of 3:1 in 0.5 ml volumes. Rosetted cells were collected using a magnetic particle concentrator, followed by disposal of the bead-free supernatant. This process was repeated five times, interspersed by 1 min washes in McCoy's/1% FCS. Partial release of beads from the cells with fucose was not performed as they became detached in culture, leading to a virtually bead-free cell population at the second passage.
Purified endothelial cells (the number of isolated cells varied between different preparations and was 5000–17 000 cells) were initially seeded into 35 mm2 culture dishes at 6000 cells/dish, precoated with 5 µg/cm2 collagen type IV (Roche Diagnostics, Lewes, UK) and supplemented with 2 ml McCoy's culture medium containing 50% human serum (HS; pooled male AB serum; Autogen Bioclear, UK), 5 ng/ml vascular endothelial growth factor (VEGF: R&D systems, Abingdon, UK), 10 000 IU heparin (Sigma), magnesium sulphate (2 mg/ml; Sigma), hydrocortisone (0.5 µg/ml; Sigma) and 3-isobutyl-1-methyl xanthine (IBMX; 3.3x10–4 mol/l; Sigma). Cultures were incubated at 37°C in a 5% CO2 humidified atmosphere and the medium replaced every 1–2 days. Cells were passaged 1:2 at confluence by release with trypsin/EDTA (Sigma). Decidual endothelial cells were isolated and cultured using the same method.
Human umbilical vein endothelial cell (HUVEC) cultures and human dermal microvascular endothelial cells (HDMEC) were obtained from Clonetics (Biowhittaker, Wokingham, UK). HUVECs were seeded either into plastic polylysine coated culture dishes or precoated with 5 µg/cm2 collagen type IV (Roche Diagnostics, Lewes, UK) (both were equally satisfactory). HDMECs were only seeded into culture dishes precoated with collagen type IV, as poly-L-lysine appeared to be detrimental to cell growth. HUVECs were grown either in Dulbecco's modified Eagle's medium (DMEM) supplemented with 20% FCS (Autogen Bioclear), 30 ng/ml ECGS (Sigma), 5 µg/ml gentamycin (Sigma), 90 µg/ml heparin (Sigma; Cat No H3149), 1 µg/ml fungizone (Gibco) or in EGM-2MV BulletKit medium (Biowhittaker, Walkersville, MD, USA). HDMECs were grown in EGM-2MV BulletKit medium (Biowhittaker).
Immunohistochemical characterization of endometrial endothelial cells
Cultured cells
Endometrial endothelial cells were grown to confluence on collagen IV precoated 8-well Permanox slides (Nunc, Naperville, IL, USA). Prior to immunohistochemistry cultures were fixed in ice-cold acetone/methanol for 3–5 min and air-dried for a minimum of 2 h. Slides were stored at –20° C prior to staining.
Cryostat sections
Fragments of uterine tissues containing both endometrium and myometrium were snap-frozen in liquid nitrogen and stored at –70°C. Frozen sections 8–10 µm were cut on a Leica Kryostat 1720 Digital cryostat, allowed to dry at room temperature and then fixed for 5 min in 4% ice-cold paraformaldehyde solution in PBS, rinsed three times for 2 min in PBS, washed for 5 min in 70% ethanol, for 5 min in absolute ethanol and stored at 4°C in absolute ethanol. Before immunohistochemistry, frozen sections were dried, washed for 5 min in distilled water and rinsed in buffer 1 (100 mmol/l Tris–HCl, 150 mmol/l NaCl pH 7.5) for 5 min.
Paraffin-embedded tissue sections
Formalin-fixed, paraffin-embedded specimens of normal uterine tissues were selected from archival files of The Department of Cellular Pathology, John Radcliffe Hospital, University of Oxford, Oxford, UK. Sections 4 µm were cut and dewaxed in two changes of xylene, rehydrated sequentially in absolute, 95%, 70% ethanol and distilled water and rinsed in buffer 1 prior to immunohistochemistry. The antigen retrieval procedure (microwaving in citrate buffer) was carried out according to the manufacturer`s instructions for anti-CD31 and anti-CD68 antibodies.
Antibodies
Endothelial cells in culture and tissue were characterized according to their staining pattern with the monoclonal and polyclonal antibodies listed in Table I. Secondary antibodies were biotinylated horse anti-mouse and horse anti-rabbit, Texas Red and fluorescein horse anti-mouse and horse anti-rabbit immunoglobulin G (IgG) (all from Vector Laboratories, Burlingame, CA, USA).
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Immunohistochemistry
After rinsing in buffer 1 sections and cultures were incubated for 30 min in buffer 2 (buffer 1 + 2% horse serum + 0.1% Triton X-100), followed by incubation with primary antibody diluted as required in buffer 3 (buffer 1 + 1% horse serum + 0.1% Triton X-100) for 60–90 min at room temperature. They were then washed twice in buffer 1 for 5 min and incubated for 60 min with the secondary antibody labelled either with biotin or fluorescent marker (Texas Red, Fluorescein). This procedure was followed by washing twice in buffer 1 for 5 min. Cultures stained with the use of fluorescent marker were then embedded in Vectashield mounting medium with 4',6-diamidino-2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, USA). When biotinylated secondary antibodies were used, cultures were washed twice in buffer 1, followed by application of streptavidin–alkaline phosphatase complex using Vectastain ABC-AP Kit (Vector Laboratories), washed twice in buffer 1, and washed briefly in buffer 4 (100 mmol/l Tris–HCl, 100 mmol/l NaCl, 50 mmol/l MgCl2, pH 9.5). Signal was revealed by colour development using Nitroblue Tetrazolium chloride and X-phosphate, 4-toluidine salt (Boehringer Mannheim, Lewes, UK). Controls included omission of primary antibodies. Sections were then counterstained with Mayer's haematoxylin, rinsed in tap water and PBS and finally embedded in Hydromount (BDH, Merck, Poole, UK).
Staining for UEA binding sites in endothelial cells was performed using biotinylated Ulex europaeus agglutinin I (Vector Laboratories), followed by the procedure described for the biotinylated secondary antibody. Visualization was performed on Leitz Diaplan microscope.
Endothelial cell growth assays
[Methyl-3H]-thymidine uptake assay
Mitogenic assays were performed in 96-well plates coated with collagen type IV (Boehringer Mannheim) Cells at fourth passage were plated at 5000 per well in culture medium in the presence of 50% HS and left for 24 h. The medium was then changed to one with 20% HS and left for another 24 h. Cells were then treated with growth factors: VEGF, basic fibroblast growth factor (bFGF), epidermal growth factor (EGF) (all from R&D Systems, UK) and adrenomedullin (Peninsula, St Helens, UK) in fresh culture medium supplemented with 10% HS. [Methyl-3H]-thymidine 1 µCi (Amersham Pharmacia Biotech, Little Chalfont, UK) was added to each well. After 24 h, cells were detached with trypsin and harvested with a Pharmacia-Wallac 96-well harvester directly onto filter mats and counted in a ß-plate scintillation counter.
Intracellular cAMP assay
Endometrial endothelial cells at fourth passage were seeded into the 96-well microtitre plates (tissue-culture grade) precoated with collagen IV (Boehringer Mannheim, Lewes, UK) at cell concentrations of 8000 per well and grown for 3 days in McCoy's 5A/50% HS medium at 37°C in a 5% CO2 humidified atmosphere and the medium was replaced daily. Intracellular cAMP measurement was performed using the non-acetylation enzyme-immunosorbent assay (Amersham Pharmacia Biotech) according to the manufacturer's instructions. Briefly, cells were supplemented with serum-deprived medium for 3 h followed by addition of serum-deprived medium containing either adrenomedullin at a range of concentrations from 0.01 to 1000 ng/ml or no peptide (4 wells for each dilution). Plates were incubated for 15 min at 37°C. Following aspiration of culture medium, lysis reagent was added and the plate agitated for 10 min at room temperature. Trypan Blue staining was used to confirm lysis. Equal amounts of standard dilutions or samples were added to the individual wells of the microtitre plate, followed by the addition of antiserum. Plates were covered and incubated at 3–5°C for 2 h. cAMP–peroxidase conjugate was pipetted into all wells except the blank and plates were incubated at 3–5°C for 60 min. After aspiration wells were washed four times with wash buffer, blotted on tissue paper and enzyme substrate was dispensed into each well. The plate was then covered and agitated on a microtitre plate shaker for 60 min at room temperature. The plates were read at 630 nm on a MRX Microplate reader (DYNEX Technologies, Chantilly, VA, USA). The average optical density (OD) was calculated for each set of duplicate standards and quadriplicate samples. The percentage bound for each standard and sample was calculated according to the formula:
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A standard curve was generated by plotting the percent B/B0 as a function of the log cAMP c concentration. The fmol/well value of samples was read from the graph using the GraphPad Prism computer program.
Results
Isolation and culture of human endometrial endothelium
A method developed for the isolation of human decidual endothelial cells was adapted to the isolation of non-pregnant human endometrial endothelial cells (HEECs). The method employs positive selection with magnetic beads coated with Ulex europeus agglutinin that selectively binds to endothelial cells in intact tissue (Figure 1a). Initially, isolated cells are seen as clumps with beads attached to them. After 24 h primary cultures appeared as discrete rounded colonies, similar to primary cultures of human decidual endothelial cells (DECs) (Grimwood et al., 1995). The method proved reliable with 80% of the attempted isolations showing success. A few preparations (<5%) showed a contamination of epithelial cells due to the fact that some of these cells occasionally show staining for UEA-1 (Figure 1a). In the culture conditions described epithelial cells did not grow and did not survive longer than the second passage leaving a pure population of endothelial cells.
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Endometrial endothelial cells showed a typical `cobblestone' morphology and reached confluence after ~1 week (Figure 1e
) in comparison with 3–5 days for decidual endothelium (Figure 1i). This could be explained by the fact that fewer cells were isolated from endometrium due to the smaller amount of tissue being available in hysterectomy specimens. Human endometrial endothelium displayed typical endothelial morphology (Figure 1e) with large rounded nuclei, marked contact inhibition and no overlapping of adjacent cells at confluence. The beads remained attached to the cells through the first passage but not later. This is similar for first passage endothelial cells from both decidua (Figure 1k) and non-pregnant endometrium (not shown).
Characterization of human endometrial endothelium
The cultured endothelial cells were characterized by immunohistochemistry using endothelial markers (Table II). CD31, von Willebrand factor and binding sites for UEA 1 were strongly expressed on normal HEECs and DECs both in intact tissue and in primary cultures and were retained after 4–5 passages (Table II). Intense punctate perinuclear staining for von Willebrand factor was seen in the cytoplasm of the HEECs (Figure 1f) and DECs (Figure 1j). At confluence, CD31 was expressed mainly at sites of cell-to-cell contact in HEECs (Figure 1g), DECs (Figure 1k), HDMVECs and HUVECs (data not shown). Furthermore HEECs and DECs in culture revealed high proliferative activity as shown by the staining of nuclei of many cells in culture with antibodies recognising the proliferative marker Ki-67 (Figure 1 h,l).
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Immunocytochemical staining with antibodies against smooth muscle actin, keratins and CD 68 (Tables I and II
) was performed to check for presence of smooth muscle, epithelial cells and macrophages respectively in endothelial cells isolates and revealed minor (<1%) contamination in some cases.
Response of endometrial endothelial cells to growth factors and mitogenic activity of adrenomedullin
Adrenomedullin induced [methyl-3H]-thymidine uptake in culture (Figure 2). The stimulation was comparable to several other growth factors, e.g. VEGF, bFGF and EGF, known to be present in endometrium (Rees and Bicknell, 1998). Adrenomedullin has a growth effect not only on human endometrium endothelial cells, but also on microvascular endothelial cells from human skin (L.Nikitenko, unpublished observations).
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Stimulation of production of cAMP by adrenomedullin in human endometrial endothelial cells
Intracellular cAMP measurement demonstrated that adrenomedullin at concentrations of >1000 ng/ml significantly increased the amount of cAMP in the human endometrial endothelial cells (Figure 3
).
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Immunocytochemical localization of adrenomedullin in endothelial cells in vivo and in vitro
Adrenomedullin immunoreactivity was also found in epithelial, stromal and endothelial cells in frozen sections of human endometrium (Figure 4a
). Immunohistochemistry also showed that adrenomedullin immunoreactivity is evenly localized throughout the cytoplasm of human microvascular endothelial cells from endometrium (Figure 4c), decidua (Figure 4e) and skin (Figure 4f) in culture.
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Discussion
Aberrant angiogenesis is now believed to be involved in benign and neoplastic endometrial pathologies such as excessive menstrual bleeding (Rees and Bicknell, 1998), endometriosis (Fujimoto et al., 1999) and endometrial cancer (Sivridis et al., 1999). It is essential to gain a better insight of the factors which regulate normal endometrial angiogenesis during the menstrual cycle in order to understand its role in the development of uterine pathologies. Thus, knowledge of which angiogenic factors are present in the uterus and to which factors the endothelium of the endometrium is responsive, is needed. The present study was undertaken to examine the responsiveness of normal endometrial endothelium to growth factors and the control of endometrial angiogenesis.
Several studies have implicated a role for different growth factors in endometrial angiogenesis, e.g. VEGF (Shifren et al., 1996; Zhang et al., 1998; Iruela-Arispe et al., 1999), thymidine phosphorylase (Zhang et al., 1997), EGF (Watson et al., 1996) and bFGF (Ferriani et al., 1993). Thus, in-situ hybridization and immunohistochemistry have shown that VEGF is expressed predominantly in the epithelial glands, where expression is weak during the proliferative phase but strong in the secretory phase (Shweicki et al., 1993; Zhang et al., 1998; Sharkey et al., 2000) There is convincing evidence that growth factor synthesis and secretion is regulated by steroid hormones. In the endometrium, VEGF has been shown to be induced by both oestrogen (Zhang et al., 1998) and hypoxia (Popovici et al., 1999). Expression in the menstrual phase is probably a response to hypoxia and the protein persists in the base of the glands that remain after endometrial shedding. Thus, in the early follicular phase (Zhang et al., 1998), VEGF may be involved in initiating angiogenesis in the new cycle.
As for many human tissues, the isolation and culture of microvascular endothelium from human endometrium has, to date, proven difficult. Endothelium from decidua is easier to handle and several papers have documented successful isolations (Peek et al., 1994; Grimwood et al., 1995). Recently, a single description of the isolation of endothelium from endometrium (Iruela-Arispe et al., 1999) and one from myometrium (Gargett et al., 2000) have been published. For the first time, these reports have permitted in-vitro studies of non-pregnant uterine endothelial cell growth and behaviour.
A method for the isolation of human endometrial endothelial cells has been developed and is based on one used for the isolation of decidual endothelium (Grimwood et al., 1995). The endometrial endothelial cells express classic endothelial cell markers in culture. CD31 was expressed mainly at the sites of cell–cell contact in HEECs and DECs. CD31 staining was similar to that obtained for HDMECs and HUVECs and data reported by others (Albeda et al., 1990). Intense punctate perinuclear staining for von Willebrand factor was seen in the cytoplasm of the HEECs (Figure 1f) and DECs (Figure 1j) and was similar to the staining obtained for HUVECs and HDMECs (L.Nikitenko, unpublished observations) and to that of endothelial cells from other organs (Hewett and Murray, 1996).
In the present study, we have found that endometrial endothelial cells in culture respond to a number of growth factors known to be angiogenic, including VEGF (Plate et al., 1994), bFGF (Ziche et al., 1997) and EGF (Schreiber et al., 1986). Each factor and its receptor are known to be present in endometrium (Cordon-Cardo et al., 1990; Smith et al., 1991; Ferriani et al., 1993; Shifren et al., 1996; Watson et al., 1996; Sangha et al., 1997; Meduri et al., 2000). The growth factor responsiveness of microvascular endothelium from endometrium (the present study), myometrium (Gargett et al., 2000) and decidua (Peek et al., 1994; Grimwood et al., 1995) clearly illustrate that endothelial cells from different vascular beds of the same organ show a differential response to growth factors. It is interesting that endometrial endothelial cells respond to a wider range of growth factors than do decidual endothelial cells, which show no response to bFGF and EGF (Grimwood et al., 1995). This is probably due to the fact that cells in the endometrium undergo structural and functional changes during pregnancy resulting in the decidualization of the tissue. The lack of response of decidual endothelial cells to certain growth factors is probably due to loss of the cognate receptors.
We have previously demonstrated that adrenomedullin is expressed in the human uterus (Zhao et al., 1998). This has been confirmed by others (Michishita et al., 1999). We have also shown that it has an angiogenic activity in vivo in the chicken chorioallantoic (CAM) assay and stimulates growth of large vessel endothelial cells (Zhao et al., 1998). In the present study we have examined the effect of adrenomedullin on human endometrial microvascular endothelial cells in vitro.
The [methyl-3H]-thymidine uptake assay has shown that adrenomedullin stimulates proliferation of endometrial endothelial cells in culture and may be a growth factor for other microvascular endothelial cells (HDMECs; L.Nikitenko, unpublished observations). It has also been proposed that adrenomedullin could be a survival factor for endothelial cells, preventing apoptosis (Kato et al., 1996). This would be analogous to the anti-apoptotic effect of other angiogenic factors, such as VEGF (Carmeleit et al., 1999) and bFGF (Kazama and Yonehara, 2000). Thus it is possible that the `anti-apoptotic' properties of adrenomedullin may also contribute to the increased rate of the proliferation observed for HEECs in culture.
Adrenomedullin not only induces the proliferation of endothelial cells in endometrium but also stimulates cAMP production by them. These findings are in accordance with data obtained on the effect of adrenomedullin on DNA synthesis in Swiss 3T3 cells (Withers et al., 1996) by a mechanism involving specific adrenomedullin receptors and an increase of cAMP/PKA. Adrenomedullin stimulated an elevation of the intracellular cAMP level in HEECs only at high doses (1000 ng/ml). This is in agreement with the dose-dependent effect of the peptide on cAMP production in rat vascular endothelial cells in vitro (Isumi et al., 1998b) and also probably reflects the properties (number and sensitivity) of adrenomedullin receptors expressed in these cells. Thus the growth promoting effect of adrenomedullin on endometrial endothelial cells appears to be associated with an elevation of cAMP.
Adrenomedullin is known to be expressed in endothelial cells of different organs (Isumi et al., 1998b). Immunohistochemistry on frozen sections of human endometrium has shown localization of adrenomedullin immunoreactivity in epithelial, stromal and endothelial cells. We have also shown the presence of adrenomedullin immunoreactivity in the microvascular endothelial cells from endometrium, decidua and skin. Although most adrenomedullin is secreted from the cell, a small amount of peptide remains inside it and can be detected by immunohistochemistry (Satoh et al., 1996; Marinoni et al., 1998; Zhao et al., 1998).
Adrenomedullin expression is induced by tamoxifen in endometrial stromal cells and is probably involved in the endometrial angiogenesis and hyperplastic response of endometrium to this drug (Zhao et al., 1998). We have shown here that adrenomedullin may also play a role in normal endometrial angiogenesis acting as an autocrine growth factor for microvascular endothelial cells.
It is still not known which factors regulate adrenomedullin production and secretion from endometrial endothelial cells. Most studies have failed to demonstrate steroid receptors on endometrial endothelial cells. Furthermore, the absence of oestrogen-response elements in the adrenomedullin promoter lead to the assumption that in uterus the expression of adrenomedullin is stimulated mainly by cytokines, e.g. interleukin (IL)-1
, IL-ß, tumour necrosis factor (TNF) and TNFß, known to be present in endometrium (Tabibizadeh, 1991; Rice and Chard, 1998) and by hypoxia (Nguyen et al., 1999; Hofbauer et al., 2000) which is thought to occur during menstruation (Markee, 1940).
The present study shows that adrenomedullin is produced by endometrial endothelial cells and promotes their growth. Taken together with its angiogenic activity in vivo, these results suggest that adrenomedullin may be a significant regulator of endometrial angiogenesis during the menstrual cycle.
Acknowledgments
The authors would like to make it clear that this work is the result of a ten year collaboration between M.C.Rees and R.Bicknell, who both contributed equally to the work. This work was supported through the Johnson & Johnson Focused Giving Program and the Imperial Cancer Research Fund. We would like to thank Dr Dave Smith (Imperial College, London) for helpful discussions and advice on cAMP assay, Dr Katya Chobotova (Nuffield Department of Obstetrics and Gynaecology, University of Oxford) for help with performing [methyl-3H]-thymidine uptake assay; Dr Herbert Weich (Department of Gene Regulation and Differentiation, Division of Molecular Biotechnology, National Research Center for Biotechnology (GBF), Braunschweig, Germany) for a generous gift of antibodies; Dr Robin Roberts–Gant (Medical Informatics Unit, Nuffield Department of Clinical Laboratory Sciences, University of Oxford, UK) for assistance with formatting figures; Angela Quantutl and Caroline Pym (Department of Cellular Pathology; John Radcliffe Hospital, Oxford, UK) for help with sectioning paraffin-embedded material used in this study and also the operating theatre personnel in the Women's Centre, John Radcliffe Hospital, for the collection of hysterectomy specimens.
Notes
3 To whom correspondence should be addressed at: Molecular Angiogenesis Laboratory, Imperial Cancer Research Fund, Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Oxford OX3 9DU, UK. E-mail: r.bicknell{at}icrf.icnet.uk 
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Submitted on March 23, 2000; accepted on June 19, 2000.
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