Abstract
Adrenomedullin (AM) is a multifunctional peptide hormone, which plays a significant role in vasodilation and angiogenesis, implicating it in hypertension as well as in carcinogenesis. AM exerts its effects via the calcitonin receptor-like receptor (CRLR, now known as CL) complexed with either receptor activity modifying protein (RAMP) 2 or 3. We have investigated the effect of AM on immortalized human microvascular endothelial cells 1, since endothelial cells are a major source as well as a target of AM actions in vivo. Cells treated with AM showed elevated cAMP in a time (5–45 min)-dependent and dose (10−6–10−14 M)-dependent manner. Pre-treatment with the AM receptor antagonist AM22–52 partially suppressed the AM-induced increase in cAMP levels. An increase in extracellular signal-regulated kinase 1/2 phosphorylation was observed after 5 min of treatment with 10−8 M AM. This phosphorylation was specific, since we were able to block the AM-induced effect with 1 μM U0126, a specific mitogen-activated protein/extracellular signal-regulated kinase kinase inhibitor. Using real-time PCR, we were able to show for the first time that AM upregulates peptide and mRNA expression of vascular endothelial growth factor (VEGF). However, AM treatment of cells did not result in increased cell proliferation. Instead, we observed that AM and VEGF induced cell migration, which could be inhibited by the AM22–52 and anti-VEGF antibody respectively. AM also significantly elevated mRNA levels of CL (after 2 and 24 h treatment) and RAMP2 (after 1 and 24 h treatment). The upregulation of the AM receptor at two time points reflects possibly different cellular responses to short- and long-term exposure to AM.
Introduction
Adrenomedullin (AM) is a 52-amino acid peptide originally isolated by Kitamura et al.(1993) from a human pheochromocytoma. It is a multifunctional peptide produced by many cells and tissue systems (Kitamura et al. 1993), reviewed by Hinson et al.(2000). AM was initially characterized by its ability to stimulate cAMP production in human platelets and exerted a potent and long-lasting vasodilative effect in the rat (Kitamura et al. 1993). AM has been proposed early as an important hormone in circulation control and maintenance of vascular tone, since it regulates endothelial permeability (Hippenstiel et al. 2002) and it contributes to the differentiation of bone marrow-derived mononuclear cells into endothelial progenitor cells (Iwase et al. 2005).
AM exerts its effects via the G-protein coupled calcitonin-like receptor (CL) complexed with a receptor accessory modifying protein known as receptor activity modifying protein (RAMP) 2 (AM1 receptor) or RAMP 3 (AM2 receptor) (McLatchie et al. 1998, Poyner et al. 2002). Although the sequence identity between both RAMPs is only 30%, AM1 and AM2 receptors are pharmacologically indistinguishable and are usually co-expressed within the same tissue (Kuwasako et al. 2002).
Even though the adrenal glands show the highest tissue AM mRNA concentration, endothelial cells show levels that are 20-fold higher than those found in the adrenal, lending evidence towards making AM an important hormone in the regulation of vascular tone. AM is considered to be most important in the paracrine control of vascular, particularly microvascular, function where it acts as a potent vasodilator (Smith et al. 2002). AM increases coronary blood flow and heart rate, as well as heart contractability (Nagaya et al. 2002). The importance of AM in vascular physiology was demonstrated by AM gene knockout mice, leading to death in utero due to severe malformations in vascular morphogenesis (Caron & Smithies 2000, Shindo et al. 2001).
Additionally, AM has also been implicated in physiological and pathological angiogenesis using different types of knockout mice, xenografted tumors, and in vitro models (Hague et al. 2000, Nikitenko et al. 2000, Oehler et al. 2001, Martinez et al. 2002, Kim et al. 2003); AM has also been reported to be expressed as an angiogenic factor in tumors. For example, in some cancers, AM expression is associated with vascular density and endothelial cell proliferation (Hague et al. 2000). In vitro studies have shown the AM receptor antagonist AM22–52 was able to abrogate tumor formation of a pancreatic tumor cell line suggesting a role for AM in this cancer (Ishikawa et al. 2003).
In the endothelial cells, AM and vascular endothelial growth factor (VEGF) act in a conjoined manner to induce angiogenic effects in vitro, but the angiogenic actions of AM appear to be independent of VEGF secretion (Fernandez-Sauze et al. 2004). Instead, AM upregulates VEGF mRNA levels in the ischemic hind limb of mice after 1 day of treatment, thereby possibly enhancing the angiogenic effect of VEGF (Iimuro et al. 2004).
The main aim of the present study was to determine the biological actions, functions, and possible target genes of AM using human microvascular endothelial cells (HMEC) 1. Most pathological events involving endothelial cells occur at the level of the microvasculature, which is also thought to be the main target tissue of AM in vivo.
Materials and Methods
Cell culture
HMECs were purchased from the Center for Disease Control in Atlanta, GA, USA. This cell line was obtained by transfecting human dermal microvascular endothelial cells with a PBR-322-based plasmid containing the coding region for the Simian virus 40T gene product, and large T antigen (Ades et al. 1992). Cells were cultured in MCDB-131 medium with 11.6 g/l l-glutamine, 5% fetal bovine serum, 10 μg/l epidermal growth factor and 1% penicillin/streptomycin mixture (all obtained from Sigma–Aldrich) in T75 cm2 tissue culture flasks (Triple Red, Thame, Oxon, UK) or six-well plates (Nunc, VWR, Leicestershire, UK). Culture medium was changed every 2–3 days and cells split when they reached 90% confluency. For the experiments, cells with passage numbers between 10 and 20 were used.
Materials
Peptides and cell culture medium including necessary supplements were purchased from Sigma–Aldrich
cAMP assay
cAMP levels were determined by ELISA following the manufacturer’s instructions (Cyclic AMP Immunoassay; R&D System, Abingdon, Oxfordshire, UK). Approximately, 250 000 cells were plated into each well of a six-well plates and deprived of serum overnight. The next day, cells were treated with AM at various concentrations (dose response) and for different time points (time-course). To prevent cAMP degradation, cell medium contained 1 mM 3-isobutyl-1-methylxanthine. To stop the reaction, 95% ethanol was added and plates were stored at −20 °C overnight. After defrosting, the following day, liquid from the wells was completely transferred to clean 1.5 ml Eppendorff tubes and the samples were dried in a vacuum centrifuge. The dried samples were reconstituted in 500 μl assay buffer ED2, provided in the kit, and stored at −20 °C until further use.
MTT assay
To determine the effects of AM on cell proliferation, approximately 60 000 cells were plated into each well of 24-well plates.
Cells were deprived of serum overnight prior to incubation with 0 (serum free, SF), different concentrations of AM or growth factor-containing medium (serum rich, SR). After 24, 48, or 72 h treatment, cells were washed with PBS and subsequently incubated for 2 h in the dark with 0.5 mg/ml MTTreagent (thiazolyl blue tetrazolium bromide). The MTT solution was removed and cells were lysed by adding 250 μl 10% DMSO/90% isopropanol mixture. Plates were centrifuged for 5 min at 13 000 r.p.m. and 200 μl of each sample transferred into wells of a 96-well plate. Plates were read using an optical density of 570 nm in a microtiter plate reader.
RNA extraction and cDNA synthesis
Total RNA was extracted from cultured cells using the RNeasy Mini Kit (Qiagen). The extractions were carried out according to the manufacturer’s instructions. cDNA was synthesized using the First-strand cDNA Synthesis kit from GE Healthcare (Amersham) following the manufacturer’s instructions.
Western blot analysis
Total protein was extracted using 60% confluent cells from T75 flasks. Prior to extraction, cells were washed with PBS (137 mM NaCl, 27 mM KCl, 43 mM Na2HPO4·7H2O, 14 mM KH2PO4, pH 7.4). After adding 300 μl CytoBuster Protein Extraction Buffer (Novagen, Windsor, Berks, UK), flasks were incubated at room temperature for 5 min. The resulting solution was transferred to a sterile 1.5 ml Eppendorff tube and centrifuged for 5 min at 16 000 g at 4 °C. The supernatant was removed to a fresh tube and sonicated for 10 s to achieve a homogenous solution. Equal amounts of protein were loaded on 10% Tris–HCl gels and subsequently transferred to a Hybond-P polyvinylidene diflouride membrane (GE Healthcare, Amersham). After blocking with 5% non-fat dry milk solution, the membranes were incubated with primary antibody at 1:1000 dilution at 4 °C overnight. Phospho-p44/42 mitogen-activated protein kinase (MAPK) (Thr202/Tyr204) (E10) monoclonal antibody and p44/42 MAPK antibody were purchased from Cell Signaling Technology (Hitchin, Herts, UK).
A peroxidase-conjugated secondary goat anti-rabbit antibody was used at 1:1000 dilution. The blots were developed using the solutions of the ECL plus Western Blotting Detection System (GE Healthcare, Amersham).
Gene quantitation
Quantitative real-time reverse transcriptase (RT)-PCR was performed using dual-labeled fluorescent taqman probes to determine CL, RAMP2, RAMP3, and VEGF-A mRNA levels as listed in Table 1. RT-PCR amplifications were carried out for all genes studied and the resulting products sequenced to ensure primer specificity. Taqman reagents were obtained from Stratagene (Amsterdam, Netherlands) and primers and probes were supplied by Sigma–Genosys.
Analysis was carried out using the MX4000 (Stratagene) starting cycle with an initial 95 °C for 10 min followed by 40 cycles consisting of 95 °C for 30 s and 55 °C for 1 min.
VEGF peptide assay
VEGF levels were determined by ELISA following the manufacturer’s instructions (Quantikine, Human VEGF Immunoassay kit, R&D System). Approximately, 250 000 cells were plated into each well of a six-well plate and deprived of serum overnight. Cells were incubated for 4 h with 10−7 and 10−9 M AM. Liquid from the wells was completely transferred to clean 1.5 ml Eppendorff tubes and centrifuged. Supernatants were stored at −20 °C until further use.
Migration assay
Chemotaxis assays were performed using a Neuroprobe ChemoTx plate with 8 μm pore size (Receptor Technologies Ltd, Adderbury, Oxfordshire, UK). HMECs were diluted to 4×106 cells/ml in SF medium and either plated directly into the wells of the polycarbonate filter or pre-incubated for 45 min with an anti-VEGF-antibody (Sigma–Aldrich) or AM22–52 (10−7 M). The bottom wells of the neuroprobe plate were loaded with 300 μl of different concentrations of AM or VEGF (Sigma–Aldrich). In the bottom wells, 300 μl serum-containing and SF media were also loaded and served as positive and negative controls for migration respectively. The polycarbonate filter was placed on top of the lower chamber and 25 μl HMECs were placed on top of the filter. To determine migrated cell numbers, a serial dilution of cells ranging from 4×106 to 12.5×104 cells was plated directly into the wells of the lower plate. As an additional control to determine whether the migratory response is genuinely chemotactic, cells were incubated with either AM or VEGF and placed on the ploycarbonate filters with either SF medium in the lower chamber or the corresponding concentration of AM and VEGF.
Plates were incubated for 5 h in an incubator at 37 °C with 5% CO2. Cells remaining on top of the filter were absorbed off and the filter tops were washed with SF medium to ensure removal of all non-migrated cells. The filters were incubated with 2 mM EDTA for 30 min at 4 °C to loosen cells from the inner filter membrane. The EDTA solution was removed and plates were subsequently spun at 400 g for 10 min. The filter was removed and 150 μl solution were taken out of the bottom wells and replaced with 150 μl 2× MTT dye. Cell quantification was carried out as described for the MTT assay above.
Statistical analysis
The statistical analysis was carried out using GraphPad PRISM software (version 3.0; Graph pad, San Diego, CA, USA), one-way ANOVA and Turkey–Kramer post hoc test. Data were expressed as means ± s.e.m.
Results
AM stimulates cAMP in HMECs
Treatment of HMECs with varying concentrations of AM (10−6–10−14 M) for 5 min resulted in a significant increase in cAMP above basal (Fig. 1). In particular, AM at 10−6–10−8 M increased cAMP levels by approximately twofold. Forskolin (FSK) was used as a positive stimulation control of AM in these cells and we found that it enhanced the cAMP response 3.5-fold above basal. Co-treatment of AM and FSK showed that AM can potentiate the FSK-induced increase in cAMP.
In time-course experiments, AM (10−8 M) significantly increased cAMP levels after 2.5 min of treatment (Fig. 2). The AM response peaked after 5 min of treatment resulting in an approximately 2.7-fold increase in cAMP above the corresponding SF control.
To determine whether the observed response is AM-specific, we pre-treated the cells for 30 min with varying concentrations of AM receptor antagonist AM22–52 (10−6–10−10 M; Fig. 3). The AM-induced increase in cAMP levels was reduced in the presence of AM22–52 at high concentrations (10−6–10−7 M), but not completely abolished. Treatment with AM22–52 (10−6 M) had no effect on the cAMP levels of the positive or negative controls, excluding short-term toxic effects of the inhibitor on the cells.
AM stimulates ERK 1/2 phosphorylation in HMECs
To further examine the AM-induced signaling cascade in HMECs, we studied phosphorylation of MAPK/ERK 1/2 using western-blotting analysis. Treatment of HMECs with AM (10−8 M) resulted in a significant increase in ERK 1/2 phosphorylation after 5 min exposure (Fig. 4). The maximum mean ERK 1/2 phosphorylation increase observed was 2.9-fold above basal and this effect returned to basal levels after 10 min of treatment. This effect of AM on ERK 1/2 was specific since use of the MEK inhibitor, U0126, completely abolished the AM-induced phosphorylation of ERK 1/2 (Fig. 5).
Activation of AM target genes
Since MAPK participates in a protein kinase cascade that plays a critical role in the regulation of cell growth, especially by the activation of ERK 1/2, we examined the effect of AM on endothelial cell proliferation by MTT assay. Cells were either treated with different AM concentrations (10−6–10−11 M) for 72 h or with 10−8 M AM for 24, 28, and 72 h. In our hands, AM did not cause an increase in cell proliferation compared with the serum deplete, negative control in either experimental set-up (Fig. 6A and B).
Quantitative real-time PCR revealed that treatment of cells with AM (10−8 M) significantly increased the mRNA levels of CL, RAMP2 as well as VEGF. After 4 h, CL mRNA was elevated 4.8-fold with a further increase after 24 h to 15-fold above basal (Fig. 7). RAMP2 mRNA levels increased 1.5-fold after 1 h and 2.9-fold after 24 h treatment with AM (Fig. 7). The expression of RAMP3 mRNA levels did not change significantly with the treatment and time points studied (data not shown).
The biggest change in mRNA levels after AM treatment was seen with VEGF (Fig. 8). After 2 h exposure to AM, the initially very low VEGF levels increased 70-fold. This was followed by a decline to basal and a second mRNA elevation after 48 h (5.8-fold above basal). Treatment of cells for 4 h with AM also increased VEGF peptide levels (Fig. 9). AM 10−7 and 10−9 M increased VEGF peptide levels 2.2- and 1.9-fold respectively. Since neither AM nor the increased VEGF levels following AM treatment caused cell proliferation, we investigated whether these two peptides have a migratory effect on HMECs (Fig. 10). Using a modified Boyden chamber method, cell migration in response to AM, VEGF, and SR positive and SF negative controls was determined. Placement of 10−10 M AM in the lower chamber induced a 1.9-fold increase in HMEC migration, equivalent to 29% of all cells. This effect was AM-specific, −7 M). VEGF since it could be blocked using AM22–52 (10 also caused a dose-dependent increase in cell migration. Treatment with 100 ng/ml VEGF increased cell migration twofold (32% of cells), while 60 and 40 ng/ml VEGF increased cell migration 1.8- and 1.7-fold (27 and 25% of cells respectively) above SF control respectively. VEGF-induced cell migration was blocked by pre-incubating cells for 45 min with 20 μg/ml anti-VEGF antibody, prior to loading into the upper chamber. Serum-containing medium caused 78% of all cells to migrate into the lower chamber, compared with 15% random migration in the serum-deplete negative control. Random cell migration using additional controls with cell suspensions containing AM (10−10 M) or VEGF (40, 60, or 100 ng/ml) with the equivalent concentration of peptide or SF medium in the bottom chamber was determined. In all cases, migration in the additional controls was not significantly different from the observed random unstimulated migration with the SF control (15%), ranging from approximately 12 to 18% of HMECs. All observed migratory effects with AM and VEGF are, therefore, genuine chemotactic responses of the cells and only 15% of total cell migration in the experiments can be attributed to chemokinetic cell movement.
Discussion
In this study, we have shown that AM induces cAMP elevation in HMECs, as has previously been reported for other endothelial cells (Isumi et al. 1998, Hippentiel et al. 2002). It was observed that AM was a very potent stimulant of cAMP in HMECs, since already very low concentrations of AM (10−10–10−14 M) were already able to elicit a significant cAMP response. The AM receptor antagonist AM22–52 did not completely block this response, even at 10−6 M concentration. This antagonist has been shown previously to be inefficient at blocking the AM receptor, therefore, the result observed in our study is not surprising (Champion et al. 1997, Nishikimi et al. 1998, Ziolkowska et al. 2003). It is clear that HMECs express at least one fully functional AM receptor sub-type, which when activated leads to an increased level of cAMP.
Further, downstream of the cellular-signaling cascade, AM has been reported to induce as well as inhibit phosphorylation of ERK 1/2 in various different cell types. AM-induced decreases of phosphorylated ERK 1/2 were observed in rat mesangial cells (Parameswaran et al. 2000), while Jiang et al.(2004) report that AM inhibited the aldosterone-stimulated ERK activity in rat cardiac fibroblasts. In both reports, treatment of cells with AM leads to increased cAMP levels, decreases in ERK 1/2, and subsequently to inhibition of cell proliferation and protein synthesis. Other studies have shown that AM increases ERK 1/2 phosphorylation, thereby exerting angiogenic and proliferative effects in rat vascular smooth muscle cells and human umbilical vein endothelial cells (HUVECs) (Iwasaki et al. 2001, Kim et al. 2003).
In our study, treatment of cells not only resulted in elevated cAMP levels, but also in an increase in ERK 1/2 phosphorylation after exposure to AM for 5 min. This effect has been shown to be specific by the complete inhibition of response with 1 μM U0126. Elevated ERK 1/2 levels are commonly associated with cell proliferation (Cobb et al. 1994, Roux & Blenis 2004). However, in our hands, AM did not increase HMEC proliferation during treatment at any of the time points or AM concentrations used in this study. Therefore, we propose that proliferation might be regulated in HMECs independent of cAMP and ERK1/2 phosphorylation by a yet unknown pathway.
We have shown for the first time that AM upregulates the mRNA levels of CL and RAMP2 in HMECs. Quantitative analysis of CL mRNA levels showed upregulation of the gene in a bi-phasic manner, 4 and 24 h after exposure. The increase in CL mRNA was higher after 24 h compared with the elevation after 4 h. RAMP2 mRNA levels also appear to be regulated in a bi-phasic manner with an increase after 1 and 24 h treatment. This could suggest that the initial increase in mRNA levels are a direct response to the AM stimulation, whereas the later CL and RAMP2 induction might be mediated by distinct second messenger system components. In rat ileum, platelet-activating factor (PAF) increases the mRNA levels of PAF receptor (PAF-R) in a bi-phasic manner after 30 min and 6 h post-stimulation (Wang et al. 1997). PAF has been known to stimulate tumor necrosis factor (TNF) secretion in rat ileum cells, which in turn increases endogenous PAF levels. The authors hypothesize that PAF induces production of endogenous TNF in the cells, which in turn increases the levels of PAF. This feedback loop then leads to the observed second peak in transcriptional activation of PAF-R mRNA. A similar yet unknown mechanism might be involved in the regulation of CL and RAMP2 expression.
Alternatively, the second peak of RAMP2/CL mRNA may reflect failure of HMECs to induce a repressor, which downregulates or destabilizes these mRNA species.
It seems apparent that AM can partially regulate the expression of its own receptor. Transcriptional regulation of receptors by their ligands has been shown in various different studies. Exposure of agonists to G protein-coupled receptors frequently results in downregulation of the receptor transcript levels, but receptor upregulation has also been demonstrated (Siegrist et al. 1994, Schanstra et al. 1998, Froidevaux & Eberle 2002, Ankö & Panula 2006). Inoue et al.(1999) found that calcitonin downregulates calcitonin receptor mRNA in mouse bone marrow cells and Dupre et al.(2003) report the ligand-induced downregulation of the PAF-R. On the other hand, gene expression of the human somatostatin receptor type I is actively upregulated by somatostatin (Hukovic et al. 1999). The regulation of CL/RAMP2 mRNA levels by AM in HMECs might reflect a physiological adaptation, which enables the cells to adjust the sensitivity of receptor-mediated processes by changes in receptor number, and, therefore, according to the level of receptor activation.
The fact that CL and RAMP2 mRNA levels are also upregulated at different time points, 4 and 1 h respectively, might indicate yet unknown functions of the proteins. If the AM-induced increase in mRNA is also translated to the protein level interactions of RAMP2 and CL with other proteins, additional signaling complexes may be formed.
Since AM did not alter the expression of RAMP3, we propose that transcriptional regulation of this gene occurs via a yet unknown mechanism.
Another novel finding in HMECs is that AM upregulates VEGF mRNA and peptide levels. It has been shown previously that AM induces VEGF mRNA levels in HUVECs as well as in the hind limb of AM-treated mice (Iimuro et al. 2004). Therefore, AM might be an influential factor in angiogenesis in HMECs, but stimulation of VEGF expression also gives AM an important role in carcinogenesis. VEGF is implicated in endothelial barrier dysfunction, which allows cancer cells to migrate across the vascular lining of vessels, one of the key events in cancer metastasis. Dysfunction of the vascular endothelial barrier might facilitate the widespread dissemination of cancer cells (Zachary & Gliki 2001).
It seems surprising that AM has no indirect effect on cell proliferation either directly or via the elevation of VEGF mRNA and peptide levels. AM has been reported previously to promote cell growth as well as inhibit proliferation. AM stimulates cell proliferation in zona glomerulosa cells, skin fibroblasts and keratinocytes (Albertin et al. 2003), gliobastoma, lung cancer cells, and endometrial tumors (Oehler et al. 2002). Cell growth is inhibited by AM in myocytes, cardiac fibroblasts, vascular smooth muscle cells, prostate cancer cells (Abasolo et al. 2004), and mesangial cells (Segawa et al. 1996). This discrepancy in biological function might be related to varying cell-type-specific differences in signal transduction pathways. In HMECs, AM and VEGF have been shown to have significant effects on cell migration. However, the migratory response of HMECs to SR control is significantly higher than to either AM or VEGF. It seems likely that other factors in addition to AM and VEGF might play a role in the complex process of migration and angiogenesis. Angiopoetins or cytokines such as pleiotrophin might be additional requirements for migration of microvascular endothelial cells, as well as signals from other adjacent cell types, such as fibroblasts and pericytes within their microenvironment.
Nevertheless, AM seems to be an important contributing factor in the complex system of angiogenesis and vascular morphogenesis, especially, since AM gene knockout mice display severe cardiovascular abnormalities and die early during embryonic development. However, further studies are required to fully understand the role of AM in physiological and pathological events concerning the vasculature. HMECs carry many traits of primary endothelial cells (Unger et al. 2002), reviewed in Bouïs et al.(2001), but they are immortalized cell lines and in vitro findings might not always reflect those of in vivo studies.
In conclusion, we have demonstrated that the treatment of HMECs with AM increases cAMP levels and phosphorylation of the ERK 1/2 pathway. In our cell line, this does not lead to an increase in cell proliferation. Instead, we have identified that the cAMP, ERK 1/2 pathway is involved in AM-induced upregulation of CL, RAMP2, and VEGF mRNA and protein levels. AM and VEGF are able to induce HMEC cell migration, suggesting a role for AM in microvascular endothelial physiology and pathology. Further studies are needed to determine whether AM influences the regulation of other genes, possibly inflammatory cytokines or other growth factors.
Funding
The work was supported by the Special Trustees of the Royal London and St Bartholomew’s Hospitals. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.
Sequences of the used primers and dual-labeled fluorescent probes for CL, RAMP2, RAMP3, and VEGF
| Primer/probe sequences | Product size (bp) | |
|---|---|---|
| bp, base pairs; S, sense strand; As, antisense strand | ||
| Gene | ||
| CL | S: aaagcacaatcaacttttctgagc As: aataagggtagaatcatgcccaac Probe: agccagttccagcacaccattgca | 118 |
| RAMP2 | S: tctccctaggacccgagtcg AS: tgtgcctgtggtgggaagag Probe: agccgtccgcctcctccttctgc | 112 |
| RAMP3 | S: caggtttctatgctgtttcttagc AS: gcaaggagtcctagtccaagc Probe: ccagcctagccttagccgcagtct | 83 |
| VEGF | S: aaccagcagaaagaggaaagagg AS: cactcactttgcccctgtcg Probe: ttcgctgctcgcacgcccgc | 121 |
cAMP levels in HMECs following incubation with AM. HMECs were incubated with increasing concentrations of AM (10−6–10−14 M), 10−4 M forskolin (FSK) or serum-free (SF) medium. Values are means ± s.e.m., n = 9. *P<0.05, **P<0.01, ***P<0.001 compared with serum-free control.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
cAMP levels in HMECs after exposure to AM or forskolin with time. HMECs were incubated with 10−8 M AM (▴) or 10−5 M forskolin (▪) for increasing amounts of time and intracellular cAMP was measured by ELISA. Values are means ± s.e.m., n = 9. ***P<0.001 compared with control (time point 0).
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Effect of AM receptor antagonist on HMEC cAMP levels. HMECs were incubated with decreasing concentrations of the AM antagonist AM22–52 for 30 min prior to being exposed to 10−8 M AM for a further 5 min. Values are means ± s.e.m., n = 9. *P<0.05, ** P<0.01, ***P<0.001 compared with serum-free control.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Western blot of ERK 1/2 phosphorylation in HMECs. HMECs were serum starved overnight and subsequently treated with 10−8 M AM for the indicated time points. Cell homogenates (15 μg protein) were tested for phosphorylated ERK 1/2 (A), stripped and re-probed for total ERK 1/2 (B). Scanning densitometry of three different experiments is shown as a bar graph. Values are means ± s.e.m. *P<0.05 compared with time 0.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Effect of an MEK inhibitor on ERK 1/2 phosphorylation in HMECs. Western-blot analysis of 15 μg protein of cell homogenates for phosphorylated (p) ERK 1/2 expression (upper panel) or total ERK 1/2 (lower panel) in HMECs exposed to 10−8 M AM, 10−5 M forskolin (FSK) or serum-free medium (SF) for 5 min in the absence (−) or presence (+) of 1 μM U0126. Scanning densitometry of three different experiments is shown as a bar graph. Values are means ± s.e.m., n = 3. ***P<0.001 compared with presence of U0126.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
MTT assay analysis of HMEC cell proliferation. HMECs were treated for 72 h with increasing concentrations of AM (10−6–10−11 M) as well as serum-rich (SR) and serum-free (SF) medium (A). For the time-course experiment, HMECs were treated for 24, 48, and 72 h with either 10−8 M AM (white bars), SF (grey bars) or SR (black bars) (B). Values are means ± s.e.m., n = 9. *P<0.05, **P<0.01, ***P<0.001 compared with serum-free control.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Expression of CL and RAMP2 mRNA expression in response to AM. HMECs were treated with 10−8 M AM for increasing lengths of time and CL (triangles) and RAMP2 (open squares) mRNA was quantified using real-time RT-PCR and normalized to the housekeeping gene glyseraldehyde-3-phosphate dehydrogenase. Values are expressed as means ± s.e.m., n = 3. *P<0.05, **P<0.01 when compared with time point 0.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Expression of VEGF mRNA expression in response to AM. HMECs were treated with 10−8 M AM for increasing lengths of time and VEGF mRNA was quantified using real-time RT-PCR and normalized to the housekeeping gene glyseraldehyde-3-phosphate dehydrogenase. Values are expressed as means ± s.e.m., n = 3. *P< 0.05, **P<0.01 when compared with time point 0.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Expression of VEGF peptide in response to AM in HMECs. Following treatment with 10−7 and 10−9 M AM for 4 h, supernatants were collected and analyzed. Values are expressed as picogram VEGF per 25×105 cells, n = 6. ***P<0.001 when compared with serum-free control.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
Adrenomedullin and VEGF treatment of HMECs induce cell migration. The lower chambers of the neuroprobe chamber were loaded with 300 μl of varying concentrations (10−9–10−12 M) AM or 40, 60, and 100 ng/ml VEGF, while serum-free (SF) and serum-rich (SR) media served as negative and positive migration controls. After loading cells into the wells of the upper chamber, plates were incubated for 5 h, prior to staining with MTT. The migratory effect of both peptides was blocked by incubating the cell −7 M) or anti-VEGF suspension for 45 min with the AM receptor antagonist AM22–52 (10 antibody (Vab) (20 μg/ml) prior to loading on the upper chamber. Migration is expressed as fold change in cell number in the bottom chamber above serum-free (basal) control. Values are expressed as means ± s.e.m., n = 6. *P<0.05, **P<0.01 when compared with serum-free control.
Citation: Journal of Endocrinology 190, 2; 10.1677/joe.1.06806
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