May 2004
Volume 45, Issue 5
Free
Retinal Cell Biology  |   May 2004
Adrenomedullin Affects Two Signal Transduction Pathways and the Migration in Retinal Pigment Epithelial Cells
Author Affiliations
  • Wei Huang
    From the Institute of Ophthalmology and
  • Lin Wang
    From the Institute of Ophthalmology and
  • Ming Yuan
    Cardiology, Xijing Hospital, The Fourth Military Medical University, Xi’an, China.
  • Jixian Ma
    From the Institute of Ophthalmology and
  • Yannian Hui
    From the Institute of Ophthalmology and
Investigative Ophthalmology & Visual Science May 2004, Vol.45, 1507-1513. doi:10.1167/iovs.03-0731
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      Wei Huang, Lin Wang, Ming Yuan, Jixian Ma, Yannian Hui; Adrenomedullin Affects Two Signal Transduction Pathways and the Migration in Retinal Pigment Epithelial Cells. Invest. Ophthalmol. Vis. Sci. 2004;45(5):1507-1513. doi: 10.1167/iovs.03-0731.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. Adrenomedullin is a multifunctional regulatory peptide known to inhibit the migration of smooth muscle cells. In vitro studies were performed to identify whether adrenomedullin (ADM) also inhibits the migration of RPE cells. The aberrant behavior of these cells is an early event in proliferative vitreoretinopathy, and these studies were designed to determine how ADM acts on RPE cells at the second-messenger level.

methods. Migration of cultured human RPE cells was determined by the Boyden chamber method, using 10% fetal calf serum (FCS) as a chemotactic factor. The attachment assay was performed on fibronectin, laminin, or poly-d-lysine–coated 96-well plates. RPE cells were incubated in PBS buffer with or without ADM for 15 minutes. Intracellular cAMP and cGMP changes were then measured by enzyme immunoassay (EIA). To determine the cytoplasmic free Ca2+ concentration ([Ca2+]i) response to ADM, fluo-3 AM–loaded RPE cells were imaged with a laser scanning confocal microscope, after stimulation with ADM (10−12–10−7 M).

results. ADM exhibited a concentration-dependent inhibition of FCS-stimulated RPE cell migration. The maximum inhibitory effect of ADM, observed at 10−7 M, on basal and FCS-induced RPE cell migration was approximately 53.8% and 43.8% of the control, respectively. Exogenously added ADM (10−9–10−7 M) had no significant effect on RPE cell attachment on all tested substrates. ADM increased intracellular cAMP and decreased intracellular cGMP levels dose dependently (10−10–10−7 M) in RPE. The maximum effect was observed at 10−7 M. ADM also induced a [Ca2+]i decrease in a dose-dependent manner (10−12–10−7 M). The maximum effect was observed at 0.1 μM, at which point the level declined to 42.9% of the control.

conclusions. ADM inhibits the migration of RPE cells in vitro by a mechanism that involves the reciprocal upregulation of cAMP and downregulation of cGMP, in association with reductions in [Ca2+]i. ADM-mediated fluctuations in [Ca2+]i, which are well known to be involved in cell migration, appear to be regulated in part by mechanisms involving cAMP synthase. Thus, it appears that ADM acts as a constitutive regulatory system to control aberrant RPE cell behavior and specific migration in response to inflammatory mediators.

Adrenomedullin (ADM) is a vasorelaxant peptide originally isolated from an extract of human pheochromocytoma tumor tissue. 1 Subsequent studies revealed that ADM is a multifunctional regulatory peptide with actions that range from regulating cellular growth and differentiation, through modulating hormone secretion, to antiapoptotic and antimigratory effects. 2 Most of these effects are exerted in a cell-type–specific manner. In Swiss 3T3 cells, ADM increases DNA synthesis in a dose-dependent manner by a mechanism involving specific ADM-receptor–mediated increases in cAMP and protein kinase A (PKA). 3 In human normal glial cells and glial cell tumors, ADM suppresses cell growth and increases intracellular cAMP. 4 The growth of human and rat astrocytomas and human glioblastomas, as well as cultured glioblastoma-derived cell lines, was also inhibited by ADM. 4 5 6 However, Moody et al. 7 reported that ADM exerts mitogenic effects on cultured C6 glioma cells that correlates with increases in cAMP and c-fos expression. To understand the mechanism of ADM function, several studies have investigated how receptor-mediated signal transduction modifies gene transcription at the nuclear level. cAMP has been found to be the major second messenger involved in ADM-induced cell responses. 2 8 9 However, the effects of ADM are not fully mimicked by forskolin, a known inducer of cAMP, suggesting a role for an additional second messenger. 
ADM transcripts are expressed in various human tissues and cells, including the eye. 10 11 In the iris sphincter isolated from cats and other mammalian species including humans, ADM is a much more efficacious activator of adenylate cyclase and a much more effective vasorelaxant than calcitonin gene-related peptide (CGRP). 12 Udono et al. 13 recently reported that human RPE cells also produce and secret ADM, and IFN-γ and IL-1β induce ADM expression in ARPE-19 cells, an immortalized RPE cell line. The same group 14 later demonstrated that hypoxia increases the expression of ADM in three human RPE cell lines, whereas the induction of endothelin (ET)-1 by hypoxia is found only in D407 cells. These findings indicate that ADM may be involved in modulating the roles of RPE in physiological and pathologic processes. 
The RPE lies between the retina and the choroid and plays a vital role in ocular metabolism. Apoptosis, degeneration, and proliferation of RPE cells are responsible for the development of a variety of blinding diseases such as retinitis pigmentosa, age-related macular degeneration (AMD), and proliferative vitreoretinopathy (PVR). The migration of activated RPE is an initial step in the development of PVR. 15 Immunoreactive ADM levels in the vitreous of patients with PVR are found at significantly higher levels than those of patients with proliferative diabetic retinopathy, AMD, and macular hole, 16 suggesting that ADM may be involved in the pathophysiology of PVR. It is noteworthy that agents that significantly increase intracellular levels of cAMP are also inhibitors of RPE cell migration. 17 Recent studies have shown that ADM inhibit platelet-derived growth factor (PDGF)-BB and fetal calf serum (FCS)–induced smooth muscle cell (SMC) migration in a concentration-dependent manner. 18  
These data suggest that ADM may be implicated in the early stages of PVR, which involve the migration of RPE cells. However, we have little information about the signal transduction pathways activated by ADM in RPE cells. The physiological role or roles of ADM and its possible pharmacologic effects on the mechanism of action in RPE cells are still unclear. In the present study, we investigated whether ADM acts on RPE cells through two independent signal-transduction pathways—cAMP and intercellular calcium ([Ca2+]) accumulation—previously known to be implicated in other cell types. We also determined whether ADM has effects on attachment and migration of RPE cells. 
Materials and Methods
Chemicals
Human ADM (1-52) 3-isobutyl-1-methylxanthine (IBMX), forskolin, N Gnitro-l-arginine methyl ester (l-NAME), trypsin, EGTA, A23187, and 8-Br-cAMP were purchased from Sigma-Aldrich (St. Louis, MO); cAMP and the cGMP EIA kit from Amersham Pharmacia Biotech, Ltd. (Amersham, UK); fluo-3 AM from Molecular Probes (Eugene, OR); phosphate-buffered saline (PBS), Dulbecco’s modified Eagle’s medium (DMEM), and FCS, from Invitrogen-Gibco (Rockville, MD). All other chemicals were of reagent grade. 
Human RPE Cell Culture
Human RPE cells were obtained from eye bank donor eyes. Human eyes were used in accordance with applicable laws and with the tenets of the Declaration of Helsinki. As previously described, 19 eyes were opened 360° posterior to the ora serrata, and the vitreous and the retinal tissues were removed. The remaining eyecup was washed with PBS, and 0.025% trypsin-EDTA was added and incubated with the eyecup for 30 minutes at 37°C. The cells were then gently scraped and seeded in DMEM containing 15% FCS in a culture dish. After proliferation, cells were retrypsinized with a 0.1% trypsin-EDTA solution for 5 minutes at 37°C. After they were washed with DMEM, the cells were plated in six-well plates (Nalge Nunc, Napierville, IL) at 2 × 104 cells/well and allowed to grow to confluence. Third- or fourth-passage cells were used for most experiments. 
Migration Assay
Migration of the cells was assessed by a modified Boyden chamber method 20 using microchemotaxis chambers (Neuro Prob Inc., Gaithersburg, MD) with polycarbonate membrane (5.7-mm diameter, 8.0-mm pore size). In all experiments, both sides of the membrane were precoated with laminin (5 μg/mL) at 37°C for 1 hour and air dried. Medium containing 10% FCS was added to the lower chambers as chemoattractant. To determine the effect of ADM on basal RPE cell migration, serum-free DMEM was added to the lower chambers. The subconfluent RPE cells were trypsinized and resuspended in 0.4% FCS to a concentration of 8 × 105 cells/mL. The cell suspension was then diluted 1:1 with a 2× concentration of test agents, and 60 μL of cell suspension (final concentration 2.5 × 104 cells/well) was added to the upper surface of the membrane. The cells were then incubated for 6 hours at 37°C in a 5% CO2 atmosphere. The upper surface of the membranes was scraped with a cotton-tripped stick to remove nonmigrated cells and then fixed with 4% paraformaldehyde for 15 minutes at room temperature. Migrated cells were counted manually in five random fields using an inverted microscope (IX70; Olympus, Tokyo, Japan) with a 400× objective after nuclear staining with hematoxylin. 
Attachment Assay
The attachment assay was performed on fibronectin, laminin, or poly-d-lysine-coated 96-well plates (Nalge Nunc) using a modified protocol by Jin et al. 21 Confluent RPE cells (2 × 105/mL) were trypsinized and resuspended in DMEM with 0.4% FCS. The cell suspension was then diluted 1:1 with a 2× concentration of test agents, and 100 μL of cell suspension (104 cells) was added to each well and allowed to attach for 60 minutes. The cells were washed twice with Hanks’ balanced salt solution, and fresh medium (150 μL) was added to each well with 20 μL MTT (3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H tetrazolium bromide, 5 mg/mL). After 5 hours of incubation the supernatants were aspirated. The formative precipitates were solubilized by the addition of 150 μL of 100% dimethylsulfoxide (DMSO) and placed on a plate shaker for 10 minutes. Absorbance at 490 nm was determined on a microplate reader (model 550; Bio-Rad, Hercules, CA). The living cell number was proportional to the absorbance of MTT at 490 nm. 
cAMP Measurement
Intracellular cAMP content was measured with the cAMP EIA kit, using novel lysis reagents. Briefly, RPE cells were seeded in 96-well plates at a density of approximately 1.0 × 105 cells/well and allowed to grow for 12 hours with 10% FCS. The medium was then replaced with fresh PBS. IBMX (1 mM), an inhibitor of a cyclic nucleotide phosphodiesterase, was added to each well 30 minutes before the addition of ADM to prevent breakdown of accumulated cAMP. After a 30-minute incubation, ADM (10−10–10−7 M) or PBS only (control) was added to the wells. The intracellular cAMP-elevating agent forskolin (10−6 M), an adenylyl cyclase activator, and 8-Br-cAMP (10−5 M), a membrane-permeable analogue of cAMP, were also added. The plate was incubated for 15 minutes at 37°C. After excess culture medium was aspirated, lysis reagent was added. The plate was shaken on a microtiter plate shaker for 10 minutes. Fifty-microliter samples of each unknown sample from the cell culture plate was transferred into the appropriate well of the immunoassay microtiter plate. An incubation time of 15 minutes was chosen based on a time course measurement of intracellular cAMP levels after ADM stimulation. cAMP was measured as described by the manufacturer’s nonacetylation protocol (Amersham Pharmacia Biotech, Ltd.). 
cGMP Measurement
Intracellular cGMP content was measured with the cGMP EIA kit as just described, with the exception that we used the manufacturer’s acetylation protocol. A 15-minute incubation time was chosen based on a time course measurement of intracellular cGMP levels after ADM stimulation. In some experiments, RPE cells were preincubated with l-NAME (10−4 M) in DMEM with 0.4% FCS for 30 minutes at 37°C. Then PBS with or without ADM (10−8 M) was added. 
[Ca2+]i Measurement
Intracellular Ca2+ was labeled by using the fluorescent dye fluo-3 AM and detected under a laser confocal scanning microscope (LCSM), which was able to acquire data in real time regarding the Ca2+ dynamics of individual cells in vitro. 22 Briefly, the fresh isolated, third- or fourth-passage, cells were grown respectively in culture medium on customized circular disks overnight at 37°C. For loading of fluo-3 AM, cells were incubated for 1 hour in the dark at 37°C with 10 mM membrane-permeant fluo-3 AM in normal physiological saline solution (N-PSS) that contained (in mM) 140 NaCl, 1 KCl, 1 CaCl2, 1 MgCl2, 10 glucose, and 5 HEPES [pH 7.4]. After the cells were rinsed twice with N-PSS, the circular disc with RPE cells attached was placed on the stage of a fluorescence microscope (Bio-Rad). ADM (10−7–10−12 M), 8-Br-cAMP (10−5 M) or isoprenaline (10−6M) dissolved in N-PSS were added. We also used N-PSS and freshly isolated cells as the control. All experiments were conducted at room temperature (20–24°C). The fluo-3 AM fluorescence was measured with a confocal microscope (model MRC-1024; Bio-Rad) using a krypton-argon laser and a fluorescein filter cartridge. The scanner and detectors were attached to an inverted microscope (Axiovert; Carl Zeiss Meditec). A Pian-Neofluar 40× (NA 1.3) oil-immersion objective was used for image acquisitions. The dye was excited at 488 nm, and fluorescent light was collected at 530 nm. 
The fluo-3 AM fluorescence signals were calibrated by addition of 10 μM A23187 to the medium (Fmax), followed by the addition of 10 mM EGTA and 20 mM NaOH to obtain Fmin. [Ca2+]i was calculated as described before using a K d of 370 nM at 37°C. 22  
Statistical Analysis
All experiments were performed in quadruplicate and repeated three times. Statistical analyses were performed with one-way ANOVA. Statistical significance was determined at P < 0.01. 
Results
Effect of ADM on RPE Migration
RPE cell migration was assayed by the Boyden technique, as described in the Methods section. In the presence of medium alone, RPE cells migrated minimally through the filter pores 1 . RPE cell migration was significantly increased when 10% FCS was added to the lower compartment of chambers. ADM inhibited both basal and 10% FCS-induced RPE migration in a dose-dependent manner at the concentrations of 10−8 and 10−7 M, respectively 1 . At 10−9 M, ADM did not significantly alter 10% FCS-induced and basal migration. The maximum inhibitory effect of ADM at 10−7 M on basal and 10% FCS-induced RPE migration was approximately 53.8% and 43.8% of control levels, respectively. Forskolin, an agent that affects cAMP levels, also inhibited FCS-induced RPE cell migration in a dose-dependent manner between 10−5 and 10−4 M and inhibited basal RPE cell migration at 10−5 and 10−4 M 1 . The viability of RPE cells was not affected by ADM at any of the doses tested (trypan blue dye exclusion test). 
Effect of ADM on RPE Cell Attachment
The effect of ADM on RPE cell migration may have been due to travail inhibition of cell attachment, a requirement for cell migration. RPE cell attachment was therefore measured on fibronectin, laminin, poly-d-lysine, and uncoated tissue culture plastic 2 . After 60 minutes of incubation, fibronectin, laminin, and poly-d-lysine significantly enhanced attachment of RPE cells. Exogenously added ADM (10−9–10−7 M) had no significant effects on RPE cell attachment to all tested substrates. Forskolin (10−6 M) also did not affect the attachment of RPE cells to all tested substrates. 
Effect of ADM and Forskolin on Intracellular cAMP Accumulation
ADM is known to exert its effect through the second messenger system involving cAMP. 2 3 4 7 3 shows the time-course of ADM effect on cAMP formation in primary cultures of RPE cells incubated in the absence and presence of IBMX. Despite the addition of a phosphodiesterase inhibitor, the time course of cAMP showed a “mountain-like” curve. The time cause curve at 15 to 20 minutes reached its maximum and subsequently declined after 30 minutes. At 60 minutes cAMP had reached control levels. To examine the effect of ADM stimulation on intracellular cAMP levels in RPE cells, we compared its effects on the cyclic nucleotide with other cAMP-elevating agents. As shown in 4 , incubation with ADM for 15 minutes increased intracellular cAMP levels in a dose-dependent manner (10−10–10−7 M). An effective response was observed at concentrations higher than 10−10 M. The maximum effect was observed at 10−7 M and was 186% of the control. In addition, forskolin (10−6 M) and 8-Br-cAMP (10−5 M) caused a marked increase in intracellular cAMP contents with 266% and 261% of control values, respectively. 
Effect of ADM and l-NAME on Intracellular cGMP
ADM is also associated with a decrease in cGMP levels under certain conditions. To investigate the change of intracellular cGMP in response to ADM, we compared the effect of ADM on intracellular cGMP with the effects of nitric oxide (NO) synthase inhibitor, also known to affect cGMP levels. 23 Even with the addition of a phosphodiesterase inhibitor, ADM induced a cGMP decline that peaked at 15 to 20 minutes 5 . Exposure of RPE cells to ADM for 15 minutes dose dependently decreased the intracellular cGMP level 6 . The maximum effect was obtained at 10−7 M, and the value was 25% of the control level. Pretreatment of RPE cells with l-NAME (10−4 M) further decreased the cGMP level, alone or when added with ADM (10−8 M), to cGMP levels of 4 ± 2 fmol/well and 3 ± 1 fmol/well, respectively. The difference was not significant (P > 0.01). ADM and l-NAME effects were neither additive nor synergistic. 
Effect of ADM on [Ca2+]i
Cell migration generally is known to involve Ca2+ signaling and changes in [Ca2+]i. 24 ADM induced a [Ca2+]i decrease in RPE cells that consisted of an initial decline within 30 seconds followed by a phase with a prolonged plateau that was parallel to the basal level 7 . ADM decreased [Ca2+]i in a dose-dependent manner 8 . The maximum response was observed at 0.1 μM, declining to 42.9% of the control. Exogenously added isoprenaline could slightly increase [Ca2+]i in RPE cells 7 . 8-Br-cAMP (10−5 M), a membrane-permeable analogue of cAMP, decreased [Ca2+]i to 55.6% of control level. The efficiency of ADM (10−7 M) and 8-Br-cAMP (10−5 M) to decrease [Ca2+]i was significantly different. No significant difference between fresh, isolated RPE cells and third- or fourth-passage cells was observed. 
Discussion
In this study, we attempted to elucidate for the first time the action of ADM on RPE cells, which occurs through two independent signal transduction pathways—cAMP accumulation and intracellular Ca2+ signaling—and is associated with inhibition of basal and 10% FCS-induced RPE cell migration without affecting cell attachment. The effect of ADM on Ca2+ flux is mediated, at least in part, by activation of a signaling pathway other than the cAMP-dependent process, because the ability of ADM to decrease the intracellular Ca2+ level is more significant than 8-Br-cAMP. We also found that the migration-inhibitory effect of ADM coincided with its ability to increase the intracellular level of cAMP. These data suggest that activation of the adenylate cyclase-cAMP system may mediate the physiological function of ADM on RPE cells. 
PVR is a well-recognized complication of retinal detachment and a major cause of failure for retinal reattachment surgery. RPE cells is the predominant cell type involved in this disease process, and migration of RPE cells is an early step in the development of PVR. 25 Several studies have shown that activators of RPE cells can be found in the vitreous during development of PVR. High levels of interleukin (IL)-1 and IL-6, 26 transforming growth factor (TGF)-β, 27 monocyte chemotactic protein (MCP)-1, 28 and PDGF 29 have been found in the vitreous humor or epiretinal membranes of patients with PVR. These inflammatory cytokines are known to be involved in the proliferation and migration of RPE cells and some are also inducers of RPE NO release. 30 In addition, ADM levels in the vitreous of patients with PVR have been found to be significantly higher than those of patients with other retinopathy. 16 This finding raises the possibility that ADM may also be involved in the pathologic processes of PVR. 
ADM is known to inhibit SMC migration in a dose-dependent manner by chemoattractants such as FCS and PDGF-BB. 18 Conversely, inhibition of FCS- and PDGF-induced SMC migration by ADM has been paralleled to an increase in the cellular level of cAMP. ADM can also inhibit PDGF-BB- and Ang II–stimulated migration of rat mesangial cells, at least in part through cAMP-dependent mechanisms. 31 We found ADM inhibited basal and 10% FCS-induced RPE migration in a dose-dependent manner at concentrations of 10−8 and 10−7 M. At 10−8 M, the inhibitory activity was 38.5% and 25.0% respectively. At 10−7 M ADM-induced inhibition was 53.8% and 43.8%, respectively. In fact, the percentage increase in cAMP level was strongly correlated with the percentage decrease in migration effect of RPE cells after treatment with ADM. An activator of adenylate cyclase, forskolin, also reduced FCS-induced and basal RPE cell migration. These data indicate that ADM inhibits the migration of RPE, probably through a cAMP-dependent mechanism. The adenylate cyclase/cAMP/PKA system may be involved in the migration-inhibitory effect of ADM in RPE cells. 
Inhibition of RPE cell migration by ADM is also correlated with a decrease in intracellular Ca2+ and cGMP. Intracellular Ca2+ increase is a central component of cellular activity during migration and is induced shortly after chemoattractant–receptor ligation. 32 In RPE cells, the increase in calcium signaling occurs after chemokine binding, and RPE cells thus appear to follow the general pattern of cellular reactivity in this context. 33 In the present study, we have also observed that ADM can block the increase in levels of intracellular cGMP in RPE cells. NO activates soluble guanylyl cyclase (sGC), and the resultant increase in cGMP is an important intracellular signaling pathway in the retina. 34 RPE NO synthase is known to be induced by inflammatory mediators such as lipopolysaccharide (LPS). 35  
We therefore propose the following mechanism for a regulatory role for ADM on RPE cell activity in the course of PVR development. PVR is known to be associated with a significant level of intraocular inflammation and the release of inflammatory mediators. Indeed, factors such as PDGF-BB released from platelets and IL-1 from monocytes activate RPE cells and induce NO release, and cell migration mediated by Ca increases, thus modulating cytoskeletal components for cell movement. 36 37 Thus, any mediator that can stimulate NO release and Ca increase is likely to induce RPE cell migration. ADM counterregulates this response by inhibiting cell migration through a mechanism that involves blocking the increase in cGMP and intracellular calcium. This mechanism itself involves reciprocal upregulation of intracellular cAMP and downregulation of cGMP and can be mimicked by other agents that alter the levels of these second messengers. The previous studies by Udono et al. 13 showed that a combination of two or three cytokines synergistically increases the ADM production in ARPE-19 cells. They also found that exogenously added ADM increased the number of F-0202 cells and ARPE-19 cells. They suggested in some inflammatory ocular disorders that ADM may stimulate the proliferation of RPE cells and other types of cells in an autocrine or paracrine fashion. In our study, we did not find significant effects of ADM on the growth of primary cultured RPE cells incubated with ADM up to 10−7 M for 24 to 48 hours. We therefore propose that the high levels of ADM found in the vitreous of eyes of patients with PVR may represent an increased production of this mediator by activated RPE cells within the eye, as a regulatory response to limit the extent of RPE cell activity. Clearly, in many clinical situations, this increased production of ADM may still be insufficient to control the PVR response. However, the data from this study open up the possibility of developing therapeutic agents that increase cAMP levels and decrease Ca signaling in RPE cells as treatments for PVR. Previous studies have shown that such agents can be effective in vitro and offer possibilities for the future. 
The pathogenesis of PVR is thought to consist of several critical steps: cell migration, cell adhesion, cell proliferation, and cell-mediated contraction of extracellular matrix components within the subretinal space and through retinal holes to form pathologic membranes on both surfaces of the neural retina. In the PVR epiretinal membrane, RPE cells predominate, but fibroblasts, macrophages, and glial cells are also found. 38 We have not found any reports about the effects of ADM on those kinds of cells. In contrast, RPE cells have been found to lose pigment and transform into fibroblast-like cells in vitro and epiretinal membranes (ERMs). Dedifferentiation of human RPE cells may induce different responses to the same stimulus. So the exact role of ADM in the pathogenesis of PVR is still a matter of discussion. Further investigation of the effects of ADM in the PVR animal model will provide insight to the pathogenetic role of ADM. 
Figure 1.
 
Effect of ADM and forskolin on the migration of RPE. ADM at concentration of 10−9 M and forskolin at 10−6 M had no significant effect on basal and 10% FCS-induced RPE migration. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 1.
 
Effect of ADM and forskolin on the migration of RPE. ADM at concentration of 10−9 M and forskolin at 10−6 M had no significant effect on basal and 10% FCS-induced RPE migration. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 2.
 
Effect of ADM and forskolin on the attachment of RPE. The value of control on uncoated tissue culture plastic was 0.38 ± 0.02. ADM and forskolin had no significant effects on RPE attachment. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 2.
 
Effect of ADM and forskolin on the attachment of RPE. The value of control on uncoated tissue culture plastic was 0.38 ± 0.02. ADM and forskolin had no significant effects on RPE attachment. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 3.
 
Time course of ADM effect on accumulation of cAMP in primary cultures of human RPE cells, incubated in the absence and presence of 1 mM IBMX. Points represent the mean ± SD of experiments performed in quadruplicate and repeated three times.
Figure 3.
 
Time course of ADM effect on accumulation of cAMP in primary cultures of human RPE cells, incubated in the absence and presence of 1 mM IBMX. Points represent the mean ± SD of experiments performed in quadruplicate and repeated three times.
Figure 4.
 
Effects of ADM, forskolin, and 8-Br-cAMP on intracellular cAMP formation in RPE. Averaged intracellular cAMP levels are shown as the mean ± SD (n = 12). The basal cAMP formation value of 1.0 × 105 RPE was 280 ± 34 fmol/well. *P < 0.01 versus control; **P< 0.001 versus control.
Figure 4.
 
Effects of ADM, forskolin, and 8-Br-cAMP on intracellular cAMP formation in RPE. Averaged intracellular cAMP levels are shown as the mean ± SD (n = 12). The basal cAMP formation value of 1.0 × 105 RPE was 280 ± 34 fmol/well. *P < 0.01 versus control; **P< 0.001 versus control.
Figure 5.
 
Time-course of the effect of ADM on accumulation of cGMP in primary cultures of human RPE cells. Points are the mean ± SD of experiments performed in quadruplicate and repeated three times.
Figure 5.
 
Time-course of the effect of ADM on accumulation of cGMP in primary cultures of human RPE cells. Points are the mean ± SD of experiments performed in quadruplicate and repeated three times.
Figure 6.
 
Effects of ADM and l-NAME on intracellular cGMP formation in RPE. Averaged intracellular cGMP levels are shown (mean ± SD, n = 12). The basal value of cGMP formation was 28 ± 4 fmol/well. After exposure to l-NAME (10−4 M) for 15 minutes, intracellular cGMP decreased markedly to 11% of the control. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 6.
 
Effects of ADM and l-NAME on intracellular cGMP formation in RPE. Averaged intracellular cGMP levels are shown (mean ± SD, n = 12). The basal value of cGMP formation was 28 ± 4 fmol/well. After exposure to l-NAME (10−4 M) for 15 minutes, intracellular cGMP decreased markedly to 11% of the control. *P < 0.01 versus control; **P < 0.001 versus control; NS, not significant.
Figure 7.
 
The response of [Ca2+]i to ADM, and isoprenaline (10−6 M). (A) Exogenously added ADM (10−10 M) induced the decrease of [Ca2+]i within 30 seconds. (B) Isoprenaline (10−6 M) initiated a slight increase of [Ca2+]i in RPE. Arrows: time point of drug administration.
Figure 7.
 
The response of [Ca2+]i to ADM, and isoprenaline (10−6 M). (A) Exogenously added ADM (10−10 M) induced the decrease of [Ca2+]i within 30 seconds. (B) Isoprenaline (10−6 M) initiated a slight increase of [Ca2+]i in RPE. Arrows: time point of drug administration.
Figure 8.
 
ADM induced a [Ca2+]i decrease in RPE in a dose-dependent manner. Averaged intracellular Ca2+ levels are shown (mean ± SD, n = 12). The value of normal RPE was 198.67 ± 24.63 nmol/L. *P < 0.01 versus control; **P < 0.001 versus control.
Figure 8.
 
ADM induced a [Ca2+]i decrease in RPE in a dose-dependent manner. Averaged intracellular Ca2+ levels are shown (mean ± SD, n = 12). The value of normal RPE was 198.67 ± 24.63 nmol/L. *P < 0.01 versus control; **P < 0.001 versus control.
 
The authors thank Qibing Mei and John V. Forrester for helpful discussions during various phases of the project; Chunmei Wang and Dan Chen for excellent technical support on intracellular Ca2+ measurement. 
Kitamura K, Kangawa K, Kawamoto M, et al. Adrenomedullin: a novel hypotensive peptide isolated from human pheochromocytoma. Biochem Biophys Res Commun. 1993;192:553–560.
Hinson JP, Kapas S, Smith DM. Adrenomedullin, a multifunctional regulatory peptide. Endocr Rev. 2000;21:138–167.
Withers DJ, Coppock HA, Seufferlein T, Smith DM, Bloom SR, Rozengurt E. Adrenomedullin stimulates DNA synthesis and cell proliferation via elevation of cAMP in Swiss 3T3 cells. FEBS Lett. 1996;378:83–87.
Yeung VT, Ho SK, Nicholls MG, Cockram CS. Adrenomedullin, a novel vasoactive hormone, binds to mouse astrocytes and stimulates cyclic AMP production. J Neurosci Res. 1996;46:330–335.
Gumusel B, Chang JK, Hao Q, et al. Adrenotensin: an adrenomedullin gene product contracts pulmonary blood vessels. Peptides. 1996;17:461–465.
Tanaka M, Kitamura K, Ishizaka Y, et al. Plasma adrenomedullin in various diseases and exercise-induced change in adrenomedullin in healthy subjects. Intern Med. 1995;34:728–733.
Moody TW, Miller MJ, Martinez A, Unsworth E, Cuttitta F. Adrenomedullin binds with high affinity, elevates cyclic AMP, and stimulates c-fos mRNA in C6 glioma cells. Peptides. 1997;18:1111–1115.
Chini EN, Choi E, Grande JP, Burnett JC, Dousa TP. Adrenomedullin suppresses mitogenesis in rat mesangial cells via cAMP pathway. Biochem Biophys Res Commun. 1995;215:868–873.
Barker S, Kapas S, Corder R, Clark AJ. Adrenomedullin acts via stimulation of cyclic AMP and not via calcium signalling in vascular cells in culture. J Hum Hypertens. 1996;10:421–423.
Takahashi K. Adrenomedullin: from a pheochromocytoma to the eyes. Peptides. 2001;22:1691.
Udono T, Totsune K, Takahashi K, et al. Increased expression of adrenomedullin mRNA in the tissues of intraocular and orbital tumors. Am J Ophthalmol. 2000;129:555–556.
Yousufzai SY, Ali N, Abdel–Latif AA. Effects of adrenomedullin on cyclic AMP formation and on relaxation in iris sphincter smooth muscle. Invest Ophthalmol Vis Sci. 1999;40:3245–3253.
Udono T, Takahashi K, Nakayama M, et al. Adrenomedullin in cultured human retinal pigment epithelial cells. Invest Ophthalmol Vis Sci. 2000;41:1962–1970.
Udono T, Takahashi K, Nakayama M, et al. Induction of adrenomedullin by hypoxia in cultured retinal pigment epithelial cells. Invest Ophthalmol Vis Sci. 2001;42:1080–1086.
Kirchhof B, Kirchhof E, Ryan SJ, Dixon JF, Barton BE, Sorgente N. Macrophage modulation of retinal pigment epithelial cell migration and proliferation. Graefes Arch Clin Exp Ophthalmol. 1989;227:60–66.
Udono T, Takahashi K, Takano S, Shibahara S, Tamai M. Elevated adrenomedullin in the vitreous of patients with proliferative vitreoretinopathy. Am J Ophthalmol. 1999;128:765–767.
Hackett S, Friedman Z, Campochiaro PA. Cyclic 3′,5′-adenosine monophosphate modulates retinal pigment epithelial cell migration in vitro. Arch Ophthalmol. 1986;104:1688–1692.
Horio T, Kohno M, Kano H, et al. Adrenomedullin as a novel antimigration factor of vascular smooth muscle cells. Circ Res. 1995;77:660–664.
Han QH, Hui YN, Du HJ, Zhang WJ, Ma JX, Wang SY. Migration of retinal pigment epithelial cells in vitro modulated by monocyte chemotactic protein-1: enhancement and inhibition. Graefes Arch Clin Exp Ophthalmol. 2001;239:531–538.
Hinton DR, He S, Graf K, et al. Mitogen-activated protein kinase activation mediates PDGF-directed migration of RPE cells. Exp Cell Res. 1998;239:11–15.
Jin M, He S, Worpel V, Ryan SJ, Hinton DR. Promotion of adhesion and migration of RPE cells to provisional extracellular matrices by TNF-alpha. Invest Ophthalmol Vis Sci. 2000;41:4324–4332.
Tortorici G, Zhang BX, Xu X, Muallem S. Compartmentalization of Ca2+ signaling and Ca2+ pools in pancreatic acini: implications for the quantal behavior of Ca2+ release. J Biol Chem. 1994;269:29621–29628.
Kogishi JI, Akimoto M, Mandai M, et al. Nitric oxide as a second messenger in phagocytosis by cultured retinal pigment epithelial cells. Ophthalmic Res. 2000;32:138–142.
Ehring GR, Szabo IL, Jones MK, et al. ATP-induced CA2+-signaling enhances rat gastric microvascular endothelial cell migration. J Physiol Pharmacol. 2000;51:799–811.
Machemer R, van Horn D, Aaberg TM. Pigment epithelial proliferation in human retinal detachment with massive periretinal proliferation. Am J Ophthalmol. 1978;85:181–191.
Limb GA, Little BC, Meager A, et al. Cytokines in proliferative vitreoretinopathy. Eye. 1991;5:686–693.
Connor TB, Jr, Roberts AB, Sporn MB, et al. Correlation of fibrosis and transforming growth factor-β type 2 levels in the eye. J Clin Invest. 1989;83:1661–1666.
Abu el-Asrar AM, Van Damme J, Put W, et al. Monocyte chemotactic protein-1 in proliferative vitreoretinal disorders. Am J Ophthalmol. 1997;123:599–606.
Robbins SG, Mixon RN, Wilson DJ, et al. Platelet-derived growth factor ligands and receptors immunolocalized in proliferative retinal diseases. Invest Ophthalmol Vis Sci. 1994;35:3649–3663.
Liversidge J, Grabowski P, Ralston S, et al. Rat retinal pigment epithelial cells express an inducible form of nitric oxide synthase and produce nitric oxide in response to inflammatory cytokines and activated T cells. Immunology. 1994;83:404–409.
Kohno M, Yasunari K, Minami M, et al. Regulation of rat mesangial cell migration by platelet-derived growth factor, angiotensin II, and adrenomedullin. J Am Soc Nephrol. 1999;10:2495–2502.
Schwab A. Function and spatial distribution of ion channels and transporters in cell migration. Am J Physiol. 2001;280:R739–R747.
Smith-Thomas L, Haycock JW, Metcalfe R, et al. Involvement of calcium in retinal pigment epithelial cell proliferation and pigmentation. Curr Eye Res. 1998;17:813–822.
Baldridge WH, Fischer AJ. Nitric oxide donor stimulated increase of cyclic GMP in the goldfish retina. Vis Neurosci. 2001;18:849–856.
Koga T, Zhang WY, Gotoh T, Oyadomari S, Tanihara H, Mori M. Induction of citrulline-nitric oxide (NO) cycle enzymes and NO production in immunostimulated rat RPE-J cells. Exp Eye Res. 2003;76:15–21.
Kutty RK, Kutty G, Hooks JJ, Wiggert B, Nagineni CN. Transforming growth factor-beta inhibits the cytokine-mediated expression of the inducible nitric oxide synthase mRNA in human retinal pigment epithelial cells. Biochem Biophys Res Commun. 1995;215:386–393.
Holtkamp GM, Kijlstra A, Peek R, de Vos AF. Retinal pigment epithelium-immune system interactions: cytokine production and cytokine-induced changes. Prog Retin Eye Res. 2001;20:29–48.
Charteris DG. Proliferative vitreoretinopathy: pathobiology, surgical management, and adjunctive treatment. Br J Ophthalmol. 1995;79:953–960.
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