Activation of endogenously expressed ion channels

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Nov 7, 2014 - addition to maxi-K channels and TRP channels absent from any insertion of a lytic pore. Keywords (4–6): complement system . Retinal pigment.

Activation of endogenously expressed ion channels by active complement in the retinal pigment epithelium Andreas Genewsky, Ingmar Jost, Catharina Busch, Christian Huber, Julia Stindl, Christine Skerka, Peter F. Zipfel, Bärbel Rohrer, et al. Pflügers Archiv - European Journal of Physiology European Journal of Physiology ISSN 0031-6768 Pflugers Arch - Eur J Physiol DOI 10.1007/s00424-014-1656-2

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Author's personal copy Pflugers Arch - Eur J Physiol DOI 10.1007/s00424-014-1656-2


Activation of endogenously expressed ion channels by active complement in the retinal pigment epithelium Andreas Genewsky & Ingmar Jost & Catharina Busch & Christian Huber & Julia Stindl & Christine Skerka & Peter F. Zipfel & Bärbel Rohrer & Olaf Strauß

Received: 4 November 2014 / Revised: 7 November 2014 / Accepted: 13 November 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract Defective regulation of the alternative pathway of the complement system is believed to contribute to damage of retinal pigment epithelial (RPE) cells in age-related macular degeneration. Thus we investigated the effect of complement activation on the RPE cell membrane by analyzing changes in membrane conductance via patch-clamp techniques and Ca2+ imaging. Exposure of human ARPE-19 cells to complementsufficient normal human serum (NHS) (25 %) resulted in a biphasic increase in intracellular free Ca2+ ([Ca2+]i); an initial peak followed by sustained Ca2+ increase. C5- or C7-depleted Electronic supplementary material The online version of this article (doi:10.1007/s00424-014-1656-2) contains supplementary material, which is available to authorized users. A. Genewsky Max-Planck Institute of Psychiatry, Munich, Germany A. Genewsky : I. Jost : J. Stindl : O. Strauß Experimental Ophthalmology, Eye Clinic, University Medical Center Regensburg, Regensburg, Germany C. Skerka : P. F. Zipfel Department of Infection Biology, Leibniz Institute for Natural Product Research and Infection Biology, Jena, Germany P. F. Zipfel Friedrich Schiller University, Jena, Germany B. Rohrer Department of Ophthalmology, Medical University of South Carolina, Charleston, SC 29425, USA B. Rohrer Research Service, Ralph H. Johnson VA Medical Center, Charleston, SC 29401, USA C. Busch : C. Huber : O. Strauß (*) Experimental Ophthalmology, Department of Ophthalmology, Charite University Medicine Berlin, Campus Virchow-Klinikum, Augustenburger Platz 1, 13353 Berlin, Germany e-mail: [email protected]

sera did not fully reproduce the signal generated by NHS. The initial peak of the Ca2+ response was reduced by sarcoplasmic Ca2+-ATPase inhibitor thapsigargin, L-type channel blockers (R)-(+)-BayK8644 and isradipine, transient-receptor-potential (TRP) channel blocker ruthenium-red and ryanodine receptor blocker dantrolene. The sustained phase was carried by CaV1.3 L-type channels via tyrosine-phosphorylation. Changes in [Ca2+]I were accompanied by an abrupt hyperpolarization, resulting from a transient increase in membrane conductance, which was absent under extracellular Ca2+- or K+-free conditions and blocked by (R)-(+)-BayK8644 or paxilline, a maxiK channel inhibitor. Single-channel recordings confirmed the contribution of maxiK channels. Primary porcine RPE cells responded to NHS in a comparable manner. Pre-incubation with NHS reduced H2O2-induced cell death. In summary, in a concerted manner, C3a, C5a and sC5b-9 increased [Ca2+]i by ryanodine-receptor-dependent activation of L-type channels in addition to maxi-K channels and TRP channels absent from any insertion of a lytic pore. Keywords (4–6): complement system . Retinal pigment epithelium . RPE . L-type channels . BK channels

Introduction Activation products of the complement system directly attack infectious microbes in the human organisms, opsonize cellular components to allow safe and inflammatory silent removal, and co-ordinate the accompanying inflammatory response, recruiting cells from the innate and adaptive immune system [45]. The active complement system generates efficient and generally toxic effector compounds and triggers secondary messenger cascades in responding cells. Despite the fact that ion channels in general and intracellular free Ca2+ more specifically are involved in the regulation of almost

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all cellular functions, little is known about the activation of ion channels in response of host cells or autologous cells to complement activation. Complement-dependent activation of ion fluxes has been reported in erythrocytes [4, 12–14, 32], leukocytes [26, 35], fibroblasts [27, 30, 31], Ehrlich ascites cells [5], macrophages [16], oligodendrocytes and glial cells of the central nervous system [17, 25, 28, 38]. A careful analysis of the type of ion channels formed by the activated complement system components suggests that specific endogenously expressed ion channels are activated: the maxiK-Ca2+-dependent K+ channels [30], voltage-dependent L-type Ca2+ channels [27] and the Kv1.3 delayed rectifier K+ channels [38]. In lung epithelial cells the involvement of Ca2+ release from intracellular Ca2+ stores has been observed for inflammasome activation [41]. Deregulation of the complement system’s alternative pathway is involved in the etiology of many degenerative diseases and plays a central or possibly even an obligatory role [39]. In age-related macular degeneration (AMD), which represents the most common cause for blindness in elderly people of industrialized countries, single nucleotide polymorphisms (SNPs) in a number of the complement regulatory proteins are associated with an increased risk for AMD. Many of these SNPs are thought to affect the regulation of the alternative pathway [34]. AMD is associated with chronic inflammation and cell loss of the retinal pigment epithelium (RPE) [37], which forms a functional unit with the light-sensitive photoreceptors in the retina [36]. In this functional unit the RPE fulfills a variety of duties which are essential for visual function [36]. Therefore a loss of the RPE leads to a secondary loss of photoreceptors which subsequently leads to blindness. Complement products have been found in drusen [11, 29], which are deposits located between the RPE and Bruch’s membrane, and represent the typical hallmark of the disease. In addition, activation of complement components such as C3a, C5a and terminal complement complex (TCC, sC5b9), also termed membrane attack complex (MAC), have been identified at the level of the RPE, Bruch’s membrane and the choroid [1, 11]. Furthermore, in these structures the endogenous complement regulators, such as Factor H, FHL1, CD55 [8] and CD59 [7], have been shown to be altered in level and localization in AMD patients. Based on the histological and genetic evidence, it is suggested that pathologic activation of the alternative pathway of complement, damages intact RPE cells and this ultimately leads to the development of AMD. RPE injury might be executed by the increased formation of the terminal sC5b-9. The soluble constituents the complement system are synthesized predominantly by hepatocytes in the liver. However recently, local production of complement components and regulators has been shown for the RPE and the choroid, thereby forming a local complement system [1, 11]. This could explain why polymorphisms in complement protein

genes affect only the retina and not the entire body. The RPE is known for its function as an active blood/retina barrier, which interacts with the immune system by either inhibition or activation [44]. The RPE secretes factor H and FHL1 as well as C3 and also expresses toll-like receptors, MHC receptors and can be stimulated by interleukin-1 (IL-1) [33]. In addition it secretes immune modulatory factors such as CFH [10, 20], cytokines [19], or TGF-beta [15]. Finally, despite the expression of endogenous complement inhibitors, upon oxidative stress the RPE is sensitized by sublytic complement components [40], leading to the activation of intracellular signaling cascades including Ras, Erk and Src signaling [23], secretion of vascular endothelial growth factor (VEGF-A) [40] and the activation of matrix metalloproteinases [2], generating a pathological environment. Complement attack upon oxidative stress results in the exposure of neoepitopes on the RPE cell surface and changes its morphology resulting in modified “selfstructures” [18]. Thus deregulation of the alternative pathway probably leads to increased local activation in the retina which results in complement-mediated changes in cell behavior. Sublytic sC5b-9 complement complexes have effector function, in addition to pore formation including cytokine like activity and regulation of inflammation. Terminal complement activation results in the assembly, surface binding or likely insertion of limited amounts of the sC5b-9 complex into the cell membrane, or the partial assembly of the terminal complement complex. None of the two scenarios causes cell lysis, but rather cause various metabolic effects. Here we continue to explore the role of sublytic sC5b-9 on complement activation and the potential involvement of C5b9 as a cellular trigger or as a pore formation. A careful analysis, comparing the activation of endogenously expressed ion channels with changes in nonspecific ion conductance in response to complement would provide further clues. Therefore, we investigated ion channel activity under the control of the complement system via the patch-clamp technique, which represents the most sensitive experimental set-up to detect changes in the membrane conductance of cells. By means of patch-clamp recordings together with intracellular Ca2+imaging we observed no indications for the insertion of a terminal complement complex (TCC) related nonspecific pore, but rather the concerted activation of endogenously expressed ion channels resulting in a finely tuned increase in intracellular free Ca2+ as a second-messenger.

Materials and methods Cell culture Human RPE cells (ARPE-19, LGC Standards/ATCC) were maintained in DMEM/Ham’s F12 (E15-813, PAA) supplemented with 10 % (v/v) fetal bovine serum (FCS) (F7524,

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Sigma-Aldrich), 20 mM sodium bicarbonate (S11-002, PAA) and 0.5 % penicillin/streptomycin (P11-010, PAA). Cells were maintained at 37 °C in a humidified atmosphere (95 % air, 5 % CO2) and subcultivated when 90 % confluency was reached. Primary porcine retinal pigment epithelial cultures were established as described previously [9]. In brief, eyes were obtained from local slaughter houses. After removing the anterior parts of the eye, RPE cells were harvested after papain digestion. Cells were seeded at a high density (1.2×106 cells per ml) so that they did not need to proliferate to form confluent monolayers and keep a high degree of differentiation. Cells were incubated in alpha-modification of MEM supplemented with non-essential amino acids, THT (hydrocortisone (20 μg/L), taurine (250 mg/L), and triiodo-thyronine (0.013 μg/L) and 10 % FCS under standard culture conditions. Prior to all experiments, ARPE-19 or porcine cells were subjected to serum-free conditions for 24 h. Electrophysiology Voltage- and current-clamp, whole-cell and cell-attached patch clamp recordings were obtained from serum-deprived cells grown on 12 mm glass cover slips (6.5×103cells/cm2) and placed into a custom-made recording chamber (500 μL bath volume). Electrodes with a resistance of 3–5 MOhms (whole cell mode) or 8–10 MOhms (cell-attached mode) were pulled from borosilicate glass (GB150T-8P, Science Products) on a Zeitz DMZ-Universal-Puller (Zeitz Instruments), generating seal resistances of 1–10 GOhms. The pipette filling solution for whole-cell recordings contained (in mM): 10 NaCl, 100 KCl, 2 MgSO4, 0.5 CaCl2, 5.5 EGTA, 10 HEPES, 0.16 nystatin, adjusted to pH 7.2 (Tris powder). The standard 1.25× extracellular solution (concentrated for the supplementation with 25 % e.g. NHS) contained (in mM): 170 NaCl, 7.25 KCl, 0.51 MgSO 4, 0.6 MgCl 2, 1.19 CaCl 2, 5.21 NaHCO3, 1.38 NaH2PO4, 31.25 HEPES, 13.88 glucose adjusted to pH 7.2 (Tris powder). For calcium-free conditions the standard extracellular solution was used without CaCl2, but supplemented with 12.5 mM EGTA. For potassium-free conditions the 1.25× extracellular solution was used, but KCl was replaced with CsCl. The pipette-filling and extracellular solution (1.25×) for cell-attached recordings contained (in mM): 167.5 KCl, 13.75 NaCl, 1.25 KH2PO4, 1.13 MgCl2, 1.25 CaCl2, 12.5 HEPES adjusted to pH 7.4 (NaOH). Whole-cell and cell-attached currents were amplified using a L/M-EPC-7 patch clamp amplifier (List Medical Electronic) and digitized at 10–50 kHz using a NI PCI-MIO-16E-4 (PCI6040E) 12-bit data acquisition board (National Instruments). Stimulation, acquisition and data analysis were carried out using WinWCP V4.3.1, WinEDR V3.2.4 (University of Starthclyde, Glasgow) and GNU Gnumeric. Fast and slow capacitive transients were canceled online by means of analog circuitry. Residual capacitive transients were canceled in

voltage clamp by the P/4 method [3]. For analysis the recording were low-pass filtered offline at 1–5 kHz. Calcium imaging To assess the intracellular free Ca2+, serum-deprived cells grown on 18 mm glass cover slips (1.5×104 cells/cm2) were incubated with 2 μg/ml fura-2/AM (F1221, Invitrogen) in serum-free medium for 45 minutes at 37 °C (humidified, 95 % air, 5 % CO2). After loading the cells with fura2/AM, the cover slips were placed in a custom-made recording chamber (filled with 250 μl of the respective bath solution, see electrophysiology section) and measured using a Zeiss Axiovert 40 CFL inverted microscope (Carl Zeiss AG) equipped with a 40× oil immersion objective, a Visichrome High Speed Polychromator System (Visitron Systems) and a high resolution CCD camera (CoolSNAP EZ, Photometrics). Analysis and control were carried out using the MetaFluor Fluorescence Ratio Imaging Software (Visitron Systems). The fluorescence intensity of Fura-2 was detected at an emission wavelength of 505 nm while the excitation wavelengths were set to 340/380 nm, respectively. Changes in intracellular free Ca2+ are all given as ratios of the fluorescence of the two excitation wavelengths (dF/F) and normalized to baseline (ddF/F). Taking into account previous work with ARPE-19 cells using the same method the NHS-induced changes in the ratio (resting values at 0.6 units and NHS ddF/F 0.4 units) are below fura-2 saturation and above fura-2 detection levels [6]. Cell viability assay To assay the cell viability, 3×104 cells/cm2 were seeded to 96well cell culture plates and after 24 h growth subjected to serum-free conditions. After additional 24 h the cells were exposed to the respective treatments for 16 h. After treatment, the medium was aspirated and replaced with serum-free medium, supplemented with 50 μmol/L resazurin (R7017, Sigma-Aldrich) and incubated for 1 h at 37 °C (humidified, 95 % air, 5 % CO2). The fluorescent dye resorufin was detected using a multi-well plate reader at 530/590 nm (ex/abs) as an endpoint measurement. Western blot For immunoprecipitation of calcium channels, ARPE-19 cells (1 well of a 6-well plate) were lysed with 100 μl denaturing lysis buffer (RIPA buffer: in [mM] 10 Tris-Cl, 1 EDTA, 0.5 EGTA, 140 NaCl; in [%] 1 Triton X-100, 0.1 sodium deoxycholate, 0.1 SDS; pH 8.0) supplemented with 1× Halt™ protease inhibitors and 1 mM phenylmethanesulfonylfluoride(Fisher Scientific). Samples were incubated with 1 μg of the anti-CaV1.3 antibody (Alamone Labs) overnight at 4 °C with agitation, followed by addition of 70 μL of ProteinA-Agarose beads (Cell Signaling

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Technology) and continued incubation for 4 hrs on ice. Immunoprecipitated complexes were collected by centrifugation (3000×g for 2 min at 4 °C) and washed 3-times by resuspension and centrifugation with lysis buffer. Finally, each pellet was resuspended in 50 μL of loading buffer, heated to 95 °C for 5 min and centrifuged (12,000×g) prior to loading. Samples were separated by electrophoresis on a 4–20 % BisTris polyacrylamide gel (Bio-rad), and proteins transferred to a PVDF membrane. The membrane was incubated with primary antibody (rabbit polyclonal, PhosphoSer/Thr; Cell Signaling Technology) overnight at 4 °C, followed by secondary antibody binding (anti-rabbit IgG, HRP-linked; Sigma). Proteins were visualized using a chemiluminescence detection kit (Immobilon Western; Millipore Corporation, Billerica, MA). The intensity of the bands was quantified using the Alpha Innotech Fluorchem 9900 imaging system running Alpha Ease FC Software 3.3 (Alpha Innotech, San Leandro, CA). Normal human serum (NHS) All experiments were conducted with commercially available human serum (C15-051, PAA), which was aliquoted and stored at −20 °C until use. Heat-inactivation was performed at 57 °C for 45 minutes. The complement proteins within the human serum were found to be highly active verified with a standard hemolysis assay conducted with porcine erythrocytes. Complement factor depleted sera were purchased from CompTech, Complement Technology Inc., Texas, USA. Statistical analysis All data are represented in mean values±SEM. Statistical significance was calculated using one-way ANOVA combined with the Bonferroni post-hoc test [p values *p60 % (ddF/F 0.09±0.004; [ddF/F]/min 0.12±0.02). Taken together, the initial Ca2+ rise included the release of Ca2+ from cytosolic Ca2+ stores and activation of L-type channels with ryanodine receptors. Second, we investigated the contribution of ion channels to the sustained phase of the NHS-induced Ca2+-response (Fig. 2). The sustained phase was fully blocked by (R)-(+)BayK8644 (10 μM), a potent L-type channel blocker (Fig. 2a–b). Long-term L-type Ca2+ channel activation is correlated with phosphorylation [40]. Phosphorylation of the L-type channel expressed in the RPE, CaV1.3, was investigated by means of western blotting 1 h after application of NHS. Here, CaV1.3 channel protein showed increased tyrosinephosphorylation 1 h after serum stimulation, which was not observed with heat-inactivated serum (Fig. 2c–d). Thus the long lasting, sustained phase of the Ca2+ increase involves an influx of Ca2+ through L-type channels of the CaV1.3 subtype, which showed sustained activity due to phosphorylation of the pore-forming subunit.

Changes in intracellular free Ca2+ of ARPE-19 cells Activation of ion channels by complement in ARPE-19 cells ARPE-19 cells were challenged by normal human serum NHS (25 %) application by extracellular solution containing 25 % normal human serum (NHS). In the first set of experiments, the effect of complement challenge was studied by Ca2+imaging of fura-2/AM loaded RPE cells. Addition of NHS resulted in elevation in intracellular free Ca2+ consisting of an initial peak (ddF/F 0.25±0.005; [ddF/F]/min 0.49±0.05) followed by a long-lasting Ca2+increase (Fig. 1a). When the RPE cells were treated with heat-inactivated HS (HINHS) no

In summary, these first data indicate a physiological response upon NHS challenge resulting in an increase in intracellular free Ca2+ involving the activation of endogenously expressed ion channels. Since most of the early and sustained Ca2+ signal could be eliminated by specific ion channel blockers, we could not find a strong indication that NHS led to the formation of a nonspecific pore expected for the insertion of sC5b-9 into the plasma membrane. To support this hypothesis

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Fig. 1 Ca2+ transients activated by normal human serum in ARPE-19 cells. a: Example of a full response in an ARPE-19 cell; changes in intracellular free Ca2+ were given as fluorescence ratio between the two excitation wavelengths 340 nm and 380 nm. The black bar indicates the time of 25 % normal human serum (NHS) application. b: Ca2+ transients at the initial peak phase induced by serum application under different experimental conditions: from left to right: 25 % NHS control, 25 % heatinactivated NHS (HINHS), NHS plus preincubation with the SERCA blocker thapsigargin (1 μM), NHS plus L-type channel blocker (R)-(+)BayK8644 (10 μM), NHS plus isradipine (5 μM), NHS plus the ryanodine receptor blocker dantrolene (1 μM) and NHS plus ruthenium-red (1 μM); total number of cells and number of independent experiments are given in D. c: Analysis of the slope of the serum induced Ca2+ transients under various experimental conditions,

from left to right: 25 % NHS control, 25 % HINHS, preincubation with the SERCA blocker thapsigargin (1 μM), L-type channel blocker (R)-(+)BayK8644 (10 μM), isradipine (5 μM), the ryanodine receptor blocker dantrolene (1 μM) and ruthenium-red (1 μM). The slope is given in difference in fluorescence ratio per time in min. d: Changes in intracellular free Ca2+ induced by human serum or under various experimental conditions where NHS was applied with indicated blockers. Insets indicate the total number of cells and the number of independent experiments. e: Changes in the slope of rising intracellular free Ca2+ induced by human serum or under various experimental conditions where NHS was applied with indicated blockers. Total number of cells and independent experiments same as in D. (Data are given as mean values±SEM significance is indicated as ***=p25 % (0.184±0.018 ddF/F), and challenge with C7-depleted serum decreases the Ca2+ peak by >30 % (0.163±0.005ddF/F) as compared to the effects of NHS application (Fig. 7a, b). The use of C3-, C5- and in particular C7-depleted human serum shows that the complement mediated effect on intracellular free Ca2+ release is –most likely- at least also mediated by the terminal components sC5b-9. The effects of human serum treatment on ARPE-19 cells challenged with oxidative stress Both Ca2+ imaging experiments and patch-clamp recordings suggest that the plasma membrane remained intact upon serum challenge and did not result in the insertion of sC5b-9 in ARPE-19 cell plasma membrane. In contrast, this cell response suggests a triggered and orchestrated Ca2+ signal which is generated by the activation of endogenously expressed ion channels. This probably may trigger specific different downstream effects. Therefore we next examined the

effect of NHS treatment on cell viability in combination with an oxidative stress challenge. Subconfluent ARPE-19 cells incubated for 16 h with serum-free medium (control), were treated with NHS (25 %) or HINHS (25 %) in combination with different concentrations of H2O2. Cell-viability was measured using the Alamar-Blue/Resazurin assay. Under control conditions and in the presence of heat-inactivated serum, cells exhibited the same concentration-response curves. However, upon challenge with NHS, the concentration-response curve was shifted to∼ten times higher concentrations of H2O2 (Fig. 8a). This can be calculated as survival rates at 1 mM H2O2 0.8±1.3 % for untreated cells, 7.5±1.4 % for HINHS treated cells and 51.4±3.5 % for NHS treated cells survived oxidative stress. Thus exposure of ARPE-19 cells with NHS does not decrease cell viability but rather has protective functions via Ca2+-dependent changes in cell metabolism.

Discussion In this study we provide evidence that active complement components induce a finely tuned Ca2+ response, which is generated by the orchestrated activation of endogenously expressed ion channels. This Ca2+ signal appears to be generated by active components of the complement system and seems to activate mechanisms that protect RPE cells against oxidative stress. These results suggest that the RPE cells directly respond to complement activation products (effector components generated by the activated complement system) and thereby the cells in combination with endogenous regulators or modulates modifying its own environment in response to complement activation. This novel mechanism for

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Fig. 8 Cell viability of ARPE-19 cells exposed to H2O2 and normal human serum. a: Plot of cell viability measured with Alamar-blue/ resazurin assay normalized to 100 μM H2O2 individually and plotted versus different concentrations of H2O2; data is from non-treated cells (serum-free), from cells treated with NHS and from cells treated with

heat-inactivated NHS. b: Comparison of cell viability at 1 mM H2O2 normalized to control: non-treated cells, NHS treated cells and heatinactivated NHS treated cells. (Data is given as mean values±SEM; n=24/2; ***=p

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