Pflugers Arch - Eur J Physiol DOI 10.1007/s00424-014-1561-8

ION CHANNELS, RECEPTORS AND TRANSPORTERS

Activation of ATP secretion via volume-regulated anion channels by sphingosine-1-phosphate in RAW macrophages Philipp Burow & Manuela Klapperstück & Fritz Markwardt

Received: 30 May 2014 / Accepted: 17 June 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract We report the activation of outwardly rectifying anion currents by sphingosine-1-phosphate (S1P) in the murine macrophage cell line RAW 264.7. The S1Pinduced current is mainly carried by anions, because the reversal potential of the current was shifted by replacement of extracellular Cl− by glutamate− but not when extracellular Na+ was substituted by Tris+. The inhibition of the current by hypertonic extracellular or hypotonic intracellular solution as well as the inhibitory effects of NPPB, tamoxifen, and glibenclamide indicates that the anion current is mediated by volume-regulated anion channels (VRAC). The S1P effect was blocked by intracellular GDPβS and W123, which points to signaling via the S1P receptor 1 (S1PR1) and G proteins. As cytochalasin D diminished the action of S1P, we conclude that the actin cytoskeleton is involved in the stimulation of VRAC. S1P and hypotonic extracellular solution induced secretion of ATP from the macrophages, which in both cases was blocked in a similar way by typical VRAC blockers. We suppose that the S1P-induced ATP secretion in macrophages via activation of VRAC constitutes a functional link between sphingolipid and purinergic signaling in essential processes such as inflammation and migration of leukocytes as well as phagocytosis and the killing of intracellular bacteria.

Keywords Volume-regulated anion channel . Voltage clamp . Macrophage . Sphingosine-1-phosphate . ATP secretion

P. Burow : M. Klapperstück : F. Markwardt (*) Julius Bernstein Institute for Physiology, Martin Luther University Halle, Magdeburger Str. 6, 06097 Halle/Saale, Germany e-mail: [email protected]

Introduction Sphingolipid metabolites play an important role in the immune and inflammatory system [5]. A key mediator is sphingosine-1-phosphate (S1P), which is produced by phosphorylation of sphingosine catalyzed by sphingosine kinase (SphK). Apart from its role in regulating lymphocyte egress from lymphoid organs, S1P influences the complex reactions of the innate immune system during the defense against infectious organisms. Furthermore, S1P is also of importance during aberrant or exacerbated production of inflammatory cytokines in autoinflammatory disorders and sepsis [6]. SPhK is activated by toll-like receptors (TLR2 and TLR4) that bind bacterial lipopeptides and lipopolysaccharide (LPS) from Gram-negative bacteria, and also by other mediators released during inflammation like platelet-derived growth factor (PDGF), tumor necrosis factor alpha (TNF-α), thrombin, IgE-bound antigen, and nucleotides [60, 70, 77, 35, 69, 26]. S1P signals to lymphocytes, macrophages, and macrophagelike cells as well as to endothelial cells via five different S1P receptor subtypes (S1PR1–S1PR5) [6, 68, 26]. S1P signaling is upregulated in autoinflammatory diseases and sepsis [21, 70]. Inhibition of SPhK was shown to protect mice from sepsis [58]. Fingolimod (FTY720) is applied as an immunosuppressant in patients suffering from multiple sclerosis. FTY720 is able to act as an agonist as well as a functional antagonist of S1PRs via desensitization of the S1P receptor [11, 41, 60]. In macrophages or macrophage-related cells such as monocytes and microglia, S1P is involved in regulation of proliferation and apoptosis [76, 22, 78, 20], chemotaxis [44, 75, 50], secretion of cytokines and NO [58, 45, 57, 51], and phagocytosis [76], as well as in killing of intracellular bacteria [47, 57, 19]. Furthermore, it was shown that S1P can induce actin assembly [42], which might take place via secretion of nucleotides and activation of P2X7 receptors [40, 39]. P2X7

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receptors also play an important role in inflammatory processes like the secretion of interleukin 1 and interleukin 18 [17, 28, 12, 8] and the killing of intracellular microorganisms [9, 56, 2]. Other nucleotide P2 receptors are involved in chemotaxis [36, 66]. Hence, S1P-dependent secretion of nucleotides may link inflammatory signaling mediated via nucleotides acting on G-protein-coupled P2Y or ionotropic P2X receptors and the many cellular responses elicited by the stimulation of S1P receptors (S1PR). We asked how the S1P-dependent release of nucleotides, which couples sphingolipid signaling to purinergic effects, may occur in macrophages. In the present report, we demonstrate in RAW macrophages that S1P activates volumeregulated anion channels (VRAC) that mediate ATP secretion in these cells. Therefore, VRAC may potentially constitute pharmacological targets for the treatment of inflammatory disorders.

Materials and methods Chemicals and solutions The standard bath solution contained (in mM) 140 NaCl, 5.4 KCl, 10 glucose, 10 Hepes, 0.5 MgCl2, and 1 CaCl2, pH 7.4 adjusted with NaOH, and had an osmolarity of 305 mOsmol/kg H2O. For current measurements, the bathing solution was exchanged to an isoosmolar solution consisting of (in mM) 140 NaCl, 10 glucose, 10 Hepes, 2 MgCl2, and 1 CaCl 2 , pH 7.4 adjusted with NaOH (osmolarity 303 mOsmol/kg H2O). K+ was omitted to suppress K+ currents and the Mg2+ concentration was increased to avoid possible effects of secreted ATP on P2X7 receptors. Hypoosmolar solution was made by partly omitting NaCl (100 mM NaCl, 230 mOsmol/kg H2O), and hypertonic solution by adding NaCl (180 mM NaCl, 390 mOsmol/kg H2O). Sphingosine-1-phosphate (Cayman Chemicals, Ann Arbor, MI, USA) was dissolved in methanol to make a 1-μM stock solution. This solution was sonicated and diluted to the 10 nM used to activate the currents. The final concentration of methanol in the ready-made solution did not exceed 1 vol%. The intracellular pipette solution consisted of (in mM) 120 Naaspartate, 10 glucose, 10 Hepes, 2 CaCl2, 5.5 MgCl2, 3 EGTA, 3 BAPTA, and 5 NaATP, pH 7.2 adjusted with NaOH (300 mOsmol/kg H2O). The free intracellular Ca2+ concentration ([Ca2+]i) was calculated to 73 nM according to Schubert et al. [65]. Hypoosmolar or hyperosmolar pipette solutions were made by decreasing or increasing Na-aspartate (for hypoosmolar: 90 mM Na-aspartate, 250 mOsmol/kg H2O; for hyperosmolar: 160 mM Na-aspartate, 375 mOsmol/ kg H2O). Osmolarities were measured using a freezing point osmometer (OM-815, Vogel GmbH, Gießen, Germany).

Electrophysiology For a detailed description of the voltage clamp technique, see Kuehnel et al. [39]. RAW 264.7 cells were grown on polylysine-coated coverslips in cell culture medium (DMEM with high glucose, Biochrom AG, Berlin, Germany) with 4 mM L-glutamine, 4.5 g/l glucose, 10 % fetal calf serum, 100 U/ml penicillin, and 100 μg/ml streptomycin to a non-confluent density for recording from single cells. For electrophysiological measurements, the coverslips were transferred to a perfusion chamber and superfused by the standard bath solution (see above). The flow rate of the bath solution did not exceed 1 ml/ min in order to reduce ion currents evoked by laminar shear stress. Patch pipettes were fabricated from borosilicate glass capillaries and had tip resistances of 4–7 MΩ after filling with pipette solution. Only cells with access resistances (measured after establishing the whole-cell configuration) below 15 MΩ were used for the experiments. For rapid change of the bathing solution, the Utube technique was used [3]. For measuring whole-cell conductance and current reversal potential, voltage ramps were used every 1 s going from −80 to +40 mV within 500 ms. The currents were recorded and filtered at 1 kHz using an Axopatch 200A amplifier (Axon Instruments, Inc., Foster City, USA) and sampled at 2 kHz. The data were stored and analyzed on a personal computer using software written in our department (Superpatch 2000, SPAnalyzer by T. Böhm). All voltage clamp experiments were done at room temperature (20–22 °C). Measurement of the intracellular Ca2+ concentration [Ca 2+ ] i changes were monitored using the calciumsensitive fluorescent indicator fluo-4. Cells were loaded with fluo-4 AM (2 μM; 1-mM stock solution in anhydrous DMSO; Molecular Probes, Eugene, OR, USA) by incubation in bathing solution (30 min, 37 °C). Cells were adhered by poly-L-lysine in a perfusing chamber mounted on an inverted phase contrast microscope (Diaphot TMD, Nikon, Düsseldorf, Germany). All measurements were done at 37 °C. The Ca indicator was excited via a×40 fluorine objective (Nikon) using a 75 W Xenon lamp at 460–490 nm. Fluorescence signals were collected at 520– 540 nm. Images were obtained by an intensified charge coupled device (ICCD) camera C 3077 and digitized by the Video-to-FireWire-Converter “The Imaging Resource” (Hamamatsu Photonics, Germany). Data acquisition of single cells was done using the Wasabi 1.5 program by Hamamatsu. The analysis of fluorescence data by regions of interest was calculated by ImageJ (http://rsb.info.nih. gov/ij). Cells were superfused with standard bath solution and images were taken every 1 s. To elicit [Ca 2+ ] i

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Fig. 1 S1P-induced increase in ion conductance in RAW macrophages. a Typical example of ion currents activated by S1P application in isoosmolar solution in mouse RAW macrophages. After establishing the whole-cell configuration, the cell membrane was clamped to a holding potential of 40 mV interrupted by voltage ramps as shown in d and g. b, e

Single ramp current records before (b) and during (e) application of S1P via the U-tube. Three to five ramp currents before U-tube-mediated application of solution were considered leak currents and were averaged and subtracted from the current (c) to visualize the application-dependent currents (f)

responses 20 s later, the perfusion solution was switched to the same buffer containing S1P as indicated in the figure. The normalized relative fluorescence (Frel) was calculated according to

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where Frel denotes the relative fluorescence and F the timedependent fluorescence, Fmin is the fluorescence of the background, and Fmax is the maximal fluorescence of each cell during application of the Ca2+ ionophore A23187. [Ca2+]i measurements were performed at 37 °C using a heatable cell chamber (Harvard Apparatus, March–Hugstetten, Germany).

These experiments were performed in phenol-red-free cell culture medium at 37 °C on a heat block. ATP released into the culture medium of RAW cells was measured using a luciferin-luciferase assay (FL-AA, Sigma). Briefly, RAW cells cultured in 35-mm Petri dishes were grown to 70–80 % confluence in cell culture medium. One hour before measurement, the cell culture medium was replaced by 2 ml of fresh medium to reduce the ATP background. The remaining ATP background for each Petri dish was measured in a 50-μl medium sample taken directly before starting the experiment. At the indicated sampling intervals, a 50-μl sample of cell supernatant was obtained and immediately delivered into liquid nitrogen. After that, the cells were mechanically removed from the Petri dish and their protein concentration was measured using

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the BCA method as described previously. For luminometric ATP measurements, samples were defrosted and briefly centrifuged at 1,000 rpm for 30 s. Ten microliters of each sample was diluted with 90 μl of ATP assay mix in a glass tube to reduce the inhibitory effect of high salt concentrations on the luciferin-luciferase reaction. Luminescence was then measured immediately by a luminometer (Lumat LB 9507, Berthold Technologies, Bad Wildbad, Germany). ATP calibration standards were run for each experiment. As revealed by appropriate control experiments, the ionic conditions and substances used in our experiments did not directly interfere with the ATP assay. Data presentation and statistics The data presentation was accomplished using the program SigmaPlot (Systat Software Inc.). Averaged data are presented as means±SEM. Test of significant differences between mean values was performed by one-way ANOVA. The statistical

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significance of differences between pairs of means was then tested using the multiple t test (Bonferroni) with the SigmaPlot program. Significance was set at p2.0 were used for data evaluation. The reversal potential shifted to positive values and the slope conductance was greatly reduced when extracellular Cl− ions were replaced by glutamate− or aspartate− (Fig. 2b, d, e), whereas substitution of Na+ by Tris+ was without effects on

Vrev and G (Fig. 2c, d, e). Therefore, it could be inferred that S1P evoked an anion conductance. Dependence of the S1P-dependent current on osmolarity and intracellular ATP The outwardly rectifying current showed characteristics of the volume-regulated anion channel (VRAC) described in other macrophage-like cells such as fibroblasts [64] and microglia [80]. We therefore examined whether manipulations of the cell volume by changing the extracellular or intracellular osmolarity could induce similar currents like S1P in normal extracellular solution, and if osmolarity has an impact on S1P-induced currents in RAW macrophages. Indeed, as shown in Fig. 3, extracellular hypoosmolarity induced currents similar to those evoked by S1P, i.e., with about the same time course and magnitude (Fig. 3c). Likewise, the currents induced by reducing the extracellular osmolarity exhibited reversal potentials (−30.5±6.5 mV) and outward rectification indices (3.5±1.0) that were both not significantly different from the S1Pinduced currents. Additionally, the use of a hypoosmolar intracellular solution inhibited the increase in the membrane conductance by hypotonicity as well as by S1P (Fig. 3b, d),

For a further characterization of the ion channels mediating the S1P-induced current, we applied various substances reported to block VRAC [53]. As shown in Fig. 4, conductances activated by both the hypoosmolar extracellular solution and S1P displayed indistinguishable action profiles to VRAC blockers, indicating that the S1P- and the hypoosmolarityactivated ion channels behave identically, i.e., S1P presumably activates VRAC in RAW macrophages.

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It is known that the actions of S1P are at least partially mediated by five subtypes of G-protein-coupled receptors (S1PR1–S1PR5). Murine macrophages express the S1P receptor subtypes S1PR1 and S1PR2 [77]. To evaluate which receptor subtype was involved in the S1P effect in the RAW cells, we applied the S1PR1 blocker W123 (Fig. 5a) and the S1PR2 blocker JTE-013 [6] (Fig. 5b). The statistical analysis shown in Fig. 5c demonstrates that W123 inhibited the stimulating effect of S1P whereas JTE-013 was without effect, indicating that the S1PR1 mediates the action of S1P on membrane conductance. To analyze the signaling pathway downstream of S1PR1, GDPβS, which blocks G protein signaling [29], was applied intracellularly via the patch pipette. Figure 5d shows that GDPβS inhibited the S1P-induced currents. This points to the involvement of G protein signaling in the action of S1P. S1P receptors may also be involved in Ca2+ signaling. S1PR2 and S1PR3 are reported to couple to phospholipase C (PLC) via the Gq protein. Furthermore, all S1P receptors can activate Gi and its βγ subunits might also activate PLC, leading to inositol trisphosphate (IP3)-mediated Ca2+ release from the endoplasmic reticulum [26]. Therefore, to test the involvement of the RAW macrophage S1P receptors in Ca2+ signaling, we measured the effect of S1P application on [Ca2+]i. As shown in Fig. 6, application of S1P was without effect on the intracellular Ca2+ concentration. S1P receptor signaling exerts several effects on the cytoskeleton [70, 39], which also modulates the activation of VRAC [43, 34]. We used a disruptor of the actin cytoskeleton,

Fig. 4 Pharmacological profile of S1P- and hypoosmolarity-induced currents in RAW cells. After stimulating the cells with S1P in isoosmolar solution or with hypoosmolar solution, the indicated substances were added to a concentration of 100 μM each (except for tamoxifen, 10 μM). The relative blocking effect was calculated by relating the S1P-induced conductance after 10-s application of blocker to the S1Pinduced conductance before addition of the blocker to the bath. All blockers except 9-AC and phloretin exerted a significant block. The relative blocking effect on S1P- or hypoosmolarity-induced conductance, respectively, was not significantly different for each blocker. All measurements are from different cells. Means from 6–10 cells for each blocker

cytochalasin D (CytD) [4], to test the involvement of the actin cytoskeleton in the activation of VRAC. Figure 7a demonstrates that CytD diminished the S1P-induced current. The statistics show that CytD inhibited VRAC activation by both S1P and hypoosmolar solution (Fig. 7b) and also reduced VRAC if applied after activation of the current by S1P or hypoosmolarity (Fig. 7c). Involvement of VRAC in ATP secretion To test if the S1P-induced ion channels can mediate ATP secretion, we performed measurements of the extracellular ATP concentration using a luciferase essay. As shown in Fig. 8a, S1P elicits an increase in the ATP concentration. This effect could be inhibited by anion channel blockers with a similar pharmacological profile as determined for the S1P- or

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hypoosmolarity-activated ion currents (see Fig. 4). Accordingly, substitution of the isoosmolar solution by a hypoosmolar one also induced ATP secretion, and this could be blocked by NPPB and tamoxifen but not by phloretin like the S1P-induced ATP increase (Fig. 8c). Although the activation of VRAC was sustained (Figs. 1a and 3a), the increase in ATP displays a partial phasic behavior (Fig. 8b). In contrast, the amount of ATP in the extracellular solution stayed constant if ATP was secreted due to a hypoosmolar challenge (Fig. 8d).

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The S1P-induced current in RAW macrophages was characterized as an anion-selective outwardly rectifying current. Furthermore, we demonstrated that this current depends on intracellular and extracellular osmolarity. Therefore, we presumed that this current was mediated by volume-regulated outwardly rectifying anion channels (VRAC, also referred to

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as volume-sensitive outwardly rectifying Cl− channel, VSOR). VRAC-mediated currents have been described in many cell types like epithelial cells, endothelium, glia cells, cardiomyocytes, keratinocytes, T lymphocytes, and fibroblasts [52]. Their molecular identity is not established to date, and possibly different ion channel species may contribute to osmolarity- or cell-volume-dependent ion currents [54]. In epithelial cells, apart from VRAC/VSOR with intermediate single-channel conductance, another anion channel has been described that is also activated by osmotic cell swelling. According to its large conductance, it was referred to as maxi anion channel [63]. Although we did not perform singlechannel current measurements, we suppose that the S1Pinduced current in macrophages is mediated mainly by VRAC. The reasons are the rather pronounced outward rectification, the block by glibenclamide [63] and tamoxifen, which also blocks VRAC in endothelial and human cervical cancer cells [25, 67] and the dependence on intracellular ATP [52]. Rather surprising is the lack of an inhibitory effect by phloretin, usually considered a specific VRAC blocker [63]. This points to different channel entities carrying the current or the involvement of cell-specific additional subunits responsible for the different sensitivities to anion channel blockers. Furthermore, connexins, pannexins, or even P2X7 receptors were reported to mediate ATP efflux [7], and pannexins and connexins can be inhibited by several anion channel blockers [79, 13]. However, the anion selectivity and the blocking

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effect of flufenamic acid on the hypotonicity- and S1Pinduced currents in macrophages make the involvement of connexins, P2X7 receptors, and pannexins unprobable [46, 33]. More detailed analysis of the S1P-dependent current is necessary to characterize the biophysical properties of the involved ion channels. Nevertheless, the use of seven different blockers demonstrated that the hypotonicity-induced and the S1P-dependent currents share the same pharmacological profile and are presumably carried by the same type(s) of ion channels. The recent identification of LRRC8A as a critical component of hypotonicity-induced anion currents will foster the development of more specific pharmacological tools for investigation of the physiological role of VRAC [59, 73]. Activation signaling of the S1P-dependent current Human macrophages express S1PR1–S1PR4, whereas murine macrophages are restricted to expressing S1PR1 and S1PR2 [77]. The activation of the ion current by S1P was inhibited by the S1PR1 blocker W123 but not by the S1PR2 blocker JTE-013. Therefore, our findings point to a key involvement of S1PR1in the activation of VRAC. S1P receptors signal via the G proteins Gi/o, Gq, and G12/13 [26]. The involvement of G proteins in VRAC activation was confirmed by the blocking effect of intracellular GDPβS. Accordingly, intracellular GTPγS, an irreversible stimulator of G proteins, enhances volume-sensitive anion currents in several cell types

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[55]. S1PR1 is the only S1P receptor coupling only via Gi/o proteins [30, 61], which therefore seem to be the responsible G proteins mediating VRAC activation. This is in accordance with our finding that Gq-mediated Ca2+ release is not involved in VRAC activation. Earlier characterizations of volumedependent anion channels [38] had already pointed out that the activation of VRAC is independent of increases in [Ca2+]i and cAMP, although a permissive [Ca2+]i of about 50 nmol/l is necessary for VRAC activation [71], which is present in our pipette solution used (see “Materials and methods” section). Interestingly, Leukotriene D4 and thrombin, which also act via Gi/o proteins, can also activate VRAC [34, 48, 32, 49]. The signaling pathway downstream of Gi/o is not completely clear, but a reorganization of the actin cytoskeleton via activation of phosphatidylinositol-3-kinase (PI3K), protein kinase B (PKB/akt), and the small G protein rac seems to be involved [70, 26], which may link S1P to the activation of VRAC. Cholesterol depletion potentiates VRAC by eliciting actin polymerization [34] and, as also demonstrated in our experiments, cytochalasin-D-induced disruption of F-actin

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Fig. 8 S1P- and hypoosmolarityinduced ATP release from RAW cells. a, c Pharmacological characterization of ATP released from RAW macrophages by application of S1P or hypoosmolar solution, respectively. The indicated substances were co-applied at a concentration of 100 μM each (except for tamoxifen, 10 μM). All blockers except 9-AC and phloretin exerted a significant block. Means from 8 to 15 measurements for each blocker. b, d Time course of the S1P- or hypoosmolarity-induced increase in the extracellular ATP concentration, respectively. Means from 12 (b) and 10 (d) measurements. Experiments were performed in cell culture medium (a, b) or isoosmolar/hypoosmolar buffer (c, d)

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integrity inhibits hypotonicity-induced anion currents [74, 15], pointing to cytoskeletal control of VRAC. Nevertheless, one also has to take into account that preincubation of cells with cytochalasins has been reported to have either no effect on [27] or even increase [43] VRAC activation. Whether different volume-dependent anion current subtypes were induced in these experiments or the exact application protocol of cytochalasins is of importance remain to be analyzed in detail. ATP secretion via S1P-activated ion channels The secretion of ATP via VRAC remains a controversial issue. Whereas in endothelial cells, ATP secretion via VRAC has been demonstrated [25], in intestinal and mammary epithelial cells, a large conductance anion channel (maxi anion channel) seems to play the major role [62]. In the macrophage cell line, we investigated, according to the pharmacological profile (see above), the S1P- and hypoosmolarity-induced ATP release which seems to be mediated by the medium conductance VRAC channels. Notably, the bacterial endotoxin LPS, which

Pflugers Arch - Eur J Physiol

activates sphingosine kinase and release of S1P [23, 77, 70, 78], induces ATP release from microglia and macrophages [16]. Accordingly, S1P induces nucleotide release into the extracellular space [39] and into phagosomes [40]. We found that the ATP concentration displayed a more phasic or sustained time course after application of S1P or hypoosmolar extracellular solution, respectively, for unknown reasons. Extracellular ATP concentrations depend on ATP secretion and ATP breakdown by ecto-ATPases. If the activity of the latter is assumed to be similar during both S1P- and hypoosmolarity-induced ATP release, then the ATP concentration mainly depends on the ATP secretion kinetics. Application of hypoosmolar solution for more than 30 s evoked a steady increase in VRAC, whereas the S1P-induced current varied considerably after application for more than 30 s. Some cells demonstrated a steady current increase but others displayed a constant or even diminished current amplitude, possibly due to desensitization of S1PR1 signaling [1]. This desensitization of S1P-induced current kinetics may have led to a partial prevailing ATP breakdown and therefore to the observed phasic time course in S1P-induced ATP secretion. Functional impact of S1P-induced currents The S1P-induced opening of VRAC with concomitant ATP release constitutes a link between sphingolipid and purine signaling systems, which play important roles under pathological conditions like defense against bacteria, hypoxia, and inflammation. ATP is considered a danger-associated signal (danger-associated molecular pattern, DAMP). Due to the induction of ATP release by macrophages recognizing a pathogen-associated molecular pattern (PAMP) like LPS, the defense mechanisms against bacteria may become amplified. That means that ATP may stimulate PAMP-recognizing macrophages in an autocrine manner without the need for DAMPs to be released from compromised or damaged cells. ATP and S1P are released from different cell types during hypoxia and inflammation [10, 12, 70, 77]. S1PRs and purinergic receptors are involved in the release of cytokines like TNF-α [28, 51]. Furthermore, signaling via S1PR and purinergic receptors and the activation of anion channels are involved in cell migration [72, 35, 70, 24, 64, 80, 36] and in phagocytosis and intracellular killing of bacteria [14, 47, 18, 57]. In the latter case, stimulation of phospholipase D by S1P or P2X7 receptors is involved [8, 31, 19]. To what extent S1P and purinergic signaling act in parallel or sequentially by S1PR1-mediated ATP release remains to be investigated.

Acknowledgments This work was supported by the Roux program of the Medical Faculty of the Martin Luther University Halle (FKZ 28/29).

Ethical standards The experiments comply with the current laws of Germany.

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