Petr Obrdlik
email: petr.obrdlik@iongate.de

Kerstin Diekert

Natalie Watzke

Christine Keipert

Ulrich Pehl

Catrin Brosch

Nicole Boehm

Inga Bick

Maarten Ruitenberg

Walter Volknandt

Bela Kelety

ELECTROPHYSIOLOGICAL CHARACTERIZATION OF ATPases IN NATIVE SYNAPTIC VESICLES AND SYNAPTIC PLASMA MEMBRANES

Keywords: V-ATPase, synaptic vesicles, electrophysiology, solid supported membrane, Na + /K + -ATPase, chloride conductance, V-ATPase, V-type ATPase, SSM, solid supported membrane, BafA1, bafilomycin A1, Dig, digitoxigenin, DIDS, 4,4'-diisothiocyanatostilbene 2,2'-

Vesicular V-type H + -ATPase (V-ATPase) and the plasma membrane-bound Na + /K + -ATPase are essential for the cycling of neurotransmitters at synapse, but direct functional studies on their action in native surroundings are limited due to the poor accessibility via standard electrophysiological equipment. We performed solid supported membrane (SSM)-based electrophysiological analyses of synaptic vesicles and plasma membranes prepared from rat brains by sucrose gradient fractionation. Acidification experiments revealed V-ATPase activity in fractions containing the vesicles but not in the plasma membrane fractions. For the SSM-based electrical measurements, the ATPases were activated by ATP concentration jumps. In vesicles, ATP-induced currents were inhibited by the V-ATPase-specific inhibitor bafilomycin A1 (BafA1) and by 4,4'diisothiocyanatostilbene-2,2'-disulfonic acid (DIDS). In plasma membranes, the currents were inhibited by the Na + /K + -ATPase inhibitor digitoxigenin. Distribution of the V-ATPase-and Na + /K + -ATPase-specific currents correlated with the distribution of vesicles and plasma membranes in the sucrose gradient. V-ATPase-specific currents depended on ATP with K 0.5 of 51 ± 7 µM and were inhibited by ADP in a negatively cooperative manner with IC 50 of 1.2 ± 0.6 µM. Activation of V-ATPase had stimulating effects on the chloride conductance in the vesicles. Low micromolar concentrations of DIDS fully inhibited the V-ATPase activity, whereas the chloride conductance was only partially affected. In contrast, 5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB) inhibited the chloride conductance but not the V-ATPase. The presented data describe electrical characteristics of synaptic V-ATPase and Na + /K + -ATPase in their native surroundings, and demonstrate the feasibility of the method for electrophysiological studies of transport proteins in native intracellular compartments and plasma membranes.

Introduction

Electrophysiological studies of transport proteins in their native environment can help to understand the interplay of different ions and solutes at membranes, and its impact on the physiology of the corresponding organs or tissues. At synaptic plasma membranes, Na + /K + -ATPase builds electrochemical Na + and K + gradients, which are the major driving forces for the transport of neurotransmitters from the synaptic cleft into the cytosol [START_REF] Blanco | Isozymes of the Na-K-ATPase: heterogeneity in structure, diversity in function[END_REF]. Loading of neurotransmitters into synaptic vesicles is driven by an electrochemical proton gradient built up by a multi-subunit V-type ATPase (V-ATPase) [START_REF] Beyenbach | The V-type H+ ATPase: molecular structure and function, physiological roles and regulation[END_REF][START_REF] Takamori | Molecular anatomy of a trafficking organelle[END_REF]. Beside the V-ATPase, chloride conducting proteins also play an important role in building the optimal proton gradient [START_REF] Cidon | Characterization of a H+-ATPase in rat brain synaptic vesicles. Coupling to L-glutamate transport[END_REF][START_REF] Schenck | A chloride conductance in VGLUT1 underlies maximal glutamate loading into synaptic vesicles[END_REF]. Both, the transport of protons via V-ATPases as well as the translocation of Na + /K + cations across membranes via Na + /K + -ATPases generate electrical currents and thus are suited for electrophysiological analysis [START_REF] Davies | Vacuolar H(+)-pumping ATPase variable transport coupling ratio controlled by pH[END_REF][START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF]. However, electric measurements of ATPases in native membranes and intracellular compartments are challenging. Particularly, synaptic vesicles are difficult to access with standard electrophysiological equipment due to their small size (~40 nm). Therefore, the existing electrophysiological studies deal mainly with vesicular channels but synaptic V-ATPase and vesicular transport proteins have not yet been electrically characterized in their native surroundings [START_REF] Ahdut-Hacohen | Hydrogen ions control synaptic vesicle ion channel activity in Torpedo electromotor neurones[END_REF][START_REF] Rahamimoff | Ion channels in synaptic vesicles from Torpedo electric organ[END_REF]. On the other hand, direct measurements on plasma membrane Na + /K + -ATPases are usually performed in excised-out giant patch clamp configuration complicating the electrophysiological analysis [START_REF] Friedrich | Na+,K(+)-ATPase pump currents in giant excised patches activated by an ATP concentration jump[END_REF]. For these reasons, knowledge of electrical properties of synaptic V-ATPase and Na + /K + -ATPases in their native surrounding is limited. During the last ten years, the solid supported membrane (SSM) technique was shown to be suited for electrical analysis of transporters, pumps and channels in different membrane environments [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF][START_REF] Schulz | SSM-based electrophysiology[END_REF]12]. For such electrical measurements, the SSM is built on a planar gold electrode by depositing an alkane-thiol layer followed by a phospholipid layer on top of it. Membrane fragments or vesicles containing the protein of interest are then adsorbed onto this SSM (Fig. 1). Thereby, a fluid compartment is formed between the SSM and the protein-containing membranes. The proteinspecific charge movements into or out of this compartment are induced by rapid exchange of a nonactivating solution for a solution containing either a substrate or a protein-activating agent [START_REF] Schulz | SSM-based electrophysiology[END_REF]. Depending on the sign of the moved charges and the direction of the transport either positive or negative transient currents are detected [START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]. The transient nature of the currents is based on the arrangement of the adsorbed membranes and SSM on the sensor, and the current detection principle [START_REF] Schulz | SSM-based electrophysiology[END_REF]. Briefly, the SSM behaves like a capacitor with the solution in the fluid compartment and the gold electrode beeing the capacitor plates (Fig. 1). According to these arrangements, the transport-dependent transient currents correspond to the charging of the sensor gold electrode and the charging kinetics depends on the transport activity of the assayed protein [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF][START_REF] Schulz | SSM-based electrophysiology[END_REF][START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]. In general, the peak current value is used to analyze the protein transport activity. We applied the SSM-based technique for electrical analysis of ATPases in native synaptic vesicles and synaptic plasma membranes. The specificity of the underlying protein activities was determined by measuring substrate dependencies and by testing V-ATPase and Na + /K + -ATPase inhibitors. Beside that, the regulation of V-ATPase by ADP was studied. To distinguish between V-ATPase and chloride-specific effects in synaptic vesicles both, the ATP-and chloride-induced currents were characterized with the respective inhibitors.

Materials and Methods

Preparation and storage of synaptic vesicles and plasma membranes. Synaptic vesicles (SV) and synaptic plasma membranes (SPM) were isolated from rat brains via percoll and sucrose gradient centrifugation as described in Morciano et al. [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF] with following modifications. One whole brain of an adult Sprague Dawley rat (Harlan, Borchen, Germany) was used per each preparation. The brain was homogenized with a glass homogenizer as described previously, the homogenate was centrifuged for 10 min at 1000 x g and 4°C, and the resulting supernatant was layered on a discontinuous percoll gradient (3%, 10%, 23% percoll (v/v) in 5 mM Tris/HCl, 320 mM sucrose, pH 7.4). After 7 min centrifugation at 31000 x g the fractions containing synaptosomes (between 3 and 10% and 10 and 23%) were collected and lysed according to the original protocol. The resulting suspension was layered onto a continuous sucrose gradient ranging from 0.3 M to 1.2 M sucrose in 0.5 mM EGTA, 10 mM HEPES/NaOH pH 7.4 of a total volume of 32 ml. The gradient was centrifuged for 2 h at 85000 x g and 30 fractions (1 ml each) were collected. Typically, SV were enriched in lower density fractions 5 to 9 and contained around 80 ng/µl total protein. SPM were enriched in fractions 22 to [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: cytoplasmic cation binding and release[END_REF] A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

each fraction were divided into 100 µl aliquots equivalent of 8-20 µg total protein, frozen in liquid nitrogen, and stored at -75°C. Preparation of biosensors and SSM-based electrical measurements. The biosensors were prepared with gold-electrode sensors from IonGate Biosciences (Germany) as described by the manufacturer. Briefly, the solid supported membrane (SSM) was built on a gold electrode by applying first an alkane-thiol followed by a phospholipid di-phytanoyl-phosphatidyl-choline. Subsequently, the SSM-coated sensors were covered with 50 µl of equilibration buffer (150 mM KCl, 50 mM Hepes, pH 7.2/NMG, 2 mM MgCl 2 , 0.2 mM DTT) and incubated at 4°C for 15 min. An aliquot of SV or SPM (5-20 µg of total protein) was thawed and sonicated with a microsonicator by applying 5 bursts with an amplitude of 30% (ultrasonic processor UP 50 H with a MS 1 tip, Dr. Hielscher, Germany). 0.5-2 µg total protein of the sonicated SV or SPM were loaded on each sensor and incubated at 4°C for 12 h. After this incubation the sensors could be stored for another 48 h at 4°C without significant reduction of the protein activities. For electrical measurements, the SV-or SPM-loaded biosensors were integrated into the fluidic system of the SURFE 2 R One setup (Surface Electrogenic Event Reader, IonGate Biosciences, Germany) and the proteins were activated via rapid exchange between pre-incubating, non-activating and activating solutions [START_REF] Schulz | SSM-based electrophysiology[END_REF][START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]. The application of activating solution induced transient rapidly decaying currents. After the current decayed, the system was set back by rinsing the sensor with the pre-incubating or non-activating solutions. The proteins were then activated again by applying the non-activating and the activating solution. Like this, different solutions and experimental conditions could be assayed on the same sensor. If required, possible changes (run down) of the protein activity during the experimental period was monitored by application of a reference activating solution. V-ATPase-mediated currents were induced by exchanging the non-activating solution (150 mM K-Asp, 50 mM Hepes, pH 7.2/NMG, 2 mM MgSO 4 , 10 mM KCl) by the activating solution (non-activating solution supplied with 5 -300 µM ATP, "ATP-concentration jump"). 0.2 mM DTT was added to stabilize the V-ATPase protein activity. Na + /K + -ATPase was activated by 5 -1000 µM ATP in Na + -containing buffers (140 mM NaCl, 30 mM HEPES, pH 7.0/NaOH, 2 mM MgCl 2 ). Chloride conductance was measured by rapid exchange of a pre-incubating solution (330 mM K-Asp, 50 mM Hepes, pH 7.2/NMG, 2 mM MgSO 4 ) for a nonactivating solution (preincubation solution with 300 µM K-ATP) followed by an activating solution (300 mM K-Asp, 50 mM Hepes, pH 7.2/NMG, 2 mM MgSO 4 , 30 mM KCl, 300 µM K-ATP). Inhibitors were prepared as DMSO stock solutions (DIDS, Baf A1, NPPB) or ethanol stock solutions (digitoxigenin) and added in equimolar amounts to all solutions. The final concentration of DMSO or ethanol in solutions never exceeded 0.1%. Acidification assays. Acidification of membrane vesicles was measured by following the quenching of acridine orange (AO) fluorescence in the fluorescence spectrophotometer (FlexStation, Molecular Devices, USA) with excitation and emission wavelengths at 485 and 530 nm. The reaction buffer contained 60 mM KCl, 10 mM MOPS, pH 7.4/KOH, 4 mM MgSO 4 , and 300 mM sucrose. Shortly before the experiment, AO (1 mM AO in 30% EtOH) was added to the final concentration of 10 µM, and SV or SPM suspensions were applied at the final total protein concentration of ~0.05 µg/µl. The reaction was started by adding 120 µM K-ATP, and stopped by adding either V-ATPase inhibitors or ionophores. Fluorescence changes were monitored at 37°C.

Results

Preparation of synaptic vesicles and membranes, and acidification assays. The synaptic vesicles (SV) and plasma membranes (SPM) were prepared from whole rat brains using adapted sucrose gradient protocols of Morciano et al. [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF]. Using this protocol SV are enriched in lower density fractions 5 to 9, whereas SPM are contained in higher density fractions 22 to 28. We tested the distribution of SV in the low-and high-density fractions by measuring the V-ATPase-mediated acidification of vesicles with the fluorescence dye acridine orange (AO) [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]. In these assays the acidification is detected as the quenching of the AO fluorescence. For the experiments the SV fractions F5 to F9, and the SPM fractions F24 to F26 were each pooled, and the AO fluorescence changes were recorded with the spectrophotometer (FlexStation, Molecular Devices, USA). Adding 120 µM ATP to the reaction mix decreased the AO fluorescence in the SV fractions, but not in the SPM fractions (Fig. 2A). The moderate increase of the fluorescence in SPM fractions was not ATP specific since it was observed also after adding other chemicals such as DMSO (data not shown). In SV the ATP-induced fluorescence decrease was inhibited by the V-ATPase inhibitor bafilomycin A1 (BafA1) at a concentration of 25 nM [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. In SPM BafA1 had no effect on the AO fluorescence. The pre-incubation of SV with the proton ionophore carbonyl cyanide-m- A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. chlorophenylhydrazone (CCCP) or with DIDS inhibited the ATP-induced effects (Fig. 2B). Moreover, supplying CCCP after the ATP application reversed the fluorescence quenching, demonstrating that the quenching was due to the accumulation of protons in SV. Electrical measurements of the V-ATPase activity. To measure the electric activity of synaptic V-ATPase, SSM sensors were prepared with the SV-containing fractions (F5 to F9) and the protein was activated by rapid exchange of a non-activating, ATP-free solution for an activating solution containing ATP (ATP concentration jumps). To suppress possible Na + /K + -ATPase activity due to residual SPM contaminations in SV fractions, the measurements were performed in Na + -free conditions. Consistent with a translocation of positive charges (protons) across the adsorbed SV membranes towards the electrode, the ATP concentration jumps induced positive currents on the SV-loaded sensors (Fig. 3A, [12,[START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]). In SSM-based electrophysiology, the protein-dependent currents correspond to the charging of a capacitor formed by the SSM, the fluid compartment and the gold electrode (Fig. 1, [START_REF] Schulz | SSM-based electrophysiology[END_REF][START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]). As expected for capacitor charging, the ATP-induced currents were transient and decaying in range of several hundred milliseconds after the application of the activating solution. The time range of the current decay correlated with the SSM-measurements of other transporter proteins [START_REF] Schulz | SSM-based electrophysiology[END_REF][12][START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]. As soon as the current decayed completely, the system was set back by equilibrating the sensor in the non-activating solution. The protein could be repeatedly activated by applying the activating solution over an extended time period (4-6 hours) and different experimental conditions could be tested on the same sensor. The ATP concentration jumps induced electrical currents on the synaptic vesicles only in presence of 2 mM Mg 2+ , whereas no ATPinduced currents were observed with solutions lacking Mg 2+ and containing 0.5 mM EDTA (Fig. 3A). The currents were fully inhibited by BafA1 (25 nM, Fig. 3B), and depended on the ATP concentration with the apparent half maximal concentration K 0.5

ATP at 51 ± 7 µM (Fig. 3C,D). Electrical measurements of the Na + /K + -ATPase. For the measurements of the plasma membraneassociated Na + /K + -ATPase, the sensors were prepared with the higher density fractions (F24 to F28) enriched in SPM. The experiments were performed as described by Pintschovius et al. [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF] with non-activating and activating solutions containing Na + but free of K + . On SPM-coated sensors, 150 µM ATP concentration jumps induced fast-decaying electric currents (Fig. 4A). The currents were reduced by the Na + /K + -ATPase inhibitor digitoxigenin (Dig, 1 µM), whereas 10 nM BafA1 had no effect on the ATP-induced currents of SPM fractions (Fig. 4A,[18]). At 1 µM concentration, Dig inhibited the ATP currents to approximately 20-30%. The remaining currents could be further inhibited by increasing Dig concentrations (data not shown). The Dig-sensitive currents depended on ATP with the calculated K 0.5 ATP of 12 ± 5 µM (Fig. 4B). Distribution of V-ATPase and Na + /K + -ATPase in sucrose gradient. The described protocol for the separation of SV and SPM by continuous sucrose gradients was originally applied for proteomic studies and the distribution of V-ATPase, Na + /K + -ATPase, and other SV and SPM marker proteins was characterized immunologically via western blot [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF]. To perform functional analysis of the V-ATPase and Na + /K + -ATPase distribution in the gradient, the representative SV fractions F6-8, the SPM fractions F24-26 and the intermediate fraction F14 were tested for the V-ATPase and Na + /K + -ATPase activity by ATP concentration jumps in Na + -free or in Na + -containing solutions, respectively, and by applying the respective inhibitors BafA1 and Dig. Fig. 4C and Suppl. Tab. 1 show the BafA1-sensitive and the Dig-sensitive ATP-induced currents in the different fractions. In SV fractions F6-8 under Na + -free conditions, the ATP-induced currents were inhibited by 10 nM BafA1 but insensitive to Dig. In Na + -containing solutions, the currents were to a minor extent (~18%) also sensitive to 1 µM Dig (Fig. 4C and Suppl. Tab. 1) suggesting some Na + /K + -ATPase activity in the SV-enriched fractions possibly originating from residual SPM contaminations. In SPM fractions F24-26 the ATP-induced currents were insensitive to BafA1 but inhibited by Dig. No ATP-induced currents were detected with the intermediate fraction F14. Overall, these results correlate with the distribution of the V-ATPase and Na + /K + -ATPase proteins in the sucrose gradient, as described by immunoblot analysis [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF]. Partial Dig sensitivity of the ATPdependent currents in F6-8 suggested that there may be some residual contamination with Na + /K + -ATPase (SPM) in SV fractions. To avoid any effects of the Na + /K + -ATPase, all further V-ATPase studies were performed under Na + -free conditions. V-ATPase and kinetics of inhibition with BafA1, DIDS and ADP. BafA1 inhibits V-ATPases in low nanomolar range, whereas other membrane-bound ATPases are not affected or sensitive only to significantly higher concentrations in the micromolar range [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. Accordingly, the V-ATPase-but not the Na + /K + -ATPase-specific currents were inhibited with 10 nM BafA1 supplied to the nonactivating and activating solutions (Fig. 4A,C). In previous studies BafA1 was described to accumulate in membranes [19]. Therefore, the effects of different BafA1 concentrations were A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

studied over an extended period of time. BafA1 at concentrations of 5 and 1 nM also inhibited the V-ATPase-specific currents but longer incubation times were needed (Fig. 5A). The incubation time necessary for the inhibition was dose-dependent with the slowest kinetics at the lowest BafA1 concentration. The BafA1 effects were either not or only poorly reversible under the given experimental conditions. Beside BafA1, 4,4'-diisothiocyanatostilbene 2,2'-disulfonic acid (DIDS), a general inhibitor of chloride conducting proteins, has also been described to inhibit V-ATPases at low micromolar concentrations [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]. To further characterize the V-ATPase-specific currents, DIDS and as a competitive inhibitor ADP were used [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]20,[START_REF] Yabe | Patch clamp studies on V-type ATPase of vacuolar membrane of haploid Saccharomyces cerevisiae. Preparation and utilization of a giant cell containing a giant vacuole[END_REF]. For testing DIDS, the inhibitor was added to the non-activating and activating solutions and the V-ATPase activity was recorded for 15 to 50 min (Fig. 5B). Addition of 10 and 1 µM DIDS resulted in a complete inhibition of the V-ATPase-specific currents. Similar to BafA1, the incubation time necessary for the complete inhibition was dose-dependent with slower kinetics at the lower DIDS concentration. After 50 min incubation with 0.1 µM DIDS the V-ATPase activity was reduced by 30% with a further falling tendency suggesting that the inhibition did not reach the maximum. For ADP inhibition experiments, V-ATPase was activated by 120 µM ATP in presence of increasing ADP concentrations in the non-activating and activating solutions. At these conditions, the IC 50 for ADP was at 1.2 ± 0.6 µM with the Hill coefficient at 0.5 ± 0.1 (Fig. 5C). The reversibility of the ADP effects depended on the applied ADP concentration (Fig. 5D). Whereas low ADP effects (≤1 µM) could not be reversed at the given conditions, the effects of 10 or 100 µM ADP were partially reversible. V-ATPase activities after the washout of 10 or 100 µM ADP both recovered to the V-ATPase activity measured in presence of ~1 µM ADP. V-ATPase and vesicular chloride conductance. Beside the inhibitory effects on V-ATPase, DIDS is a general inhibitor of chloride-conducting proteins [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF][START_REF] Jentsch | Molecular structure and physiological function of chloride channels[END_REF]. To test if DIDS may also act on the V-ATPase activity through the inhibition of chloride-conducting proteins in SV, protocols were adapted for measuring chloride fluxes on SSM-based biosensors. On SV-loaded sensors, 30 mM Cl -concentration jumps induced negative electric currents (Fig. 6A). The negative sign of these currents is in agreement with the movement of negative charges (chloride anions) toward the sensor electrode, as opposed by the positive currents induced by the movement of positive charges via the V-ATPase and Na + /K + -ATPase [START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF]. Chloride transport into the synaptic vesicles is linked to the V-ATPase activity [START_REF] Cidon | Characterization of a H+-ATPase in rat brain synaptic vesicles. Coupling to L-glutamate transport[END_REF][START_REF] Schenck | A chloride conductance in VGLUT1 underlies maximal glutamate loading into synaptic vesicles[END_REF][START_REF] Stobrawa | Disruption of ClC-3, a chloride channel expressed on synaptic vesicles, leads to a loss of the hippocampus[END_REF]. Accordingly, the activation of the V-ATPase with 0.3 mM ATP in non-activating and activating solutions resulted in an increase of chloride-induced currents by ~49% (Suppl. Fig. 1, Fig. 6A). The positive currents induced by the ATP concentration jump from pre-incubating to non-activating solution mark the activation of the V-ATPase. DIDS and NPPB are blockers of anion channels and transporters [START_REF] Jentsch | Molecular structure and physiological function of chloride channels[END_REF]. To test the influence of these inhibitors on chloride-induced currents, either DIDS or NPPB were supplied to the solutions and the measurements were performed as described above. Like in the previous experiments (Fig. 5B) the V-ATPase signal was inhibited by DIDS at low micromolar concentrations (Fig. 6B). The chlorideinduced peak currents were reduced by 32 and 45% at 1 and 10 µM DIDS and the level of reduction remained the same between 10 µM and 100 µM DIDS (Fig. 6B,D). In contrast to DIDS, 10 µM NPPB inhibited the chloride currents by 80% but did not affect the V-ATPase-specific currents (Fig. 6C,D).

Discussion

Electrophysiological measurements are a standard tool for studying the function and the physiology of transporters, pumps and channels at plasma membranes. However, electrophysiological data on intracellular proteins such as V-ATPases, as well as data on electrical properties of plasma membrane-bound Na + /K + -ATPases in their native surroundings are limited. In this study we used SSM-based biosensors to electrically analyze V-ATPase and Na + /K + -ATPase in purified synaptic vesicles (SV) and synaptic plasma membranes (SPM). Quality of the purified SV and SPM. In the applied membrane purification protocol, SV accumulate in the low-density sucrose gradient fractions, whereas SPM are concentrated in the higher-density fractions at the bottom of the gradient [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF]. The quality of the purified SV was first tested via measuring V-ATPase-dependent acidification using acridine orange (AO) as a pH-sensitive fluorescent dye [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]. On SV-containing low density fractions, ATP induced a decrease of the AO fluorescence that was inhibited by DIDS, by the proton ionophore CCCP and by the V-ATPase inhibitor BafA1. This demonstrates that the measured fluorescence changes were due to the V-ATPase-dependent proton accumulation in the vesicles, thus showing that the low-density fractions are suited for functional studies of V-ATPase activity [START_REF] Beyenbach | The V-type H+ ATPase: molecular structure and function, physiological roles and regulation[END_REF][START_REF] Takamori | Molecular anatomy of a trafficking organelle[END_REF][START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF][START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. No significant AO fluorescence decrease could be detected testing the high density fractions, showing that these A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

fractions were largely free of vesicle contaminations. The separation of SV and SPM was further tested by analyzing the distribution of the V-ATPase-and Na + /K + -ATPase-specific electrical activities in the sucrose gradient fractions. Here, the distribution of V-ATPase and Na + /K + -ATPase activities correlated with the distribution of V-ATPase and Na + /K + -ATPase proteins in previously published immunoblot assays [START_REF] Morciano | Immunoisolation of two synaptic vesicle pools from synaptosomes: a proteomics analysis[END_REF][START_REF] Morciano | The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels[END_REF]. In SV-enriched fractions, the minor effects of the Na + /K + -ATPase inhibitor Dig on the ATP-induced signals suggested some Na + /K + -ATPase activity due to residual contamination with SPM. However, the use of Na + -free solutions fully suppressed the Dig effects (i.e. Na + /K + -ATPase activity) and selected specifically for the V-ATPase activity (see also below). Hence, the use of Na + -free conditions should be generally applicable for selective studies of the Na + -independent vesicular transport proteins in SV fractions. All together, the results demonstrate that the applied preparation and the SSM measurement protocols are well suited for functional studies of synaptic vesicles as well as plasma membranes. Specificity of the V-ATPase-related currents. Low-density sucrose gradient fractions containing the purified SV were used for the electrical analysis of synaptic V-ATPase. Several features show that the ATP-induced currents correspond to the V-ATPase activity. First, like other membrane-bound ATPases, V-ATPase needs Mg 2+ as a cofactor for an efficient ATP hydrolysis [START_REF] David | The catalytic cycle of the vacuolar H(+)-ATPase. Comparison of proton transport in kidney-and osteoclast-derived vesicles[END_REF]. Accordingly, the ATP-induced currents were Mg 2+ -dependent. Second, the currents were sensitive to the inhibitor BafA1 at low nanomolar concentrations, which are highly specific for V-ATPases [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. In contrast, P-type ATPases are either not affected or are inhibited by BafA1 only at >1000 times higher micromolar concentrations [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. Third, the absence of Na + in the experiments demonstrates that the ATP signals cannot originate from the P-type Na + /K + -ATPase via possible plasma membrane contamination. Fourth, the calculated K 0.5 ATP of 51 µM is in range of K 0.5 ATP for other V-ATPases [START_REF] Yabe | Patch clamp studies on V-type ATPase of vacuolar membrane of haploid Saccharomyces cerevisiae. Preparation and utilization of a giant cell containing a giant vacuole[END_REF][START_REF] David | The catalytic cycle of the vacuolar H(+)-ATPase. Comparison of proton transport in kidney-and osteoclast-derived vesicles[END_REF][START_REF] Johnson | H+ ATPase of chromaffin granules. Kinetics, regulation, and stoichiometry[END_REF]. Together with the fact that the applied SV showed a well-detectable V-ATPase activity in acidification assays, these features demonstrate that the detected electrical currents are V-ATPase-specific. Characterization of the Na + /K + -ATPase-specific currents. In presence of K + , the Na + /K + -ATPase pumps three Na + into the cytosol and two K + out, thus being electrogenic [START_REF] Blanco | Isozymes of the Na-K-ATPase: heterogeneity in structure, diversity in function[END_REF][START_REF] Friedrich | Na+,K(+)-ATPase pump currents in giant excised patches activated by an ATP concentration jump[END_REF]. In addition, Na + /K + -ATPase can also conduct electrical currents in K + -free solutions as demonstrated in black lipid membrane (BLM) and SSM experiments [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF][START_REF] Gropp | K+-dependence of electrogenic transport by the NaK-ATPase[END_REF]. The SSM-based measurements of the synaptic Na + /K + -ATPase were performed with SPM-enriched sucrose gradient fractions in absence of K + . According to the previously published SSM-based experiments with the pig kidney Na + /K + -ATPase [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF], fast decaying, transient currents were obtained on SPM-loaded sensors upon rapid ATP concentration jumps under K + -free conditions. The ATP-induced currents were inhibited with the membrane-permeable Na + /K + -ATPase-specific inhibitor digitoxigenin (Dig) demonstrating that the detected currents correspond to the Na + /K + -ATPase activity [18]. Moreover, the calculated K 0.5 ATP of ~12 µM was in good agreement with the K 0.5 ATP of ~3 µM determined for a pig Na + /K + -ATPase in K + free solutions [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF]. Since ATP can bind to the Na + /K + -ATPase only from the intracellular side of the protein, these results demonstrate that the applied SPM adsorbed -at least partially -in the inside-out configuration to the sensors. For the Na + /K + -ATPase, a low affinity and a high affinity ATP binding had been described [START_REF] Scheiner-Bobis | Shift to the Na+ form of Na+/K+transporting ATPase due to modification of the low-affinity ATP-binding site by Co(NH3)4ATP[END_REF]. ATP binds with low affinity to the K + -bound protein, whereas the high-affinity ATP binding was observed in absence of K + [START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: cytoplasmic cation binding and release[END_REF][START_REF] Post | Activation by adenosine triphosphate in the phosphorylation kinetics of sodium and potassium ion transport adenosine triphosphatase[END_REF]. Interestingly, the K 0.5

ATP values determined at K + -free conditions with the SSM-based technique were significantly lower than the K m ATP of 70 -460 µM determined for different rat Na + /K + -ATPase αβ oligomers in presence of Na + and K + (this work, [START_REF] Blanco | Isozymes of the Na-K-ATPase: heterogeneity in structure, diversity in function[END_REF][START_REF] Pintschovius | Charge translocation by the Na+/K+-ATPase investigated on solid supported membranes: rapid solution exchange with a new technique[END_REF]). This suggests that the K 0.5 ATP determined in this work reflects at least partially the high-affinity binding of ATP to the Na + /K + -ATPase. Inhibition of the V-ATPase-specific currents with BafA1 and DIDS. BafA1 is a highly hydrophobic macrolide antibiotic binding to the membrane spanning domain of the V-ATPase protein [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]. Several independent groups have reported on the reversibility of BafA1 effects on the V-ATPase activity in vivo [START_REF] Saurin | Inhibition of mitogeninduced DNA synthesis by bafilomycin A1 in Swiss 3T3 fibroblasts[END_REF]31]. In such in vivo studies, the de novo synthesis of the enzyme was postulated to be responsible for the recovery of the activity [19]. In vitro studies using reconstituted V-ATPase in liposomes have shown that BafA1 does not bind covalently and that the binding is reversible [START_REF] Xie | Salicylihalamide A inhibits the V0 sector of the V-ATPase through a mechanism distinct from bafilomycin A1[END_REF][START_REF] Zhang | Proton conduction and bafilomycin binding by the V0 domain of the coated vesicle V-ATPase[END_REF]. However, in these assays the BafA1 binding was reversible only in presence of lipid excess provided by addition of protein-free liposomes. Because of its highly lipophilic nature BafA1 was suggested to accumulate and/or tightly bind to the membranes [START_REF] Bowman | Bafilomycins: a class of inhibitors of membrane ATPases from microorganisms, animal cells, and plant cells[END_REF]19]. This high hydrophobicity might explain why the inhibitory effect of BafA1 on the V-ATPase-specific currents could not be reversed under the here described experimental conditions. Similarly, the time dependence of the BafA1 effect may reflect the accumulation of the inhibitor on the sensors. A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

The stilbene derivate DIDS had been described to inhibit the synaptic V-ATPase at low micromolar concentrations [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]. The inhibition of proteins by DIDS is often irreversible as DIDS can form covalent bonds with amino groups of amino acids [START_REF] Guizouarn | Multiple transport functions of a red blood cell anion exchanger, tAE1: its role in cell volume regulation[END_REF][START_REF] Wilson | Studies on the dids binding site of monocarboxylate transporter 1 suggest a homology model of the open conformation and a plausible translocation cycle[END_REF]. In our experiments DIDS inhibited the V-ATPase-specific currents at low micromolar concentrations and the inhibition was dose-and time-dependent, and irreversible. The time dependency and the poor reversibility of the DIDS effect on ATP-induced currents suggest that the stilbene derivate may interact with the V-ATPase via covalent bonding. Inhibition of the V-ATPase-specific currents with ADP. Compared to batch-based methods such as acidification assays, the SSM-based approach allows real-time V-ATPase measurements without accumulation of the ATP-hydrolysis products in the experimental solutions. Therefore, the approach is well suited for studying the regulation of the V-ATPase by ADP. The sensitivity of the V-ATPase-specific currents to ADP is in agreement with the general mode of action of V-ATPases, and with ADP being important for the feedback control of V-ATPases [20, [START_REF] Yabe | Patch clamp studies on V-type ATPase of vacuolar membrane of haploid Saccharomyces cerevisiae. Preparation and utilization of a giant cell containing a giant vacuole[END_REF][START_REF] David | The catalytic cycle of the vacuolar H(+)-ATPase. Comparison of proton transport in kidney-and osteoclast-derived vesicles[END_REF]36]. The Hill coefficient of ~0.5 suggests a negative cooperativity of the ADP binding to the V-ATPase protein. This is consistent with the existence of a high-affinity (K d ~66 nM) and a low-affinity (K d ~17 µM) ADP binding site in the V-ATPases [START_REF] Webster | Analysis of nucleotide binding by a vacuolar proton-translocating adenosine triphosphatase[END_REF]. In addition to the Hill coefficient, the characteristics of the ADP washout in the SSM-based experiments also suggest that multiple ADP binding sites exist. In these experiments, the inhibition with up to 1 µM ADP was only poorly reversible, which correlates with the dynamic range of the high-affinity ADP binding [START_REF] Webster | Analysis of nucleotide binding by a vacuolar proton-translocating adenosine triphosphatase[END_REF]. In contrast, the recovered V-ATPase-specific currents obtained after the washout of higher ADP concentrations (10 or 100 µM) approximated to values corresponding to the currents measured in presence of 1 µM ADP. This suggests that the high-affinity binding of ADP to the enzyme was poorly reversible under the respective experimental conditions, whereas ADP at the low-affinity binding site(s) could be well washed out. V-ATPase activity and chloride conductance. Chloride serves as the counter anion of protons and plays an important role in acidification of SV and endosomes [START_REF] Cidon | Characterization of a H+-ATPase in rat brain synaptic vesicles. Coupling to L-glutamate transport[END_REF][START_REF] Jentsch | Molecular structure and physiological function of chloride channels[END_REF]. In agreement with the H + and Cl -ions being electrochemically coupled, the chloride-induced currents of SV were significantly increased in presence of an activated V-ATPase. DIDS is a general inhibitor of chloride conducting proteins, but it also inhibits synaptic V-ATPase at low micromolar concentrations [START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]. The inhibition profile of ATP-and chloride-induced currents suggested that DIDS may have affected the chloride-induced currents indirectly by inhibiting the V-ATPase activity. Therefore experiments using another general inhibitor of chloride-conducting proteins NPPB were performed. In contrast to DIDS, NPPB inhibited the vesicular chloride currents but not the V-ATPase-related currents. Despite these observations, a direct inhibition of the chloride conductance with DIDS is possible. Such a direct inhibition is especially interesting in the light of the new finding that the DIDS-sensitive glutamate transporter VGLUT1 is essential for the acidification of synaptic vesicles [START_REF] Schenck | A chloride conductance in VGLUT1 underlies maximal glutamate loading into synaptic vesicles[END_REF][START_REF] Hartinger | An anion binding site that regulates the glutamate transporter of synaptic vesicles[END_REF]38]. This finding suggests that VGLUT1 represents the major chloride permeation pathway in synaptic vesicles. However, other chloride conducting proteins have also been demonstrated to play a role in the acidification of synaptic vesicles [START_REF] Jentsch | Molecular structure and physiological function of chloride channels[END_REF][START_REF] Stobrawa | Disruption of ClC-3, a chloride channel expressed on synaptic vesicles, leads to a loss of the hippocampus[END_REF][START_REF] Li | Biophysical properties of ClC-3 differentiate it from swelling-activated chloride channels in Chinese hamster ovary-K1 cells[END_REF]. Therefore, the chloride-induced currents detected with the SSM-based approach may represent activities of different vesicular chloride-conducting proteins. Further combined studies using the SSM-based technology and acidification assays may help to elucidate the significance of these currents for the acidification of the vesicles in vivo. Conclusions. During the last decade SSM-based electrophysiological studies on different transporters, pumps and channels have accumulated demonstrating the wide applicability of the technology for the transport protein research [START_REF] Schulz | SSM-based electrophysiology[END_REF][START_REF] Kelety | Transporter assays using solid supported membranes: a novel screening platform for drug discovery[END_REF][START_REF] Ganea | Bacterial transporters: Charge translocation and mechanism[END_REF]41]. Here, the described approach expands the applicability of the SSM-based electrophysiology to ATPases of synaptic vesicles and plasma membranes. The protocols allowed detailed mechanistic and pharmacological characterization of the vesicular V-ATPase as well as plasma membrane Na + /K + -ATPase, and enabled discriminating between the activities of these two proteins in their native surroundings. Together with protocols for measuring neurotransmitter transporters such as the glutamate transporter EAAC1 [START_REF] Krause | An automatic electrophysiological assay for the neuronal glutamate transporter mEAAC1[END_REF] future SSM studies can help to understand the complex interplay of these proteins in synaptic signalling. Acidification of synaptic vesicles (SV, ▼) was prevented by pre-incubating the vesicles with 25 nM bafilomycin A1 (BafA1, ∇). No acidification was detected with synaptic plasma membranes (SPM, ■ and □). SV correspond to sucrose gradient fractions F5 to F9, SPM to fractions F24 to F26. B: Inhibition of SV acidification with DIDS and CCCP. For the two upper traces, 10 µM DIDS (○) or 20 µM CCCP (▲) were applied prior to the addition of 120 µM ATP. In the lower trace (∆), the acidification was reverted by the application of 20 µM CCCP 10 min after ATP application. Each trace is representative of ≥4 independent experiments.

  and contained around 200 ng/µl total protein. SV or SPM of Biochemical Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

Biochemical

  Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

  Biochemical Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

Biochemical

  Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

Biochemical

  Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

Fig. 1 :

 1 Fig. 1: Arrangement of vesicles (left) and membrane fragments (right) on the sensors and the corresponding equivalent electrical circuits. "TR" is the transport protein embedded in the vesicle membrane (M) or in the membrane fragment (M). The arrows indicate the translocation of positive charges into the fluid compartments between the biological membranes M and the SSM. The charging of the compartments results in charging of the gold electrode (E). The SSM consists of an upper phospholipid layer and a lower alkane thiol layer residing on E.

Fig. 2 :

 2 Fig.2: ATP-induced acidification of synaptic vesicles measured as the quenching of acridine orange (AO) fluorescence. The acidification was induced by 120 µM ATP (arrow). A: Acidification of synaptic vesicles (SV, ▼) was prevented by pre-incubating the vesicles with 25 nM bafilomycin A1 (BafA1, ∇). No acidification was detected with synaptic plasma membranes (SPM, ■ and □). SV correspond to sucrose gradient fractions F5 to F9, SPM to fractions F24 to F26. B: Inhibition of SV acidification with DIDS and CCCP. For the two upper traces, 10 µM DIDS (○) or 20 µM CCCP (▲) were applied prior to the addition of 120 µM ATP. In the lower trace (∆), the acidification was reverted by the application of 20 µM CCCP 10 min after ATP application. Each trace is representative of ≥4 independent experiments.

Fig. 3 :

 3 Fig.3: V-ATPase-specific currents induced by ATP-concentration jumps on sensors covered with SV (fractions F5-F9). The currents were induced by fast exchange of an ATP-free solution for a solution containing 0-1000 µM ATP. The upper bar indicates the solution exchange. If not stated, the solutions contained 2 mM Mg 2+ . All experiments were performed in Na + -free conditions. A:ATP (300 µM) induced magnesium-dependent electrical currents. For Mg 2+ -free conditions 0.5 mM EDTA was added to the solutions. B: Inhibition of the ATP-induced currents with the V-ATPase-inhibitor BafA1. C and D: Dependence of the signal amplitudes on the applied ATP concentration. The amplitudes (peak currents) were normalized relative to the peak currents at 1000 µM ATP. The K 0.5 was calculated by fitting the current amplitudes at different ATP concentrations using Michaelis-Menten kinetics. Concentrations are shown at logarithmic scale. N = 3-5 (± SEM) for data points at each concentration.

Fig. 4

 4 Fig.[START_REF] Cidon | Characterization of a H+-ATPase in rat brain synaptic vesicles. Coupling to L-glutamate transport[END_REF] Na + /K + -ATPase-specific currents on sensors covered with SPM (A and B) and the distribution of V-ATPase and Na + /K + -ATPase activities in sucrose gradient fractions (C). A: Currents were induced by rapid solution exchange using 150 µM ATP on SPM sensors under Na +containing but K + -free conditions. The currents were inhibited by digitoxigenin (Dig) but not by BafA1. The upper bar indicates the solution exchange. B: ATP dependence of the Dig-sensitive currents in SPM at K + -free conditions. The K 0.5 was determined by fitting a sigmoidal dose response kinetic to the data (n = 6 ± SEM for each concentration). C: Distribution of BafA1-(white) and Dig-sensitive (striped) ATP signals (150 µM) in sucrose gradient fractions F6 to F8 (SV), F14 (intermediate fraction) and F24 and F26 (SPM). Black bars correspond to ATP-induced signals without inhibitors. The white and stripped bars correspond to the residual currents after the application of the respective inhibitor. No significant ATP-induced currents were detected in F14. BafA1 was 10 nM, Dig 1 μM. Bars are mean (±SEM) of 3-5 independent experiments.

Fig. 5 :Fig. 6 :

 56 Fig.5: Kinetics of V-ATPase inhibition with BafA1, DIDS and ADP. Sensors were prepared with SV (fractions F5-F8). The graphs show peak currents normalized to the peak currents recorded at 100 µM ATP (A and B) or 120 µM ATP (C and D) in absence of BafA1, DIDS or ADP. A: Timeand dose-dependence of the BafA1 inhibition. The currents were repeatedly induced by 100 μM ATP. After 30 min, 1 nM (■) or 5 nM (○) BafA1 were applied and the currents were recorded for another 40-50 min. Finally, inhibitor-free solutions were used again (washout). Each data point represents the mean of two independent experiments. The inset shows sequential application of increasing BafA1 concentrations on one SV-loaded sensor. B: Inhibition of V-ATPase with DIDS. The currents were repeatedly induced by 100 µM ATP concentration jumps. After 20 min, 0.1 (○), 1 (▲), or 10 (■) µM DIDS was supplied to the solutions, and the V-ATPase activity was recorded for another 15 to 50 min. C: Inhibition of V-ATPase with ADP. V-ATPase was activated by 120 µM ATP. The IC 50 for ADP was determined by fitting a sigmoidal dose response kinetic to the data (n = 5 ± SEM for each concentration). D: Washout of ADP. The graph shows V-ATPasespecific currents at 120 µM ATP before ADP application (left black bar), in presence of 1, 10 or 100 µM ADP (striped, dotted and white bars), and after the washout of the corresponding ADP

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© 2010 The Authors Journal compilation © 2010 Portland Press Limited

Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.© 2010 The Authors Journal compilation © 2010 Portland Press Limited

Acknowledgements

We gratefully acknowledge Klaus Fendler and Karin Schumacher for their critical reading of the manuscript, and Wolf Berger for his excellent technical assistance. We thank Robin Krause for providing Fig. 1. Biochemical Journal Immediate Publication. Published on 25 Jan 2010 as manuscript BJ20091380 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091380

A c c e p t e d M a n u s c r i p t

Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.


Stephanie Engel

Silvia Scolari

Bastian Thaa

Nils Krebs

Thomas Korte

Andreas Herrmann

PhD Michael Veit
email: mveit@zedat.fu-berlin.de

FLIM-FRET and FRAP reveal association of influenza virus hemagglutinin with membrane rafts

Keywords: hemagglutinin, raft, palmitoylation, transmembrane region, budding 2

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Introduction

Membrane rafts are very small, dynamic liquid-ordered assemblies enriched in cholesterol and sphingolipids, which can selectively incorporate proteins, most notably GPI-anchored proteins to the outer leaflet, doubly-acylated proteins to the inner leaflet as well as several transmembrane proteins. Rafts recruit certain membrane proteins, which should favour interactions between the raft components. Rafts are also considered as point of entry as well as assembly and budding sites of a wide range of viruses. Published results demonstrating the occurrence of most proteins in rafts were obtained biochemically using cold extraction with Triton. It is, however, highly controversial whether partitioning of proteins into these detergent-resistant membranes (DRMs) reflects their association with rafts inside living cells [START_REF] Edidin | The state of lipid rafts: from model membranes to cells[END_REF][START_REF] Jacobson | Lipid rafts: at a crossroad between cell biology and physics[END_REF][START_REF] Mayor | Rafts: scale-dependent, active lipid organization at the cell surface[END_REF][START_REF] Mukherjee | Membrane domains[END_REF][START_REF] Simons | Model systems, lipid rafts, and cell membranes[END_REF]. If molecules are present in rafts, they should form cholesterol-sensitive clusters largely independent of their cell surface density. Fluorescence resonance energy transfer (FRET) is exceptionally well suited to demonstrate clustering of proteins, because it occurs only if two molecules tagged with a fluorophore are in close proximity (2-5 nm), which is in the range of the size of a membrane raft. However, mixed results were obtained when the FRET-technique was applied to probe cells for the existence of rafts, i.e. for the clustering of proteins which had been shown to partition into DRMs. Significant energy transfer between fluorophorelabelled antibodies against GPI-anchored proteins was detected. However, FRET correlated with the surface density of the raft marker suggesting that FRET was due to random interaction of mobile proteins in the membrane, but not to clustering of proteins [START_REF] Kenworthy | Distribution of a glycosylphosphatidylinositol-anchored protein at the apical surface of MDCK cells examined at a resolution of <100 A using imaging fluorescence resonance energy transfer[END_REF][START_REF] Kenworthy | High-resolution FRET microscopy of cholera toxin B-subunit and GPI-anchored proteins in cell plasma membranes[END_REF][START_REF] Glebov | Lipid raft proteins have a random distribution during localized activation of the T-cell receptor[END_REF]. Using a homo-FRET approach others have shown that GPI-anchored proteins form cholesterolsensitive clusters independent of their expression levels. Only a small, but significant (20-40%) fraction of GPI-anchored proteins form high density clusters of nanometer size (4-5nm), each consisting of only a few molecules [START_REF] Sharma | Nanoscale organization of multiple GPI-anchored proteins in living cell membranes[END_REF]. Zacharias et al. fused the yellow-and cyan fluorescent protein (YFP and CFP) to peptide sequences directed to membranes by dual acylation. FRET independent of the expression level was observed at the inner leaflet of the membrane and was sensitive to cholesterol depletion [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. The hemagglutinin (HA) of influenza virus was the first transmembrane protein described as a component of DRMs [START_REF] Fiedler | Glycosphingolipid-enriched, detergent-insoluble complexes in protein sorting in epithelial cells[END_REF]. Hydrophobic amino acids in the outer leaflet of the transmembrane region (TMR) and palmitoylation at cytoplasmic and transmembrane cysteines are required for partitioning of HA into DRMs [START_REF] Scheiffele | Interaction of influenza virus haemagglutinin with sphingolipid-cholesterol membrane domains via its transmembrane domain[END_REF][START_REF] Melkonian | Role of lipid modifications in targeting proteins to detergent-resistant membrane rafts. Many raft proteins are acylated, while few are prenylated[END_REF]. HA acquires detergent resistance at a late stage during its transport to the cell surface, probably in the trans-Golgi-network [14]. It was postulated that cholesterol-sphingolipid enriched domains form vesicles in the trans-Golginetwork (TGN), which serve as carriers for lipids and entrapped proteins to the apical membrane in epithelial cells [START_REF] Simons | Lipid sorting in epithelial cells[END_REF]. Not only does HA require rafts for its transport to the plasma membrane, but influenza virus also buds from raft-domains and HA needs to be concentrated in membrane rafts to perform its fusion activity efficiently [START_REF] Scheiffele | Influenza viruses select ordered lipid domains during budding from the plasma membrane[END_REF][START_REF] Takeda | Influenza virus hemagglutinin concentrates in lipid raft microdomains for efficient viral fusion[END_REF][START_REF] Wang | The interferon-inducible protein viperin inhibits influenza virus release by perturbing lipid rafts[END_REF]. However, attempts to demonstrate the association of HA with membrane rafts with methods other than Triton extraction have produced ambiguous results: Using fluorescence recovery after photobleaching (FRAP) it was demonstrated that wild-type HA diffused more slowly compared to a non-raft HA mutant, but its diffusion rate was elevated to non-raft HA values after disruption of rafts by depletion of cholesterol [19]. In contrast, authors performing largescale confocal FRAP assays have shown that the diffusion coefficients of proteins localized in DRMs (including HA) do not differ from those of proteins present in the bulk phase of the membrane [START_REF] Kenworthy | Dynamics of putative raft-associated proteins at the cell surface[END_REF]. More consistent data for clustering of HA were obtained by quantitative immunoelectron microscopy and by fluorescence-photoactivation-localization microscopy (FPALM). It was shown that HA is not randomly distributed in the plasma membrane, but forms irregular clusters ranging in size from a few nanometers up to many micrometers. However, only clusters at the nanometer length scale, i.e. with the size of rafts, could be disintegrated by extraction of cholesterol [START_REF] Hess | Quantitative electron microscopy and fluorescence spectroscopy of the membrane distribution of influenza hemagglutinin[END_REF][START_REF] Hess | Dynamic clustered distribution of hemagglutinin resolved at 40 nm in living cell membranes discriminates between raft theories[END_REF][START_REF] Leser | Influenza virus assembly and budding in raftderived microdomains: a quantitative analysis of the surface distribution of HA, NA and M2 proteins[END_REF].

Using FLIM-FRET we have shown recently that a construct consisting of the cytoplasmic tail and the transmembrane region of HA fused to variants of GFP clusters with a GPI-linked peptide, a marker for outer-leaflet rafts [START_REF] Scolari | Lateral distribution of the transmembrane domain of influenza virus hemagglutinin revealed by time-resolved fluorescence imaging[END_REF]. Here we have analyzed whether authentic HA equipped at its cytoplasmic tail with a fluorophore associates with a doubly-acylated peptide, a marker for inner-leaflet rafts [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF].

Materials and methods

Cloning of fluorescent proteins

The plasmids pEYFP (Clontech) and pECerulean [START_REF] Rizzo | An improved cyan fluorescent protein variant useful for FRET[END_REF] were used for cloning. The A206K monomeric mutation [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF] was made in YFP using overlap extension PCR. The HA gene from influenza virus strain A/FPV/Rostock/34 (H7N1) was inserted into the plasmid such that the resulting construct contains the amino acid sequence LRPEAPRARDPPVAT between the end of HA and the start of YFP. For FLIM-FRET experiments we used HA which contains glycine instead of arginine at position 339 at the cleavage site of HA 1 [26]. The non-acylated HA contains serine instead of cysteine at postions 551, 559 and 562 at the C-terminus of HA 2 [START_REF] Veit | Site-specific mutagenesis identifies three cysteine residues in the cytoplasmic tail as acylation sites of influenza virus hemagglutinin[END_REF]. The amino acids VIL at position 527 to 530 were converted to alanines to delete the other signal for association of HA with DRMs [START_REF] Scheiffele | Interaction of influenza virus haemagglutinin with sphingolipid-cholesterol membrane domains via its transmembrane domain[END_REF][START_REF] Takeda | Influenza virus hemagglutinin concentrates in lipid raft microdomains for efficient viral fusion[END_REF]. Myr-Pal-YFP contains the amino acid sequence MGCIKSKRKDNLNDDEPPVAT derived from the N-terminus of the Lyn kinase before the start of YFP.

Transfection, metabolic labeling and treatment of cells CHO-K1 cells were transfected using Lipofectamine 2000 (Invitrogen) 24 hours prior to performing the experiments. Metabolic labelling, immunoprecipitation with 2 µl anti-fowl plaque virus (FPV) antiserum, SDS-PAGE and fluorography were carried out as described previously [START_REF] Veit | Site-specific mutagenesis identifies three cysteine residues in the cytoplasmic tail as acylation sites of influenza virus hemagglutinin[END_REF]. Endoglycosidase-H and PNGase-F digestions of immunoprecipitated samples were performed as described by the manufacturer (New England Biolabs). For cholesterol depletion, cells were incubated for 30 min at 37 °C with 5 mM or with 10 mM methyl-Ecyclodextrin (Sigma) as indicated in the figure legends. To estimate the amount of extracted cholesterol, cells were labelled with >1D, 2D(n)-3 H@-cholesterol (1mCi/ml, GE-Healthcare) for about 4 h. Subsequent to extraction, the supernatant was incubated with scintillation cocktail (Lumasafe Plus, Lumac Lsc.) and cells were lysed with NaOH (0,1M, 20 min) and also incubated with scintillation cocktail. Radioactivity (in counts per minute, CPM) was measured with a luminescence counter (1600 RT, Packard) and the percentage of cholesterol extracted was calculated as: CPM s/(CPMs + CPMc) where CPMs are the CPM in the supernatant and CPMc are the CPM in the cell lysate.

Confocal fluorescence microsocopy

Confocal microscopy with live cells was performed using an Olympus FluoView 1000 microscope equipped with a UPLSAPO 60x/1.35 NA oil objective (Olympus) at 25 °C. Images were obtained by sequential excitation at 440 nm (Cerulean) using a laser diode and at 515 nm (YFP) using an argon laser. Emission was recorded between 460 and 490 nm for Cerulean and between 535 and 575 nm for YFP.

FLIM-FRET

FLIM images were acquired using the time-resolved LSM-upgrade Kit (PicoQuant) which uses the time-correlated single photon counting (TCSPC) technique. Cerulean was excited at 440 nm using a pulsed laser diode. The emitted fluorescence was detected by a single photon avalanche photodiode and a 470 ± 15 nm filter. Electrical signals were processed by the TimeHarp 200 PC card. FLIM pictures of cells were accumulated for 90 seconds using the SymPhoTime software. A TCSPC-histogram containing the accumulated decay signals only from the plasma membrane was created and fitted to calculate the average lifetime. For the fitting the instrument response function was taken into account. Since Cerulean has been reported to show either mono-or biexponential fluorescence decay [START_REF] Rizzo | An improved cyan fluorescent protein variant useful for FRET[END_REF][START_REF] Millington | High-precision FLIM-FRET in fixed and living cells reveals heterogeneity in a simple CFP-YFP fusion protein[END_REF], both fits were tested. We achieved the best fitting by assuming a biexponential decay, which yields a short and a long lifetime. The actual lifetime was calculated as the average of the two lifetimes. To calculate the lifetime of the donor fluorophore in the absence of acceptor, ten cells expressing only the donor were analyzed on each day an experiment was performed.

The FRET efficiency (E) was calculated using the equation:

av D DA E W W / 1
where IJ DA is the lifetime of a single cell co-expressing donor and acceptor and IJ Dav is the average lifetime of the ten cells expressing only the donor. FLIM measurements and the acquisition of the picture to determine the fluorescence intensity of the acceptor (Myr-Pal-YFP) and of the donor (HA-Cer) were performed on the same confocal picture. To determine the amount of acceptor and donor at the plasma membrane, the mean intensity of their fluorescence was calculated with ImageJ. A cell, where HA-Cer was clearly visible at the plasma membrane (see Fig. 1B) was selected and the software recognized the plasma membrane as a continuous structure with (almost) the same fluorescence intensity. To ensure that the amount of fluorophore is directly related to its fluorescence intensity, we included the instrument settings, i.e., laser intensity and gain settings in the analysis as follows:

Ac = { [( M -bg) / laser intensity ] / gain }
Ac is the acceptor concentration, M represents the mean of the measured fluorescence intensity at the plasma membrane and bg the background. Intensities of HA-Cer and of its mutants were in the same range, i.e. the variation in their fluorescence intensity was larger between individual cells expressing the same construct than between different HA-constructs. The FRET efficiency E of every cell co-expressing donor and acceptor was plotted against the fluorescence intensity of the acceptor F in this cell. The data points were fitted according to the equation:

D K F F E E / * % max
assuming that E is a hyperbolic function of the amount of acceptor. E% max is the maximal FRET efficiency calculated from the fitting. The equation gives a dissociation constant K D as a parameter to assess the associative properties of donor and acceptor [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. K D values should be compared with the acceptor intensities within the same experiment rather than to K D ´s from other experiments whose absolute intensities scales are not directly comparable as described above. Results for each acceptor/donor pair were compiled from one to nine different transfections, either sister cultures, i. e. cells grown in different dishes but transfected and measured in parallel or (mostly) cells transfected on different days, therefore representing completely independent experiments. The actual data for each graph are listed in the figure legends. In each transfected cell culture dish 10 to 20 different cells were measured. Each cell was included in the evaluation, no "outstanders" were eliminated.

Fluorescence recovery after photobleaching (FRAP)

We first established the laser intensity at 515 nm to photobleach about 60-70% of the fluorescence of YFP. FRAP was then performed at the plasma membrane above the nucleus of cells expressing HA-YFP. After 10 scans at low laser intensity, a circle of 15.11 µm 2 was bleached for 100 ms. Subsequently, scanning of the bleached circle was continued at 515 nm at low laser intensity for three minutes. Analysis was performed in Origin 7.5 (OriginLab Corp.). Recovery curves were fitted with an approximation of the theoretical recovery curve:

W t t F F t F r / ) ( 0
where t is the time after bleach, F(t) is the fluorescence as a function of t, F 0 is the fluorescence immediately after bleach, F r is the amount of fluorescence recovery, and IJ is the time of half-maximal recovery [START_REF] Axelrod | Mobility measurement by analysis of fluorescence photobleaching recovery kinetics[END_REF][START_REF] Kwon | Lateral mobility of tetramethylrhodamine (TMR) labelled G protein alpha and beta gamma subunits in NG 108-15 cells[END_REF]. The fits accurately match the experimental curves. The diffusion coefficient (D) was calculated using the equation: where Ȗ is related to the bleaching depth and is approximately 1.3 under our experimental conditions [START_REF] Yguerabide | Lateral mobility in membranes as detected by fluorescence recovery after photobleaching[END_REF], Z 2 is the radius, and IJ is the calculated time for half-maximal recovery. The mobile fraction (M f ) was calculated using the equation: 0

W Z J
0 1 / F F F M r f

Results and Discussion Construction and testing of fluorescent HA and raft marker

As an established constituent of membrane rafts of the inner leaflet of the plasma membrane, we created Myr-Pal-YFP. This construct consists of a myristoylated and palmitoylated peptide from the Lyn kinase fused to the N-terminus of the yellow-fluorescent protein (YFP), which can serve as an acceptor in FRET experiments [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. As a donor fluorophore we fused Cerulean (Cer), a variant of CFP with improved quantum yield and a higher excitation coefficient [START_REF] Rizzo | An improved cyan fluorescent protein variant useful for FRET[END_REF], to the cytoplasmic tail of HA (H7 subtype). We used HA with a mutation at its cleavage site since proteolytical cleavage of the HA-subtype H7 by a cellular enzyme causes instability of the protein [32,[START_REF] Takeuchi | Influenza virus M2 protein ion channel activity stabilizes the native form of fowl plague virus hemagglutinin during intracellular transport[END_REF]. Two mutants were generated by removing either of the known signals for incorporation of HA into DRMs. The three palmitoylated cysteines were exchanged by serines (HA-Cer-C3S) and three bulky, hydrophobic side chains (VIL) present at the N-terminal end of the external leaflet of the transmembrane region were converted to alanines (HA-Cer-VILA3). A similar mutation in the transmembrane region of a H3-subtype HA renders the molecule completely soluble in cold detergent [START_REF] Takeda | Influenza virus hemagglutinin concentrates in lipid raft microdomains for efficient viral fusion[END_REF]. In another mutant (HA-Cer-C3S+VIL3A) both raft-targeting signals were deleted simultaneously (see Fig. 1A for a schematic view of all proteins used in this study). All our constructs contain the A206K mutation in the fluorescent proteins. This prevents their dimerization and thus avoids the occurrence of a false-positive dimerization-induced FRET signal [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. Upon expression in CHO cells all constructs received Endo-H resistant carbohydrates (Fig. 1C) and were transported to the cell surface as visualized by confocal microscopy (Fig. 1B). Thus, attachment of additional sequences to the cytoplasmic tail of HA does not interfere with its processing and its transport to the plasma membrane.

FLIM-FRET is suitable to demonstrate clustering of proteins in the membrane

We used FLIM to study possible energy transfer between HA-Cer (donor) and the raft marker Myr-Pal-YFP (acceptor). FLIM-FRET has several advantages compared to conventional FRET methods, such as sensitised emission or acceptor photobleaching. FRET measurements done by FLIM are more robust and quantitative and no corrections are needed for donor fluorophore emission bleed through in the acceptor emission channel [START_REF] Wallrabe | Imaging protein molecules using FRET and FLIM microscopy[END_REF]. If FRET occurs, the lifetime of the excited state of the donor will be shortened in the presence of the acceptor in close proximity. From these measurements the FRET efficiency, defined as the fraction of donor excitation events that result in energy transfer to the acceptor, can be calculated. However, analyzing FRET in a two-dimensional system, such as the plasma membrane, requires a more systematic evaluation of the data [START_REF] Silvius | Fluorescence-quenching and resonance energy transfer studies of lipid microdomains in model and biological membranes[END_REF]. Since HA-Cer as well as Myr-Pal-YFP are mobile in the plane of the membrane, energy transfer can simply occur by random collision of both molecules. In this case the FRET efficiency is a linear function of the concentration of the acceptor protein at the membrane, i.e. it is absent at very low expression levels, but FRET-efficiencies increase linearly with increasing protein concentration. In contrast, if FRET is due to clustering of HA with the raft marker, the FRET efficiency is largely independent of the concentration of the acceptor protein. To evaluate our FLIM data we used a model applied by Zacharias et al. to demonstrate clustering of the double-acylated raft marker [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. In this model, which describes the binding of a ligand to its receptor, the FRET efficiency is a hyperbolic function of the concentration of the acceptor protein, i.e. its fluorescence intensity. The respective equation yields a dissociation constant K D as a parameter to assess the associative properties of donor and acceptor. If K D is very small compared to the intensity of the acceptor, clustering of acceptor and donor occurs, but if K D is in the same range as or larger than the acceptor intensity, FRET is due to random collision of both molecules. However, K D values cannot be compared between different protein pairs, even if they are attached to the same acceptor and donor fluorophore. The FRET efficiency depends on the distance between the donor and the acceptor and on the relative orientation of the donor emission dipole moment and the acceptor absorption dipole moment, parameters which can not be measured [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. To analyze whether this method is suitable to evaluate FLIM-FRET data we first analyzed a donor-acceptor pair which is known to form stable clusters. We co-expressed HA-Cer with HA fused in identical manner to YFP. As known, HA trimerizes in the ER, which is a prerequisite for its intracellular transport along the exocytic pathway [36]. If mixed trimers containing HA-Cer and HA-YFP will be present at the plasma membrane, both fluorophores should be in close proximity, since the cytoplasmic tail of HA consists of only eleven amino acids. Hence a significant FRET signal would be observed, which should be independent of the expression level of HA. CHO cells transfected with HA-Cer alone and cells co-transfected with HA-Cer and HA-YFP were selected and the lifetime of the donor fluorophore and the intensity of the acceptor fluorophore were analyzed at the plasma membrane. From these data the FRET efficiency was calculated and plotted against the intensity of the acceptor (Fig. 2A). The FRET-efficiencies were clearly independent of the intensity of the acceptor fluorescence and hence the expression level of HA-YFP. Fitting the data points to the described equation revealed a very low K D of 1×10 -16 indicating the situation of strong clustering and thus the presence of mixed trimers. We then asked whether FLIM-FRET can also be used to detect clusters such as those formed by the double-acylated raft marker with itself. Cells coexpressing Myr-Pal-Cer and Myr-Pal-YFP showed significant FLIM-FRET independent of their expression levels with a K D of 120, which is clearly lower than the fluorescence intensity of the acceptor (Fig. 2B). Incubation of cells with cyclodextrin to remove cholesterol increased the K D by an order of magnitude suggesting that clustering is reduced by disintegration of membrane rafts (Fig. 2C). Thus, we could confirm by FLIM-FRET clustering of Myr-Pal-XFP at the plasma membrane as has been described by others using acceptor photobleaching FRET [START_REF] Zacharias | Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells[END_REF]. This clearly demonstrates that FLIM-FRET is suitable for studying raft clustering.

FLIM-FRET reveals clustering of HA with the raft marker dependent on DRMtargeting signals and intact rafts

As a negative control, we measured FLIM-FRET between HA-Cer and YFP, which is localized all over the cell. In individual CHO cells transfected with HA-Cer alone and cells co-transfected with HA-Cer and YFP, the lifetime of the donor fluorophore and the intensity of the acceptor fluorophore were measured at the plasma membrane. From these data the FRET efficiency was calculated and plotted against the fluorescence intensity of the acceptor. Although a mean FRET efficiency of 12% was determined, the almost linear shape of the curve and the high K D of about 4500 indicate no clustering, as expected (Fig. 3A + B). We then measured FLIM-FRET of HA-Cer-wt with Myr-Pal-YFP to assess the clustering of HA with membrane rafts. We observed a high FRET efficiency (52%, Fig. 3A) clearly independent of the intensity of the acceptor fluorescence and hence the expression level of Myr-Pal-YFP (Fig. 3C). Fitting the data points to the described equation revealed a very low K D of 1.6×10 -16 . Several experiments were performed to analyze whether clustering is indeed due to partitioning of HA-Cer into membrane rafts. Extraction of cholesterol with methyl-Ecyclodextrin led to a minor, but statistically significant lowering of the FRET efficiency (46%, Fig. 3A) and to considerable reduction of clustering since the K D calculated for this association was increased significantly (Fig. 3D). Likewise, expression of non-acylated HA-Cer-C3S (Fig. 3E), of the HA mutant with a deletion of the DRM-targeting signal in its transmembrane region (Fig. 3F) and of the HA mutant devoid of both DRM-targeting signals (Fig. 3G) caused substantial disintegration of clustered HA arrangement with the raft marker. It should be noted that even HA with both DRM-targeting signals deleted showed high FRETefficencies with the raft marker (Fig. 3A) and that a few cells in individual graphs showed FRET even at very low acceptor intensities. Measurements of the fluorescence intensities of donors and acceptors revealed that Myr-Pal-YFP was approximately 10-fold overexpressed compared to HA-Cer. The overexpression of the acceptor renders a FRET pair prone to high transfer rates, since only a few interactions of HA-Cer with the raft marker yield a high FRET signal. On the other hand, since the acceptor concentrations are not limiting, plotting of the FRET-efficencies against the acceptor concentration is a suitable method to evaluate the data.

FRAP demonstrates transient interactions of HA with rafts

Next, we attempted to verify our conclusions with a second method, fluorescence recovery after photobleaching (FRAP). Proteins present in rafts should have a lower diffusion rate compared to non-raft proteins, since under conditions where viscosity is limiting, lateral diffusion varies inversely with the logarithm of the radius of the transmembrane portion of the diffusing species, for example the accumulated radii of all transmembrane proteins diffusing together in a raft [START_REF] Lippincott-Schwartz | Studying protein dynamics in living cells[END_REF]. An exception are apical membranes of polar cells, where the raft domain is the percolating phase in the membrane [38]. For the FRAP-experiments HA fused to YFP was used, since its fluorescence can be bleached more easily. HA-YFP-wt exhibited a diffusion coefficient of 0.14 µm 2 /sec and a mobile fraction of more than 80% (Fig. 4A), which is in agreement with published studies on other HA-subtypes not fused with a fluorescent protein [START_REF] Veit | Site-specific mutagenesis identifies three cysteine residues in the cytoplasmic tail as acylation sites of influenza virus hemagglutinin[END_REF][START_REF] Millington | High-precision FLIM-FRET in fixed and living cells reveals heterogeneity in a simple CFP-YFP fusion protein[END_REF]. Successive removal of the DRM-targeting signals moderately increased the diffusion coefficient of HA from 0.18 µm 2 /sec (HA-YFP-C3S) to 0.20 µm 2 /sec (HA-YFP-VIL3A) up to 0.30 µm 2 /sec if both signals are deleted (Fig. 4A). Likewise, the mobile fraction decreases to slightly over 70% suggesting that HA outside rafts might be prone to aggregation. Finally, we analyzed the temporal stability of the association of HA with membrane rafts. If HA and the raft marker are both present in a stable raft complex, they should diffuse together for minutes [39] and thus both molecules should exhibit a similar mobility in FRAPexperiments. However, Myr-Pal-YFP exhibited a diffusion coefficient of 0.7 µm 2 /sec, which is consistent with previous studies on proteins anchored by fatty acids to the inner leaflet of membranes [START_REF] Kenworthy | Dynamics of putative raft-associated proteins at the cell surface[END_REF], but much faster compared to HA-YFP (Fig. 4B). Thus, HA and the raft-maker did not diffuse together for minutes, the time period in which their mobilities are observed in FRAP experiments. Indeed, it has been shown that the type of membrane anchorage, i. e. by a transmembrane region versus fatty acids, has a much greater effect on the mobility of a protein compared to its distribution in rafts or non-raft domains of the membrane [START_REF] Kenworthy | Dynamics of putative raft-associated proteins at the cell surface[END_REF]. Our results are consistent with a model of dynamic partitioning of proteins into and out of raft domains, which permits raft proteins to transiently populate raft domains as well as to undergo diffusion outside of rafts [START_REF] Sharma | Nanoscale organization of multiple GPI-anchored proteins in living cell membranes[END_REF][START_REF] Kenworthy | Dynamics of putative raft-associated proteins at the cell surface[END_REF]. Since wild-type HA associates with rafts more strongly than HA lacking DRM-targeting signals (as evidenced in this study by FLIM-FRET), it probably remains inside rafts for longer periods of time, which would retard its overall mobility in the membrane. It remains to be shown how such a rather transient interaction of HA with rafts leads to the formation of the viral budzone, a nanodomain of the plasma membrane where assembly and finally budding of virus particles occurs [40].

6 Biochemical

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. 7 Biochemical

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Biochemical

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  Meder, D., Moreno, M. J., Verkade, P., Vaz, W. L. and Simons, K. (2006) Phase coexistence and connectivity in the apical membrane of polarized epithelial cells. Proc Natl Acad Sci U S A. 103, 329-334 39 Pralle, A., Keller, P., Florin, E. L., Simons, K. and Horber, J. K. (2000) Sphingolipidcholesterol rafts diffuse as small entities in the plasma membrane of mammalian cells. J Cell Biol. 148, 997-1008 40 Chen, B. J. and Lamb, R. A. (2008) Mechanisms for enveloped virus budding: Can some viruses do without an ESCRT? Virology. 372, 221-232 15 Biochemical Journal Immediate Publication. Published on 04 Nov 2009 as manuscript BJ20091388 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091388A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

Figure Legends Figure 1 :

 1 Figure LegendsFigure 1: Characterization of HA-Cer constructs expressed in CHO cells. A: Scheme and abbreviations of the constructs used in this study. B: Confocal fluorescence microscopy demonstrates transport of constructs to the plasma membrane. Scale bars: 10 µm. C: Glycosylation: CHO cells expressing the indicated construct were labeled for four hours with [ 35 S]-methionine/cysteine. Samples were then subjected to immunoprecipitation with anti-GFP antibodies, digestion with Endo-H (H), PNGase-F (F) or were left undigested (-) prior to SDS-PAGE (10% gel) and fluorography. C: untransfected control cells. Numbers on the left represent the mobility of molecular weight markers. HA-Cer*: deglycosylated HA.

Figure 2 :

 2 Figure 2: FLIM-FRET to demonstrate clustering within HA trimers and between raft markers. For each CHO cell the calculated FRET efficiency (FRET-E) in percent is plotted against the fluorescence intensity of the acceptor. Continuous line: Data were fitted to the saturable onesite binding model as described in "materials and methods". The dashed line is the 95% confidence interval. K D : dissociation constant, describing the associative properties. A low K D relative to the acceptor intensity indicates high clustering. A: HA-Cer-wt and HA-YFP-wt were coexpressed. n: cells analyzed: 66, K D : 1×10 -16 ± 7. B: Coexpression of Myr-Pal-Cer with Myr-Pal-YFP. n: 58, K D : 120 ± 57. C: Coexpression of Myr-Pal-Cer with Myr-Pal-YFP. Prior to FLIM-FRET cholesterol was extracted with cyclodextrin (MECD, 5 mM, 30 min, 37°C). This treatment removed about 50% of cholesterol from cellular membranes (see "materials and methods"). n:12, K D : 1180 ± 1188.

Figure 3 :

 3 Figure 3: FLIM-FRET reveals clustering of HA-Cer with raft markers dependent on DRM-targeting signals. A: Mean of the FRET-efficiencies for each experiment described in B to G. Error bars indicate the standard error of the mean (SEM). Statistics: Results are statistically significantly different (three asterisks: p < 0.001, two asterisks: p < 0.01) from HA-Cer-wt as calculated with a one-tailed unpaired Student's t-test. B: Coexpression of HA-Cer-wt and YFP. K D : 4442 ± 856, number of analyzed cells (n): 41, number of cell culture dishes (d): 4, number of different experiments, i.e. independent transfections (t): 1. C-D: Coexpression of HA-Cer-wt and Myr-Pal-YFP. C: untreated cells. K D : 1.6×10 -16 , n: 39, d: 9, t: 8. D: Cholesterol was extracted with 10 mM cyclodextrin (MECD, 30 min, 37°C) prior to FLIM-FRET. K D : 784 ± 139, n: 48, d: 3, t: 3. E: Coexpression of HA-Cer-C3S and Myr-Pal-YFP, K D : 872 ±287, n: 53, d: 8, t: 4. F: Coexpression of HA-Cer-VIL3A and Myr-Pal-YFP, K D : 422 ± 180, n: 35, d: 4, t: 3. G: Coexpression of HA-Cer-C3S+VIL3A and Myr-Pal-YFP, K D : 739 ± 460, n: 38, d: 4, t: 3.

Figure 4 :

 4 Figure 4: FRAP indicates transient interaction of HA with rafts. FRAP was performed at the plasma membrane of CHO cells expressing HA-YFP (wt or mutants) or Myr-Pal-YFP as indicated. Left part: Calculation of the diffusion coefficient (µm 2 /sec) and right part: Calculation of the mobile fraction. Bars represent the mean ± SEM of at least 30 individual cells. Statistics: Results are statistically significantly different (three asterisks: p < 0.001, two asterisks: p < 0.01, one asterisk: p < 0.05) from HA-Cer-wt as calculated with a two-tailed unpaired Student's t-test.

Biochemical Journal Immediate Publication. Published on 04 Nov 2009 as manuscript BJ20091388 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091388 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited

© 2009 The Authors Journal compilation © 2009 Portland Press Limited

Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited

Acknowledgement

We thank Dave Piston (Vanderbilt) for supplying the Cerulean plasmid, Hans-Dieter Klenk and Wolfgang Garten (Marburg) for antisera and Claudia Tielesch for technical assistance.

Funding

The project was funded by the German Research Foundation [grants SPP 1175 and SFB 740].

Abbreviations

Cer: Cerulean, DRM: detergent-resistant membrane, FLIM: Fluorescence lifetime imaging microscopy, FRET: fluorescence (or Förster´s) resonance energy transfer, FRAP: Fluorescence recovery after photobleaching, HA: hemagglutinin, MECD: methyl-betacyclodextrin, Myr-Pal-YFP: myristoylated and palmitoylated peptide fused to YFP, SEM: standard error of the mean, XFP: any variant of the green fluorescent protein, YFP: yellow fluorescent protein.