Protonophore properties of hyperforin are essential for its pharmacological activity

Hyperforin is a pharmacologically active component of the medicinal plant Hypericum perforatum (St. John's wort), recommended as a treatment for a range of ailments including mild to moderate depression. Part of its action has been attributed to TRPC6 channel activation. We found that hyperforin induces TRPC6-independent H+ currents in HEK-293 cells, cortical microglia, chromaffin cells and lipid bilayers. The latter demonstrates that hyperforin itself acts as a protonophore. The protonophore activity of hyperforin causes cytosolic acidification, which strongly depends on the holding potential, and which fuels the plasma membrane sodium-proton exchanger. Thereby the free intracellular sodium concentration increases and the neurotransmitter uptake by Na+ cotransport is inhibited. Additionally, hyperforin depletes and reduces loading of large dense core vesicles in chromaffin cells, which requires a pH gradient in order to accumulate monoamines. In summary the pharmacological actions of the “herbal Prozac” hyperforin are essentially determined by its protonophore properties shown here.

due to TRPC6 activation. However, other effects of hyperforin are described which might also participate in its pharmacological actions. For example, hyperforin attenuates various voltage-and ligand-gated ionic conductances in isolated hippocampal neurons or cerebellar Purkinje neurons [29][30][31][32] , it changes the fluidity of membranes 33 and collapses the mitochondrial membrane potential leading to the release of Zn 21 and Ca 21 into the cytosol 12 .
In the present study we used fluorescent imaging to monitor intracellular pH, cytosolic sodium changes and neurotransmitter release, and the whole-cell patch clamp technique to identify and to characterize the hyperforin-induced conductance in four independent systems, in HEK-293 cells, primary mouse microglia and chromaffin cells, and lipid bilayer membranes. Our data show that hyperforin itself acts as a protonophore and thereby mediates a significant proton conductance. This conductance does not require the presence of channel proteins like TRPC6 and its direction depends on the existing driving forces such as membrane potential and pH gradient. In line with these results, accumulation of neurotransmitters is abolished in primary mouse chromaffin cells in the presence of hyperforin.

Results
OAG-induced TRPC6 versus hyperforin-induced currents. Diacylglycerol (DAG), generated by G q/11 -coupled signaling pathways, or directly applied DAG analogs, are known to stimulate TRPC3, TRPC6, TRPC7 and mouse TRPC2 channels, independent from a DAG-mediated activation of protein kinase C (PKC). Using the whole-cell patch clamp technique we measured the development of inward and outward currents, extracted from voltage ramps at 280 and 180 mV, respectively, upon external application of 100 mM 1oleoyl-2-acetyl-sn-glycerol (OAG), an analog of DAG, in HEK-293 cells stably expressing TRPC6 cDNA (HEK-TRPC6 cells; Fig. 1a, black trace). 100 mM flufenamic acid (FFA), also shown to activate TRPC6, revealed the same current as OAG in HEK-TRPC6 cells (Fig. 1a, blue trace). Figure 1b shows the corresponding current-voltage relationships (IVs) revealing the typical double-rectifying TRPC6 currents. External application of 1, 3 and 10 mM hyperforin dosedependently activated a different current in HEK-TRPC6 cells (Fig. 1c). The corresponding IVs (Fig. 1d) did not represent the typical shape of the TRPC6-mediated current (see Fig. 1b). When applying hyperforin onto non-transfected HEK-293 cells, not expressing TRPC6 (HEK; Fig. 1e), or on primary mouse cortical microglial cells (Fig. 1g) we found the same current with the same IV (Fig. 1f,h) as seen in HEK-TRPC6 cells. Thus, the current induced by hyperforin was not mediated via TRPC6. The same current appeared using hyperforin supplied as dicyclohexylammonium (DCHA) salt (Sigma; Fig. 1a-f), or as free acid in methanol (Sigma or Cayman Chemical; Fig. 1g,h).
Hyperforin-induced currents in microglial cells. The hyperforininduced current depends on the dose of hyperforin, as shown in mouse microglial cells (Fig. 2a,b), with half-maximal concentrations (EC 50 ) of 9.3 and 8.7 mM hyperforin for the inward (Fig. 2d) and outward current (Fig. 2e), respectively. In parallel the cellular capacitance increased with higher hyperforin concentrations as long as hyperforin was present, and decreased thereafter again (Fig. 2c). Current and capacitance did not reach a plateau during hyperforin application, but the rate of increase decelerated. Apparently, the capacitance changes are due to an incorporation of the lipophilic hyperforin into the lipid bilayer of the plasma membrane. Microglial cells isolated from TRPC3/TRPC6-deficient mice revealed the same hyperforin-mediated current (Fig. 2f) with the same IV (Fig. 2g) as microglial cells isolated from wild-type mice, and neither cells developed a specific current upon OAG application (Fig. 2h,i). RT-PCR of 50 sorted microglial cells from wild-type mice did not reveal any transcript for TRPC6 (Fig. 2j), proofing that the hyperforin-induced current, appearing in all so far tested cells, does not depend on the presence of TRPC6.
To test which ions are responsible for the hyperforin-induced current, we modified the ion composition of the external solution. But, neither external substitution of Na 1 and K 1 (replaced by the non-permeable cation NMDG 1 ), or removal of divalent cations (nominal 0Ca0Mg), nor substitution of Cl 2 (replaced by the nonpermeable anion aspartate) significantly changed the development of the hyperforin-induced current in microglial cells (Fig. 2k). The normalized IVs of the hyperforin-mediated currents in the presence and absence of Na 1 , Ca 21 /Mg 21 and Cl 2 all reveal virtually the same shape and reversal potential, indicating that the hyperforin-induced current neither is driven by the typical monovalent and divalent cations, nor the typical anions (Fig. 2l).
Hyperforin mediates a proton conductance. We next changed the pH of the external solution, since the only remaining possibly permeable ions were protons. Applying hyperforin at more acidic or more basic external pH than 7.2 (intracellular always pH 7.2) either way resulted in a significant decrease of the current amplitude (Fig. 3a). The reversal potentials obtained from the IVs (Fig. 3b) significantly shifted towards more positive and more negative values upon extracellular acidification and alkalization, respectively (Fig. 3c). Since the development of the hyperforininduced current at acidic or basic external conditions was significantly reduced, we activated the current at external pH 7.2 and changed the pH after the current had already developed (Fig. 3d). Changing to pH 8.9 resulted in an immediate decrease of the current amplitude towards similar values as seen for the development of the hyperforin-induced current in pH 8.9. In contrast, after an initial increase of inward current, switching to pH 5.4 only led to a small decrease of the current amplitude. However, as already seen in figures 3b and c, the reversal potentials of the hyperforin-induced currents shifted towards more positive values upon external acidification and more negative values upon external alkalization (Fig. 3e,f). Figure 3g shows the plot of all measured reversal potentials in dependence of the external pH (black dots; intracellular pH 7.2). The red dotted line depicts the theoretical H 1 reversal potentials, calculated by the Nernst equation under the given intra-and extracellular pH, i.e. proton concentrations. The experimentally detected and calculated reversal potentials are similar, and the slight difference is most probably due to the substantial proton flux, which might change the H 1 concentration close to the membrane, and thus the expected reversal potential. Changing the external pH from 5.4 to 8.9 without applying hyperforin did not evoke similar currents in microglial cells ( Fig. 3h; see Fig. 3i for IVs). In summary, the results so far suggest that hyperforin mediates a proton conductance in HEK as well as microglial cells.
Carbonyl cyanide m-chlorophenylhydrazone (CCCP), a protonophore mainly used as mitochondrial un-coupler, activated currents in microglial cells (Fig. 3j) with similar IV (Fig. 3k; black trace), but significantly smaller amplitude than hyperforin. Upon external acidification to pH 5.4 the CCCP-mediated inward current increased (Fig. 3j,k), and the reversal potential changed exactly the same way as in hyperforin towards more positive values, as expected from the proton distribution ( Fig. 3l; see red trace in Fig. 3f for comparison). The red trace in figure 3k shows the initial IV of the hyperforinmediated current directly after switching to pH 5.4. The orange trace depicts the IV at the maximal current amplitude in pH 5.4, where most probably an additional mainly outward current contributed, which was not further investigated.
Assuming that hyperforin mediates a proton conductance the cytosol of intact cells is expected to become more acidic upon hyperforin application, since the negative membrane potential presents a driving force for protons into the cell. To proof this, we loaded intact www.nature.com/scientificreports SCIENTIFIC REPORTS | 4 : 7500 | DOI: 10.1038/srep07500 HEK cells with the pH-sensitive fluorescent dye BCECF-AM, applied different concentrations of hyperforin, and measured the fluorescence ratio F 490 /F 450 as indicator for intracellular pH changes. With increasing concentrations of hyperforin the BCECF fluorescence ratio dose-dependently decreased (Fig. 4a). The BCECF ratio revealed a constant rundown (see also Fig. 4c). Therefore, we calculated the hyperforin-induced relative changes of the BCECF ratio in respect to the control application of 1% DMSO, which did not significantly alter the signal. The sigmoidal fit of the relative changes of the BCECF ratio at different external hyperforin concentrations, extracted at 200 s in figure 4a, revealed an EC 50 of 8.5 mM hyperforin (Fig. 4b), which is virtually the same as for the hyperforin-induced currents in microglial cells (see Fig. 2d,e). To validate the pH measurements, NH 4 Cl (20 mM) was extracellularly applied. NH 3 enters the cells and binds H 1 elevating the intracellular pH (alkalization) resulting in an increase of the BCECF ratio (Fig. 4c), proofing that a decrease, as seen for hyperforin, represents an intracellular acidification. The relatively small negative membrane potential of HEK cells (250 mV 34 ) represents a limited driving force for protons into the cell. Therefore, we performed BCECF fluorescence imaging experiments in combination with the whole-cell patch clamp technique, to control the membrane potential of the HEK cells and thus the driving force for H 1 into and out of the cell. BCECF (free acid) was added into the patch pipette solution. After reaching a stable fluorescence ratio (Fig. 4f), the membrane potential (holding potential, V h ) was clamped to different values (Fig. 4d). Figures 4e, f, g, and h represent the normalized capacitance, intracellular pH changes (BCECF F 490 / F 450 ), inward and outward currents at 280 mV and 180 mV,  respectively, and the reversal potential before and during the application of 10 mM hyperforin. Only in the presence of hyperforin (indicated by the box), when the putative proton current appears (Fig. 4g), holding potentials of 280 mV and 180 mV mediate intracellular acidification and alkalization (Fig. 4d,f), respectively, as expected from the driving force for protons into the cell at 280 mV and out of the cell at 180 mV. As consequence the reversal potential shifts to more positive values as calculated by the Nernst equation for intracellular decrease (alkalization), and more negative values as calculated for intracellular increase (acidification) of the proton concentration (Fig. 4h). Figures 4i and j show the IVs extracted at the indicated time points (arrowheads in Fig. 4g), and thus the reversal potentials before (Fig. 4i) and during (Fig. 4j) 10 mM hyperforin application, respectively, at a holding potential of 280, 0 and 180 mV.
Hyperforin acts as a protonophore. All whole-cell patch clamp and pH imaging experiments so far clearly indicate that hyperforin mediates a proton conductance in microglia and HEK cells. To proof whether this conductance is due to proton channels in the cellular membranes of these cells, activated by hyperforin, or if hyperforin itself reveals protonophore activity, we performed patch clamp experiments on lipid bilayers 35 , devoid of any ion channel proteins. The tip of a patch pipette was dipped into the lipid film on top of the bath solution, and after establishment of a tightly sealed lipid bilayer we ran the same voltage protocol as for the whole-cell patch clamp experiments, and applied 10 mM hyperforin, 100 mM CCCP or 1% DMSO to the lipid bilayer at the tip of the patch pipette (Fig. 5a). A gigaohm-resistance and a small basic current (IV see Fig. 5b) revealed the successful formation of a tight lipid bilayer. Upon application of 10 mM hyperforin a significant current appeared, showing, that hyperforin itself is able to mediate an ion conductance. The IV of the hyperforin-mediated current was similar as in microglia and HEK cells (Fig. 5c). The small shift of the reversal potential towards more negative values (black trace Fig. 5c) most probably results from an accumulation of protons, and thus pH decrease, in the very small volume inside the tip of the patch pipette, driven by the negative potential inside the pipette at the beginning of the ramp. This is a similar situation as the intracellular acidification in whole-cell experiments during hyperforin application, driven by a negative holding potential, which also results in a shift of the reversal potential of the hyperforin-mediated current towards more negative values (see Fig. 4h,j). The protonophore CCCP (100 mM) resulted in a much smaller current than hyperforin, and 1% DMSO did not change the basic current at all (Fig. 5a,c). In summary, hyperforin itself reveals Hyperforin induces proton currents and depletes and reduces loading of monoamine-containing vesicles in chromaffin cells.
To evaluate the action of hyperforin on neurotransmitter storage and release we prepared primary mouse chromaffin cells, in which catecholamines are stored in large dense core vesicles (LDCVs) possessing the vesicular monoamine transporter 1 (VMAT1) 19 . They are excitable neuroendocrine cells, have the same neural crest precursor cells as postganglionic sympathetic neurons, and various aspects of neurotransmitter metabolism, storage and release have been extensively studied with these cells.
As expected, application of 10 mM hyperforin induced the same whole-cell current with the same IV in chromaffin cells (Fig. 6a,b) as seen in HEK cells, microglial cells and lipid bilayers. To visualize effects on vesicular monoamine storage and release, we incubated chromaffin cells from wild-type and TRPC6-deficient mice with the fluorescent ''false'' neurotransmitter FFN511 36 . Like endogenous monoamines, FFN511 is carried by the plasma membrane monoa- www.nature.com/scientificreports SCIENTIFIC REPORTS | 4 : 7500 | DOI: 10.1038/srep07500 mine transporter (MAT) into the cytosol and accumulates in a pattern consistent with LDCVs (Fig. 6c). As shown in figure 6d, e, f and g the cellular FFN511-dependent fluorescence gradually decreased under control conditions (0.1% DMSO) indicating the basic rate of FFN511 destaining, which was similar in cells from wild-type and TRPC6-deficient mice, and which apparently depended on lightactivated bleaching and spontaneous release of FFN511. However, upon application of 10 mM hyperforin or membrane depolarization (70 mM KCl) the destaining of the FFN511-dependent fluorescence was significantly accelerated. Similar results were obtained in the presence of the protonophore CCCP (10 mM; see Fig. 6e). Hyperforin-induced destaining, i.e. release of FFN511, was similar in cells isolated from wild-type or TRPC6-deficient mice (Fig. 6d,e,f). The initial increase of the FFN511-dependent fluorescence during hyperforin and CCCP application might be due to a dequenching effect after reduction of the massive accumulation of FFN511 in the monoamine vesicles or due to the hyperforin-mediated pH changes. However, finally hyperforin resulted in a decrease of the FFN511dependent fluorescence. As control we used HEK cells, which do not contain transporters for neurotransmitters and accordingly did not accumulate FFN511 (Fig. 6h).
In addition to an accelerated FFN511 destaining, loading of FFN511 into monoamine vesicles of chromaffin cells was significantly reduced in the presence of 10 mM hyperforin (Fig. 6i). Cellular (synaptic) monoamine uptake is arranged by the plasma membrane monoamine-sodium symporter (MAT; Fig. 7c (1)), driven by the Na 1 gradient and the negative membrane potential (E M ). The hyperforin-induced current did not depend on extracellular sodium (see Fig. 2k,l), however, imaging experiments on HEK cells, loaded with the Na 1 -sensitive fluorescent dye SBFI, revealed a significant increase of SBFI ratio, i.e. increase in intracellular sodium, upon hyperforin application (Fig. 7a). Using NMDG 1 instead of Na 1 in the external solution proved that the increase in SBFI ratio was not due to hyperforin-mediated pH-dependent effects on SBFI. However, the hyperforin-induced decrease of BCECF fluorescence ratio (see Fig. 4a), i.e. intracellular acidification, was accelerated in HEK cells in the absence of extracellular Na 1 (Fig. 7b), suggesting a sodium-dependent pH regulating mechanism via e.g. the plasma membrane sodium-proton exchanger (NHE), which transports protons out of the cell and, in exchange, Na ions along their concentration gradient into the cell. The protonophore action of hyperforin moves H 1 into the cell (synapse) (Fig. 7c (3)) and out of the vesicles (Fig. 7c (4)) due to the negative E M and the vesicular H 1 gradient, respectively, resulting in an intracellular acidification. Thus, along with the intracellular acidification, hyperforin leads to an increase of the intracellular Na 1 concentration, probably due to stimulation of the NHE (Fig. 7c (5)), and thereby reduces the driving force for the cellular MAT-dependent monoamine uptake (Fig. 7c (1)). Vesicular monoamine uptake is arranged by the vesicular monoamine-proton antiporter (VMAT; Fig. 7c (2)), driven by the huge H 1 gradient and the vesicular membrane potential, both established by the vesicular proton pump (H 1 -ATPase). The hyperforin-induced dissipation of the vesicular H 1 gradient (see above) ruins the driving force for the vesicular VMAT-dependent monoamine uptake.
Taken together these data show that release of FFN511 was enhanced and accumulation of FFN511 was reduced in the presence of hyperforin, which, due to its protonophore activity, collapses the vesicular pH and increases intracellular Na 1 by enhanced activation of the sodium-proton exchanger.

Discussion
The present study shows that hyperforin, the major active ingredient of St. John's wort extract, induces a prominent ion conductance in primary mouse cortical microglia and chromaffin cells, HEK-293 cells and lipid bilayers. This ion conductance is independent of the presence of TRPC6 channels, neither mediated by Na 1 , K 1 , Ca 21 , and Mg 21 nor by Cl 2 , but significantly depends on the extracellular and intracellular pH. Performing whole-cell patch clamp and pH imaging experiments we could show that the hyperforin-induced current is carried by protons. The presence of the same hyperforin-mediated conductance in lipid bilayers like in the three cell types indicates that hyperforin does not require a channel protein for its proton conducting activity, but rather itself acts as a protonophore. The non-saturating current and capacitance increase (see Fig. 2c) indicates accumulation of the lipophilic hyperforin in the plasma membrane, which might explain the effects of even low doses of hyperforin during St. John's wort extract treatment 37 .
Several models describe the H 1 translocation by protonophores across lipid bilayers. The simplest model bases on a weak acidic molecule, which is absorbed in its anionic form at the solution/membrane interface on the positive side of the membrane and there binds a H 1 . Due to the interaction with the H 1 the molecule becomes neutral and diffuses towards the negative side of the membrane. Here the H 1 leaves the weak acid again, which then, in its anionic form, moves back electrophoretically to the more positive side of the membrane and the cycle restarts 38 . The necessary driving forces are provided by the membrane potential and the proton concentration on both sides of the membrane. Since hyperforin reveals a weak acidic character 7 and seems to accumulate in the cellular membrane (see Fig. 2c), this model might also apply for the protonophore mode of action of hyperforin.
The uncoupling protein 1 (UCP1) which transports protons across the inner membrane of brown fat mitochondria is activated by long-chain fatty acids (LCFAs) 42 . It also mediates proton currents after reconstitution in planar lipid bilayers upon activation by LCFAs 43 . Recently, Fedorenko et al. 44 suggested a mechanism where the LCFAs bind to UCP1, catch protons on one side of the membrane and guide them trough UCP1 to the other side of the membrane. The protons are then released and the LCFAs stay bound at the UCP1. The mechanism of the hyperforin-mediated proton conductance might be more related to the transport mechanism of other weak acids (see above) rather than UCP1. However, the current-voltage relationship of the LCFA-activated UCP1-mediated proton current in mitoplasts, isolated from mouse brown adipose tissue 44 , looks exactly the same as the IV of the hyperforin-induced currents in the present study. In addition, the reversal potentials of the currents shift in a very similar manner upon pH changes.
Chatterjee et al. 29 reported about a dose-dependent hyperforinactivated inward current in isolated rat hippocampal neurons. The sigmoidal fit revealed an EC 50 of 9.1 mM, very close to the EC 50 for the hyperforin-induced inward and outward currents (9.3 mM and 8.7 mM, respectively) and the intracellular acidification (8.5 mM) measured in the present study. However, they claimed that some preliminary data on the voltage and ionic dependence of the hyperforin-activated inward current, which was not shown in their paper, suggested a predominant role of anions.
Our data clearly show that hyperforin itself acts as a strong protonophore mediating significant proton currents. These currents can occur at the plasma membrane, leading to H 1 influx due to the negative membrane potential, and at intracellular organelles. Most organelles keep an acidic intraorganellar environment and protonophore activity leads to efflux of protons into the cytosol. Thus, either way, the cytosolic pH becomes more acidic. This intracellular acidification and/or the dissipation of the mitochondrial proton gradient with subsequent Ca 21 and Zn 21 release 12 might directly contribute to the various effects of hyperforin, including its antiinflammatory, proapoptotic, antiproliferative and antibiotic action 1 . St. John's wort has been shown to alleviate symptoms of mild to moderate depression 6,45 , and recent meta-analysis shows that St. John's wort extracts are also effective antidepressants for major depression 46 . The most probably major active ingredient of St. John's wort, hyperforin is a potent inhibitor of 5-HT, dopamine, norepinephrine, acetylcholine, GABA and glutamate uptake in synaptosomal preparations [13][14][15]17,18 , and part of its antidepressant actions has been attributed to changes in intracellular H 1 and Na 1 concentrations and of neurotransmitter storage in synaptic vesicles 17,[22][23][24][25][26] . Cellular neurotransmitter uptake by presynaptic transporters relies on Na 1 cotransport 47 , and conditions that decrease the Na 1 gradient of the neuronal membrane, either by lowering extracellular or elevating intracellular Na 1 , significantly reduce the efficiency of these transporters. Singer et al. 23 suggested already that hyperforin inhibits serotonin uptake by elevating the intracellular Na 1 concentration in human platelets, but the mechanism how hyperforin increases the cytosolic Na 1 was not known at that time. In 2007, Leuner et al. 26 proposed that hyperforin mediates Na 1 increase, and thus effects neurotransmitter re-uptake, by specifically activating the non-selective cation channel TRPC6. However, by cytosolic acidification, as shown in figure 4, hyperforin fuels the plasma membrane sodium-proton exchanger (NHE), thereby increases free intracellular Na 1 concentration, as shown in figure 7a, which inhibits the neurotransmitter uptake, depending on Na 1 cotransport. At the same time hyperforin dissipates the H 1 gradient in neurotransmitter vesicles 16,22 by its protonophore activity and thereby impairs vesicular uptake, storage and release of neurotransmitters (see Fig. 6). Our data indicate that hyperforin's protonophore activity is under-lying both processes (see Fig. 7c) and explain why hyperforin in contrast to reserpine or the classical antidepressant monoamine reuptake inhibitors is unique in that it exerts its effects not only on monoamines but also on other neurotransmitters including acetylcholine, GABA and glutamate.

Methods
Microglia preparation and culture. Microglial cells were obtained from the cerebral cortex of newborn mice (postnatal days 0-3) as previously described 48 . In brief, pups were decapitated and their brains were removed and collected under sterile conditions in Dulbecco's Modified Eagle Medium (DMEM, Invitrogen) supplemented with 1% penicillin/streptomycin (Sigma) and 1% L-alanyl-Lglutamine (GlutaMAX, Invitrogen). Cortices were dissected, the meninges removed, and triturated after enzymatic digestion with 0.25% trypsin/EGTA and washing in medium. The homogenate was then forced through a 40 mm filter and the resulting suspension was plated in 75 cm 2 poly-L-lysine-coated culture flasks containing medium with 10% fetal bovine serum (FBS). The cells were kept in an incubator at 37uC and 5% CO 2 , and the medium was changed 3 days following preparation and once per week thereafter. After about 2-3 weeks in culture, the flasks showed a confluent layer of astrocytes with microglial cells growing on top. Microglial cells were harvested and isolated by shaking the flasks to release the loosely attached microglia into the supernatant. Microglia from the supernatant were washed, plated on poly-L-lysine-coated glass coverslips (1 cm in diameter) and kept in medium with 10% FBS. To allow the cells to adapt to the subculture conditions plated cells were used for patch clamp experiments 3-7 days after harvest.  1)). The negative membrane potential and low intracellular Na 1 concentration provide the driving force for cellular Na 1 and MA 1 uptake. Vesicular monoamine uptake is arranged by the vesicular monoamine-proton antiporter (VMAT; (2)), moving MA 1 into the vesicle in exchange for two protons. The huge H 1 gradient and the vesicular membrane potential, both established by the vesicular proton pump (H 1 -ATPase) drives H 1 out of the vesicle and thereby promotes vesicular MA 1 uptake. Effects of hyperforin on monoamine uptake (right side of the model): The protonophore action of hyperforin moves H 1 into the cell (synapse) due to the negative membrane potential (3). In addition, hyperforin moves H 1 out of the vesicles, driven by the huge H 1 gradient and positive vesicular membrane potential (4). The dissipation of the vesicular H 1 gradient ruins the driving force for the vesicular VMAT-dependent MA 1 uptake (2). The cytosolic acidification, mediated by the hyperforin-dependent H 1 influx and H 1 release from vesicles, increases the activity of the plasma membrane sodium-proton exchanger (NHE; (5)), which is driven by the Na 1 gradient and the negative membrane potential, and moves one H 1 out of the cell in exchange of one Na 1 into the cell. This results in an increase of cytosolic Na 1 concentration, reducing the driving force for the MAT-dependent cellular MA 1 uptake (1).
Chromaffin cell preparation and culture. Chromaffin cells were obtained from the adrenal glands of newborn mice (postnatal days 0-1). Pups were decapitated and their adrenal glands were dissected, collected in Locke's Medium (in mM: 154 NaCl, 5.6 KCl, 3.6 NaHCO 3 , 5.6 glucose, 5 HEPES, pH 7.3), and connective tissue was removed. After 45 min of enzymatic digestion with 10 units/ml papain in DMEM containing 0.02% L-cysteine, 1 mM CaCl 2 and 0.5 mM EDTA the enzymatic reaction was stopped by DMEM supplemented with 10% FBS, 0.25% albumin and 0.25% trypsininhibitor (Type II-O, chicken egg). Then glands were washed in ''enriched'' DMEM (4.5 g/l glucose, 0.1% Pen/Strep, 1% Insulin-Transferrin-Selenium-X) and gently triturated with a 200 ml pipette tip until getting a cell suspension. Cells were then plated on glass cover slips (1 cm in diameter) and kept in ''enriched'' DMEM in the incubator at 10% CO 2 and 37uC. Experiments were performed within 1-5 days after preparation.
Mice. TRPC6 2/2 and TRPC6 2/2 /TRPC3 2/2 mice have been described elsewhere 49,50 . Single TRPC6-and TRPC3-deficient mice were generated and kindly provided by the laboratory of Lutz Birnbaumer (NIEHS/NIHR, Research Triangle Park, NC 27709, USA). All animal care and experimental procedures were carried out in accordance with the approved guidelines. Guidelines and all experimental protocols were approved by the animal welfare committee of the University of Saarland School of Medicine.
HEK cell culture. HEK-293 cells (ATCC, CRL 1573) obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA) and HEK cells stably expressing the TRPC6 cDNA (kindly provided by Dr. M.X. Zhu, University of Texas Health Science Center, Houston) were cultured in MEM and DMEM, respectively, with 10% FBS. The cells were kept in an incubator at 37uC and 5% CO 2 , and the medium was changed every third day. Cells were plated on glass coverslips of 1 cm in diameter coated with poly-L-lysine.
Electrophysiological recordings. Membrane currents were recorded in the tight seal whole-cell patch clamp configuration using an EPC-9 amplifier (HEKA Electronics, Lambrecht, Germany). Patch pipettes were pulled from glass capillaries GB150T-8P (Science Products, Hofheim, Germany) at a vertical Puller (PC-10, Narishige, Tokyo, Japan) and had resistances between 2 and 4 MV when filled with standard internal solution (in mM: 120 Cs-glutamate, 8 NaCl, 1 MgCl 2 , 10 HEPES, 10 Cs-BAPTA, 3.1 CaCl 2 (100 nM free Ca 21 , calculated with WebMaxC (http://www.stanford.edu/ ,cpatton/webmaxcS.htm)), pH adjusted to 7.2 with CsOH). Standard external solution contained (in mM): 140 NaCl, 2.8 KCl, 2 MgCl 2 , 1 CaCl 2 , 10 HEPES, 10 glucose, pH adjusted to 7.2 with NaOH. For some experiments the pH was adjusted to 5.4 or 6.5 and 7.9 or 8.9 by adding HCl and NaOH, respectively. In monovalent cation-free solution Na 1 and K 1 were replaced by NMDG 1 (N-methyl-d-glucamine), and for nominally divalent cation-free solution CaCl 2 and MgCl 2 were omitted. Chloride-free external solution comprised of (in mM): 140 Na-aspartate, 1 Cagluconate, 2 MgSO 4 , 10 HEPES and 10 glucose, pH adjusted to 7.2 with NaOH. Hyperforin, carbonyl cyanide m-chlorophenylhydrazone (CCCP), 1-oleoyl-2-acetylsn-glycerol (OAG) or flufenamic acid (FFA) were added to the standard external solution from a 100 mM or 50 mM stock solution in DMSO to reach final concentrations as indicated. OAG and FFA experiments on TRPC6-expressing HEK cells were performed at 200 mM external CaCl 2 . All modified solutions were directly applied onto the patch-clamped cell via a pressure-driven application pipette. In some experiments a second application pipette was used. Osmolarity of all solutions ranged between 290 and 310 mOsm. Voltage ramps of 50 ms duration spanning a voltage range from -100 to 100 mV were applied at 0.5 Hz from a holding potential (V h ) of 0 mV over a period of up to 500 s using the PatchMaster software (HEKA). All voltages were corrected for a 10 mV liquid junction potential. Currents were filtered at 2.9 kHz and digitized at 100 ms intervals. Capacitive currents and series resistance were determined and corrected before each voltage ramp using the automatic capacitance compensation of the EPC-9. Basic currents before an application were subtracted to get the net developing current. Inward and outward currents were extracted from each individual ramp current recording by measuring the current amplitudes at 280 and 180 mV, respectively, and plotted versus time. Currentvoltage (IV) relationships were extracted at indicated time points. Currents were normalized to the initial size of the cell to obtain current densities (pA/pF). For some experiments changes of the cell size (normalized capacitance as measured and extracted from the automatic capacitance compensation of the EPC9) and the reversal potential of currents were plotted versus time.
pH imaging. Intracellular live cell pH imaging experiments were performed using a Polychrome II and photomultiplier (MEA1530SF-V2DN, SMT, Seefeld, Germany)based imaging system from TILL Photonics (Martinsried, Germany) at a Zeiss Axiovert 135 M fluorescence microscope equipped with a Zeiss 403/0.65 Achroplan objective. Control of the polychrome II and data acquisition was accomplished via the EPC-9 and PatchMaster software. Prior to the experiments cells were incubated in media supplemented with 2 mM of the pH-sensitive fluorescent dye BCECF-AM for 30 min in the dark at 37uC, and washed with standard external solution to remove excess BCECF-AM. The BCECF-loaded cells, growing on 1 cm glass coverslips, were transferred to a bath chamber containing standard external solution, and BCECF fluorescence of a single cell was monitored with the photomultiplier at .510 nm after excitation at 490 and 450 nm for 15 ms each at a rate of 0.5 Hz for up to 600 s. Hyperforin was added to the standard external solution from a 100 mM stock (DMSO) to reach final concentrations as indicated and directly applied onto single cells via a pressure-driven application pipette (see Fig. 4a-c). For some experiments 20 mM NH 4 Cl was added to the standard external solution and pressure applied to the measured cell. The ratio of the background-corrected BCECF fluorescence at 490 and 450 nm (F 490 /F 450 ) was plotted versus time. For some experiments BCECFdependent fluorescence was detected by a camera (SensiCam) -based imaging system (Polychrom V, TILL Photonics) at a Zeiss Axiovert 100 M fluorescence microscope via a Zeiss Fluar 203/0.75 objective, and hyperforin was added into the bath as indicated (see Fig. 7b). Data acquisition and analysis were performed by the software TILLvisION (TILL Photonics).
Combined electrophysiological recordings and pH imaging. In order to determine the hyperforin-induced changes of the intracellular pH under defined membrane potentials, the photometric technique (see pH imaging) was used in combination with the whole-cell patch clamp technique (see electrophysiological recordings). The patch pipette contained standard internal solution supplemented with 100 mM of BCECF (free acid). BCECF was excited at 490 and 450 nm for 25 ms each at a rate of 0.5 Hz for up to 800 s. Hyperforin was applied to the cell via a pressure driven application pipette as described for the pH imaging and the patch clamp experiments. The holding potential between the voltage ramps was changed from 0 mV to 280 or 180 mV as indicated (see Fig. 4d-h). Cellular currents and fluorescence changes were analyzed as described above.
Sodium imaging. Intracellular live cell Na 1 imaging experiments were performed using the same camera-based imaging system (Polychrom V, TILL Photonics) as for pH imaging (see above). Prior to the experiments cells were incubated in media supplemented with 2 mM of the Na 1 -sensitive fluorescent dye SBFI-AM for 30-45 min in the dark at 37uC, and washed with standard external solution to remove excess SBFI-AM. The SBFI-loaded cells, growing on 1 cm glass coverslips, were transferred to a bath chamber containing external solution, and SBFI fluorescence of single cells was monitored at .510 nm after excitation at 340 and 380 nm for 100 ms each at a rate of 0.5 Hz for up to 800 s. Hyperforin and DMSO were added into the bath as indicated. The ratio of the background-corrected SBFI fluorescence at 340 and 380 nm (F 340 /F 380 ) was plotted versus time (see Fig. 7a). Data acquisition and analysis were performed by the software TILLvisION (TILL Photonics).
FFN511 imaging experiments. Mouse chromaffin and HEK-293 cells, plated on glass coverslips, were incubated at 37uC for 15 min in culture media containing 10 mM of the fluorescent false neurotransmitter FFN511 (trifluoroacetate salt hydrate). For some experiments 10 mM hyperforin or 0.1% DMSO (as control) was already present during incubation with FFN511. Cells were washed with standard external ringer solution (see electrophysiological recordings) and kept in the dark for another 15 min. During the experiment every 5 s FFN511 was excited at 400 nm for a period of 10 ms and the FFN511-dependent fluorescence (.510 nm) of single cells was detected using the same camera-based imaging system (Polychrom V, TILL Photonics) as for pH and Na 1 imaging (see above). Hyperforin, CCCP, high KCl or DMSO were applied into the bath as indicated. Data acquisition and analysis were performed by the software TILLvisION (TILL Photonics).
Confocal microscopy. Mouse chromaffin cells were loaded the same way as for FFN511 imaging experiments (see above). In addition to its peak excitation at around 400 nm FFN511 also shows a substantial excitation at 488 nm. Thus, the 488 nm line of an argon laser was used to visualize the FFN511-dependent fluorescence (GFP filter settings) with a Zeiss LSM780 confocal microscope controlled by the software ZEN System 2012 (both Carl Zeiss, Oberkochen, Germany).
Lipid bilayer experiments. Lipid bilayers on the tip of patch pipettes were made from phospholipid monolayers on the surface of the bath solution (see Ref. 35). In brief, 10 mg lipids (Ionovation, Osnabrück, Germany) were dissolved in 100 ml n-decan, and 1 ml was added gently onto the surface of 1 ml bath solution in which a patch pipette was already immerged. Bath and pipette solution comprised of 150 mM KCl and 10 mM HEPES, and the pH was adjusted to 7.0 with KOH. Within about 10 minutes a lipid monolayer was formed on top of the bath solution. The patch pipette was gently moved out of the bath with a lipid monolayer forming at its tip. By immerging the patch pipette back into the lipid monolayer-covered bath solution a tight lipid bilayer (sometimes) forms at the tip of the patch pipette, indicated by a gigaohm resistance. Voltage ramps were applied and currents were measured and analyzed as described for the whole-cell patch clamp experiments above. 10 mM hyperforin, 100 mM CCCP or 1% DMSO were applied onto the lipid bilayer at the tip of the patch pipette.
Data Analysis. Initial analysis of electrophysiological and photomultiplier-based pH imaging experiments were performed with FitMaster (HEKA), and the camera-based FFN511, SBFI and pH imaging experiments were analyzed using TILLvisION (TILL Photonics). IGOR Pro (Wave Metrics, Lake Oswego, OR, USA), ImageJ (NIH, USA) and Adobe Illustrator were used for further analysis and for preparing the figures.

www.nature.com/scientificreports
Where applicable the data was averaged and given as means 6 S.E.M for an indicated number of cells. Two tailed two-sample equal variance Student's t-tests (Microsoft Excel) were used to test for significant differences. The theoretical reversal potentials for proton currents were calculated via the Nernst equation Chemicals. Hyperforin-dicyclohexylammonium (-DCHA) salt was obtained from Sigma, and hyperforin dissolved as free acid in methanol from Sigma and Biomol (Cayman Chemical). BCECF-AM and the free acid of BCECF were obtained from Molecular Probes (Eugene, OR, USA), and SBFI-AM from TEFLabs (Austin, TX, USA). The lipids for the lipid bilayer were kindly provided by Dr. Martin Jung, Medizinische Biochemie und Molekularbiologie, Homburg. FFN511 and all other chemicals were obtained from Sigma.