1. Cell Biology
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A FRET sensor of C-terminal movement reveals VRAC activation by plasma membrane DAG signaling rather than ionic strength

  1. Benjamin König
  2. Yuchen Hao
  3. Sophia Schwartz
  4. Andrew JR Plested
  5. Tobias Stauber  Is a corresponding author
  1. Freie Universität Berlin, Germany
  2. Humboldt Universität zu Berlin, Germany
  3. Leibniz Forschungsinstitut für Molekulare Pharmakologie (FMP), Germany
  4. Charité Universitätsmedizin, Germany
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Cite this article as: eLife 2019;8:e45421 doi: 10.7554/eLife.45421

Abstract

Volume-regulated anion channels (VRACs) are central to cell volume regulation. Recently identified as hetero-hexamers formed by LRRC8 proteins, their activation mechanism remains elusive. Here, we measured Förster resonance energy transfer (FRET) between fluorescent proteins fused to the C-termini of LRRC8 subunits. Inter-subunit FRET from LRRC8 complexes tracked VRAC activation. With patch-clamp fluorometry, we confirmed that the cytoplasmic domains rearrange during VRAC opening. With these FRET reporters, we determined VRAC activation, non-invasively, in live cells and their subcompartments. Reduced intracellular ionic strength did not directly activate VRACs, and VRACs were not activated on endomembranes. Instead, pharmacological manipulation of diacylglycerol (DAG), and protein kinase D (PKD) activity, activated or inhibited plasma membrane-localized VRACs. Finally, we resolved previous contradictory reports concerning VRAC activation, using FRET to detect robust activation by PMA that was absent during whole-cell patch clamp. Overall, non-invasive VRAC measurement by FRET is an essential tool for unraveling its activation mechanism.

https://doi.org/10.7554/eLife.45421.001

Introduction

Volume regulation is essential for virtually all cell types, not only to counteract osmotic swelling and shrinkage but also during various processes like proliferation, migration and apoptosis (Hoffmann et al., 2009). Volume-regulated anion channels (VRACs) are ubiquitously expressed and involved in regulatory volume decrease (RVD) of vertebrate cells (Chen et al., 2019; Jentsch, 2016; Stauber, 2015). These channels open slowly following osmotic cell swelling and mediate the extrusion of chloride and organic osmolytes, leading to the efflux of water and hence cell shrinkage.

VRACs are composed of LRRC8 heteromers containing the essential subunit LRRC8A and at least one other LRRC8 protein, LRRC8B-E (Qiu et al., 2014; Voss et al., 2014). The subunit composition determines various biophysical properties and substrate permeability (Planells-Cases et al., 2015; Syeda et al., 2016; Voss et al., 2014). Although unlikely to be of physiological relevance, LRRC8A homomers can form anion channels, but with weak volume-sensitivity and very low conductance (Deneka et al., 2018; Syeda et al., 2016; Yamada and Strange, 2018). LRRC8 proteins form hexamers with the N-terminal halves of the six LRRC8 proteins constituting the transmembrane domain of the channel complex with a single pore. The C-terminal halves contain at least 16 leucine-rich repeats that form horseshoe-like structures reaching into the cytoplasm and their apparent flexibility has been hypothesized to be related to channel gating (Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019).

Prior to cloning, the biophysical properties of VRAC-mediated chloride currents (ICl,swell) and osmolyte conductance were studied extensively using mainly electrophysiological and pharmacological methods (König and Stauber, 2019; Pedersen et al., 2016; Strange et al., 2019); but their activation mechanism remained enigmatic for decades, in the absence of an unequivocal molecular identity for the channel. Numerous studies proposed contributions from membrane tension, the actin cytoskeleton and signaling pathways involving reactive oxygen species, calcium, G-proteins and phosphorylation cascades (Akita and Okada, 2014; Chen et al., 2019; Pedersen et al., 2016). Notably, the reduction of intracellular ionic strength (Γi) was reported to be crucial for VRAC activation upon osmotic cell swelling (Voets et al., 1999) and reconstituted LRRC8 complexes in lipid bilayers could indeed be activated by low ionic strength (Syeda et al., 2016). However, VRACs can also be activated under iso-osmotic conditions that are unlikely to encompass any changes in Γi (Pedersen et al., 2016; Strange et al., 2019).

Here, we use intra-complex Förster resonance energy transfer (FRET) to monitor VRAC activity in a spatio-temporal manner. This method demonstrates movement of the LRRC8 C-termini and provides insights into the activation mechanism of VRACs.

Results

Inter-subunit FRET shows C-terminal movement during VRAC activation

Following the hypothesis of a conformational rearrangement of the LRRC8 C-terminal domains during VRAC activation, we aimed to monitor VRAC activity by inter-subunit FRET (Figure 1A). We first used acceptor photo-bleaching to test for FRET between cyan- and yellow-fluorescent proteins (CFP/YFP) fused to the C-termini of VRAC subunits. In HeLa cells co-expressing CFP- and YFP tagged LRRC8A (A-CFP/A-YFP), bleaching of YFP (the FRET acceptor) robustly increased the intensity of CFP (the FRET donor), compared to unbleached cells in the same field of view (Figure 1B,C). The same was observed when LRRC8A was combined with the VRAC subunit LRRC8E (A-CFP/E-YFP or A-YFP/E-CFP), but not when combined with plasma membrane-localized CD4-YFP, used as control for FRET by membrane crowding (Figure 1C). Therefore, irrespective of VRAC subunit composition, the C-termini are within FRET distance for the fluorescent protein adjuncts (~5 nm).

Figure 1 with 2 supplements see all
FRET between fluorescent proteins fused to LRRC8 C-termini and its changes with VRAC activity.

(A) Schematic view of two VRAC subunits of a hexamer fused with the FRET pair CFP and YFP at their C-termini. Hypothetical movement of the intracellular C-terminal domain leads to FRET changes. (B) Wide-field images of HeLa cells expressing A-CFP/A-YFP before and after bleaching YFP. White circles indicate the size of the closed field diaphragm to bleach this part of the FOV. Scale bar, 20 µm. (C) Quantification of acceptor photo-bleach experiments. For each experiment, CFP intensity was measured before and after bleaching for individual cells within (bleached, black diamonds) and outside (control, red diamonds) of the bleached part of the FOV. ER: overexpressed proteins were trapped in the endoplasmic reticulum by 5 µg/ml BFA during expression. Data represent percentage of CFP intensity change for individual cells (diamonds) and mean ± s.e.m. (D) Representation of an seFRET experiment with an A-CFP/E-YFP-expressing cell. Left: images of the acquired channels (donor CFP, acceptor YFP, FRET) with the calculated cFRET map for two time points in isotonic and hypotonic buffer below. Scale bar, 10 µm. Images were acquired in 10 s intervals with 8 × 8 binning. Right: normalized cFRET values during buffer exchange experiment. (E) Measurements of cFRET, normalized to the value in isotonic buffer, when switching to hypotonic buffer for cells expressing A-CFP/E-YFP (n = 9 dishes with 47 HeLa cells; n = 8 with 29 HEK293 cells), A-CFP/A-YFP (n = 8, 24 HeLa cells), glutamate receptor GluA2-6Y-10C (n = 4, 13 HEK293 cells) and CFP-18aa-YFP (n = 4, 11 HeLa cells). Data represent individual cells (diamonds) and mean ± s.e.m. (F) Simultaneous measurements of whole-cell current at −80 mV (open circles) and normalized cFRET values (solid circles) during buffer exchange experiment with A-CFP/E-YFP-expressing HEK293 KO cells. Data represent mean of 4 cells ± s.d. Above, representative current traces from voltage-step protocols at the indicated time points (a, b, c). (G) Left: normalized cFRET in a time-course for a single cell with application of decreasing osmolarities (as indicated above in mOsm). Right: summary of experiments with different osmolarity steps to cover the titration curve from 50 to 400 mOsm. In total, 68 cells were measured for the titration curve, n = 7–21 depending on osmolarity. Data represent mean ± s.e.m. Statistics: *p<0.05; ***p<0.0005; n.s., not significant, Student’s t-test, comparing bleached and control (C) or hypotonic with isotonic (E).

https://doi.org/10.7554/eLife.45421.002

Next, we explored whether FRET changes during VRAC activation. We used sensitized-emission FRET and monitored the corrected FRET (cFRET, see Materials and methods) values in HeLa cells expressing A-CFP/E-YFP (Figure 1D). Switching from isotonic (340 mOsm) to hypotonic (250 mOsm) buffer led to a robust decrease in cFRET by 5–10% within 90 s in both HeLa and HEK293 cells (Figure 1E). In contrast, the intra-molecular cFRET of a cytosolic tandem FRET pair construct, CFP-18aa-YFP (Elder et al., 2009), and of a plasma membrane-localized ionotropic glutamate receptor (Zachariassen et al., 2016) were unaffected by hypotonic buffer (Figure 1E, Figure 1—figure supplement 1). These observations rule out FRET artifacts from changes of osmolarity or ion concentrations. To better understand the cFRET changes with the activation of VRAC, we expressed fluorescently-labelled A-CFP/E-YFP (in this case Cerulean/Venus, see Materials and methods) in HEK293 cells deficient for all five LRRC8 genes (HEK293 KO) (Lutter et al., 2017) and monitored cFRET and whole-cell currents simultaneously using patch-clamp fluorometry (Zheng and Zagotta, 2003). The kinetics of the cFRET drop and its reversibility correlated with the activation and inactivation of hypotonicity-induced VRAC-mediated anion currents (Figure 1F), demonstrating that the cFRET decrease indeed reflects VRAC opening. The current was in addition to any potential residual activity at isotonicity due to the fluorescent tags (Gaitán-Peñas et al., 2016). A progressive decrease in cFRET accompanied additional hypotonicity but, again mirroring VRAC, cFRET did not increase with hypertonic treatment (Figure 1G, Figure 1—figure supplement 2A). We frequently observed varying extents of intracellular retention of LRRC8A or LRRC8E. We did not see a correlation between intensity of CFP or YFP and absolute cFRET values (not shown), neither did relative changes in cFRET depend on the A-CFP/E-YFP expression ratio (Figure 1—figure supplement 2B). Remarkably, cFRET also decreased for A-CFP/A-YFP constructs (Figure 1E, Figure 1—figure supplement 1), providing evidence for activation of homomeric LRRC8A complexes in a cellular context.

Overall, these cFRET measurements show that the C-termini of LRRC8 proteins in VRAC complexes rearrange during activation and that this can be exploited to monitor their activity in living cells via FRET microscopy.

Reduced ionic strength does not activate VRAC on endomembranes

Reduced ionic strength is sufficient to activate reconstituted VRACs in lipid bilayer droplets (Syeda et al., 2016). To measure intracellular ionic strength, we used a FRET-based sensor that reports the electrostatic attraction between arginine and aspartate residues in opposed α-helices (‘RD’ sensor) (Liu et al., 2017). As expected, this sensor indicated a reversible decrease in Γi in HeLa cells during hypotonic treatment (Figure 2A). Notably, the decrease was homogeneous over the entire cell volume (Figure 2B, Figure 2—video 1). If a decline in Γi were sufficient to activate VRAC, then we hypothesized that it should already do so in the secretory pathway. To test for this, we trapped FRET-endowed VRACs in the endoplasmic reticulum (ER) by treatment with brefeldin-A (BFA) during expression (Figure 2C). FRET and hence complex formation in the ER of A-CFP/E-YFP (but not A-CFP/CD4-YFP) was confirmed by acceptor bleaching (Figure 1C). However, ER-trapped VRACs surprisingly did not respond to hypotonic treatment (Figure 2D). This was likely not due to incomplete glycosylation since mutation of the LRRC8A N-glycosylation sites (Voss et al., 2014) did not impinge on the activation-dependent cFRET drop at the plasma membrane (Figure 2D).

Figure 2 with 1 supplement see all
Hypotonicity reduces ionic strength over the whole cell, but does not activate intracellular VRACs.

(A) Changes of ionic strength measured in HeLa cells expressing the ratiometric RD sensor during hypotonic treatment. FRET intensity was divided by Cerulean intensity and normalized to basal values, with increasing values corresponding to decreasing ionic strength. Data represent mean of 10 cells ± s.d. (B) Ratio maps of two representative cells in isotonic (green star in panel A) and hypotonic (magenta star in panel A) buffer. Right: intensity profile along the white line shown in the ratio maps. (C) Epifluorescence images of A-GFP and E-RFP in live cells without (top row) and with (bottom row) 5 µg/ml brefeldin-A (BFA) treatment. Images of BFA-treated cells were background-subtracted by rolling-ball algorithm (radius = 50 pixels) and contrast increased by an unsharp mask (sigma = 3, weight = 0.7). Scale bar, 20 µm. (D) Normalized cFRET of A-CFP/E-YFP trapped in the ER by presence of BFA during expression (ER; n = 16 dishes with 23 cells) and AN66A,N83A-CFP/E-YFP without BFA (noGlyc PM; n = 7, 21 cells). Data represent individual cells (diamonds) and mean ± s.e.m. Statistics: ***p<0.0005; n.s., not significant, Student’s t-test, comparing hypotonic with isotonic.

https://doi.org/10.7554/eLife.45421.006

Intrigued by this finding, we set out to follow VRACs along the secretory pathway. To this end, we exploited a reverse aggregation system and fused two self-dimerizing mutated FK506-binding protein (FM) domains (Rollins et al., 2000) to the C-terminus of A-CFP. Inter-molecular dimerization of FM domains should lead to aggregation of VRACs in the ER, and upon addition of D/D solubilizer they should disaggregate and travel in bulk along the secretory route to the plasma membrane (Figure 3A). Indeed, initially aggregated A-CFP-FM2 expressed alone (Figure 3B, Figure 3—video 1) or with E-YFP (not shown) spread homogenously over the ER within 10 min in presence of D/D solubilizer, peaked in the Golgi apparatus at 80 min and displayed clear plasma membrane localization after 165 min. Concurrent sensitized FRET measurements showed that neither ER-localized (at 10 min) nor Golgi-localized (80 min) A-CFP-FM2/E-YFP VRAC responded to application of hypotonic buffer. On the other hand, as expected, cFRET was reduced for A-CFP-FM2/E-YFP that had reached the plasma membrane after 165 min (Figure 3C). Thus, despite hypotonic treatment reducing Γi homogenously over the whole cell, VRACs are only activated when they reach the plasma membrane.

Figure 3 with 3 supplements see all
Plasma membrane localization is required for VRAC activation.

(A) Scheme depicting the different subcellular localizations of VRACs using the reverse aggregation system. Upon addition of the D/D solubilizer, VRACs carrying FM dimerization domains disaggregate in the ER and travel through the Golgi complex to the plasma membrane. Indicated time points were deduced from experiments shown in (B) and (Figure 3—video 1). (B) Still images of HeLa cells expressing LRRC8A-CFP-FM2 and GalNAcT2-RFP as Golgi marker from Figure 3—video 1 at time points indicated in (A). Scale bar, 20 µm. (C) cFRET measured in HeLa cells expressing A-CFP-FM2/E-YFP. VRACs localizing to the ER (n = 4, 10 cells), Golgi (n = 5, 26 cells) and plasma membrane (PM; n = 5, 11 cells) were measured at 10, 80 and 165 min after addition of D/D solubilizer. Data in time traces represent mean ± s.d. Right: average normalized cFRET in of the first seven time points in isotonic and last three time points in hypotonic buffer. Data represent individual cells (ER, PM) or FOVs (Golgi) and mean ± s.e.m. Statistics: ***p<0.0005; n.s., not significant, Student’s t-test, comparing hypotonic with isotonic.

https://doi.org/10.7554/eLife.45421.009

Robust VRAC activation by PMA is blocked by whole-cell patch clamp

Next, we asked which plasma membrane-specific factors might be required for VRAC activation. First, we tested the influence of the actin cytoskeleton and cholesterol content, both of which have previously been implicated in cell volume regulation (Byfield et al., 2006; Klausen et al., 2006). In agreement with earlier reports on ICl,swell in lymphocytes (Levitan et al., 1995), depolymerization of the actin cytoskeleton by latrunculin B did not alter VRAC activation by ~25% hypotonicity (Figure 3—figure supplement 2A,B). In contrast, cholesterol depletion by methyl-β-cyclodextrin potentiated VRAC activation, as judged by the reduction in cFRET (Figure 3—figure supplement 2B,C), as was previously shown for VRAC-mediated ICl,swell in bovine aortic endothelial cells (Levitan et al., 2000). However, absence of cholesterol is unlikely to underlie plasma membrane localization as a prerequisite for VRAC activation, since the ER membrane usually has lower cholesterol concentrations than the plasma membrane.

Phospholipase C (PLC) activity was reported as a determinant of ICl,swell activation, but contradictory results abound for the involvement of protein kinase C (PKC), activated by the PLC product diacylglycol (DAG) is required (Hermoso et al., 2004; Roman et al., 1998; Zholos et al., 2005). PKC contains a C1 domain and can hence be activated by phorbol-12-myristate-13-acetate (PMA). In isotonic conditions, PMA evoked a robust decrease in cFRET in HeLa and HEK293 cells expressing A-CFP/A-YFP or A-CFP/E-YFP (Figure 4A), but not CFP-18aa-YFP (Figure 4—figure supplement 1A). This reflects gating of the LRRC8 complex, because subsequent inactivation of VRAC by treatment with hyper-osmotic solution restored basal FRET levels (Figure 4—figure supplement 1B). Like hypotonicity-driven VRAC activation, activation by PMA required cell surface localization, as VRACs released from aggregation were immune to PMA in the ER and Golgi, but they were activated at the plasma membrane (Figure 4—figure supplement 1C). Surprisingly, recording HEK293 KO cells expressing A-CFP/E-YFP in whole-cell voltage-clamp configuration, we could no longer detect any FRET reduction, nor did PMA activate VRAC currents (Figure 4B). To test whether this discrepancy was due to the absence of endogenous LRRC8 proteins, or to the dialysis of the cell contents during whole-cell patch, we first incubated HEK293 KO cells expressing A-CFP/E-YFP (Figure 4C, left panel) and wild-type (WT) HEK293 (Figure 4C, right panel) with PMA for at least five minutes before membrane breakthrough. We then established the whole-cell configuration for immediate current measurements. Indeed, with this experimental design we observed a reliable increase in classical VRAC currents in HEK293 KO cells expressing A-CFP/E-YFP incubated with PMA. The increase was robust compared to cells in isotonic buffer alone, or untransfected HEK293 KO cells incubated with PMA (Figure 4C). This result demonstrates that the whole-cell voltage-clamp configuration affects signaling pathways involved in activation of VRAC in a time-dependent manner, an experimental caveat that can be overcome with our non-invasive FRET sensor.

Figure 4 with 2 supplements see all
VRAC activation by PMA.

(A) Addition PMA (1 µM, indicated by arrows) activated VRACs composed of A-CFP/A-YFP (left; 11 cells), A-CFP/E-YFP (middle; 12 cells) in HeLa cells or A-CFP/A-YFP in HEK293 cells (right; 12 cells). Data represent mean ± s.d. (B) Simultaneous measurements of whole-cell current at −80 mV (open circles) and normalized cFRET values (solid circles) in A-CFP/E-YFP-expressing HEK293 KO cells with addition of 1 µM PMA (indicated by arrow). Data represent mean of 3 cells ± s.d. Above, representative current traces from voltage-step protocols at the depicted time points (a, b, c). (C) Whole-cell currents at −80 mV of A-CFP/E-YFP-expressing (and untransfected as control, n = 8 cells) HEK293 KO cells (left) and current densities of wild-type HEK293 cells (right) with whole-cell configuration established after pre-incubation with indicated buffers: isotonic buffer without (n = 9) or with 1 µM PMA (n = 8) (left), isotonic (n = 7) and hypotonic (n = 7) buffers without and isotonic buffer with 1 µM PMA (n = 8) (right). Data represent individual cells (diamonds) and mean ± s.e.m. Above, representative current traces from voltage-step protocols for each condition.

https://doi.org/10.7554/eLife.45421.014

Protein kinase D activity, rather than ionic strength, determines VRAC activation

Using the PKC inhibitor Gö6983 not only prevented the cFRET reduction upon hypotonicity, it also gradually increased cFRET over the course of the experiment (Figure 4—figure supplement 2A). However, in the whole-cell patch-clamp configuration, the hypotonicity-induced current even persisted in isotonic buffer when supplemented with Gö6983 (Figure 4—figure supplement 2B, left panel). The cFRET of the same cells, on the other hand increased to high values (Figure 4—figure supplement 2B, middle panel). Again, we observed different results for the cell being recorded in whole-cell mode and neighboring non-dialyzed cells in the same field of view (FOV). For neighboring cells, cFRET changes matched those seen for HeLa cells expressing A-CFP/E-YFP (Figure 4—figure supplement 2A,B, right panel). Together, these data do not support a role for PKC in VRAC activation.

However, because DAG and PMA also recruit C1 domain-containing protein kinase D (PKD) to the plasma membrane (Rozengurt, 2011), we next assessed a possible involvement of PKD. The presence of the PKD inhibitor CRT 0066101 during hypotonic treatment abolished cFRET changes and strongly reduced VRAC currents in HEK293 KO cells expressing A-CFP/E-YFP (Figure 5A). To exclude the potential confounder of cell dialysis during voltage clamp, we also measured steady-state currents in cells that were incubated in hypotonic buffer before membrane breakthrough, with and without CRT 0066101 or Gö6983. Similar to incubation during the whole-cell patch-clamp experiment, the presence of CRT 0066101 readily diminished currents following breakthrough after pre-incubation in hypotonic buffer, whereas Gö6983 did not (Figure 5B). This suggests an essential role of DAG-activated PKD in hypotonicity-induced VRAC activation.

Figure 5 with 1 supplement see all
DAG and PKD as determinants of VRAC activity.

(A) Left: simultaneous measurements of whole-cell current at −80 mV (open circles) and normalized cFRET values (solid circles) during buffer exchange experiment with A-CFP/E-YFP-expressing HEK293 KO cells, after 15 min pre-incubation with 5 µM CRT 0066101 in the presence of CRT 0066101. Data represent mean of 5 cells ± s.d. Right: average currents of the last five time points in hypotonicity with (data from left panel) and without (data from Figure 1F) CRT 0066101; data represent individual cells (diamonds) and mean ± s.e.m. (B) Whole-cell currents of HEK293 KO cells expressing A-CFP/E-YFP clamped at −80 mV after pre-incubation with indicated buffers: isotonic buffer (n = 9), hypotonic buffer (n = 7), hypotonic buffer with 5 µM CRT 0066101 (n = 4) or 1 µM Gö6983 (n = 5). Data represent individual cells (diamonds) and mean ± s.e.m. (C) Recovery of whole-cell currents in HEK293 cells expressing A-CFP/E-YFP (left) and wild-type HEK293 (right) at −80 mV to basal levels after switching from extracellular hypotonicity to isotonicity in the absence or presence of the DAG kinase inhibitor dioctanoylglycol (100 µM DOG). In HEK293 KO, n = 3 for untreated (from measurements in Figure 1F) and n = 3 for DOG-treated; in wild-type HEK293, n = 3 for untreated and n = 4 for DOG-treated. Data represent individual cells (diamonds) and mean ± s.e.m. (D) cFRET was measured for HeLa cells expressing either VRACs containing A-CFP/E-YFP (left, n = 7 dishes with 9 cells) or the RD sensor for ionic strength (right, n = 6, 23 cells) consecutively in isotonic buffer, hypotonic buffer supplemented with 100 µM DOG and isotonic buffer with DOG. Average cFRET of the last five time points per condition. Data represent individual cells and mean ± s.e.m. Statistics: *p<0.05; ***p<0.0005; n.s., not significant, Student’s t-test.

https://doi.org/10.7554/eLife.45421.018

DAG is converted to phosphatidic acid by DAG kinases (DGKs), and so inhibition of DGKs by dioctanoylglycol (DOG) should lead to accumulation of DAG. According to our hypothesis concerning PKD signaling, following VRAC activation by hypotonicity, DOG should support persistent activity of VRAC even after returning to isotonic conditions. Indeed, in the presence of DOG, we observed a persistent VRAC current after switching from extracellular hypotonicity to normal, isotonic buffer for A-CFP/E-YFP expressed in HEK293 KO cells (Figure 5C, left panel, Figure 5—figure supplement 1A, left panel), as well as for endogenous VRAC in WT HEK293 cells (Figure 5C, right panel, Figure 5—figure supplement 1A, right panel). For the A-CFP/E-YFP VRAC expressed in either HEK293 KO cells in whole-cell patch clamp (Figure 5—figure supplement 1A, left panel) and non-dialyzed HeLa cells (Figure 5D, left panel, Figure 5—figure supplement 1B) the DGK inhibitor DOG impaired the recovery of cFRET to basal values when cells were returned to isotonicity after hypotonic treatment. While cFRET in HeLa cells without DOG recovered by >80% (recovery of normalized cFRET from 95.2 ± 2.5% in hypotonic buffer to 99.4 ± 4.0% in isotonic buffer for 47 cells), cFRET in DOG-treated HeLa cells recovered by less than 40% (from 89.6 ± 8.0% to 93.6 ± 5.9%, for 9 cells). In striking contrast, Γi fully recovered to the normal levels irrespective of the presence of DOG (Figure 2A, Figure 5D, right panel). These results demonstrate that opening of VRACs is not directly coupled to Γi, but rather to the DAG pathway, likely through activation of PKD in HeLa and HEK293 cells.

Discussion

Here we present an intra-complex FRET sensor to monitor VRAC activity in live cells with subcellular resolution. The relationship of the observed FRET changes to VRAC gating are clear from numerous common features (Akita and Okada, 2014; König and Stauber, 2019; Pedersen et al., 2016; Strange et al., 2019): (i) the reversible decrease in FRET matches reversible VRAC activation upon treatment with hypo-osmotic solutions, and with the same kinetics as simultaneously-measured whole-cell VRAC currents, (ii) the intensity of the FRET decrease correlates with the applied extracellular hypotonicity; (iii) the modulation of VRAC currents, for example by cholesterol depletion, is mirrored by the observed FRET alteration; (iv) after activation with PMA, FRET rises in hypertonicity as VRAC is inactivated by cell shrinkage. FRET sensors unrelated to VRAC are not affected by hypo-osmotic solutions, demonstrating that the observed changes with our FRET sensor are indeed specific for VRAC activation. Moreover, our optical method to monitor VRAC activity has some clear advantages over electrophysiological approaches, including its application to study subcellular profiles of VRAC activity, and its non-invasive nature. The activation of low-conducting VRACs composed of only LRRC8A (Deneka et al., 2018; Kefauver et al., 2018; Syeda et al., 2016; Yamada and Strange, 2018) could also be detected by FRET. Importantly, the non-invasive, genetically-encoded FRET sensor enables the study of VRAC in connection with signaling pathways that are altered (presumably by dialysis of critical signaling molecules) by the whole-cell patch-clamp configuration. This way we could observe VRAC activation by PMA, which was otherwise blocked by whole-cell patch clamp of cells.

Cryo-EM structures of VRAC complexes suggest a certain flexibility of their C-terminal leucine-rich domains (LRRDs) on the intracellular side. Related conformational rearrangements might be related to channel gating (Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019). In support of this idea, we observe a decrease in FRET between fluorophores at the C-termini of LRRC8 proteins upon VRAC activation, which may reflect a displacement of LRRDs or possibly a simple change in the average orientation of the fluorophores to each other. Thus, as has previously been shown for cyclic nucleotide-gated (CNG) channels (Zheng and Zagotta, 2000), slow gating of CLC chloride channels (Bykova et al., 2006), BK potassium channels (Miranda et al., 2013) and ionotropic glutamate receptors (Zachariassen et al., 2016), changes in FRET demonstrate movement of the cytosolic domains during VRAC activation.

VRACs were previously shown to be directly activated by low ionic strength in lipid bilayer reconstitution (Syeda et al., 2016). However, we find that reduced Γi is neither sufficient to activate VRAC on intracellular compartments, nor is it indispensable to keep plasma membrane-localized VRACs active. When cells were treated with the DAG kinase inhibitor DOG, VRAC remained active in isotonic buffer after activation by hypotonicity, even though Γi recovered to normal levels. The notion that ionic strength directly activates VRACs (Sabirov et al., 2000; Voets et al., 1999) has been controversial (König and Stauber, 2019; Strange et al., 2019). Several studies reported VRAC activation when isotonic fluid was injected into, or inactivation when fluid was withdrawn from the cells without altering the intracellular ionic strength (Best and Brown, 2009; Cannon et al., 1998; Poletto Chaves and Varanda, 2008; Zhang and Lieberman, 1996). Moreover, the extents of low ionic strength required for VRAC activation (Cannon et al., 1998; Deneka et al., 2018; Nilius et al., 1998; Syeda et al., 2016) are unlikely to occur in physiological conditions or during whole-cell current measurements routinely used to study hypotonicity-induced VRAC currents (Strange et al., 2019). Instead, a putative role for ionic strength in controlling the volume set point for VRAC activation (Best and Brown, 2009; Cannon et al., 1998; Jackson et al., 1996) would be consistent with our findings.

Rather than ionic strength, we observe a specific and potent action of the DAG-PKD pathway in VRAC activation. Application of PMA led to VRAC opening even under isotonic conditions and VRACs did not inactivate completely in isotonic buffer after activation by hypotonicity when the consumption of DAG by DAG kinase was inhibited. A role for DAG, which is generated by the activity of phospholipases C (PLCs), in the signaling for VRAC activation has been reported previously, for example (Roman et al., 1998; Rudkouskaya et al., 2008; Zholos et al., 2005). However, there are multiple contradictory reports on the involvement of DAG-activated PKCs. In some studies, PKCs were shown to be required for regulatory volume decrease (Hermoso et al., 2004), activation of swelling-induced VRAC currents (Roman et al., 1998) and ATP-induced VRAC-mediated glutamate release (Hyzinski-García et al., 2014; Rudkouskaya et al., 2008). These results are somewhat compromised by variations in the efficacy of pharmacological PKC inhibitors (Haskew-Layton et al., 2005; Mongin and Kimelberg, 2005). Other studies found no role for PKC despite a VRAC-activating effect of DAG (Zholos et al., 2005), or even reduction or prevention of hypotonicity-induced VRAC-mediated currents by pharmacological PKC inhibition (Ben Soussia et al., 2012; Shlyonsky et al., 2011; von Weikersthal et al., 1999). While our data speak against a core role for PKC in VRAC activation, they do not rule out a regulatory role for this kinase family. Instead, our experiments reveal that protein kinase D (PKD) activity is critically involved in hypotonicity-induced activation of VRAC. LRRC8 proteins possess several putative and some confirmed phosphorylation sites for various kinases in their intracellular loops and C-terminal domains (Abascal and Zardoya, 2012; Mongin, 2016). As opposed to direct phosphorylation of LRRC8 subunits, PKD may act on another, as yet unknown, third party in the process of VRAC activation.

The central role for DAG and PKD, however, does not explain the observed plasma membrane requirement for VRAC activation by hypotonicity as PKD is also recruited to endomembranes of the secretory pathway by DAG (Baron and Malhotra, 2002; Fu and Rubin, 2011). Plasma membrane-intrinsic properties such as membrane stiffness, the lipid composition, including the amount of cholesterol and different phosphatidylinositol phosphates (PIPs), or the transmembrane potential may play a role. Besides, further limiting factors are likely to be involved, potentially explaining why the overexpression of functional LRRC8 heteromers does not increase ICl,swell above wild-type levels (Voss et al., 2014) and similar VRAC subunit expression levels displayed different activities in different cell types (Okada et al., 2017). Such limiting factors at the plasma membrane could include substrates of PKD. Although we also observed PMA-mediated activation of LRRC8A homomers, regulatory proteins could require particular VRAC subunit combinations to contribute to VRAC activation. Alternatively, cell type-specific factors may exist, explaining inconsistent findings on the signal transduction pathways impinging on VRAC activation. This could explain divergent results related to PKC in different cell lines (Ben Soussia et al., 2012; Hermoso et al., 2004; Roman et al., 1998; von Weikersthal et al., 1999; Zholos et al., 2005).

In summary, activation-dependent inter-subunit FRET in VRACs demonstrated movement of their C-termini and enabled subcellular monitoring of VRAC activity. Importantly, this noninvasive FRET sensor supersedes some aspects of electrophysiological measurements because it leaves intracellular signaling pathways untouched. Measurement of FRET can also be used for non-invasive structure-activity studies. The genetic encoding of this sensor also makes it highly amenable to studying VRAC activation during various cell physiological processes in situ.

Materials and methods

Key resources table
Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional
information
Cell line (Homo sapiens)HeLaRRID: CVCL_0030ACC no. 57Obtained from Leibniz Forschungsinstitut DSMZ (Deutsche Sammlung von Mikroorganismen undZellkulturen GmbH, Germany)
Cell line (Homo sapiens)HEK293RRID: CVCL_0045ACC no. 305Obtained from LeibnizForschungsinstitut DSMZ (Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, Germany)
Cell line (Homo sapiens)HEK293 KO (LRRC8-/-)Lutter et al., 2017 (PMID:28193731)Gift from T.J. Jentsch (MDC and FMP, Berlin, Germany)
Transfected constructA-GFP (LRRC8A-GFP)Voss et al., 2014 (PMID: 24790029)Gift from T.J. Jentsch (MDC and FMP, Berlin, Germany)
Transfected constructA-CFP (LRRC8A-CFP)This paperLRRC8A-codingsequence subcloned from A-GFP (Voss et al., 2014)
Transfected constructA-CFP-FM2(LRRC8A-CFP-FM2)This paper2 FM domains (Rollins et al., 2000) cloned in LRRC8A-CFP with insertion sites generated by Q5-mutagenesis
Transfected constructAN66A,N83A-CFP (LRRC8AN66A,N83A-CFP)This paperLRRC8A-coding sequence subcloned from AN66A,N83A-GFP (Voss et al., 2014; gift from T.J. Jentsch, MDC and FMP, Berlin, Germany)
Transfected constructA-CFP (LRRC8A-Cerulean)This paperLRRC8A-coding sequence subcloned from A-GFP (Voss et al., 2014)
Transfected constructA-YFP (LRRC8A-YFP)This paperLRRC8A-coding sequence subcloned from A-GFP (Voss et al., 2014)
Transfected constructE-CFP (LRRC8E-CFP)This paperLRRC8E-coding sequence subcloned from LRRC8E-GFP(Voss et al., 2014; gift from T.J. Jentsch, MDC and FMP, Berlin, Germany)
Transfected constructE-YFP (LRRC8E-YFP)This paperLRRC8E-coding sequence subcloned from LRRC8E-GFP (Voss et al., 2014)
Transfected
construct
E-YFP (LRRC8E-Venus)This paperLRRC8E-coding sequence subcloned from LRRC8E-GFP (Voss et al., 2014)
Transfected constructE-RFP (LRRC8E-RFP)This paperLRRC8E-coding sequence subcloned from LRRC8E-GFP (Voss et al., 2014)
Transfected constructCFP-18AA-YFPElder et al., 2009 (PMCID: PMC2706461)Gift from C.F. Kaminski (University of Cambridge, UK)
Transfected constructGluA2-6Y-10CZachariassen et al., 2016 (PMID:27313205)
Transfected constructRD sensor (ionic strength sensor)Liu et al., 2017 (PMID: 28853549)Gift from B. Poolman and A.J. Boersma (University of Groningen, The Netherlands)
Transfected
construct
ER-YFP (pEYFP-ER)Clontech, TaKaRaCatalog no. 6906–1
Transfected constructGalNAc-T2-RFPThis paperCoding sequence of stalk region of GalNAc-T2 subcloned from GalNAc-T2-GFP (Le Bot et al., 1998; gift from I. Vernos, CRG, Barcelona, Spain)
Transfected constructCD4-YFPThis paperhCD4-coding sequence subcloned from CD4-GFP(Leisle et al., 2011; PMID: 21527911)
Chemical compound, drugBrefeldin-A; BFASigma-AldrichCatalog no. B5936Ready-made solution of 10 mg/ml in DMSO-based solution, used at final concentration of 5 μg/ml
Chemical compound, drugD/D solubilizerClontech, TaKaRaCatalog no. 635054Obtained as solution, used at final concentration of 0.5 µM in growth medium
Chemical compound, drugLatrunculin B; LatBSigma-AldrichCatalog no. 428020Stock prepared in DMSO, used at final concentration of 2 μM
Chemical compound, drugAlexa Fluor 546-phalloidinThermo Fisher ScientificCatalog no. A22283Diluted 1:1000 in PBS + 1% BSA
Chemical compound, drugMethyl-β-cyclodextrin; MbCDSigma-AldrichCatalog no. C4555Dissolved in DMEM and sterile-filtered before use, used at final concentration of 5 mM
Chemical compound, drugFilipinSigma-AldrichCatalog no. SAE0088Ready-made filipin complex solution of 5 mg/ml in DMSO-based solution, used at final concentration of 125 μg/ml in PBS
Chemical compound, drugPhorbol-12-myristat-13-acetat; PMATocris, Bio-TechneCatalog no. 1201/1Stock prepared in DMSO, used at final concentration of 1 μM
Chemical compound, drugGö6983AbcamCatalog no. ab144414Stock prepared in DMSO, used at final concentration of 1 μM
Chemical compound,
drug
CRT 0066101Tocris, Bio-TechneCatalog no. 4975/10Stock prepared in H2O, used at final concentration of 5 μM
Chemical compound, drugDioctanoylglycol; DOGTocris, Bio-TechneCatalog no. 0484Stock prepared in DMSO, used at final concentration of 100 μM
Software, algorithmFijiSchindelin et al., 2012 (PMID: 22743772)
Software, algorithmPixFRET pluginFeige et al., 2005 (PMID: 16208719)
Software, algorithmµManagerEdelstein et al., 2014 (PMID: 25606571)
Software, algorithmPySerial libraryhttps://pythonhosted.org/pyserial/index.html

Expression constructs

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Expression plasmids for human LRRC8A-GFP, LRRC8AN66A/N83A-GFP and LRRC8E-RFP (Voss et al., 2014) were kindly provided by T.J. Jentsch. To generate FRET constructs, the coding sequences were subcloned with EcoRI and KpnI into pECFP-N1 and pEYFP-N1, resulting in 12-amino acid linkers LVPRARDPPVAT for LRRC8A or WVPRARDPPVAT for LRRC8E. For electrophysiological experiments, CFP and YFP were replaced with Cerulean and Venus by adding AgeI and NotI sites and insertion into the respective CFP- or YFP-tagged versions without altering the linker region. Cerulean and Venus are also referred to as CFP and YFP throughout. For expression of CD4-YFP, human CD4 was subcloned from CD4-GFP (Leisle et al., 2011) into pEYFP-N3. For the generation of A-CFP-FM2, two FM domains (Rollins et al., 2000) were inserted into XbaI-SpeI restriction sites 3’ of A-CFP that were generated using the Q5 sited directed mutagenesis kit (New England Biolabs) with forward primer 5´ATCACTAGTAGCGGCCGCGACTCTAGA and reverse primer 5´ATCTCTAGACTTGTACAGCTCGTCCATGCC. The FRET-based RD sensor for ionic strength (Liu et al., 2017) was kindly provided by B. Poolman and A.J. Boersma, CFP-18AA-YFP (Elder et al., 2009) by C.F. Kaminski. The glutamate receptor construct GluA2-6Y-10C has been described previously (Zachariassen et al., 2016). For expression of GalNAcT2-RFP, the stalk region of N-acetylgalactosaminyltransferase 2 (GalNAcT2) was subcloned from pGalNAcT2-GFP (Le Bot et al., 1998) into pmRFP-N1. For expression of ER-localized YFP (ER-YFP), we used the plasmid pEYFP-ER (Clontech).

Cell lines

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HeLa (RRID: CVCL_0030) and HEK293 (RRID: CVCL_0045) cells were obtained from Leibniz Forschungsinstitut DSMZ and regularly tested for mycoplasma contamination. LRRC8-/- HEK293 (HEK293 KO) cells deficient in all five LRRC8 subunits (Lutter et al., 2017) were kindly provided by T.J. Jentsch. Cells were grown in DMEM (Pan-Biotech) supplemented with 10% fetal calf serum at 37°C in 5% CO2. For imaging experiments without simultaneous electrophysiology, cells were plated in 35 mm glass bottom dishes (MatTek), coated with poly-L-lysine 0.01% solution (Sigma-Aldrich) for HEK293 cells. For electrophysiology, cells were plated on poly-L-lysine-coated 25 mm coverslips. Cells were transfected with FuGENE 6 (Promega) according to the supplier’s manual. For co-expression, constructs were co-transfected at equimolar ratios.

Drug treatment

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Brefeldin-A (BFA, Sigma-Aldrich, 10 mg/ml in DMSO) was added at 5 µg/ml to culture medium during transfection. To depolymerize the actin cytoskeleton, 2 µM latrunculin B (Sigma-Aldrich, dissolved in DMSO) was added to the growth medium for 1 hr in normal growth conditions. For cholesterol depletion, 5 mM methyl-β-cyclodextrin (MbCD, Sigma-Aldrich), dissolved in DMEM and stirred for 30 min at RT before sterile filtration, was applied for 1 hr in normal growth conditions. 1 µM phorbol-12-myristat-13-acetat (PMA, Bio-Techne, dissolved in DMSO), 1 µM Gö6983 (Abcam, DMSO), 5 µM CRT 0066101 (Bio-Techne, H2O) and 100 µM dioctanoylglycol (DOG, Bio-Techne, DMSO) were added during or before measurements as indicated.

Imaging of intracellular localization, actin cytoskeleton and cholesterol staining

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Images of the intracellular localization of LRRC8A-GFP/LRRC8E-RFP and of LRRC8A-CFP-FM2 with the organelle markers ER-YFP, GalNAc-T2-RFP and CD4-YFP were acquired with a 64x/1.4 oil objective at a Leica Dmi8 microscope equipped with a Hamamatsu OcraFlash4.0 controlled by LAS X. Images of the actin cytoskeleton and cholesterol content were acquired with a 20x/0.7 dry objective at the same microscope. For cholesterol staining, cells were washed with PBS, fixed with 4% paraformaldehyde (PFA) for 15 min at room temperature, washed with PBS and incubated with DMEM +10% FCS for 10 min at 37°C, and, after washing with PBS, stained with 125 µg/ml filipin (Sigma-Aldrich, dissolved in DMSO and diluted in PBS) for 45 min at room temperature. After washing with PBS, images were acquired with a DAPI filter set (EX: 360/40, DC: 400, EM: 425 lp). To stain the actin cytoskeleton, PFA-fixed cells (as above) were permeabilized with 0.1% Triton-X100 in PBS + 1% BSA for 5 min at room temperature and incubated with Alexa Fluor 546-phalloidin (Thermo Fisher Scientific) diluted 1:1000 in PBS + 1% BSA at room temperature for 1 hr, washed with PBS, air dried and mounted on glass cover slides for imaging with a rhodamine filter set (EX:540/23, DC:580,EM:590 lp). Cells co-expressing LRRC8A-CFP-FM2 with organelle markers were fixed and mounted as described above 24 hr after transfection and incubation with D/D solubilizer for the indicated times. Images where acquired with filter sets for CFP (EX:436/20, DC:455 EM:480/40), YFP (EX:500/20, DC:515, EM:535/30) and rhodamine (as above).

Acceptor photo-bleaching, sensitized-emission and ratiometric FRET assays

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FRET experiments were performed on a high-speed setup of Leica Microsystems (Dmi6000B stage, 63x/1.4 objective, high speed external Leica filter wheels with Leica FRET set filters (11522073), EL6000 light source, DFC360 FX camera, controlled by Las AF). All experiments were executed at room temperature. Isotonic imaging buffer (340 mOsm) contained (in mM): 150 NaCl, 6 KCl, 1 MgCl2, 1.5 CaCl2, 10 glucose,10 HEPES, pH 7.4. NaCl was adjusted for osmolarity titration in hypotonicity; standard hypotonic buffer (250 mOsm) contained 105 mM NaCl. Hypertonic buffers were as isotonic buffer supplemented with 60 mM (for 400 mOsm) or 160 mM (for 500 mOsm) mannitol.

Acceptor bleaching experiments were conducted on field-of views (FOVs) with at least two cells expressing both fluorescent proteins. An image in both CFP and YFP channels was taken per time point. To bleach only part of the FOV, the field diaphragm was closed until control cells were no longer illuminated. Cells were bleached with 515–560 nm light at full intensity (Leica filter cube N2.1) for 2.5–5 min per time point until YFP signals reached below 10% of initial intensity. Intensities of CFP and YFP for respective time points were measured in Fiji in background-subtracted images and the percentage change of CFP intensity calculated for bleached and control cells.

Sensitized-emission FRET (seFRET) images were recorded with the same settings for donor, acceptor and FRET channels (8 × 8 binning, 100 ms exposure, gain 1) every 10–20 s. Images were processed with Fiji (Schindelin et al., 2012). cFRET maps were calculated with PixFRET plugin (Feige et al., 2005) (threshold set to 1, Gaussian blur to 2) with a self-written macro to process movies according to following equation (Jiang and Sorkin, 2002)

cFRET=IDA-IDD*β-IAA*γIAA

with emission intensities of IDA (FRET channel), IDD (donor channel), IAA (acceptor channel). Correction factors (β = bleed through of donor emission; γ = cross excitation of acceptor by donor excitation) were calculated from acceptor- and donor-only samples that were measured on three different days with at least three FOVs per day. cFRET maps were measured by manually-drawn regions of interest (ROIs) over whole cells, excluding intracellular aggregates that were oversaturated in any of the three channels. Because of the variability in absolute FRET values between individual cells, cFRET values of individual cells were normalized to their mean cFRET in isotonic buffer.

For monitoring ionic strength in living cells, we used the ratiometric FRET-based RD sensor (Liu et al., 2017). Sequential images of the CFP and FRET channels were acquired using the same filters as for seFRET experiments every 10 s. Images were acquired with 2 × 2 binning, 500 ms exposure, gain 3.0 and intensity four for hypotonicity experiments and with 8 × 8 binning, 127 ms exposure, gain 3.0 and intensity two for DOG experiments. Ratio maps of I_FRET/I_Cerulean were created from background-subtracted images in Fiji and mean ratio values were taken from hand-drawn ROIs of individual cells. To account for ionic strength differences between individual cells, ratios were normalized to baseline values in isotonic buffer.

Electrophysiology and simultaneous seFRET measurements

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For whole-cell patch-clamp recordings, the isotonic bath solution was composed as follows (in mM): 150 NaCl, 6 KCl, 1 MgCl2, 1.5 CaCl2, 10 glucose, and 10 HEPES, pH 7.4 with NaOH (320 mOsm). We used an intracellular (pipette) solution with the following composition (in mM): 40 CsCl, 100 Cs-methanesulfonate, 1 MgCl2, 1.9 CaCl2, 5 EGTA, 4 Na2ATP, and 10 HEPES, with pH adjusted to 7.2 with CsOH (290 mOsm). For VRAC activation, cells were perfused with drugs and/or a hypotonic buffer containing (in mM) 105 NaCl, 6 CsCl, 1 MgCl2, 1.5 CaCl2, 10 glucose, 10 HEPES, pH 7.4 with NaOH (240 mOsm). The osmolarities of all solutions were measured with a vapor pressure osmometer (VAPRO 5600, Elitech). Pipettes had a resistance of 3–5 MΩ when filled with intracellular solution, and we used ISO-type pipette holders (G23 Instruments) to minimize pipette drift. For cells overexpressing exogenous, fluorescently-labelled VRAC constructs, the success rate for obtaining a gigaseal and whole-cell configuration were <20%, compared to >50% for wild-type untransfected HEK293 cells. Whole-cell recordings from cells overexpressing VRAC were also less stable. For normalizing current to cell capacitance, we used the capacitance from the manual compensation of transients from the Axopatch 200B amplifier (Molecular Devices). After whole-cell configuration was obtained, cells were held at –30 mV and the currents were recorded using Axograph X (Axograph Scientific) via an Instrutech ITC-18 D-A interface (HEKA Elektronik). The standard protocol for VRAC current recordings consisted of 9 episodes of a 0.5 s step to –80 mV every 12 s, followed by a 3 s voltage step protocol (300 ms steps to –40, 0, 40, 80, 100 mV in turn). This protocol was repeated throughout the recording. The voltage-step protocol confirmed VRAC-typical properties of outward rectification and depolarization-dependent inactivation for LRRC8A/E-containing VRAC (König and Stauber, 2019; Ullrich et al., 2016; Voss et al., 2014). Excitation by 445 and 514 nm diode lasers (iChrome MLE, Toptica Photonics) for CFP and YFP was directed through a manual total internal reflection fluorescence (TIRF) input to an Olympus IX81 microscope. We used a 40 × Olympus objective (NA 0.6) for all recordings. For simultaneous ratiometric FRET recordings, filters specific to donor and acceptor excitation and emission wavelengths were alternated using the fluorescence turret. We controlled the microscope turret control with an in-house script written in PYTHON. Briefly, the turret was synchronized to the electrophysiology acquisition computer using voltage output from the digitizer via an Arduino microcontroller to a reverse-engineered serial connection (using the PySerial library; https://pythonhosted.org/pyserial/index.html) from a second computer to the IX81 microscope. Fluorescence intensities in response to 445 and 514 nm excitation were recorded sequentially with 100 ms exposure time and 2 × 2 binning on a Prime 95B CMOS camera (Photometrics) fitted to Cairn Optosplit-II. CFP and YFP emission were split with a T495lpxr dichroic mirror, and recorded on the same frame after passing through ET520/40 m (YFP) and ET470/24 m (CFP) filters (all mounted within the Optosplit, all Chroma). Exposures were timed to precede the –80 mV voltage steps. Laser emission and camera exposure were triggered in hardware directly from the digitizer. Images were recorded with Micromanager (Edelstein et al., 2014) and analyzed with FIJI as described above.

Reverse aggregation system

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A-CFP-FM2- or A-CFP-FM2/E-YFP-expressing cells were measured 24 hr post-transfection. Aggregates were released by adding 0.5 µM D/D solubilizer (TaKaRa) to the growth medium and incubation for 10, 80 or 165 min at 37°C in 5% CO2 before seFRET measurements. All buffers used during the measurement contained 0.5 µM D/D solubilizer. Mean cFRET was acquired from manually-drawn ROIs over the whole cell for ER- and plasma membrane-localized VRAC. For plasma membrane, only cells with clear plasma membrane localization were selected by visual inspection. To determine the cFRET of Golgi-localized VRAC, masks were created on CFP images by applying a threshold over the high-intensity, juxta-nuclear regions and applying these masks on the respective cFRET maps.

Statistical analysis

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Data are represented as mean of individual cells ± standard deviation (s.d.) or mean of n (number of independent experiments) ± standard error of the mean (s.e.m.) as indicated. P values were determined by two-tailed Student’s t-test and are indicated in all figures according to convention: n.s. = not significant, *p≤0.05, **p≤0.005 and ***p≤0.0005.

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    6. T Stauber
    (2019)
    Biological Chemistry, 10.1515/hsz-2019-0189.
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Decision letter

  1. Leon D Islas
    Reviewing Editor; Universidad Nacional Autónoma de México, Mexico
  2. Richard Aldrich
    Senior Editor; The University of Texas at Austin, United States
  3. Leon D Islas
    Reviewer; Universidad Nacional Autónoma de México, Mexico

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: the authors were asked to provide a plan for revisions before the editors issued a final decision. What follows is the editors’ letter requesting such plan.]

Thank you for sending your article entitled "FRET changes by C-terminal movement reveal VRAC activation by plasma membrane PKC signaling rather than ionic strength" for peer review at eLife. Your article is being evaluated by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation is being overseen by a Reviewing Editor and Richard Aldrich as the Senior Editor.

Given the list of essential revisions, including new experiments, the editors and reviewers invite you to respond within the next two weeks with an action plan and timetable for the completion of the additional work. We plan to share your responses with the reviewers and then issue a binding recommendation.

The three reviewers agree that the main concern in this paper is that there isn't a validation by electrophysiology of the FRET data. It is essential that you perform simultaneous experiments measuring FRET and current in response to the activating stimuli. If this is not within your technical possibilities, at least the same kind of experiments to measure FRET should be repeated by electrophysiology. It is also essential that the FRET and imaging data is presented in a clearer manner and that the PKC FRET experiments be also validate by electrophysiology. Please see the reviewers report below for a more detailed list of comments.

Reviewer #1:

This paper by the Stauber group presents results from experiments in which they have measured a FRET signal between an attached fluorescent protein pair in the LRRC8A or E proteins which form the volume-regulated VRAC channels. The measurements show that the FRET signal is modulated by application of hypotonic solutions. The authors assert that the signal is related to activation of the VRAC channel and present further experimental evidence that activation is also mediated by activation of PKC phosphorylation. The experimental methods are adequate and in general the conclusions are supported by the evidence.

There are a few points that need to be presented in a clearer manner. In particular, while there is evidence that the FRET signal is specific of LRRC8, the authors cite a couple of reviews as evidence that the FRET signal correlates with the time course of activation of VRAC by the same stimulus. However, the evidence in those references shows activation of VRAC by voltage clamp stimuli occurs in an ms time scale and no data is shown on the time course of responses to a hypertonic stimulus. The authors need to show a parallel experiment showing the magnitude of the VRAC current produced by LRRC8A/A and LRRC8A/E in their particular expression system.

Voltage-clamp experiments show that VRAC currents inactivate in response to sustained applied stimulus. It is important to discuss the sustained nature of the FRET signal if it is related to channel activation. Is it thought that there are distinct gates for each modality of channel activation?

Reviewer #2:

For editors

The authors developed a highly interesting assay to monitor activity of LRRC8 mediated VRAC channels using a purely optical FRET based assay. Such an assay could be useful for structure-function studies, as well as for studies of VRAC function in physiological settings. However, a major criticism is that the assay has not been validated by parallel electrophysiological measurements. In case the authors can provide such validation I would be in favor of publication.

For authors

The manuscript by König et al., reports a method to monitor VRAC/LRRC8 volume sensitive channel activity in a time- and space-resolved manner by measuring intra-complex FRET between C-terminally fused fluorescent proteins. Two major aims are pursued: first, to demonstrate the flexibility of the leucine-rich-repeat domains (LRRDs) and their involvement in the gating of the channel; second, to provide evidence that low intracellular ionic strength is not the physiologically relevant parameter for the opening of VRACs. Rather, the authors propose that activation under hypotonic stimulation is coupled to PKC activity.

In addition, the authors use the assay to show that LRRC8 mediated VRAC is not activatable in endomembranes, but only in the plasma membrane.

The crucial questions driving the work are laudable and potentially very important for the field. In particular, the method of introducing an intramolecular FRET sensor of subunit distance is highly innovative and potentially very useful. The possibility to monitor LRRC8 activity with a purely optical assay is very intriguing.

However, several technical controls are necessary to fully validate the assay. Most importantly, it is mandatory that the assay itself, and other results, are corroborated by electrophysiological data.

All the experiments should be complemented with electrophysiological recordings. In principle, it would be most convincing if the FRET assay could be performed on the same cell that is patch-clamped to verify that FRET changes reflect current activation. Since this maybe technically too demanding, it is at least necessary to validate the constructs used by electrophysiological recordings, under the same conditions as the FRET assay. Attaching fluorescent proteins to the C-terminus might alter the "volume-sensitivity" by itself. In fact, the addition of fluorescent proteins to the C-terminus resulted in constitutive anion channel activity in Xenopus oocytes (Gaitán-Peñas et al., 2016).

The authors propose a central role for PKC in the gating of the channel and indicate that its pharmacological inhibition prevented hypotonicity-driven VRAC opening. Again, this result should be verified by electrophysiological recordings employing the same constructs, and constructs without tag – a quantitative comparison of the current density with and without application of PMA / Gö6983 / staurosporine is necessary.

Reviewer #3:

Volume-regulated anion channels (VRACs) play a central role in the modulation of cell volume in animals, by enabling the flow of chloride ions and larger osmolytes out of the cell upon activation by hypotonic conditions, and therefore counteracting cell-swelling. These channels are heterohexamers composed of the obligatory subunit LRRC8A and additional LRRC8-B-E subunits. The mechanism of activation of VRAC in not understood, but it has been proposed to be related to conformational changes at its C-terminus, and has been shown to be triggered by reduced ionic strength in a reconstituted system. In the present manuscript, Benjamin Koenig and collaborators construct a FRET-based VRAC-activity sensor by fusing the C-termini of different subunits with either of two fluorescence proteins capable of FRET. The authors show changes in FRET in response to hypotonic media that are reversible upon return to isotonic conditions, with kinetics and pharmacology that is consistent with the signal originating from VRAC activation. They also show that hypotonicity-driven FRET changes occur only for channels in the plasma membrane, whereas those located at intracellular compartments (ER and golgi) don't exhibit changes in FRET. They demonstrate that the requirement for activity of localization in the plasma membrane does not involve differences in cholesterol concentration or the integrity of the actin cytoskeleton. In contrast, the authors find that stimulation of PKC activity leads to prominent FRET changes similar to those produced by hypertonic media, whereas inhibition of PKC prevents FRET changes in response to hypotonic media. The authors have developed a set of clever tools to follow VRAC activity without requiring electrophysiological measurements, and provide relevant information on the cell-physiological conditions that enable VRAC activity and on the role of the C-terminus in channel activation. However, I think the manuscript would be much stronger if the authors provided more direct evidence that the FRET changes are indeed connected with channel activation. In addition, they should address some apparent inconsistencies within their observations and also relative to published data, and provide additional information on some experiments.

I have the following specific concerns:

1) All conclusions in the manuscript depend on the assumption that the observed FRET changes are reflecting channel activation. However, the evidence supporting this is not direct, but rather depends on correlative observations of the effects of non-specific treatments that could also affect other processes in the cell. It is possible that the FRET changes reflect a conformational change that is independent or at least partially independent of activation. This issue is particularly important given that the role of the C-terminus for activation has not been well established. I think the manuscript would be significantly stronger if the authors could provide more direct evidence for this, by performing electrophysiological measurements in combination with FRET measurements, or by using additional specific VRAC inhibitors or performing experiments with loss-of-function VRAC subunits.

2) One of the main observations of the paper is that localization in the plasma membrane is required for the observed FRET changes and channel activation. However, there is a significant amount of intracellularly located LRRC8 subunits even without treatment with BFA (see LRRC8E-RFP in Figure 2C, for example), and the hypotonicity-dependent FRET signals shown in Figure 1 appear to originate from the entire cell (see Figure 1D). The authors should address this apparent contradiction, and provide detailed information on the way they performed their analysis to measure signals mostly originating from the plasma membrane. The observation that LRRC8E appears to be more heavily retained in the ER that LRRC8A (see Figure 2C) seems to counter the observation that the 8A/8E ratio does not correlate with the observed changes in FRET, except if the majority of the measured FRET signal originates from the plasma membrane. The authors should discuss these issues.

3) The authors should discuss the apparent contradiction of their data with the observations in a reconstituted system where decreased ionic strength is sufficient to activate VRAC channels.

4) From the images in Figure 3B, it is impossible for me to determine which channels are localized at the ER vs golgi, even though the authors used a marker for golgi expression. The authors should use ER, golgi and plasma membrane markers, and provide a semi-quantitative analysis to demonstrate the location of the different channel populations, and provide a more detailed description of how they selected their ROIs for performing group analysis of channels in each of the cellular compartments.

5) In Figure 1—figure supplement 1, the authors should show the net FRET rather than the normalized values, to allow readers to see how much variability there was in FRET between the controls and the VRACs.

6) The authors should show whether the treatments with LatB or MbCD affected net FRET levels, rather than just the normalized values.

7) The Abstract, Title and the overall text seem to imply that PKC activation is connected with the physiological activation of VRACs upon exposure to hypotonic solutions. However, the presented data does not really establish a link between VRAC stimulation by PKC and changes in extracellular osmolarity. It might very well be that PKC-stimulated VRACs are indeed activated by ionic strength, and that this sensitivity is obscured by PKC inhibition. The authors should provide a more lengthy discussion on how PKC might be affecting channel activity, even if based only on speculation.

https://doi.org/10.7554/eLife.45421.023

Author response

[Editors' note: the authors’ plan for revisions was approved and the authors made a formal revised submission.]

Given the list of essential revisions, including new experiments, the editors and reviewers invite you to respond within the next two weeks with an action plan and timetable for the completion of the additional work. We plan to share your responses with the reviewers and then issue a binding recommendation.

The three reviewers agree that the main concern in this paper is that there isn't a validation by electrophysiology of the FRET data. It is essential that you perform simultaneous experiments measuring FRET and current in response to the activating stimuli. If this is not within your technical possibilities, at least the same kind of experiments to measure FRET should be repeated by electrophysiology. It is also essential that the FRET and imaging data is presented in a clearer manner and that the PKC FRET experiments be also validate by electrophysiology. Please see the reviewers report below for a more detailed list of comments.

As we describe in the point-by-point response below, we have aimed to address all these points. We have conducted patch-clamp fluorometry to measure FRET and currents simultaneously, demonstrating the relation of the change in FRET signal with VRAC activation. We improved the presentation of our data and experimental methods. Furthermore, we have performed electrophysiological experiments and simultaneous FRET/current measurements to investigate the activation mechanism of VRAC. By doing so, we have confirmed the role of diacylglycerol. Instead of PKC, we were able to identify a role for PKD in VRAC activation by using the combined power of electrophysiology and our FRET sensor. In addition, we have surprisingly found whole-cell configuration-related artifacts that have would have remained uncovered without our optical tool.

Reviewer #1:

This paper by the Stauber group presents results from experiments in which they have measured a FRET signal between an attached fluorescent protein pair in the LRRC8A or E proteins which form the volume-regulated VRAC channels. The measurements show that the FRET signal is modulated by application of hypotonic solutions. The authors assert that the signal is related to activation of the VRAC channel and present further experimental evidence that activation is also mediated by activation of PKC phosphorylation. The experimental methods are adequate and in general the conclusions are supported by the evidence.

We thank the reviewer for these positive comments.

There are a few points that need to be presented in a clearer manner. In particular, while there is evidence that the FRET signal is specific of LRRC8, the authors cite a couple of reviews as evidence that the FRET signal correlates with the time course of activation of VRAC by the same stimulus. However, the evidence in those references shows activation of VRAC by voltage clamp stimuli occurs in an ms time scale and no data is shown on the time course of responses to a hypertonic stimulus. The authors need to show a parallel experiment showing the magnitude of the VRAC current produced by LRRC8A/A and LRRC8A/E in their particular expression system.

Following the reviewers’ suggestions, we have now performed patch-clamp fluorometry to simultaneously measure FRET and currents of LRRC8A-CFP/E-YFP in HEK293 cells deficient in endogenous VRAC. These measurements demonstrated that the time course of FRET changes correlated with the current activation and inactivation when applying hypotonic buffer and consecutively isotonic buffer (new Figure 1F). This replaces the citation of the reviews at this position. So far, we were not able to measure currents from LRRC8A/A homomeric channels – perhaps due to the extremely low conductance (Gaitán-Peñas et al., 2016; Deneka et al., 2018).

Voltage-clamp experiments show that VRAC currents inactivate in response to sustained applied stimulus. It is important to discuss the sustained nature of the FRET signal if it is related to channel activation. Is it thought that there are distinct gates for each modality of channel activation?

As also seen in our simultaneous FRET/current measurements, the FRET signal (reduced FRET, correlating to active VRAC) follows the currents (which we measure at -80mV). Sustained VRAC activity in hypotonicity is not unusual both in whole-cell voltage-clamp mode (e.g. reviewed by Pederson et al., 2016, and shown in their Figure 1A) and cells not subjected to whole-cell patch clamp (as seen in numerous experiments investigating VRAC-mediated osmolyte efflux). Time-dependent inactivation by non-physiological depolarized voltages is typical for VRAC currents (with the inactivation kinetics and voltage-dependence varying with LRRC8 subunit composition – Voss et al., 2014, Ullrich et al., 2016). We observed this characteristic inactivation in our voltage-step protocol measurements that we routinely conducted to confirm that the currents we measured were mediated by VRAC (new Figure 1F and new Figure 4C). There are likely distinct gates for activation/inactivation by osmolarity and depolarization-induced inactivation, but a discussion on this is beyond the scope of this study.

Reviewer #2:

For editors

The authors developed a highly interesting assay to monitor activity of LRRC8 mediated VRAC channels using a purely optical FRET based assay. Such an assay could be useful for structure-function studies, as well as for studies of VRAC function in physiological settings. However, a major criticism is that the assay has not been validated by parallel electrophysiological measurements. In case the authors can provide such validation I would be in favor of publication.

We thank the reviewer for the generally positive comments. Following the reviewers’ suggestions, we have now performed a series of electrophysiological measurements, also in parallel to our FRET assay.

For authors

The manuscript by König et al., reports a method to monitor VRAC/LRRC8 volume sensitive channel activity in a time- and space-resolved manner by measuring intra-complex FRET between C-terminally fused fluorescent proteins. Two major aims are pursued: first, to demonstrate the flexibility of the leucine-rich-repeat domains (LRRDs) and their involvement in the gating of the channel; second, to provide evidence that low intracellular ionic strength is not the physiologically relevant parameter for the opening of VRACs. Rather, the authors propose that activation under hypotonic stimulation is coupled to PKC activity.

In addition, the authors use the assay to show that LRRC8 mediated VRAC is not activatable in endomembranes, but only in the plasma membrane.

The crucial questions driving the work are laudable and potentially very important for the field. In particular, the method of introducing an intramolecular FRET sensor of subunit distance is highly innovative and potentially very useful. The possibility to monitor LRRC8 activity with a purely optical assay is very intriguing.

We thank the reviewer for these positive comments. Indeed, we now see a major advantage of a purely optical assay – we do not dialyze the cell and lose signaling pathways.

However, several technical controls are necessary to fully validate the assay. Most importantly, it is mandatory that the assay itself, and other results, are corroborated by electrophysiological data.

All the experiments should be complemented with electrophysiological recordings. In principle, it would be most convincing if the FRET assay could be performed on the same cell that is patch-clamped to verify that FRET changes reflect current activation. Since this maybe technically too demanding, it is at least necessary to validate the constructs used by electrophysiological recordings, under the same conditions as the FRET assay. Attaching fluorescent proteins to the C-terminus might alter the "volume-sensitivity" by itself. In fact, the addition of fluorescent proteins to the C-terminus resulted in constitutive anion channel activity in Xenopus oocytes (Gaitán-Peñas et al., 2016).

As suggested by the reviewers, we have now performed patch-clamp fluorometry and measured FRET and current in the same cells. We found that (as previously described by Gaitán-Peñas et al., 2016) the tagged constructs yielded residual currents in isotonicity. Nonetheless, the currents were clearly potentiated by hypotonicity (as shown with measurements in oocytes by Gaitán-Peñas et al., 2016). This rendered the measurements indeed technically demanding (with a gigaseal rate below 20% overexpressing cells versus more than 50% success rate in wild-type cells), as also stated by Gaitán-Peñas et al., 2016: “However, we found this to be extremely difficult because of a very low success rate of giga-seal formation. In the few successful recordings, constitutive VRAC-like channel activity could be observed (data not shown).” Nonetheless, by using patch-clamp fluorometry we could convincingly confirm the relationship of the observed FRET changes with the activation of VRAC.

The authors propose a central role for PKC in the gating of the channel and indicate that its pharmacological inhibition prevented hypotonicity-driven VRAC opening. Again, this result should be verified by electrophysiological recordings employing the same constructs, and constructs without tag – a quantitative comparison of the current density with and without application of PMA / Gö6983 / staurosporine is necessary.

We first confirmed that PMA also induced a reduction in cFRET in HEK293 in addition to HeLa cells (new right panel in Figure 4A) and that cFRET changes were specific for VRAC, since an unrelated FRET sensor (CFP-18aa-YFP) did not respond to PMA (new Figure 4—figure supplement 1A). Subsequently, we have performed electrophysiological experiments on cells overexpressing our fluorescently-tagged LRRC8 constructs and on wild-type cells with endogenous VRAC to investigate the role of DAG (and PMA)-activated kinases.

Remarkably, and at first in apparent contradiction to our results from the FRET measurements, we found that PMA did not activate VRAC in clamped cells (new Figure 4B). Neither the current, nor –surprisingly- the cFRET were affected by PMA application. However, when we incubated the cells with PMA before establishing the whole-cell configuration, we detected activation of fluorescently-tagged VRAC and of endogenous VRAC, as shown in new Figure 4C. Therefore, we could validate our cFRET data by electrophysiological methods and could show for the first time that whole-cell patching of cells affects signaling in the VRAC activation pathway, possibly by cell dialyzes. Electrophysiological measurements further verified the inhibiting effect of DOG (inhibitor of the DAG kinase) on VRAC inactivation after hypotonicity for both fluorescently tagged LRRC8s and for endogenous VRAC (new Figure 5C), corroborating the role of DAG signaling in VRAC activation.

Our attempts to confirm the effect of Gö6983 on cFRET by electrophysiological measurements led to paradoxical results. In cells held in the whole-cell patch-clamp configuration, we observed a tremendous increase in cFRET by a factor of four, which we simply cannot explain. In parallel, Gö6983 did not inhibit VRAC currents, in fact falsifying the proposed role of PKC in VRAC activation. Indeed, we would have missed this point without performing the electrophysiological control experiments suggested by the three reviewers. We have changed all respective statements in the manuscript accordingly. Instead, application of Gö6983 reduced the inactivation of currents when we changed buffers from hypotonic to isotonic. We show these data in new Figure 4—figure supplement 2, but avoid drawing strong conclusions.

Having ruled out PKC as a core mediator of VRAC activation by diacylglycerol (or PMA), we tested PKD, which can also be recruited to the membrane and activated by DAG (and PMA). In contrast to Gö6983, the PKD inhibitor CRT 0066101 reduced hypotonicity-induced VRAC currents both during clamping and when applied before membrane breakthrough (new Figure 5A,B).

Reviewer #3:

Volume-regulated anion channels (VRACs) play a central role in the modulation of cell volume in animals, by enabling the flow of chloride ions and larger osmolytes out of the cell upon activation by hypotonic conditions, and therefore counteracting cell-swelling. These channels are heterohexamers composed of the obligatory subunit LRRC8A and additional LRRC8-B-E subunits. The mechanism of activation of VRAC in not understood, but it has been proposed to be related to conformational changes at its C-terminus, and has been shown to be triggered by reduced ionic strength in a reconstituted system. In the present manuscript, Benjamin Koenig and collaborators construct a FRET-based VRAC-activity sensor by fusing the C-termini of different subunits with either of two fluorescence proteins capable of FRET. The authors show changes in FRET in response to hypotonic media that are reversible upon return to isotonic conditions, with kinetics and pharmacology that is consistent with the signal originating from VRAC activation. They also show that hypotonicity-driven FRET changes occur only for channels in the plasma membrane, whereas those located at intracellular compartments (ER and golgi) don't exhibit changes in FRET. They demonstrate that the requirement for activity of localization in the plasma membrane does not involve differences in cholesterol concentration or the integrity of the actin cytoskeleton. In contrast, the authors find that stimulation of PKC activity leads to prominent FRET changes similar to those produced by hypertonic media, whereas inhibition of PKC prevents FRET changes in response to hypotonic media. The authors have developed a set of clever tools to follow VRAC activity without requiring electrophysiological measurements, and provide relevant information on the cell-physiological conditions that enable VRAC activity and on the role of the C-terminus in channel activation. However, I think the manuscript would be much stronger if the authors provided more direct evidence that the FRET changes are indeed connected with channel activation. In addition, they should address some apparent inconsistencies within their observations and also relative to published data, and provide additional information on some experiments.

We thank the reviewer for the generally positive comments. We have addressed the reviewer’s concerns as described below.

I have the following specific concerns:

1) All conclusions in the manuscript depend on the assumption that the observed FRET changes are reflecting channel activation. However, the evidence supporting this is not direct, but rather depends on correlative observations of the effects of non-specific treatments that could also affect other processes in the cell. It is possible that the FRET changes reflect a conformational change that is independent or at least partially independent of activation. This issue is particularly important given that the role of the C-terminus for activation has not been well established. I think the manuscript would be significantly stronger if the authors could provide more direct evidence for this, by performing electrophysiological measurements in combination with FRET measurements, or by using additional specific VRAC inhibitors or performing experiments with loss-of-function VRAC subunits.

As suggested by the reviewers, we have now performed electrophysiological measurements simultaneously with cFRET measurements. The new results demonstrate a clear correlation of cFRET changes with VRAC currents. This shows that the FRET sensor is a useful tool to study VRAC activity and that the C-termini move during VRAC activation. We do not claim a strict role of the C-terminus in VRAC activation.

2) One of the main observations of the paper is that localization in the plasma membrane is required for the observed FRET changes and channel activation. However, there is a significant amount of intracellularly located LRRC8 subunits even without treatment with BFA (see LRRC8E-RFP in Figure 2C, for example), and the hypotonicity-dependent FRET signals shown in Figure 1 appear to originate from the entire cell (see Figure 1D). The authors should address this apparent contradiction, and provide detailed information on the way they performed their analysis to measure signals mostly originating from the plasma membrane. The observation that LRRC8E appears to be more heavily retained in the ER that LRRC8A (see Figure 2C) seems to counter the observation that the 8A/8E ratio does not correlate with the observed changes in FRET, except if the majority of the measured FRET signal originates from the plasma membrane. The authors should discuss these issues.

We thank the reviewer for bringing up this issue. Indeed, a portion of either LRRC8 subunit is frequently stuck in the secretory pathway, as we observed previously for exogenously expressed LRRC8 combinations (Voss et al., 2014). The amount of signal that is not from the plasma membrane showed a high variability, with some cells showing much LRRC8A and some cells LRRC8E intracellularly. Usually, there is a tendency towards LRRC8E being more stuck in the endomembrane system, but importantly, we did not observe a correlation with the expression levels. As VRACs can adopt varying subunit stoichiometries, this suggests that the amount of correctly paired (and trafficked) subunits is unaffected by the expression levels as well. Consistently, we did not observe a correlation between absolute acceptor-normalized cFRET values and LRRC8E expression levels (not shown). Indeed, the YFP signal originating from a part of the cell with little CFP (LRRC8A) will not contribute to the real FRET signal of correctly formed VRAC channels (see lack of co-localization in Figure 2C), but it would indeed lower the acceptor-normalized cFRET (see equation in Material and methods section). However, as we did not observe a correlation between expression levels and intracellular localization, there is no correlation between the expression ratio of 8A/8E and the decrease in cFRET (Figure 1—figure supplement 2B). We agree with the reviewer that this is an important issue that deserves clarification in the manuscript, and have therefore added a corresponding statement (subsection “Inter-subunit FRET shows C-terminal movement during VRAC activation”).

3) The authors should discuss the apparent contradiction of their data with the observations in a reconstituted system where decreased ionic strength is sufficient to activate VRAC channels.

Indeed, Syeda et al., (2016) observed activation of LRRC8 complexes reconstituted in membrane bilayers by reduced ionic strength. So far, we have no explanation for this apparent contradiction, except for the environment of the VRACs, i.e. artificial membrane bilayer system versus cellular context. In the cellular context, however, there are several studies providing evidence for VRAC activation without alteration of the ionic strength. Furthermore, alterations of ionic strength required to activate VRACs are usually not reached in experiments using the whole-cell configuration to study VRAC. This has recently been reviewed in detail by Strange et al., (2019). Following the reviewer’s suggestion, we have now expanded our discussion on this, also adding further citations of important studies on the regulatory role of ionic strength on VRAC activation (Discussion section).

4) From the images in Figure 3B, it is impossible for me to determine which channels are localized at the ER vs golgi, even though the authors used a marker for golgi expression. The authors should use ER, golgi and plasma membrane markers, and provide a semi-quantitative analysis to demonstrate the location of the different channel populations, and provide a more detailed description of how they selected their ROIs for performing group analysis of channels in each of the cellular compartments.

We have now added images (new Figure 3—figure supplement 1) showing the co-localization of LRRC8A-CFP-FM2 with the ER marker ER-YFP, the Golgi marker GalNAc-RFP, and with CD4-YFP as plasma membrane marker after release from the aggregation block. We provide a semi-quantitative analysis for a representative cell in which we follow the LRRC8A-CFP-FM2 construct through the secretory pathway (Figure 3—video 1; quantifications shown for each time frame), for which we measure the CFP fluorescence intensity in the Golgi area and a ROI for plasma membrane (depicted in the video). We have now added a more detailed description of the ROI selection in subsection “Reverse aggregation system”.

5) In Figure 1—figure supplement 1, the authors should show the net FRET rather than the normalized values, to allow readers to see how much variability there was in FRET between the controls and the VRACs.

Net FRET values with our fluorescently-labelled VRAC subunits varied tremendously, with measured FRET efficiencies ranging between 10% and 80%. This is likely due to the flexible subunit stoichiometry of hexameric VRAC complexes, at least when expressed exogenously LRRC8 (e.g. Gaitán-Peñas et al., 2016). Accordingly, cFRET (i.e. the corrected FRET value normalized to the acceptor level) varied within the cell population, making a normalization of the values necessary. The relative reduction in cFRET during VRAC activation was independent from the net FRET of a given cell, just as it neither correlated with the subunit expression ratio (see point 2 and Figure 1—figure supplement 2B). Therefore, we decided not to show net FRET values, since this would not add valuable information. However, we now state the variability of FRET as the reason for the normalization (subsection “Acceptor photo-bleaching, sensitized-emission and ratiometric FRET assays”). The net FRET values for the FRET control constructs (CFP-18aa-YFP and GluA2-6Y-10C), on the other hand showed little variability between cells, but we also present them as normalized values for clarity.

6) The authors should show whether the treatments with LatB or MbCD affected net FRET levels, rather than just the normalized values.

Net FRET values vary extremely between cells explained (see our answer to point 5). The mean of this highly variable value was not altered by the treatment with LatB or MbCD: we observed mean (non-normalized) cFRET of 52 ± 15% (untreated cells in isotonic buffer), 45 ± 17% (LatB) and 47 ± 12% (MbCD). We opt not to present these data in the manuscript, since we think it will not provide further useful information.

7) The Abstract, Title and the overall text seem to imply that PKC activation is connected with the physiological activation of VRACs upon exposure to hypotonic solutions. However, the presented data does not really establish a link between VRAC stimulation by PKC and changes in extracellular osmolarity. It might very well be that PKC-stimulated VRACs are indeed activated by ionic strength, and that this sensitivity is obscured by PKC inhibition. The authors should provide a more lengthy discussion on how PKC might be affecting channel activity, even if based only on speculation.

This aspect of the manuscript has changed substantially following new patch-clamp measurements and the realization that PKC is not as central as we suspected. Instead, we show a much stronger connection to DAG-PKD signaling, because we can activate, inhibit or sustain VRAC currents in the absence of, or in opposition to, hypotonicity.

https://doi.org/10.7554/eLife.45421.024

Article and author information

Author details

  1. Benjamin König

    Institute of Chemistry and Biochemistry, Freie Universität Berlin, Berlin, Germany
    Contribution
    Conceptualization, Data curation, Software, Formal analysis, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
  2. Yuchen Hao

    1. Institute of Biology, Humboldt Universität zu Berlin, Berlin, Germany
    2. Leibniz Forschungsinstitut für Molekulare Pharmakologie (FMP), Berlin, Germany
    3. NeuroCure, Charité Universitätsmedizin, Berlin, Germany
    Contribution
    Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
  3. Sophia Schwartz

    Institute of Chemistry and Biochemistry, Freie Universität Berlin, Berlin, Germany
    Contribution
    Investigation, Visualization
    Competing interests
    No competing interests declared
  4. Andrew JR Plested

    1. Institute of Biology, Humboldt Universität zu Berlin, Berlin, Germany
    2. Leibniz Forschungsinstitut für Molekulare Pharmakologie (FMP), Berlin, Germany
    3. NeuroCure, Charité Universitätsmedizin, Berlin, Germany
    Contribution
    Conceptualization, Resources, Software, Supervision, Visualization, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6062-0832
  5. Tobias Stauber

    Institute of Chemistry and Biochemistry, Freie Universität Berlin, Berlin, Germany
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    tobias.stauber@fu-berlin.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0727-6109

Funding

Bundesministerium für Bildung und Forschung (031A314)

  • Tobias Stauber

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Jonas Engelke for pilot experiments and help with the programming, Helge Ewers for providing experimental equipment, Iva Lučić for discussions and Arnold Boersma, Francesca Bottanelli, Thomas Jentsch, Clemens Kaminski, Bert Poolman and Isabelle Vernos for plasmids and cell lines.

Senior Editor

  1. Richard Aldrich, The University of Texas at Austin, United States

Reviewing Editor

  1. Leon D Islas, Universidad Nacional Autónoma de México, Mexico

Reviewer

  1. Leon D Islas, Universidad Nacional Autónoma de México, Mexico

Publication history

  1. Received: February 14, 2019
  2. Accepted: June 14, 2019
  3. Accepted Manuscript published: June 18, 2019 (version 1)
  4. Version of Record published: June 27, 2019 (version 2)

Copyright

© 2019, König et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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