Phosphatidylinositol 4,5-bisphosphate (PIP₂) and Ca²⁺ are both required to open the Cl⁻ channel TMEM16A

Abstract

Transmembrane member 16A (TMEM16A) is a widely expressed Ca²⁺-activated Cl⁻ channel with various physiological functions ranging from mucosal secretion to regulating smooth muscle contraction. Understanding how TMEM16A controls these physiological processes and how its dysregulation may cause disease requires a detailed understanding of how cellular processes and second messengers alter TMEM16A channel gating. Here we assessed the regulation of TMEM16A gating by recording Ca²⁺-evoked Cl⁻ currents conducted by endogenous TMEM16A channels expressed in Xenopus laevis oocytes, using the inside-out configuration of the patch clamp technique. During continuous application of Ca²⁺, we found that TMEM16A-conducted currents decay shortly after patch excision. Such current rundown is common among channels regulated by phosphatidylinositol 4,5-bisphosphate (PIP₂). Thus, we sought to investigate a possible role of PIP₂ in TMEM16A gating. Consistently, synthetic PIP₂ rescued the current after rundown, and the application of PIP₂ modulating agents altered the speed kinetics of TMEM16A current rundown. First, two PIP₂ sequestering agents, neomycin and anti-PIP₂, applied to the intracellular surface of excised patches sped up TMEM16A current rundown to nearly twice as fast. Conversely, rephosphorylation of phosphatidylinositol (PI) derivatives into PIP₂ using Mg-ATP or inhibiting dephosphorylation of PIP₂ using β-glycerophosphate slowed rundown by nearly 3-fold. Our results reveal that TMEM16A regulation is more complicated than it initially appeared; not only is Ca²⁺ necessary to signal TMEM16a opening, but PIP₂ is also required. These findings improve our understanding of how the dysregulation of these pathways may lead to disease and suggest that targeting these pathways could have utility for potential therapies.

Introduction

Transmembrane protein 16A (TMEM16A)² (also known as ANO1, DOG1, ORAOV2, and TAOS2) (1) is a Ca²⁺-activated Cl⁻ channel common to evolutionarily diverse organisms. The channel plays functionally varied roles including signaling contraction in cardiovascular smooth muscle cells (2), facilitating transepithelial water transport vital for mucociliary clearance in pulmonary epithelial cells (3), and transmitting pain detection in sensory neurons (as reviewed in Ref. 4). Disrupting TMEM16A function has serious implications exemplified by the perinatal lethality phenotype in TMEM16A-null mice (5). As a key regulator in multiple physiological processes, this channel could be a target for novel therapeutics to treat chronic conditions such as hypertension or cystic fibrosis.

Despite being identified only 10 years ago (6–8), several structural and functional studies revealed a wealth of information regarding how TMEM16A operates. To date, TMEM16A is known to be activated by elevated intracellular Ca²⁺ (6–8) which binds to a membrane-embedded domain located near a Cl⁻ conducting pore (9, 10). The functional TMEM16A channel is a dimer comprised of two identical subunits that each have 10 transmembrane domains and an independent Cl⁻-conducting pore (11, 12). When Ca²⁺ binds its cognate domain on TMEM16A, it induces a conformational rearrangement of the α-helix in the sixth transmembrane domain that physically opens the anion-conducting pore (10). Each Ca²⁺-binding site in the TMEM16A channel can accommodate two Ca²⁺ cations (13), and it appears as though the channel gates differently depending on whether one or two cations are bound (14). Intriguingly, the recent cryo-EM structures of TMEM16A in a Ca²⁺-bound state revealed that the anion pore was not wide enough for Cl⁻ permeation (9, 10), suggesting Ca²⁺ alone may not be sufficient to activate TMEM16A channels. Another signaling molecule that may regulate TMEM16A is the acidic phospholipid phosphatidyl 4,5-bisphosphate (PIP₂) (15–17).

PIP₂ is a minor component of the membrane, yet it is a master regulator of membrane function (18). Importantly, PIP₂ serves as the substrate for phospholipase C cleavage to produce inositol triphosphate (IP₃) and diacylglycerol; pathways that transduce extracellular signals to intracellular signaling events. However, PIP₂ also serves as a signaling molecule in its own right by regulating endocytosis and exocytosis, actin polymerization, establishment of basolateral polarity, and regulation of ion channels (19). A possible role for PIP₂ in TMEM16A regulation has already been explored by other groups, however, the interpretation of how PIP₂ alters TMEM16A currents remains disputed. One study reported that PIP₂ inhibits TMEM16A (16), another states that it does not regulate the channel (20), and yet two others indicate that PIP₂ promotes TMEM16A activity (15, 17). We speculate that the disparity among the experimental interpretations may stem from the use of indirect methods.

The objective of the research reported in this manuscript was to determine whether the phospholipid PIP₂ regulates TMEM16A channels. For these experiments, we made electrophysiology recordings from Xenopus laevis oocytes, cells that endogenously and abundantly express TMEM16A channels (7). Using excised inside-out patch clamp together with methods that directly alter the available PIP₂ content of the patch, we found that TMEM16A currents decayed following patch excision. By changing the available PIP₂ content, we altered the kinetics of this rundown. Moreover, depleting the membrane PIP₂ rendered TMEM16A channels unable to conduct current even in saturating concentrations of intracellular Ca²⁺. Together these findings establish that TMEM16A channels are potentiated by PIP₂.

Results

TMEM16A currents recorded from excised inside-out patches decayed over time

To study TMEM16A currents, we made recordings of the endogenous channel in X. laevis oocytes using the inside-out configuration of the patch clamp technique. Notably, the prominent Ca²⁺-activated current recorded from these cells is conducted by TMEM16A channels (7). During 100–150–ms steps to −60 and +60 mV, we observed robust, Ca²⁺-activated TMEM16A currents at both voltages (Fig. 1A). Surprisingly, these currents decayed over time despite the continued presence of a saturating concentration of intracellular Ca²⁺ (Fig. 1B) (9). Fig. 1A depicts typical currents recorded at −60 and +60 mV, at 30, 60, 120, and 180 s following 2 mm Ca²⁺ application. Loss of current from excised patches is a phenomenon known as rundown (21). We next sought to characterize the rundown of TMEM16A-conducted currents after patch excision.

Figure 1.

TMEM16A Ca²⁺-evoked Cl⁻ currents rundown in excised inside-out patches. Inside-out patch clamp recordings were conducted on macropatches excised from X. laevis oocytes. A, example currents recorded during 100–150–ms steps to −60 and +60 mV, at indicated times following patch excision. B, representative plot of current measured at −60 (bottom) or +60 mV (top) versus time, after patch excision. 2 mm Ca²⁺ was applied at 10 s as denoted by the gray bar. The dashed gray line represents 0 nanoamperes (nA). C, normalized plot of current measured at −60 mV versus time, fit with a single exponential (red line). D, box plot distribution of the rate of current decay (τ), measured by fitting plots of relative current versus time with single exponentials (n = 15). The central line denotes the median, the box denotes the distribution of 25–75% of the data, and the whiskers represent 10–90% of the data. The τ at +60 mV and −60 mV were not significantly different (p = 0.29) as determined by t test.

We first explored whether the rundown of TMEM16A currents was voltage dependent. To do so, plots of the currents recorded during steps to −60 mV or +60 mV, versus time, were fit with single exponential functions and enabled us to quantify the kinetics of rundown (Equation 1) (Fig. 1C). In 15 independent experimental trials, the average rate of rundown was 68.9 ± 7.1 s at −60 mV, and 63.8 ± 5.9 s at +60 mV (Table 1 and Fig. 1D). Similar kinetics measured at the two voltages reveal that the current decay is voltage independent.

Table 1.

Biophysical properties of TMEM16A excised inside-out patch currents

The mean ± S.E. for the indicated measurements taken from excised inside-out patches. The τ was obtained by fitting plots of the currents versus time with single exponential functions.

Condition	Concentration	Tau, τ (s)
Control (Ca2+)	2 mm	68.9 ± 7.1 (n = 15)
Anti-PIP2	15 μg/ml	48.6 ± 7.4 (n = 4)
Neomycin	50 mm	32.0 ± 8.8 (n = 5)
Mg-AMP	1.5 mm	57.7 ± 12.6 (n = 5)
Mg-ATP	1.5 mm	143.4 ± 17.2 (n = 5)
βGP	50 mm	166.8 ± 14.7 (n = 5)

To explore a possible relationship between the rate of rundown and the number of channels in an excised patch, we plotted the rate of current decay recorded at −60 mV versus the peak steady state current (Fig. S1). In 15 independent trials, we found that there was no apparent relationship between these two metrics. These data suggest that TMEM16A currents ran down regardless of pipette size or the number of channels opened by Ca²⁺ across the different trials.

PIP₂ recovered TMEM16A currents in inside-out excised patches

Rundown of currents recorded in the inside-out configuration of the patch clamp technique is characteristic of channels regulated by PIP₂ (19). We therefore hypothesized that if TMEM16A current rundown occurred because of the depletion of PIP₂ in excised patches, exogenous PIP₂ application should recover TMEM16A currents following their decay. To do so, we applied a water-soluble analog of PIP₂ (22–25), dicotanoylglycerol-PIP₂ (diC8-PIP₂), to inside-out patches excised from X. laevis oocytes (Fig. 2A). For these experiments, we recorded TMEM16A-conducted currents at −60 mV before and during the application of 2 mm Ca²⁺. Once the Ca²⁺-activated currents ran down to a steady state, we applied 100 μm diC8-PIP₂ (17) in the presence of 2 mm Ca²⁺. Fig. 2B depicts an example plot of TMEM16A-conducted currents recorded at −60 mV versus time, before and during 100 μm diC8-PIP₂ application in the presence of Ca²⁺. In seven independent trials, we observed that 100 μm diC8-PIP₂ application recovered an average 38% of the peak current. Because we did not observe full TMEM16A current recovery with 100 μm diC8-PIP₂, we asked if this was because of the short fatty acid tail or the concentration of diC8-PIP₂ applied. We first conducted a similar experiment where we applied 100 μm natural PIP₂ (Fig. S2, A and B). Surprisingly, we observed that natural PIP₂ recovered an average of 23% of the peak current, which was less than the current recovered by 100 μm diC8-PIP₂. We proceeded with further experimentation only using the diC8-PIP₂. Next, we sought to estimate the concentration response for diC8-PIP₂. We conducted additional TMEM16A current recovery experiments using 10 μm diC8-PIP₂ (Fig. S2C) and 30 μm diC8-PIP₂ (Fig. S2D). We observed that 10 μm or 30 μm diC8-PIP₂ incompletely recovered the current compared with 100 μm diC8-PIP₂ (Fig. 2C). Altogether, the data suggest that PIP₂ recovers TMEM16A currents and that the currents are recovered in a concentration-dependent manner.

Figure 2.

PIP₂ analog recovered TMEM16A-conducted currents following rundown when applied with intracellular Ca²⁺. The soluble synthetic analog of PIP₂ (diC8-PIP₂) was applied to excised inside-out patches once current had stably run down. Currents were recorded at −60 mV. A, schematic depiction of natural PIP₂ and the diC8-PIP₂. B, representative plot of normalized currents versus time, before and during application of 100 μm diC8-PIP₂ with 2 mm Ca²⁺. C, plot of the average -fold of current recovered by diC8-PIP₂ at 10, 30, and 100 μm diC8-PIP₂ addition in the presence of 2 mm Ca²⁺. The -fold change in current recovered was calculated as change in current upon application of diC8-PIP₂ with Ca²⁺.

After demonstrating that PIP₂ recovers TMEM16A currents, we sought to further characterize the relationship between PIP₂ and TMEM16A. We tested whether 100 μm diC8-PIP₂ (Fig. 3A) was sufficient to recover TMEM16A current following rundown, or if diC8-PIP₂ and Ca²⁺ were both required. Application of diC8-PIP₂ in the absence of Ca²⁺ had a nominal effect on TMEM16A, changing the current by 1.1 ± 0.1–fold (n = 5) (Fig. 3, B and C). These data demonstrate that although Ca²⁺ activates TMEM16A, PIP₂ is required for these channels to conduct Cl⁻ currents. Moreover, PIP₂ potentiates TMEM16A currents only in the presence of intracellular Ca²⁺.

Figure 3.

PIP₂ and Ca²⁺ are both required for TMEM16A-conducted currents. diC8-PIP₂, PI (diC8-PI), and PI(4)P (diC8-PI(4)P) were applied to excised inside-out patches once current had stably run down. Currents were recorded at −60 mV. A, schematic depiction of the various diC8 analogs used. B, box plot distribution of the -fold current recovered after the application of diC8-PIP₂ with Ca²⁺, diC8-PIP₂ without Ca²⁺, diC8-PI with Ca²⁺, or diC8-PI(4)P with Ca²⁺. The -fold change in current recovered was calculated as change in current upon application of diC8-PIP₂ with Ca²⁺ (n = 7), diC8-PIP₂ without Ca²⁺ (n = 5), diC8-PI (n = 6), or diC8-PI(4)P (n = 9) addition to the peak current observed after each synthetic analog addition. C–E, representative plots of normalized currents versus time, before and during application of 100 μm diC8-PIP₂ with no added Ca²⁺ (C), 100 μm diC8-PI with 2 mm Ca²⁺ (D), or 100 μm diC8-PI(4)P with 2 mm Ca²⁺ (E). ** denotes p < 0.01 and * denotes p < 0.05 as determined by t test. -Fold current recovered by diC8-PIP₂ without Ca²⁺ compared with diC8-PI was not significantly different (p = 0.20).

The diC8-PIP₂ compound could theoretically regulate TMEM16A channels by interactions mediated by its lipid tail group or its phosphoinositol head group. Thus, we tested the hypothesis that the dicotanoylglycerol-phosphoinositol backbone (diC8-PI) (Fig. 3A) alone was sufficient to recover current. Following TMEM16A current rundown, we applied 100 μm diC8-PI with Ca²⁺ and quantified the proportion of current recovered. Fig. 3D shows an example plot of TMEM16A currents recorded at −60 mV versus time, before and during diC8-PI application. In six separate trials, we observed that 100 μm diC8-PI application nominally altered the TMEM16A currents, recovering 1.4 ± 0.2–fold current, which was significantly lower than the 3.6 ± 0.5–fold current recovered by diC8-PIP₂ with Ca^2+, but not significantly different from the current recovered by diC8-PIP₂ in the absence of Ca²⁺ (Fig. 3B). This result demonstrates that without the phosphorylated head groups, diC8-PI was unable to recover TMEM16A currents, thereby suggesting that the phosphates on the inositol head group mediate the TMEM16A-PIP₂ interaction. To determine the role of the phosphate head group in mediating the TMEM16A-PIP₂ interaction, we also applied 100 μm diC8-PI(4)P (Fig. 3A) with Ca²⁺ and quantified the proportion of current recovered. Fig. 3E shows an example plot of TMEM16A currents recorded at −60 mV versus time, before and during diC8-PI(4)P application. In nine separate trials, we observed that 100 μm diC8-PI(4)P application recovered 2.1 ± 0.2–fold current (Fig. 3B), which represented an average of 13% of the peak current. The intermediate recovery (lying between 3.6 ± 0.5–fold current recovered by diC8-PIP₂ with Ca²⁺ and 1.4 ± 0.2–fold current recovered by diC8-PI with Ca²⁺) suggests that the phosphate head group is important for mediating TMEM16A-PIP₂ interaction. Altogether, the data reveal that in addition to Ca²⁺, PIP₂ regulates TMEM16A gating.

Scavenging PIP₂ sped up TMEM16A current rundown

Our finding that diC8-PIP₂ restored TMEM16A currents following rundown suggested that rundown may be the result of PIP₂ depletion in excised patches. Thus, we reasoned that we should be able to speed up rundown by applying compounds that compete with TMEM16A for binding to PIP₂. We tested this hypothesis by quantifying the rate of TMEM16A current decay during application of two compounds known to scavenge PIP₂: A PIP₂-targeting antibody (anti-PIP₂) (26) and neomycin (27, 28) (Fig. 4A). We began these experiments by recording TMEM16A currents at −60 mV before and during treatment with 2 mm Ca²⁺ applied with 15 μg/ml anti-PIP₂. In four independent trials, we observed that TMEM16A current ran down with an average rate of 48.6 ± 7.4 s in the presence of anti-PIP₂ compared with 68.9 ± 7.3 s from patches treated with 2 mm Ca²⁺ alone (n = 15) (Fig. 4, B and C and Table 1).

Figure 4.

Scavenging PIP₂ sped up TMEM16A current rundown in excised inside-out patches. A, schematic depicting membrane-embedded TMEM16A with the PIP₂ scavengers anti-PIP₂ or neomycin. B, box plot distribution of the rate of rundown observed from currents recorded at −60 mV from inside-out patches exposed to 2 mm Ca²⁺ with 15 μg/ml anti-PIP₂ (n = 4) or 50 mm neomycin (n = 5). The rate of current rundown was obtained by fitting the plots of the current versus time with a single exponential. The gray lines denote the box plot distribution of the rate of rundown measured from patches recorded under control conditions of 2 mm Ca²⁺-only application, where the solid gray line represents the median value and the dashed lines represent the control data distribution from 25 to 75%. C and D, representative plots of Ca²⁺-evoked Cl⁻ currents recorded at −60 mV, versus time, from inside-out patches in the presence of 2 mm Ca²⁺ with 15 μg/ml anti-PIP₂ (C) or 50 mm neomycin (D); both conditions sped up current rundown when compared with the control denoted by the gray dashed line. * denotes p < 0.05 determined by t test.

In a parallel series of experiments, we quantified the rate of TMEM16A current rundown in the presence of the PIP₂ scavenger neomycin. Neomycin is an antibiotic that also scavenges PIP₂ when applied to the intracellular membrane at high concentrations (27). Fig. 4D depicts an example plot of the TMEM16A currents recorded at −60 mV versus time, before and during 2 mm Ca²⁺ and 50 mm neomycin application. In five independent trials, neomycin sped up current rundown from 68.9 ± 7.3 s to 32.0 ± 8.8 s (Fig. 4B).

Altogether, we observed that both anti-PIP₂ and neomycin application sped up TMEM16A rundown (Fig. 4). Moreover, the rate of rundown was not significantly different in the presence of either anti-PIP₂ or neomycin (Fig. 4B and Table 1) consistent with the hypothesis that each compound was acting by scavenging PIP₂ rather than exerting nonspecific effects on the channel. Together, these data support the hypothesis that PIP₂ is required for TMEM16A to conduct Cl⁻ currents.

Slowing PIP₂ depletion slowed TMEM16A rundown

Phosphatases and kinases work together to maintain stable levels of PIP₂ in whole cells (Fig. 5A) (29). In excised patches, however, membrane-anchored kinases lack access to the ATP required to fuel phosphorylation and regeneration of PIP₂ (19, 29). Consequently, continued activity of phosphatases without counteracting kinases leads to PIP₂ dephosphorylation. We reasoned that if TMEM16A currents decayed in excised patches because of the dephosphorylation of PIP₂, then enabling rephosphorylation or inhibiting phosphatases should slow current loss. We tested the hypothesis by determining whether enabling rephosphorylation of PIP₂ with application of magnesium-adenosine triphosphate (Mg-ATP) (30, 31) would slow TMEM16A current rundown in excised inside-out patches. As a control for these experiments, we first recorded TMEM16A currents at −60 mV before and during the application of magnesium adenosine monophosphate (Mg-AMP), which can bind to, but not activate, kinases. Thus, Mg-AMP application should not affect current rundown. Indeed, we observed no difference in current rundown between Mg-AMP and control condition of 2 mm Ca²⁺ (Fig. 5, B and C). We then recorded TMEM16A currents at −60 mV before and during application of 2 mm Ca²⁺ with 1.5 mm Mg-ATP. An example plot of normalized current versus time is shown in Fig. 5D. We observed that in the presence of 1.5 mm Mg-ATP and 2 mm Ca²⁺, TMEM16A currents ran down over a longer time course, with a time constant of 143.4 ± 17.2 s (n = 5) (Fig. 5D and Table 1).

Figure 5.

Enabling rephosphorylation or inhibiting phosphatases slowed current decay in excised inside-out patches. A, schematic depicting the role of phosphatases and kinases in generating various phosphoinositide species. B, the box plot distribution of the rate of rundown (τ) observed from currents recorded at −60 mV from inside-out patches exposed to 2 mm Ca²⁺ with 1.5 mm Mg-AMP (n = 5), 2 mm Ca²⁺ with 1.5 mm Mg-ATP (n = 6), and 2 mm Ca²⁺ with 50 mm sodium βGP (n = 6). Single exponentials were fitted to current traces to obtain the rate of current rundown. Solid gray line denotes median rate of rundown measured from patches recorded under the control conditions of 2 mm Ca²⁺-only application, and the dashed lines represent the control data distribution from 25 to 75%. C–E, representative plots of normalized currents versus time made before and during application of 1.5 mm Mg-AMP with 2 mm Ca²⁺ (compared with 2 mm Ca²⁺-only application with the gray dashed line) (C), 1.5 mm Mg-ATP with 2 mm Ca²⁺ (compared with 1.5 mm Mg-AMP with the orange dashed line) (D), and 50 mm βGP with 2 mm Ca²⁺ (compared with 2 mm Ca²⁺-only application with the gray dashed line) (E). ** denotes p < 0.01 and * denotes p < 0.05 when compared with 2 mm Ca²⁺-only application or between indicated pairs determined by t test.

Next, we reasoned that if phosphatase-mediated PIP₂ depletion causes TMEM16A current rundown in excised patches then inhibiting phosphatase activity would also slow rundown. We tested this hypothesis by quantifying the kinetics of current rundown in the presence of Ca²⁺ and the general phosphatase inhibitor sodium β-glycerophosphate pentahydrate (βGP) (32). Fig. 5E depicts an example plot of TMEM16A-conducted currents recorded at −60 mV versus time, before and during application of 50 mm βGP and 2 mm Ca²⁺. In five independent trials, we observed that TMEM16A currents ran down slower in the presence of the phosphatase inhibitor, with a time constant of 166 ± 14.7 s (n = 5) (Fig. 5E and Table 1). Together, these data suggest that TMEM16A currents run down in excised patches as the result of PIP₂ depletion via its dephosphorylation. Moreover, these data are consistent with the hypothesis that the phosphates found on the inositol head group of PIP₂ are responsible for the interaction between TMEM16A and PIP₂.

Discussion

By recording TMEM16A currents while modifying the membrane PIP₂ content, here we demonstrate that these channels require both Ca²⁺ and PIP₂ to conduct current. A possible role for PIP₂ in TMEM16A regulation was initially suggested by rundown of these currents recorded from X. laevis oocytes despite the continued presence of a saturating concentration of Ca²⁺ (Fig. 1). Rundown results from the removal of a membrane patch from the cytosol that includes ATP (29). This ATP is required to fuel the phospholipid kinases which rephosphorylate the phospholipids present in the membrane (29). As such, current rundown in excised patches is a hallmark of channels regulated by PIP₂.

Application of the soluble PIP₂ analog diC8-PIP₂ recovered TMEM16A-conducted currents following rundown, but only when applied with Ca²⁺ (Figs. 2 and 3). Not all TMEM16A current was recovered with diC8-PIP₂ application. 100 μm diC8-PIP₂ applied without Ca²⁺ had a nominal effect on TMEM16A currents revealing that both Ca²⁺ and PIP₂ are required for the phospholipid to potentiate these channels. We speculate that the incomplete recovery with 100 μm diC8-PIP₂ applied with Ca²⁺ may reflect a slow rate of PIP₂ integration into the membrane. This is consistent with our observation that the current recovery was diminished when longer acyl chains were applied to patches (Fig. S2). Alternatively, the incomplete recovery may reflect the role of a Ca²⁺-dependent recovery of PIP₂. PIP₂ recovery can be more pronounced at lower Ca²⁺ than at higher Ca²⁺ (17). This complex relationship between PIP₂ and Ca²⁺ needs to be explored further in our system.

Phospholipid-mediated recovery of TMEM16A currents following rundown requires phosphorylation of the inositol ring. Accordingly, we found that 100 μm diC8-PI, the phospholipid lacking the negatively charged phosphate head groups, was unable to recover TMEM16A currents (Fig. 3). In contrast to the recent report that suggests that the neutral acyl chain of fatty acids is sufficient to regulate TMEM16A (15), our findings suggest that TMEM16A potentiation by PIP₂ requires the presence of phosphate heads which are lacking in lipids like oleic acid or cholesterol. Our results do not preclude the possibility that fatty acids change PIP₂ membrane content, but they do suggest that just a fatty acid tail or backbone like that from PIP₂ is not sufficient for channel potentiation. Consistent with this idea, we observed that adding PI(4)P, the precursor for PIP₂, containing a singular phosphate at position 4 of the phosphoinositol head group, recovered TMEM16A-conducted Cl⁻ currents (Fig. 3).

Our data suggested that TMEM16A current rundown in the excised patch was caused by PIP₂ depletion. We therefore reasoned that we ought to be able to speed TMEM16A current rundown by scavenging PIP₂ present in the patch. Indeed, application of one of two different PIP₂-scavanging compounds, anti-PIP₂ or neomycin, sped up TMEM16A current rundown (Fig. 4). Other groups have shown that anti-PIP₂ and neomycin effectively scavenged PIP₂ without disrupting ion conductance during single-channel recordings (33) or in excised macropatches expressing the inwardly rectifying K⁺ channels (34, 35).

PIP₂ depletion occurs in excised patches because of the continued activity of membrane-associated phosphatases to dephosphorylate the inositol head group, without the activity of the counteracting kinases. If TMEM16A current rundown in excised patches is the result of PIP₂ dephosphorylation, we predicted that by enabling kinases to rephosphorylate the lipid or by inhibiting phosphatase activity, we ought to be able to slow current rundown. Indeed, providing patches with Mg-ATP to fuel membrane-anchored kinases to make PIP₂, resulted in TMEM16A currents that ran down significantly more slowly compared with control recordings (Fig. 5). Although it is possible that Mg-ATP could bind to and alter the activity of other proteins in the excised patches, in the context of the other data included in this manuscript, it is likely that Mg-ATP is fueling kinases needed to rephosphorylate PIP₂. In a parallel series of experiments, the broad-spectrum phosphatase inhibitor βGP (32) similarly slowed rundown by at least 3-fold. Together, these data demonstrate that the dephosphorylation of the inositol head group of PIP₂ with patch excision leads to TMEM16A current decay. These data provide additional support that the phosphates on the inositol head group mediate the PIP₂ potentiation of TMEM16A.

Our data directly demonstrate that PIP₂ and Ca²⁺ are both required for TMEM16A activation, yet previous studies of phospholipid regulation of this channel came to varied conclusions. Although agreeably demonstrating that PIP₂ modifies native TMEM16A current in isolated rat pulmonary cells, one study reported a decrease in whole cell Ca²⁺-activated Cl⁻ currents with the application of diC8-PIP₂ via a pipette solution (16). This study found that including the PIP₂ scavenger polylysine in the pipette solution increased TMEM16A currents and therefore concluded that PIP₂ inhibits TMEM16A currents (16). Using HEK293 cells exogenously expressing mouse TMEM16A, another group reported that neither siRNA-mediated knockdown of PTEN nor wortmannin application altered TMEM16A currents (20). Because both treatments should reduce PIP₂ content by diminishing the dephosphorylation of phosphatidylinositol 3,4,5-triphosphate (PIP₃) into PIP₂, this report concluded that PIP₂ does not regulate TMEM16A (20). The final two studies on the subject also used TMEM16A-transfected HEK293 cells to conclude that mouse TMEM16A is potentiated by PIP₂ (15, 17). One group reported that although TMEM16A-mediated currents did not run down in excised inside-out patches, application of diC8-PIP₂ increased the current (17). In this same study, activation of a co-expressed voltage-sensing phosphatase reduced, but did not abolish, TMEM16A currents (17). A final study reported that fatty acids, including PIP₂, potentiate TMEM16A-conducted currents (15). Our data, by contrast, indicate that the lipid tail alone is unable to regulate TMEM16A currents. A possible explanation for the difference between our data and the finding that other fatty acids potentiate TMEM16A is that stearic acids, including those used during these particular experiments, can associate with PIP₂ (36). Consequently, stearic acids could alter membrane PIP₂ and therefore alter TMEM16A-conducted currents. We sought to resolve these seemingly conflicting results by recording from natively expressed TMEM16A channels in X. laevis oocytes to determine whether PIP₂ regulates TMEM16A gating. Using direct experimental methods, here we demonstrate that TMEM16A requires both membrane PIP₂ and intracellular Ca²⁺ to conduct currents in X. laevis oocytes.

Despite our demonstration that TMEM16A gating is regulated by PIP₂, the exact mechanism involved is yet to be determined. PIP₂ regulates other ion channels by diverse mechanisms (19). For example, PIP₂ may regulate TMEM16A currents by acting on an accessory protein, exemplified by PIP₂ binding to KCNE1 to regulate currents conducted by the voltage-gated potassium channel KCNQ1 (37). Alternatively, PIP₂ can form electrostatic interactions between a cluster of positive charges on a channel and the negative charges on the phosphoinositol head of PIP₂ as has been observed for transient receptor potential vanilloid 5 (TRPV5) (38), inward-rectifier K⁺ (Kir2.2) (39), and G protein–gated K⁺ (GIRK2) (40). Unlike KCNQ1, no accessory protein has been revealed for TMEM16A. Although CLCA1 has been identified as an accessory protein for TMEM16A channels (41), neither this protein nor the RNA are found in mature X. laevis eggs or the fertilization-incompetent oocytes (42–44). Our data, however, do not preclude an indirect mechanism for PIP₂ regulation of TMEM16A by another accessory protein for TMEM16A. We speculate that TMEM16A and PIP₂ may form electrostatic interactions similar to those revealed by the TRPV5, Kir2.2, and GIRK2 PIP₂-bound structures.

Electrostatic interactions mediate PIP₂ binding to the closely related cation channel, TMEM16F (45). The putative PIP₂-binding site in TMEM16F is comprised of positively charged residues, and neutralizing these residues perturbed PIP₂'s ability to potentiate TMEM16F currents (45). It is possible that a conserved domain mediates PIP₂ interactions with all TMEM16 family proteins including TMEM16A channels. Although the binding domain may be shared, the effects on proteins will most certainly differ. For example, the Ca²⁺-activated Cl⁻ channel TMEM16B is inhibited by PIP₂ (17). Yet, a shared PIP₂ regulation among TMEM16 family proteins is perhaps not surprising given that this protein family includes several lipid scramblases whose physiologic function requires their interaction with charged lipids.

A requirement for PIP₂ in TMEM16A activation is intriguing because the PIP₂ cleavage into IP₃ to signal Ca²⁺ release from the ER would seemingly oppose the ability of increased Ca²⁺ to activate the channel. Yet several independent studies have revealed that an IP₃-evoked Ca²⁺ release from the ER opens TMEM16A channels to signal diverse physiologic processes including signaling contraction of lymphatic vessels (46, 47) to activation of the fast polyspermy block (43, 48). We propose that in systems that use an IP₃ pathway to activate TMEM16A channels, the amount of PIP₂ cleaved to generate IP₃ matters greatly. For example, stimuli that evoke the cleavage of only a moderate amount of PIP₂ to increase IP₃ and evoke a Ca²⁺ release from the ER will ultimately signal the opening of more TMEM16A channels compared with other stimuli that signal the cleavage of most of the membrane PIP₂ to increase IP₃ and evoke an even larger Ca²⁺ release from the ER. Perhaps TMEM16A discriminates its own Ca²⁺ signal from Ca²⁺-signaling cascades that exclude it by seeking interactions with PIP₂ at the plasma membrane. Moreover, these studies reveal that the physiologic mechanisms underlying TMEM16A opening are more complicated than simply the presence or absence of intracellular Ca²⁺, thereby enabling TMEM16A to play diverse roles in different cell types.

Understanding how the channel is regulated lays the conceptual framework for drugging this novel interaction in disease. In hypertension, the overconstriction of vessels can be alleviated by inhibiting Ca²⁺-activated Cl⁻ channels like TMEM16A (49). In cystic fibrosis, a condition arising from a dysfunctional Cl⁻ channel, increasing TMEM16A activity could rescue the defects caused by poor Cl⁻ transport (50). By understanding the interaction between PIP₂ and TMEM16A, drugs targeting the PIP₂-TMEM16A interaction site can be designed to either inhibit TMEM16A activity with the benefit of lowering blood pressure or increase its activity to promote Cl⁻ transport.

Experimental procedures

Reagents

diC8-PIP₂, diC8-PI, diC8-PI(4)P, PIP₂ 18:0/20:4, and anti-PIP₂ IgM (catalog no. Z-P045, lot no. 080416) were obtained from Echelon Biosciences. MgCl₂ was obtained from Sigma. Unless otherwise noted, all other reagents were purchased from Thermo Fisher Scientific.

Solutions

All inside-out patch clamp recordings were made in HEPES-buffered saline solution (in mm): 130 NaCl and 3 HEPES, pH 7.2, and filtered using a sterile, 0.2-μm polystyrene filter. For Ca²⁺-free recordings, this solution was supplemented with 0.2 μm EGTA as indicated. For solutions used during Ca²⁺ application, the HEPES-buffered saline solution was supplemented with 2 mm CaCl₂ and with indicated reagents. For current recovery experiments, one of the diC8 analogs or natural PIP₂ was added to the HEPES-buffered saline solution supplemented with 2 mm CaCl₂. Natural PIP₂ was dissolved and sonicated prior to use because of its longer acyl chain.

Oocyte wash and storage solution were made as follows: Oocyte Ringers 2 (in mm): 82.5 NaCl, 2.5 KCl, 1 MgCl₂, and 5 mm HEPES, pH 7.2. ND96 (in mm): supplemented with 5 sodium pyruvate and 100 mg/liter gentamycin, pH 7.6, and filtered with a sterile, 0.2-μm polystyrene filter.

Animals

Animal procedures were conducted using accepted standards of humane animal care and approved by the Animal Care and Use Committee at the University of Pittsburgh. X. laevis adult, oocyte-positive females were obtained commercially (RRID:NXR_0031, NASCO, Fort Atkinson, WI) and housed at 18 °C with 12/12-h light/dark cycle.

Oocyte collection

Oocytes were collected from X. laevis females anesthetized by immersion in 1.0 g/liter tricaine, pH 7.4, for 30 min. Ovarian sacs containing the oocytes were removed from the female, manually pulled apart, and incubated for 90 min in 1 mg/ml collagenase diluted in the ND96 solution. Collagenase was removed by several Oocyte Ringers 2 rinses, and healthy oocytes were stored at 14 °C in ND96 for up to 14 days.

Patch clamp recordings

Patch clamp recordings were made on X. laevis oocytes following the manual removal of the vitelline membrane. Current recordings were made in the inside-out configuration of the patch clamp technique (51) with an EPC-10 USB patch clamp amplifier (HEKA Elektronic). Briefly, after formation of a gigaseal (greater than 1 gigaohm), inside-out patches were excised in HEPES-buffered saline solution lacking EGTA (resistances often decreased to 20–200 megaohm in solutions lacking EGTA but returned to greater than 1 gigaohm with EGTA application). Data were collected at a rate of 10 kHz. Glass pipettes were pulled from borosilicate glass (outer diameter 1.5 mm, inner diameter 0.86 mm; Warner Instruments), fire polished (Narshige microforge), and had a resistance of 0.4–1.5 megaohm. Lipids were applied to excised inside-out patches in a RC-28 chamber (Warner Instruments). All other solutions were applied using a VC-8 fast perfusion system (Warner Instruments). Experiments were initiated within 10 s of patch excision.

Data analysis

Patch clamp data were acquired with PATCHMASTER (HEKA Elektronic) and analyzed with Igor Pro (RRID:SCR_000325, WaveMetrics), Patchers Power Tools (RRID:SCR_001950), and Excel (RRID:SCR_016137, Microsoft). Currents were normalized such that the basal currents recorded in Ca²⁺-free conditions were equated to 0 and the peak currents obtained with 2 mm intracellular Ca²⁺ were normalized to 1.

Data from various experimental conditions are displayed in Tukey box plot distributions where the central line represents the median value, the box depicts 25–75% of the data range, and the whiskers span 10–90%.

To facilitate comparison between the kinetics of current rundown recorded under experimental conditions to controls, the normalized current for each condition is plotted together with the averaged rundown for associated control, either the 2 mm Ca²⁺ or the 1.5 mm Mg-AMP. These controls are plotted with dashed lines, and represent an idealized averaged normalized current versus time calculated using the single exponential Equation 1:

$$Y\left(x\right)=Y_0\times e^{\frac{-x}{\tau }}$$ (Eq. 1)

where Y(x), Y₀, x, and τ represent the current at time x, initial current, time, and rate of current rundown, respectively. Briefly, traces were collected under either condition and fitted with the single exponential equation above to derive the Y₀, x, and τ. These values where collected for each individual plot. To create averaged plots of current rundown for control and Mg-AMP conditions, single exponential functions using the averaged variables were plotted along with the normalized current versus time graphs. The predicted current traces were then plotted as the dashed lines.

To compare the magnitude of current recovered using the synthetic lipid analogs (diC8-PIP₂, diC8-PI, diC8-PI(4)P) and natural PIP₂, the -fold change in current recovered was calculated by dividing the peak current after diC8-PIP₂ or diC8-PI by baseline current. The peak current was defined as the highest current obtained after diC8-PIP₂, diC8-PI, diC8-PI(4)P, or natural PIP₂ addition. The baseline current was equated to 1 and defined as the current observed at point of diC8-PIP₂ or diC8-PI addition.

All experimental conditions include trials that were conducted on multiple days with oocytes collected from different females. Two-tailed analyses of variance (ANOVAs) were used to report differences between experiments conditions, followed by post hoc t tests to discern differences between particular experimental treatments.

Author contributions

M. T. and A. E. C. conceptualization; M. T. and A. E. C. data curation; M. T. and A. E. C. formal analysis; M. T. and A. E. C. funding acquisition; M. T., K. L. W., R. E. B., and A. E. C. investigation; M. T., K. L. W., R. E. B., and A. E. C. methodology; M. T. and A. E. C. writing-original draft; M. T. and A. E. C. project administration; M. T., K. L. W., R. E. B., and A. E. C. writing-review and editing; A. E. C. supervision.