Specific PIP2 binding promotes calcium activation of TMEM16A chloride channels

TMEM16A is a widely expressed Ca2+-activated Cl− channel that regulates crucial physiological functions including fluid secretion, neuronal excitability, and smooth muscle contraction. There is a critical need to understand the molecular mechanisms of TMEM16A gating and regulation. However, high-resolution TMEM16A structures have failed to reveal an activated state with an unobstructed permeation pathway even with saturating Ca2+. This has been attributed to the requirement of PIP2 for preventing TMEM16A desensitization. Here, atomistic simulations show that specific binding of PIP2 to TMEM16A can lead to spontaneous opening of the permeation pathway in the Ca2+-bound state. The predicted activated state is highly consistent with a wide range of mutagenesis and functional data. It yields a maximal Cl− conductance of ~1 pS, similar to experimental estimates, and recapitulates the selectivity of larger SCN− over Cl−. The resulting molecular mechanism of activation provides a basis for understanding the interplay of multiple signals in controlling TMEM16A channel function.

while others can be delegated to the SI. Was there a single PIP2 per subunit in the simulations? Or only one PIP2? This has not been stated explicitly in the manuscript. Based on the previous paper by the authors, it seems that they have two copies of PIP2, but that is only my guess, and needs to be clarified. Related to this, why is it "intriguing" that only one subunit shows opening in each simulation? Given that the opening is rather fast and starts in the very beginning of the simulation, it is actually worrisome as to why not all subunits show opening in the presence of PIP2 bound.
If I understand correctly, TMEM16A channel activation (as well as the PIP2 role) is voltage-dependent, with the TM2-TM3 linker and TM6 potentially involved in voltage sensing (Xiao et al, PNAS 2011;Peters et al., Neuron 2018). However, the simulations here starting from a nonconducting conformation were performed without any membrane potential. Moreover, the whole cytoplasmic domain (including the TM2-TM3 linker), which is not only dynamic and flexible but also functionally important to the channel opening, was restrained during all the simulations. This is an important problem and deserves some discussion in the manuscript.
A clear definition of the "open" conformation needs to be provided. In Fig. 2 and 2S, where the L547-I641 distance and the number of pore water are used to measure the openness of the pore, the main criterion defining open and closed pores needs to be spelled out. For example, compared to the snapshots colored as open pores, some snapshots with larger distance or more pore water are colored as closed pore ( Fig. 2A).
If I understand correctly, during the free energy simulations, the channel is kept in its open state. If that is the case, there is a problem with the calculated conductance value, as it probably represents only the upper value of the macroscopic conductance measured experimentally. The channel can easily continue to flicker between open and closed microscopic states in experiments, and the measured conductance would be a weighted average of these microstates. That is not what we have in the simulations. On the other hand, the opening observed in the simulations here can represent only the beginning of a larger opening with a larger conductance. These all can affect the measured conductance and how it compares with experimental data and should be briefly mentioned.
I also note that there is a large assumption that D of the ion is half of its bulk value in the calculation of conductance. As such, perhaps we should refrain from calling the captured state a "fully activated" state.
The clusters in Fig. S4 are not clearly defined and separated. Especially, the yellow cluster, which is defined as the open and conductive state, is overlapping with the other three clusters. Please clarify the measure for the separation of the clusters.
Spontaneous diffusion of a Cl-ion through the pore was captured in the equilibrium simulation, which could be used to seed the umbrella sampling simulations. Why was SMD used to pull a Cl-ion through the pore?
When pulling SCN through the pore, what orientation of SCN was used and why? Please also provide details of the SMD simulation, such as the pulling velocities.
How does one explain the absence of a major barrier at the narrowest region (the neck) in the case of SCN permeation? This is really curious.
What are the protonation states of the amino acids in the Ca2+-bound (with or without PIP2) and Ca2+-free systems? Was there any difference between the two systems? How was it determined?
In Fig 1, the three amino acids (two of them are mentioned in the figure legend) shown in yellow sticks in the neck region should be labelled. In Figs. 1 and 5, the coloring scheme of the key TMs in the molecular structure and carton representation are not consistent.
In Fig 2, scale the pore radius profile in panel B to match the molecular structures in panels C and D for better illustration.
Use one unit for energy (kcal/mole or kJ/mole) and distance (nm or A). The present presentation is confusing.
μS is the unit for conductance and should be replaced with μs (time) in Fig. 2 labels.
I suggest that most movies (if not all) be best smoothed so things can be seen better. Water can be shown as O only, if they look strange after smoothing.
In Reference 75, the first and last names of the authors are transposed. Fix. This manuscript explores the mechanisms of PIP2 regulation of the TMEM16A chloride channel using computational methods. TMEM16 has been shown to have a number of crucial physiological functions and the mechanisms of its regulation are poorly understood, so these studies are relevant and address an important physiological/biophysical problem. Overall, the approach yields some intriguing conclusions and insights and seems to be consistent with other published data. However, there are several major issues that dampen my enthusiasm considerably.
(1) This study has no controls. If the authors wish to make the argument that PIP2 binding to this site in the presence of Ca2+ stabilizes an open conformation of the channel, they need to show that other phospholipids do not produce the same effect. They should at least test PI(4)P and phosphatidylserine. Further, in their Nature Communications paper, they show that the effect of PIP2 is absent in the K567A mutant. They should show that the open conformation does not occur with this mutant and other mutants studied in this paper.
(2) This manuscript is entirely computational. Although I appreciate the power of molecular dynamic simulations, my philosophy is that computational solutions should be viewed as a predictive tool and they do not stand alone without experimental verification. I see two solutions to this problem. Preferably the authors should perform experimental tests of the model in Figure 5 or at least propose in the discussion a specific experiment that would show that this model is testable and not a pipedream. Alternatively, the authors could explore predictions made in the Le et al. paper (for example predictions about the K579-E564 salt bridge). (4) In Fig. 2A, there is surprisingly little correspondence between the N(pore water) trace and the L547-I641 distance. For example, the channel appears to be in a closed conformation between 0.6-0.7 us while pore water remains high. Conversely, between 1.8 -2.3 us, pore water is low, but the channel is in the open conformation. While I understand that these two measures might be expected to be temporally separated to some degree, if one is going to argue that pore waters are an indication that the channel is open, it seems necessary to show that these two are correlated. Further, what is the correspondence between PIP2 coordination by K567, R451, R575, and R482 and channel open conformation? Do the transient channel closures correspond to PIP2 unbinding? Finally, if the open pore can contain lipid instead of water, the authors should discuss this finding in relationship to the proposal by Whitlock et al. Pflugers Arch. 2016;468: 455-473. (5) I don't doubt that PIP2 can bind to this site, but I am concerned that binding in the MD simulations is simply caused by non-specific electrostatic attraction. Simulations that were performed in the Le et al. paper showed that PIP2 would spontaneously bind to its binding site within 50ns when PIP2 was placed "near" the binding pocket (the example in Fig. 4b shows this distance as <10A). However, no binding events were observed when PIP2 was placed further away. Positive charge density of the putative binding site will attract PIP2 electrostatically, especially if PIP2 is not initially complexed with counterions like K+, Mg2+ (typically 1 -3 mM), and Ca2+ (probably >100 uM under conditions required to activate the channel). These divalent cations will compete with protein binding to PIP2. I would like the authors to try a less biased approach to test that this site is "the" binding site, especially because, as the authors acknowledge, other investigators have reported somewhat divergent results.
In addition to these major concerns, there are a number of specific (although not less serious) issues that require attention.
(1) On page 9, the authors conclude that Cl ions remain well hydrated during permeation and then they imply that CLC channels are similar ("This is similar to what has been observed for Clpermeation through ClC channels, in which the number of hydrated water also drops to <5."). In fact, in CLC channels, Cl is almost completely dehydrated during permeation and virtually all of the coordination is provided by protein. If Cl permeates TMEM16 partially hydrated, this suggests that the selectivity mechanisms of ion permeation of CLC and TMEM16A are completely different. Perhaps more important is the question: Is there any experimental data supporting the idea that Cl permeates TMEM16A channels partially hydrated?
(2) Abstract: The statement that "we show that specific binding of PIP2 to TMEM16A can lead to spontaneous opening…." is not precise because it suggests that this was determined by experimental, not computational means. The abstract should be rewritten to include methodology.
(3) Page 2, the statement that the lower half of TM6 occludes the lower pore and blocks the entry of permeating ions is incorrect. While it is probably true that the lower half of TM6 unfolds during pore opening, there is no direct evidence that I know that supports the statement the authors make.
(4) Figure 1. Helices are not labelled in C. Color coding of helices in D is inconsistent with other panels.
(5) Figure 2. The y-axis is labelled N(pore water) but the blue line is lipid. Methods state that the results were determined without considering the side chains of K588 and K645. I presume this statement applies only to B-D, but the authors should be more precise and show as supplementary data the calculations with these side chains. Also, the legend states that panel A plots the "distance between the centers of mass of L547 and I641". I used centerofmass in PYMOL to calculate the center of mass of these two amino acids in 5OYB and it shows the distance is 7.3A, not 2-3 A as plotted here.
Exactly what was measured?
(6) Insufficient methodological detail is provided. ProMod3 requires a template. What templates were used? What information is used to determine that the models are reasonable? Also, why is water only clustered around the protein in the movies and not present in the extracellular space?
(7) The simulations were all performed with POPC bilayers, which does not mimic mammalian plasma membrane that has a significant fraction of POPS.
(8) It is stated on p7 that "The state representing the opened pore conformation is only found in simulations with PIP2", but Figure S4B shows a significant number of red dots (simulations without PIP2) in the yellow area. Further, the criteria used to define the yellow area as an "OPEN" conformation in Methods is vague. Please specify what inter-residue distances and number of waters were used.

Author Note: The original comments from the reviewers are quoted in bold fonts.
Key changes to the manuscripts are noted throughout the responses.

While capturing the conformational transition of the channel by the simulations is interesting and it provides information on the nature of the open state, there are several questions that need to be addressed before I can recommend the paper for publication. 1 Most importantly, the mechanism provided at the end of the paper for PIP2 activation comes at a bit of surprise and needs to be elaborated with some data supporting the idea. We don't see any data on the nature of TM4 motion until the end where it is proposed to be the main change resulting in pore opening in a semi-schematic figure. The nature and extent of TM4 movement need to be established to be related to lipid binding (similar to pore radius and hydration of the pore) by data. If limited by space, the network analysis can be pushed to the SI, as it does not really make a strong point in the current form; perhaps it is not well explained.
Responses: We agree with the reviewer's suggestion and have performed additional analysis of the movements and distributions of all TM helices in the closed and predicted open states. The result is summarized in a new Fig S5; it shows that TM4 is the only helix that undergoes significant movement during PIP2-induced activation.
We have also included a short discussion to the revised manuscript on P7, stating "The movements of TMs can be further visualized by comparing the distributions of their centers of mass (CMs) in the closed and open states (Fig. S5A). The result show that there are ~4 and ~1 Å movements of the upper pore segments of TM4 (T539:L547) and TM3 (G510:A523) during activation, respectively, while the other TMs show minimal movements. Note that the structures of all TM helices are very stable as reflected in the small root-mean-squared fluctuation profiles (Fig. S5B)."

The manuscript needs a more extensive comparison of the results with previous studies on some of core specific aspects. In particular the paper by Yu et al. (PNAS 2019) on PIP2-TMEM16A interaction is very close in scope to the present study, and some of the results of the two studies need to be compared more explicitly.
Responses: Please see the response to Question 3 below.

Responses:
We agree with the reviewer (and reviewer 3; see below) that there are important unanswered questions regarding the molecular basis of PIP2 activation of TMEM16A. There is strong experimental evidence to support the role of the specific binding site investigated in the current work (Le et al, Nature Communications 2019), as well as multiple other PIP2 binding sites that potentially form a dynamic network to regulate the gating of TMEM16A (Yu et al PNAS 2019). These observations likely reflect the complexity of TMEM16A regulation and existence of multiple activated states accessible under different experimental conditions (e.g., Ca 2+ concentration). Reconciling these results requires additional experiments (and simulations) beyond the scope of this study. Instead, motivated by the identification of the specific binding site, the objective of this computational study is to test if binding a single specific PIP2 would be sufficient to activate the channel and if so what are the structure features of the predicted activated state.
We fully agree with the reviewer that these complexities should be further discussed, particularly in the context of results reported in Yu et al PNAS 2019. We have followed the suggestion and included an extended discussion of these remaining issues and how our observations are consistent and different from those reported in Yu et al 2019 study (see P14-15): "Besides the specific PIP2 binding in the TMs 3-5 (Fig. 1), other PIP2 putative binding sites have also been identified, such as near the dimer interface, intracellular loop between TMs 2, 3 and the cytoplasmic end of TM6.25,28 Among these, only one at the dimer interface is near the specific PIP2 binding31, albeit with different sets of contacting basic residues. Furthermore, binding of PIP2 to these sites has been proposed to be dynamic and form a regulatory network to modulate the channel activation. At present, it is not clear how to reconcile these important differences in the molecular basis of PIP2 regulation of TMEM16A function. It has been suggested that TMEM16A can access multiple open states under different activation conditions (e.g., Ca 2+ concentration and membrane potential) 34 and that these functional states may show different responses to PIP2.25,28,31 Another important difference between the current study and Yu et al 25 is that the previous simulations identified TM6 as the key helix that moved in response to PIP2 binding. This is most likely due to distinct PIP2 binding configurations investigated. The specific binding site investigated in this work locates in the back side of TM3-5 from the pore. It is probably not a total surprise that TM4 and 5 are observed to the main helices that respond to PIP2 binding (Fig. S5A)......"

Was there a single PIP2 per subunit in the simulations? Or only one PIP2? This has not been stated explicitly in the manuscript. Based on the previous paper by the authors, it seems that they have two copies of PIP2, but that is only my guess, and needs to be clarified. Related to this, why is it "intriguing" that only one subunit shows opening in each simulation? Given that the opening is rather fast and starts in the very beginning of the simulation, it is actually worrisome as to why not all subunits show opening in the presence of PIP2 bound.
Responses: We would like to clarify that the specific binding site actually locates at TM3-5 behind the pore and is far from TM6 (Fig 1). PIP2 directly interacts with TM3/4 but not TM6.
We have slightly revised the manuscript (see P17) to clarify that: "Specifically, there is one PIP2 per subunit, directly coordinated by R455, R486, K571 and R579." Previous experimental study suggests the two subunit is activated independently (P2 and ref 14,  15). We suspect that it is coincidental that we did not observe both subunits opening in our three simulations, due to the limited timescale and transient nature of the opening transition.

If I understand correctly, TMEM16A channel activation (as well as the PIP2 role) is voltagedependent, with the TM2-TM3 linker and TM6 potentially involved in voltage sensing (Xiao et al, PNAS 2011; Peters et al., Neuron 2018). However, the simulations here starting from a nonconducting conformation were performed without any membrane potential. Moreover, the whole cytoplasmic domain (including the TM2-TM3 linker), which is not only dynamic and flexible but also functionally important to the channel opening, was restrained during all the simulations. This is an important problem and deserves some discussion in the manuscript.
Responses: We thank the reviewer for pointing out the potential pitfalls of restraining the cytosolic domains implied in voltage gating. The simulations with PIP2 (or other lipids bound) were initiated with the Ca 2+ bound structure and thus mimicked the condition under saturating Ca 2+ where membrane depolarization is not required for activation. Therefore, we believe that the weak restraints on the structured portion of the cytosolic domain, imposed to minimize the effects of missing loops, should not have a significant impact on observing PIP2-induced activation.
The following notes have been made in Methods: "In all simulations, no external electric field has been applied, as membrane potential is not required to fully activate TMEM16A under saturating Ca 2+ ." (on P17), and later "We note that voltage-dependent gating of TMEM16A has been proposed to involve the intracellular TM2-3 linker and possibly TM6.36,43 Nonetheless, the current simulations aim to capture voltage-independent PIP2-induced activation under saturation Ca 2+ . Therefore, the restraints on the structured region of the cytosolic domain is not expected to affect the activation transitions." (on P18) Fig. 2 and 2S, where the L547-I641 distance and the number of pore water are used to measure the openness of the pore, the main criterion defining open and closed pores needs to be spelled out. For example, compared to the snapshots colored as open pores, some snapshots with larger distance or more pore water are colored as closed pore (Fig. 2A).

Responses:
We have added to following discussion to the revised manuscript to clarify how the open and close states were identified (on P7): "As the pore opening involves dilating in the whole upper pore region, a single residue-residue distance or the number of pore water molecules alone could not clearly sperate the open and closed state. Here, we performed cluster analysis based on distances between pore-lining residues and the number of pore waters in the neck region (See Method part for details)." The pathway properties of the resulting clusters were then inspected to assign each cluster to open, closed or other (transient) states.

If I understand correctly, during the free energy simulations, the channel is kept in its open state. If that is the case, there is a problem with the calculated conductance value, as it probably represents only the upper value of the macroscopic conductance measured experimentally. The channel can easily continue to flicker between open and closed microscopic states in experiments, and the measured conductance would be a weighted average of these microstates. That is not what we have in the simulations. On the other hand, the opening observed in the simulations here can represent only the beginning of a larger opening with a larger conductance. These all can affect the measured conductance and how it compares with experimental data and should be briefly mentioned.
Responses: We generally agree with the reviewer's points but want to point out that the maximum single channel conductance measurements reported in the literature should have been corrected for less than 100% open probabilities. We have revised the manuscript to further clarify the pitfalls in comparing the calculated conductance and experimental measurements (see P10-11), "We note that the theoretical estimate of maximum single channel conductance does not consider larger scale conformational fluctuation within the activate state and should only be considered semi-quantitative. It is possible that the open state captured in the current simulation only reflects some early stage of a large opening."

I also note that there is a large assumption that D of the ion is half of its bulk value in the calculation of conductance. As such, perhaps we should refrain from calling the captured state a "fully activated" state.
In Reference 75, the first and last names of the authors are transposed. Fix. Fig. 1 Fig. 1 caption: TMs 6-9 Fig. 3: stick-and- TMs 4 and 6 ... is slightly separated -> ... are slightly separated It's members -> its members Methods: Fig. S ??? Responses: Thank you for the careful reading. These typos and errors have been corrected.

Author Note: The original comments from the reviewers are quoted in bold fonts.
Key changes to the manuscripts are noted throughout the responses.

Reviewer #3 (Remarks to the Author):
This manuscript explores the mechanisms of PIP2 regulation of the TMEM16A chloride channel using computational methods. TMEM16 has been shown to have a number of crucial physiological functions and the mechanisms of its regulation are poorly understood, so these studies are relevant and address an important physiological/biophysical problem. Overall, the approach yields some intriguing conclusions and insights and seems to be consistent with other published data. However, there are several major issues that dampen my enthusiasm considerably.

(1) This study has no controls. If the authors wish to make the argument that PIP2 binding to this site in the presence of Ca2+ stabilizes an open conformation of the channel, they need to show that other phospholipids do not produce the same effect. They should at least test PI(4)P and phosphatidylserine. Further, in their Nature Communications paper, they show that the effect of PIP2 is absent in the K567A mutant. They should show that the open conformation does not occur with this mutant and other mutants studied in this paper.
Responses: We appreciate the reviewer's suggestion of additional controls to show that other lipids binding to the same site would not induce pore opening. We first want to note that the same site is always occupied by POPC in existing control simulations without PIP2, which do not induce pore opening. We followed the reviewer's suggestion and perform 6 additional control simulations with PI(4)P and POPS lipids docked to the specific binding site using the same pose as in PIP2 simulations (sim 10-15, SI Table S1). Setup of these new control simulations is described in the revised Methods section (P17). The results are summarized in a revised SI Fig.  S2 and discussed in P7: "It should be noted the binding pockets are always occupied with POPC lipids even in simulations without PIP2. These observations are consistent with the experimental results showing that PIP2 is required for maintaining the conductive state of TMEM16A even under saturating Ca 2+ concentrations.26, 27, 31 As additional controls, we further examined the effects of two negatively charged lipids, POPS and PI(4)P, bound to the same pockets. Previous experimental studies have suggested that POPS could not activate TMEM16A, while PI(4)P only show minor effects on inhibiting channel rundown.26, 31 As summarized in Fig. S2, the binding of these negatively charged lipids only slightly increased the numbers of pore water (sim 10-15) compared to those without PIP2 (and with POPC) (sim 7-9). Interestingly, an transient pore-opening event was observed in one of the simulations with PI(4)P (sim 13), apparently consistent with some capacity of PI(4)P in re-activating the channel." For K567A mutant, it likely abolishes PIP2 specific binding and thus activation. Our current simulations do not attempt to make de novo predictions on what kinds of lipids can bind to the binding site or how pocket mutations may perturb the binding properties. Such prediction requires binding free energy analysis, which is beyond the scope of this work and extremely challenging due to the size and flexibility of lipid molecules. As also explained in our response to Reviewer #1, Question #3 above, our current study was motivated by the identification of the specific binding site and the objective is to test if binding a single specific PIP2 would be sufficient to activate the channel and if so what are the structure features of the predicted activated state.

(2) This manuscript is entirely computational. Although I appreciate the power of molecular dynamic simulations, my philosophy is that computational solutions should be viewed as a predictive tool and they do not stand alone without experimental verification. I see two solutions to this problem. Preferably the authors should perform experimental tests of the model in Figure 5 or at least propose in the discussion a specific experiment that would show that this model is testable and not a pipedream. Alternatively, the authors could explore predictions made in the Le et al. paper (for example predictions about the K579-E564 salt bridge).
Responses: We wholeheartedly agree with the notion that computation must be integrated with experiment to derive reliable understanding of complex channel function. We want to note that our simulations are motivated by and build on our previous experimental and computational studies (with Huanghe Yang's lab). Throughout this study, we compare key structural and functional properties to published experimental data (mutagenesis, conductance, selectivity etc).
We have followed the reviewer's advice and propose a set of specific experiments that could be used to test key predictions from this computational study (see P10): "It can be expected that removing of one or both basic residues (K588 and K645) could significantly decrease the maximum conductance of the channel and/or increase the activation barrier. Conversely, replacing the ring of hydrophobic residues above the inner gate (V543, I640 and P595) with either polar or charged residues may have similar effects as inner gate residue mutations in modulating the activation of TMEM16A CaCC." We further hypothesize that (see P16-17): "A possible way to test this is to introduce appropriate mutations to the ring of hydrophobic residues above the inner gate (V543, P595 and I640), which is responsible for giving rise to the maximum free energy barrier of Cl-permeation. If a fully dilated scramblase shows similar conductance and ion selectivity with these mutations, it may suggest that ions mainly follow the lipid pathway."  TMEM16A.6,8,32,33,34" (4) In Fig. 2A

Responses:
We agree that the number of pore water should in principle provide an intuitive indicator of the state of the channel. However, the channel can undergo relatively small conformational fluctuations and lead to transient increase in pore hydration. We observe that no single metrics (e.g., pore water, helix distance etc) can fully resolve the conformational state. As such, we performed clustering analysis to first identify major conformational states and then assigned them to open, close or other states. This provides a superior approach to resolve the open and close states. We have included a short clarification in the revised manuscript (see P7): "As the pore opening involves dilating in the whole upper pore region, a single residue-residue distance or the number of pore water molecules alone could not clearly sperate the open and closed state. Here, we performed cluster analysis based on distances between pore-lining residues and the number of pore waters in the neck region (See Method part for details) ..." The revised manuscript also includes the probabilities of PIP2 contacts with the coordinating basic residues in the binding pocket (see P5): "During the simulations, PIP2 maintained stable contacts with the coordinating residues, mainly R455, R486, K571 and R579. The contact probabilities of PIP2 to these four residues are 0.40, 0.49, 0.97 and 0.99, respectively. The average RMSD of the PIP2 headgroup and these basic residues from the initial conformation is 3.5±0.7 Å" We thank the reviewer for bringing to our attention of the nice discussion on how lipid tails contribute to the permeation pathway. The revised manuscript now includes an expanded discussion on this aspect (see P12): "We also note that it has been previously proposed that lipids likely contribute to the formation of the ion permeation pathways in TMEM16 family proteins. 34 Results from the current simulations clearly support this idea, showing that lipids line the ion permeation pathway even for TMEM16A with limited dilation (Fig. S7A) and that the presence of lipids contribute to ion permeation properties (Fig. S6)."

(5) I don't doubt that PIP2 can bind to this site, but I am concerned that binding in the MD simulations is simply caused by non-specific electrostatic attraction. Simulations that were performed in the Le et al. paper showed that PIP2 would spontaneously bind to its binding site within 50ns when PIP2 was placed "near" the binding pocket (the example in Fig. 4b shows this distance as <10A). However, no binding events were observed when PIP2 was placed further away. Positive charge density of the putative binding site will attract PIP2 electrostatically, especially if PIP2 is not initially complexed with counterions like K+, Mg2+ (typically 1 -3 mM), and Ca2+ (probably >100 uM under conditions required to activate the channel). These divalent cations will compete with protein binding to PIP2. I would like the authors to try a less biased approach to test that this site is "the" binding site, especially because, as the authors acknowledge, other investigators have reported somewhat divergent results.
Responses: We appreciate the reviewer's comments on our published work (Le et al, Nature Communications 2019). As explained above in our responses to Question #1 and Reviewer #1