Swapped and non-swapped TRAAK states co-exist in membranes at a ratio influenced by temperature

Domain: chemistry
Published: 2026-03-05
Editorial outcome: major_revision

Abstract
The potassium two-pore domain (K2P) ion channel TRAAK is expressed in the nervous system and regulates the fast action potential in membranes. Like all K2P channels, TRAAK possesses a distinct extracellular cap, which adopts a swapped and a non-swapped conformation1,2. However, the proportional representation of these two species within native membranes – and the trigger or stimulus associated with this conformational transition – are unknown. Here, we utilise Pulse Dipolar EPR Spectroscopy combined with heterologous single-subunit spin-labelling and monitor the complete conformational ensemble of TRAAK’s cap domain in membranes. We demonstrate the coexistence of the swapped and non-swapped states, quantify their populations within the TRAAK ensemble, and show that the swapped conformation dominates, with the ratio being influenced by temperature. Native lipid analysis shows that TRAAK selectively associates with and is activated by signalling lipids to the exclusion of membrane-dominant phosphatidylcholine lipids from its vicinity, forming a distinct microdomain. Our approach can identify the immediate lipid environment, detect and quantify cap state populations in homo-/hetero-K2P channels and link domain swapping to specific triggers.

Introduction
Two-pore domain potassium (K2P) channels maintain the background leak currents for the resting membrane potential in mammalian cells. K2P channels are subject to modulation by various triggers and stimuli, such as fatty acids, lipids, chemicals, temperature, pH, tension, and anaesthetics3-6. They could be also regulated by other proteins, such as Piezo1/2 mechanosensitive channels and kinases, while they exhibit distinct responses to G proteins, which play a crucial role in neuronal excitability and astrocytic glutamate release7-12. The expression of K2P channels is widespread across vital organs such as the heart13-15, brain16,17, retina18, and pancreatic duct19, making them valuable drug targets20,21, important in anaesthesia22,23, and central to disorders20,24,25.

K2P channels are structurally distinct to all other tetrameric potassium channels and unlike them, K2Ps form dimeric structures. Further, K2Ps possess a distinct extracellular cap, which protrudes approximately 3.5 nm from the membrane and bifurcates the transport pathway of potassium. Pore-blocking venom-derived peptides, which typically bind to the external entrance of potassium channel pores, cannot block K2P channels26,27, suggesting the cap may sterically obstruct the pore entrance to the extracellular side.

The first two K2P channel structures of the human TWIK-related arachidonic acid-stimulated potassium (TRAAK) (PDB 3UM7)1 and the tandem of pore domains in a weak inward rectifying potassium (TWIK-1) (PDB 3UKM)28 channel were modelled with their cap domain in a non-swapped (NS) configuration, namely with the outer transmembrane helix (TM1) interacting with the inner helix of the same subunit (Fig. 1a). Later, through the use of stabilising antibodies or the introduction of gain-of-function mutations, TRAAK’s cap was solved in a swapped (S) state, that is with the outer TM1 interacting with the inner-helix of another subunit in the dimeric complex2,17,25,29,30. Since then, the S conformation has been reported in all 52 out of 54 × K2P channel structures solved by cryoEM or x-ray crystallography and across different K2P channels, e.g. TRAAK, TWIK-Related K+ Channel 1 (TRE K1), TWIK-Related K+ Channel 2 (TREK2), TWIK-Related Acid-Sensitive K+ Channel 1 (TASK1), TWIK-Related Acid-Sensitive K+ Channel 2 (TASK2), TWIK-Related Acid-Sensitive I K+ Channel 3 (TASK3) and TWIK11,2,17,25,28-39 (Supplementary Table 1).

Pulse Dipolar electron paramagnetic resonance Spectroscopy (PDS) is a powerful technique to monitor dynamics, folding, oligomerisation and conformational ensembles in integral membrane proteins40-54. This ensemble method commonly involves the incorporation of paramagnetic spin labels through cysteine-based modifications on selected protein sites to determine inter-spin distance distributions55-58. We utilised an approach which combines Å resolution PDS distance measurements in ion channels40,41,54,59,60 with heterologous single-subunit spin labelling (HSS-SL). This strategy, we purposely developed here, was essential to distinguish between the two cap states of TRAAK (Fig. 1b,c). We unbiasedly monitored the complete conformational ensemble of the TRAAK cap domain. We studied human TRAAK, for which both the S and NS conformations have been reported1,2,29 and endeavoured to explore the (in)existence of the two states in eukaryotic lipid membranes, as well as identifying triggers (or stimuli) which would hint towards the biological (ir)relevance of K2P cap swapping. We investigated the full-length channel devoid of any truncations of functional domains, such as the C-terminus, which in K2P channels acts as a polymodal sensor through interactions with free fatty acids61, kinases7,9 and G-proteins10, and in TRAAK mediates axon initial segment localisation in brain excitatory neurons62. Further, by using detergent-free Styrene-Maleic Acid (SMA) copolymer technology63,64 we eliminated any detergent-induced artefacts, which often occur during membrane solubilisation. We also conducted lipidomics analysis65-67 on the parental cell plasma membranes surrounding the SMA-encapsulated TRAAK. Our native lipid membrane analysis shows that TRAAK selectively favours phosphatidylinositol (PI) and various phosphorylated -PIs; PIP1, PIP2 and PIP3 signalling lipids, previously associated with functional effects on several K2P channels68, while the normally dominant phosphatidylcholine (PC) lipids are excluded from TRAAK’s immediate lipid environment.

We demonstrate the coexistence of the S and NS cap states and quantify their proportional representation within TRAAK’s channel population in membranes. We find that the S state is the dominant cap conformation, aligning with its high prevalence in previous structural reports. PDS measurements show that the proportional representation of the two species in eukaryotic plasma membranes is altered, when the temperature changes. We conclude that the NS cap state in the human TRAAK (and by extrapolation a possibility in TWIK1) channels represents a sparsely represented, but biologically relevant state, in a thermo-dependent S/NS ratio. Our PDS-based approach is not limited to TRAAK, can detect and quantify cap states also within other K2P channels and thus link cap states to physiologically relevant conditions. This is applicable to both homo and hetero K2P dimers, within the same (or different) K2P subfamilies, a process which is known to enhance K2P channel functional diversity69,70.

Results
A platform for heterologous single-subunit spin labelling (HSS-SL) in K2P channels

All 54 × K2P high resolution structures reported to date have been obtained from truncated constructs, missing the crucial C-terminus, which is involved in K2P multimodal regulation7,9,10,61 and in TRAAK encodes an ankyrin G-binding motif that mediates periodic co-distribution with ankyrin G62. All these K2P constructs used previously in structural studies only produce two thirds of the protein’s length (Supplementary Table 1). Here, we heterologously expressed the full-length human TRAAK channel and performed both structural and functional analysis. We further isolated the channel in detergent-free conditions, using SMA-encapsulated lipid particles, designed to carry one channel per disc. This approach diminishes any unspecific inter-dimer distance contributions from within the same membrane disc, which could obscure our PDS experiments.

We first exchanged the five native non-disulphide-bond forming cysteine residues in human TRAAK to serine residues, creating a construct template containing only inaccessible cysteine residues (inCysTRAAK hereafter), and subsequently introduced Cys at desired sites. We retained the native Cys78 which forms a stable disulfide bond with the equivalent Cys of the second TRAAK subunit (thus not amenable to spin labelling) and is essential to stabilise the cap (Fig. 1a). We expressed the inCysTRAAK construct and evaluated its function by electrophysiology following reconstitution in giant unilamellar vesicles (GUVs) (Supplementary Fig. 1g-l). Proteoliposomes were induced to form membrane blisters, which allowed the creation of high-resistance GOhm seals between the membrane and patch pipette. This setup enabled electrophysiological recordings in the ‘inside-out’ configuration under voltage clamp conditions. The Nernst equilibrium potential for K+ (E ) in a 10-fold concentration gradient of K+ is -59 mV. The reverse potential closely matches E (Supplementary Fig. 1i, l), confirming that the channel is functional and potassium selective. To assess tension sensitivity, incremental pressure steps were applied to the patch of proteoliposomes, resulting in progressively increasing currents (Supplementary Fig. 1g). Furthermore, a single pressure step of -50 mmHg was provided while voltage steps were incremented to further examine potassium selectivity of the mechanically induced current. Both the baseline and mechanically stimulated currents displayed a reverse potential close to E (Supplementary Fig. 1h, i). Under these conditions, inCysTRAAK exhibited activation at 1.54 ± 0.10 -fold above the basal current level (mean ± SEM, −50 mmHg, Vh = 0 mV, n = 4 patches, Supplementary Fig. 1i). inCysTRAAK could also be activated by arachidonic acid (AA), a known activator of TRAAK, by 1.69 ± 0.06 -fold (mean ± SEM, Vh = 0 mV, n = 3 patches, Supplementary Fig. 1j-l). In summary, we found that inCysTRAAK is potassium selective, can be activated by AA and is responsive to membrane tension, with activities comparable to those of the wild-type TRAAK (Supplementary Fig. 1a-f, p, q).

To test whether inCysTRAAK presents any unspecific or specific labelling of the native Cys78, we incubated the purified protein with a very large excess of the spin label MTSSL ((1-Oxyl-2,2,5,5-tetramethyl-3-pyrroline-3-methyl) methanethiosulfonate)71-73 and could not detect any signal by continuous wave (CW) EPR (Supplementary Fig. 2b). After confirming our protein template cannot be spin labelled and forms a functional channel, we performed screening of multiple sites to identify candidates for Cys insertion for MTSSL labelling (delivering the R1 side chain) and generated in silico distance distributions for multiple sites spanning across the cap (67R1, 70R1, 93R1) and the transmembrane region (144R1). For spin labelling of a single engineered site per subunit in homodimeric TRAAK, the architecture of the cap results in highly similar predicted distances for the two states. Therefore, PDS would not have distinguished between the two cap conformations (Fig. 1b, Supplementary Fig. 3). To circumvent this, we performed an in silico heterologous single-subunit spin labelling analysis, by placing two spin labels on a single TRAAK monomer, while leaving the other subunit unlabelled. Among the pair combinations tested, the 67R1/144R1 pair showed the largest distance separation between the S and NS TRAAK cap states, providing the highest resolution (Fig. 1c, Supplementary Fig. 4). The predicted difference in the most probable distances between S and NS states is around 2 nm (mean ≈ 5 nm, SD ≈ 0.3 nm for S state; mean ≈ 3.3 nm, SD ≈ 0.5 nm for NS state) allowing unambiguous separation and quantification of the two states74. (Fig. 1). This separation is significantly greater than twice the length of the R1 side-chain, thus beyond and excluding any biased spin label conformation that could obscure our PDS measurements. The predicted distances also fall within the optimal distance range for PDS, which is between 2 to 6 nm, allowing the two states to be distinguished.

To identify the presence of distinct cap state(s) and quantify the cap state populations within the TRAAK ensemble we developed the HSS-SL protocol. This allowed us to ensure that one subunit of the expressed dimeric protein was spin-labelled while the other subunit remained unlabelled (i.e., EPR silent). Our protocol tailored for TRAAK could also be used for other dimeric membrane protein transporters, ion channels and pumps expressed in yeast75. The HSS-SL pipeline is illustrated in Fig. 2a. In a nutshell, the monomer carrying 2 × Cys (i.e., E67C/L144C) has a V5 tag and was co-transformed with the label-free inCysTRAAK, which contains a His-tag, in a 1:1 ratio. Integrated DNA sequencing confirmed that the two Cys mutants were present only in the fragment containing the V5-tag. The protocol was optimised for the EPR-active heterodimeric TRAAK population and was confirmed by Western blot analysis (Supplementary Fig. 2e). This led to three distinct isoform TRAAK species being expressed in our eukaryotic cells; a homodimer of inCysTRAAK (i.e., Cys inaccessible for spin labelling), a homodimer with two Cys pairs on each subunit (4 × Cys per dimeric channel) and a heterodimer with one Cys pair on only one of its subunits (2 × Cys per channel, i.e., our targeted species). During purification, only isoforms containing a His-tag would bind to the Ni2+ resin, while the homodimer, which contains 2 × Cys pairs on each subunit and carries no His-tag would not. The EPR silent inCysTRAAK homodimer and the targeted heterodimer, which contains only one Cys-accessible pair on one of the TRAAK subunits are present in our EPR samples. Thus, only the latter species could account for the total EPR signal detected and contribute to PDS distance measurements.

Prior to initiating EPR distance measurements we assessed the effect of E67C/L144C a) mutations and b) MTSSL modifications on TRAAK’s function. To this end, we expressed, purified and reconstituted TRAAK E67C/L144C into GUVs and performed patch clamp electrophysiology under the same conditions as for inCysTRAAK (Fig. 2b-e, g, Supplementary Fig. 1m-o). We then tested whether there is an effect on channel activity, due to spin label(s) attachment (i.e., on the E67C/L144C sites) and found that MTSSL does not alter TRAAK function (Fig. 2e-g). Our recordings revealed that the modified full-length channel exhibited potassium selectivity, tension sensitivity and AA activation similar to inCysTRAAK and wild-type TRAAK (Supplementary Fig. 1p,q), showing 1.60 ± 0.04-fold activation by membrane tension over basal current levels (mean ± SEM, −50 mmHg, Vh = 0 mV, n = 4 patches. Fig. 2d) and 1.45± 0.14-fold activation by AA over basal current levels (mean ± SEM, Vh = 0 mV, n = 3 patches. Supplementary Fig. 1m-o). The reversal potential of the current was also near E , indicating potassium selectivity of the modified channel (Fig. 2g)1.

TRAAK associates with and is activated by PI and signalling PIP lipids and excludes PC

We investigated which native lipids, originating from the parental eukaryotic plasma membrane used for expression, constitute the immediate microdomain environment of TRAAK. To this end, we performed electrospray mass spectrometry ( ES-MS) analysis on the SMA-encapsulated channels to identify the lipids associated with the full-length human TRAAK. These lipids were selected by and co-residing with TRAAK immediately prior to membrane solubilisation and could hint structural and functional roles.

We detected relatively high amounts of a range of phosphatidylinositol (PI) and signalling phosphatidylinositol phosphates (PIP) lipids, in particular PIs (34:1, 34:2 and 34:3) and PIP, PIP2 and PIP3 (42:6-10) (Fig. 3a). The strikingly high amounts of PIPs found compared to their natural distribution in the plasma membrane suggests that PIPs detected here could serve as an important binding partner of full-length TRAAK. PIPs is found in the inner leaflet of the plasma membrane, waiting to be hydrolysed by phospholipase C and similar. PIPs have also been reported to bind TRAAK and modulate other K2P channels, including the TREK, THIK, TALK, and TASK subfamilies6,68,76.

We further detected several other lipids with phosphatidylethanolamine (PE), phosphatidylserine (PS) headgroups and variable acyl-chain saturation levels. In particular, we detected PE (36:2, 36:3 and 36:4) and PS (34:1). PS is normally found in the inner-leaflet of the plasma membrane unless the cell is apoptotic, when PS is flipped to the outer-leaflet. Both PE and PS are found in human neuronal cells77 where TRAAK natively expresses in17 and affect the function of TRAAK6.

Despite extensive efforts we were unable to detect any PC lipids in the positive ion mode survey scans (Supplementary Fig. 5), even though PC is by far the most abundant lipid species in both Pichia pastoris78 and human neuronal membranes79. This is also evident in the inositol phosphorylceramides (IPC) lipids we detected and particularly (36:2;O3 and 36:3;O3). IPC is not present in human membranes, as it is a yeast specific lipid and is biosynthesised from PI, with both lipid species found in the plasma membrane. The equivalent of IPC in human membranes is sphingomyelin, which has a choline headgroup, and would not be present in the Pichia pastoris membranes, hence undetected in our analysis. Notably, the exclusion of highly abundant PC containing membranes by TRAAK is not an artefact of SMA extraction, as SMA does not have a preference for specific lipids65, can efficiently extract PC lipids67 and lipidomics analyses on integral membrane proteins expressed in Pichia pastoris and purified in SMALPs, have previously detected high amounts of PC lipids80. Another membrane component that is surprisingly missing from our lipidomics analysis is any sterol, which would also be present at relative high levels in the plasma membrane, so for yeast this would be ergosterol, rather than cholesterol for mammalian cells.

We then tested the effect of multiple, most abundant and representative lipids detected in our ES-MS analysis on TRAAK function using electrophysiology. We first tested each one of our patches for presence of functional TRAAK channels by applying controlled membrane tension and subsequently incubated with variable lipids and recorded channel activities. PI and PI(4,5)P₂ activated TRAAK (n=3 patches) with ~1.5-fold, similar to arachidonic acid (AA) (Fig. 3b,c, Supplementary Fig. 1r). In contrast, DOPE (PE 36:2) inhibited TRAAK (n=3 patches) generating a 2.00 ± 0.25-fold decrease (mean ± SEM, Vh = 0 mV, n = 3 patches; Fig. 3d). Finally, when our GUVs loaded with TRAAK, were incubated with POPS (PS 34:1) we observed no detectable affect (n = 4 patches; Fig. 3e) on TRAAK’s ion current.

In summary the lipids present are either neutral or negatively charged, PC (positive charge headgroup) and sterol are distinctly excluded/absent, and the PIPs and PS are likely to be on the inner-leaflet, while the PI and PE (involved in membrane curvature) will likely be on the outer-leaflet of the membrane.

PDS combined with HSS-SL enables probing of the entire TRAAK cap state ensemble

Prior to PDS, continuous-wave EPR (Lcw-EPR) confirmed that the mutated sites in each expressed protein were efficiently spin-labelled and that any excess labelling reagent was quantitatively removed (Supplementary Fig. 6). We then conducted Q-band (34 GHz) PDS distance measurements on the 67R1/144R1 pair (Fig. 4a, b) captured in native cell lipid membranes (SMALPs) at near ambient temperature (19oC). The time traces exhibited strong visual oscillations, which led to robust distance distributions of two populations centred at 3.3 nm and 5 nm respectively with different weights (Fig. 4c, d). Widths and shapes of these distance populations are in full agreement with the in silico predictions for the cap NS and S states, suggesting both states exist within the conformational ensemble of TRAAK in the lipid membrane. These measurements were reproduced for a different protein batch yielding almost identical PDS results (Supplementary Fig. 7). To further validate our findings are not dependent upon a single spin pair combination81, we performed PDS on a second heterologously spin labelled pair (70R1/93R1). For this pair we also observed strong oscillations in the raw traces, indicating reliability of the resulting distance distributions (Supplementary Fig. 8). However, the separation between S and NS modelled distances for this pair is not as ideal as for 67R1/144R1 (Supplementary Fig. 8d), due to limitations by the structural architecture of the K2P cap. Modelled distance distributions show significant overlap, thus the experimentally obtained trace and corresponding distance distribution cannot resolve the presence of two distinct distance components. Nevertheless, the width of the observed distribution almost perfectly matches the combined width of the two modelled distributions, while the presence of only one of the states should have yielded a substantially narrower distribution with a shift in distance for the maximum probability. These data further support the presence of both S and NS species in the TRAAK ensemble.

To ensure that only intramolecular TRAAK distances contribute to the resulting distance distributions for both of our HSS-SL pairs and avoid any intermolecular dipolar couplings arising from adjacently localised TRAAK channels, we designed certain features and implemented experimental controls as follows.

We used a SMA type which forms lipid discs ranging from 10 to 16 nm in diameter, depending upon the lipid environment and the presence of proteins82,83. Such adaptability of SMA is crucial for maintaining the functional activity of TRAAK in the membrane. As the TRAAK’s cross sectional TM region being approximately 8 nm, means only one TRAAK channel could fit in each lipid disc. We performed PDS for all four homo-dimeric single Cys TRAAK mutants for all the individual sites included in our heterodimeric pairs (i.e. 67R1, 70R1, 93R1, L144R1) to demonstrate efficient formation of TRAAK homodimers and rule out intermolecular distance contributions. We subsequently modified different variants of homodimeric TRAAK with MTSSL and performed PDS distance measurements in SMA (Supplementary Fig. 9, 10). For all single Cys homodimeric TRAAK mutants tested we observed strong dipolar coupling interactions between the singly labelled homo-dimeric channels and obtained reliable distance distributions. Distances are highly consistent with the in silico predictions for the two species which are expected to overlap, though with different mean values for the different sites and with no additional distance contributions present. We also designed single Cys hetero-TRAAK constructs (67C and 144C), but despite multiple attempts their production has not been possible. However, any potential intermolecular distances appearing in there should also be present in our single-Cys homodimeric TRAAK distributions, but this was not the case (Supplementary Fig. 9, 10). We further constructed a 67/144 homo-dimer containing four single Cys within a single TRAAK channel to ensure that the PDS signal originates only from a single subunit, with no contribution from any other homo-oligomeric formation within the E67/L144 sample. Once again, we observed clear oscillations in the raw data of doubly homo-labelled TRAAK, which were very different from the ones recorded for hetero-labelled (67R1/144R1) TRAAK (Supplementary Fig. 11, 12).

Our multi-layered controls taken together, demonstrate the absence of any intermolecular or unwanted intramolecular distances contributing to our heterologously labelled TRAAK distributions.

NS and S states co-exist in membranes and their ratio is influenced by temperature

The experimental PDS distance distributions are in excellent agreement with predictions for the NS and S cap states (Fig. 4, Supplementary Fig. 7, 8). For quantification of populations the shape of the distance distribution must be reliable (highest reliability range, green colour bars in Fig. 4d, f)81. To increase the reliability of the distance distributions we combined short time traces with high time resolution and excellent signal-to-noise with longer time traces and analysed them globally84,85. While usually two traces are sufficient, we decided to include all data available (up to 4 traces) to avoid bias by selecting certain subsets of data. Despite multiple different analysis workflows and controls consistently reproducing these two peaks in the non-parametric distributions, the peak representing the S state lies outwith the range where the shape of the distribution can be directly interpreted (this being ultimately limited by PDS trace length and thus echo lifetime). To overcome this technical limitation, we fitted these non-parametric distance distributions with a bimodal Gaussian model. While the quantification is having inherently higher uncertainty for longer distances, we found highly consistent results using technical and biological replicates. Estimating the relative populations of NS and S cap states we found that the majority of TRAAK is in the S state, with a substantial minority population being in the NS state S/NS: (57±4 and 43±4)% (with the uncertainty representing 95% confidence or 2). The dominance of the S state within the TRAAK ensemble aligns with the overwhelmingly frequent occurrence in the previous x-ray crystallography and cryoEM structures of TRAAK and other K2P channels. Our findings are also consistent with previous computational analysis suggesting the S state is energetically favourable86, thereby more stable and amenable to structural studies.

Following quantification of the cap state ensemble, we next wondered whether the S/NS ratio could be influenced by any physiologically relevant triggers or stimuli. Previously, TRAAK’s activity has been shown to increase by ∼10-fold with a temperature increase from 17 to ∼40°C87. We thus decided to express TRAAK under identical conditions, we incubated the membranes with SMA at two distinct temperatures, 19°C and 40°C. We then performed PDS on the TRAAK heterodimer 67R1/144R1 and found that at 40°C there was a substantial change in the relative ratio between the two cap states with a large decrease in the NS state, meaning that it becomes even rarer upon temperature increase (Fig. 4e, f). In particular, at 40°C the S state’s dominance becomes significantly greater S/NS (66±3/34±3) % vs (57±4/43±4) % at 19°C ambient temperature, with 95% confidence interval. The initial and subsequent enhancement of this dominance of the S state within the TRAAK ensemble upon temperature increase, suggests the S state has greater thermodynamic stability than the NS state. This could explain the overwhelming prevalence of the S state in previous structural reports for TRAAK in particular, and by expansion to TWIK1 and possibly other K2P channels.

To explore the effect of temperature on channel activity we recorded electrophysiology on TRAAK reconstituted in our GIUVs’ set up at 24 °C and 37 °C. Considering that cap domain swapping may be a multi-step slow process, we monitored TRAAK’s activity for over 1 hour at 37 °C and for shorter time periods at higher temperatures, due to patches being fragile and rupturing. We did not observe any channel activation at increasing temperatures over 37 °C (Supplementary Fig. 13). This is consistent with previous functional studies showing that TRAAK activation upon heating only occurs in a whole-cell configuration, but not in inside-out patches, suggesting that cell integrity is required for temperature sensitivity of TRAAK5,88.

To better understand the shift of the equilibrium towards the S state, we performed independent atomistic molecular dynamics simulations of the NS state and the S state of TRAAK embedded in lipid bilayers at 19 °C and 40 °C and calculated the energies between the channel and its entire environment (Supplementary Fig. 14a, 15, Supplementary Table 4). We first ensured systems in all four conditions are in equilibrium by monitoring the RMSD of the proteins through the trajectories (Supplementary Fig. 14b). This approach enabled a direct comparison of the non-bonded interaction energy between the protein (accounting for its distinct structural architecture) and its environment (lipid bilayer and solvent) at two distinct states and different temperatures. For the S state, the temperature dependence of the protein–environment interaction is relatively small (ΔΔG ≈ -979 ± 724 kcal mol⁻¹) and more favourable at 19 °C. The NS state exhibits a larger energy difference (ΔΔG ≈ -2689 ± 819 kcal mol⁻¹), also favouring 19 °C. Considering the fluctuations in the energy profiles and the resulting large errors (Supplementary Fig. 14a, Supplementary Table 4), S seems to be the most stable and the least temperature-sensitive state, whereas NS becomes less stable at elevated temperatures.

To further dissect the origin of the temperature-dependent interaction energies, we analysed individual membrane properties. For the NS state, the bilayer shows larger thickness fluctuations at 19 °C than at 40 °C (Supplementary Fig. 16), indicating stronger local deformations that allow better adaptation to the protein and more favourable protein–membrane interactions at the lower temperature. In contrast, thickness fluctuations in the S state appear similar at both temperatures. We also examined the distribution of negatively charged PS lipids in the inner leaflet for all trajectories (Supplementary Fig. 17). For the NS state, PS is enriched near the channel at 19 °C compared to 40 °C. This is evident in the extended PS-rich patches of the 2D maps and the higher value in the radial distribution function g(r) of PS around the protein (Supplementary Fig. 17). For the S state, the difference in g(r) between the two temperatures is small., indicating that the PS environment is weakly dependent on temperature (Supplementary Fig. 17).

Taken together, these results suggest that the stronger temperature sensitivity of the NS state arises from a combination of factors, including temperature-dependent PS (re)organisation around the channel and changes in local bilayer mechanics.

Methods
Cloning and expression

The human KCNK4 gene (K2P4.1b, residues 1-419, GenBank: AAK49390.1) with mutated surface-exposed cysteine residues (C146S, C206S, C218S, C335S, and C385S) was codon-optimised for expression in Pichia pastoris and synthesized by Integrated DNA Technologies. Subsequently, the gene was subcloned into a pPICZ-B expression vector (kindly provided by Dr. Helder Ferreira, University of St Andrews) using the BstBI/AgeI restriction sites. Following that, the gene encoding a His10 tag followed by cysteine-free GFP (cfGFP) (Addgene, plasmid number 54737) and a tobacco etch virus (TEV) with a short linker (SNS) were incorporated upstream of the K2P4.1b (C146S, C206S, C218S, C335S, and C385S) gene. This resulted in the generation of the construct His10-cfGFP-TEV-SNS-K2P4.1b (C146S, C206S, C218S, C335S, and C385S), referred to inCysTRAAK. The wild-type TRAAK was also codon-optimised and generated in an identical vector.

Mutations for the homodimer TRAAK were introduced into the inCysTRAAK construct through site-directed mutagenesis, using primer sequences provided in Supplementary Table 2. The His10 tag sequence of inCysTRAAK was replaced with a V5 tag sequence to facilitate purification of heterodimeric TRAAK mutants. Double mutant pairs—E67C_L144C and E70C_D93C—were then separately introduced into this modified construct, generating two new constructs: V5-cfGFP-TEV-SNS-K2P4.1b_E67C_L144C (C146S, C206S, C218S, C335S, and C385S), referred to V5-EL, and V5-cfGFP-TEV-SNS-K2P4.1b_E70C_D93C (C146S, C206S, C218S, C335S, and C385S), referred to V5-ED.

The vectors containing inCysTRAAK and V5-EL (or V5-ED) were linearized and amplified by PCR with the PmeI primers (Supplementary Table 2) and Q5® High-Fidelity DNA Polymerase (New England Biolabs). The linearized plasmids were transformed into Pichia pastoris (SMD1163) with similar ratio by electroporation and plated on a 1 mg/ml zeocin yeast extract peptone dextrose sorbitol (YPDS) plate for 3 days at 30 °C until colonies have grown.

Screening for multi-integrated genes was performed using a combination of both, fluorescent and Western blot analysis methods. On the first day, around 50 of the biggest colonies were selected and grown in 0.5 ml YPD media per colony overnight. Next day, each colony was pelleted down and the YPD media discarded before exchange to induction medium (2× yeast nitrogen base, 0.4 mg/l biotin, 40 mg/l L-histidine, 100 mM potassium phosphate pH 6.0, 0.5% (v/v) methanol) and further incubated for 8 hours at 28 °C.

After the induction, 200 μl of each colony were inoculated into a 96 well black plate. The fluorescence of each colony was detected under the EVOS fluorescence microscope (Thermofisher). The 6 brightest colonies were selected and normalized to the same amount of cell pellets to approximately 2 g. Those cell pellets were resuspended in around 1 ml PBS, combined with 600 mg of glass beads in 2 ml ground bottom tubes before beating with vortex for 3 minutes per cycle. Each colony was beaten for 5 cycles and kept on ice between each cycle for 5 mins. After lysis, the beads and cell debris were separated by centrifugation at 17,000 × g for 10 minutes. The supernatant of each colony was collected for two Western blots. The primary antibodies were anti-GFP (Antibodies, A283795) and anti-V5 tag (Antibodies, A269661) for each blot, and the secondary antibody was anti-rabbit HRP (Cambridge Bioscience, AC2114) for both blots. ECL detection reagent (Merck Life Science Limited, GERPN2105) was used, and the blots were detected using a Bio-Rad imager. Colonies that showed strong bands in both blots were selected for large-scale expression.

For the large-scale expression of heterodimer TRAAK, the selected colony was inoculated into 10 ml YPD media with 200 μg/ml zeocin and incubated overnight at 30°C and 200 rpm. Subsequently, the 10 ml overnight culture was transferred to a 2 l baffled flask containing 500 ml of YPD medium and grown for another 8 hours before inoculating into a 4 l minimal medium (2× yeast nitrogen base, 0.4 mg/l biotin, 40 mg/l L-histidine, 100 mM potassium phosphate pH 6.0.) with 1% glycerol. Cells were grown in a shaker at 30°C and 200 rpm for approximately 24 hours, reaching an OD600 of ~10. The cells were then harvested by centrifugation (3,000 × g at 20°C for 10 min) and resuspended in fresh minimal medium with 0.5% (v/v) methanol to induce protein expression. The cells were further incubated at a lower temperature of 28°C for another 48 hours. An additional final concentration of 0.5% methanol was added to the minimal culture at 24 hours after induction. Cell pellets were harvested by centrifugation (3,000 × g at 4°C for 30 min), snap-frozen in liquid nitrogen, and stored at -80°C until further use.

Isolation of genomic DNA and verification of the integrated gene sequence

The protocol for isolating genomic DNA was adapted from previous reports96. Briefly, a 10 mL culture of the selected colony was grown in the presence of 200 μg/ml zeocin overnight. A 200 μl aliquot of the liquid culture was spun down, and the supernatant was discarded. The pellet was resuspended in 100 μl of 200 mM lithium acetate with 1% SDS solution and incubated for 5 minutes at 70°C. After incubation, the supernatant was discarded, and 300 μl of 96–100% ethanol was added to the pellet and vortexed thoroughly. The pellet was spun down at 15,000 × g for 3 minutes and washed with 70% ethanol. After centrifugation to remove the ethanol, the pellet was dissolved in 100 μl of H₂O, and the solution was centrifuged to remove cell debris (15 seconds at 15,000 × g). A 2 μl aliquot of the supernatant was used as the template for PCR. The integrated gene fragment containing the V5 tag was amplified from the isolated genomic DNA using the primer pair "V5 tag fragment" listed in Supplementary Table 2. Similarly, the integrated gene fragment containing the His tag was amplified using the primer pair "His-tag fragment" listed in Supplementary Table 2.

Protein purification and site-directed spin labelling

The frozen cell pellets were disrupted by cryo-milling (Retsch model MM400) in 5 cycles of 3 minutes at 30 Hz and dissolved in the lysis buffer (50 mM Tris pH 8.0, 500 mM KCl, 0.1 mg/ml soy trypsin inhibitor, 1 mM benzamidine, 2 mM EDTA, 1 mM PMSF) at a ratio of 4 ml of lysis buffer per gram of cell pellet. After cryo-milling, centrifugation at 4696 × g at 4°C for 50 minutes was performed to remove cell debris and unbroken cells. The resulting supernatant underwent a further centrifugation step at 193,000 × g at 4°C for 1 hour to obtain membrane pellets. These membrane pellets were resuspended in a solubilisation buffer at a ratio of 1 g pellets per 2 ml solubilisation buffer (composed of 50 mM Tris pH 8.0, 500 mM KCl, protease inhibitor cocktail, 1 mM PMSF, 1 mM benzamidine, 0.1 mg/ml AEBSF). A final concentration of 2.5 mM MTSSL was added to the resuspended membrane pellets, and the mixture was incubated for 1 hour at room temperature. Subsequently, the spin-labelled membrane pellets were subjected to ultracentrifugation at 193,000 × g at 4°C for 1 hour, followed by a gentle rinse of the pellets with 50 ml of high salt buffer (50 mM Tris, 500 mM KCl). The labelled membrane pellets, resuspended in solubilisation buffer at a concentration of 80 mg/ml, were mixed with an equal volume of solubilisation buffer pre-dissolved with 5% (w/v) SMA co-polymer. This mixture was incubated at room temperature for 1.5 to 2 hours. For encapsulating proteins at 40°C, the membrane resuspended in solubilisation buffer was placed in a water bath at 40°C for 30 minutes at a concentration of 80 mg/ml. Following this, it was mixed with a pre-warmed solubilisation buffer containing 5% (w/v) SMA co-polymer and incubated for an additional 30 minutes in the water bath. The non-solubilized material was then separated by ultracentrifugation at 193,000 × g at 4°C for one hour.

The supernatant post-centrifugation was supplemented with a final concentration of 20 mM imidazole and incubated with 1.5 ml Ni2+-NTA beads at 4°C for 4 hours before loading onto a gravity-flow column. The beads were washed with 50 ml of wash buffer (50 mM Tris pH 8.0, 300 mM KCl, 30 mM imidazole) and eluted with elution buffer (50 mM Tris pH 8.0, 400 mM KCl, 300 mM imidazole). Finally, the elution was subjected to size-exclusion chromatography (SEC) using a Superose6 Increase 10/300 column (Cytiva) equilibrated with SEC buffer (50 mM Tris pH 8.0, 200 mM KCl). Collected fractions of protein were then concentrated to around 60 µM monomer concentration for subsequent EPR sample preparation. For proteins used for reconstitution, TRAAK in SMALPs was exchanged into DDM solution. SMALP-encapsulated TRAAK, following elution from SEC, was incubated with 1% DDM in SEC buffer for 30 minutes. A final concentration of 4 mM MgCl₂ was then added to the mixture (final buffer conditions: 50 mM Tris pH 8.0, 200 mM KCl, 1% DDM, 4 mM MgCl₂), followed by incubation at 4 °C for 2 hours to precipitate the SMA. After the incubation, the aggregated protein and precipitated SMA were pelleted by ultracentrifugation at 100,000 × g.

Continuous Wave (CW) EPR spectroscopy

The purified spin labelled sample was loaded to a 3 mm quartz EPR tube for measurement. All cwEPR experiments in solution were measured using a Bruker Magnettech EPR spectrometer ESR5000 operating at X-band and occupied with an integrated temperature controller. The sweep width was set from 330 mT to 345 mT, and the microwave power was set at 10 mW. The modulation frequency was set at 100 kHz, operating at 9.47 GHz frequency, and the modulation amplitude was set to 0.1 mT. The temperature was set to 4 °C to protect the sensitive TRAAK from temperature damage, during the multiple accumulation scans were being performed. 20-50 accumulation scans were collected per sample to acquire high quality signal-to-noise ratio spectra.

In silico spin labelling and distance modelling

Pymol-embedded MtsslWizard97 was used to analyse the labelling sites of TRAAK prior to mutagenesis. All amino acid residues of interest were mutated to cysteine using the Wizard function in PyMoL. MtsslWizard labelled the selected cysteines with the MTSSL spin labelled side chain R1. A "thorough search" was performed with "VDW restraints" set to "loose" by default. MtsslWizard calculates potential spin label rotamer positions, and an ensemble of 200 rotamer conformations (the maximum number of conformations that can be generated) is presented for each labelling PyMol-embedded site. The distance between two spin labels was calculated by selecting "Measure Mode". In this mode, two ensembles were selected and all possible spin-spin distance vectors between the two spin clusters were calculated. PDB entries 3UM7 (NS) and 4WFF (S) were used to generate the in silico distance distributions for each labelling site under different states.

PDS distance measurements and data analysis

PDS measurements were performed at an ELEXYS E580 EPR spectrometer from Bruker operating at Q-band (34 GHz) microwave frequency and equipped with a 3 mm cylindrical resonator (QT-II) and a cryogen-free variable temperature cryostat (Cryogenic Ltd, London, United Kingdom). Pulses were amplified by a traveling wave tube amplifier (150 W) (Applied Systems Engineering, United States).

For PDS measurements the 4-pulse π/2(ν )-τ -π(ν )-(τ + t)-π(ν )-(τ - t)-π(ν )-τ -echo sequence was used98. Measurements were performed at 50 K with the field position corresponding to the maximum of the field swept spectrum of the nitroxide. A frequency-offset (pump – detection frequency difference) of +80 MHz and a shot repetition time (SRT) of 2.5 ms were used. τ was set to 380 ns and τ was chosen as transverse dephasing allowed up to 7.5 µs. Pulse lengths were set to 12 ns, 16 ns, and 32 ns for π(ν ), π/2(ν ) and π(ν ), respectively, and optimised on maximum signal. Measurements were performed using an arbitrary waveform generator with a 16 step phase cycle99. The pump pulse was set to the maximum of the spectrum and resonance frequency of the resonator.

PDS data was analysed using DeerAnalysis2022100 in MATLAB (MathWorks, Natick, MA, United States) using the automated processing under default parameters. To estimate populations of the two distances a fit of 2 Gaussians was performed in DEERLab85.

For quantification of the two populations (NS and S state), PDS data of each sample were recorded with different dipolar evolution times and subjected to global fitting of multiple signals from different experiments in DeerLab using Tikhonov regularisation and strong robust generalised cross-validation for the selection of the regularisation parameter. We generally find that two time traces give very good results but as we had iteratively pushed the dipolar evolution to beyond 7 µs we had up to 4 traces per sample available and decided to include all of them into the global fit for not introducing bias by deciding which data to discard. Systematic omission of single traces in the global fits of 4 traces leads to no significant changes in the distributions confirming the self-consistency and robustness of this analysis. A 9 nm distance axis was chosen. Resulting non-parametric distance distributions were further subjected to fitting with a bimodal Gaussian model with peak widths limited to maximum 1 nm. The relative populations were derived from the weights (amplitudes) in the DeerLab fit reflecting the peak areas. These weights have uncertainties derived from the model fit in the range of 2-3%representing their 95% (2σ) confidence interval. However, by systematically varying processing parameters (distance axes of 8, 9 or 10 nm), we observed a slightly larger spread in results amounting to ~1/10th of the weight value of p1 over multiple samples. We thus, estimate the true uncertainty to be in this range.

The data analysis for Supplementary Fig. 8-11 was performed using Tikhonov regularisation as implemented in the MATLAB-based DeerAnalysis2022 package. Raw data were loaded into DeerAnalysis2022 and prior to analysis, data points at the end of the time trace were cut off to remove any “2+1” end artefacts56 and data were validated as previously described41.

Reconstitution in GUVs for electrophysiology

200 μl of a 10 mg/ml stock of 20% L-α-Phosphatidylcholine (PC) extracted from soybean (Avanti, 541601G-50g) in chloroform was transferred to a glass vial and dried to form a thin film under nitrogen gas. Subsequently, 500 μl of dehydration/rehydration (DR) buffer (5 mM HEPES, pH 7.2 (adjusted with KOH), and 200 mM KCl) was added to the lipid film, followed by bath sonication until the solution became transparent. A final concentration of 0.2% DDM (n-dodecyl-β-D-maltoside) was introduced to the solution and incubated at room temperature for 30 minutes. TRAAK protein was then added to the 2 mg lipids at a protein-to-lipids ratio of 1:100 (wt:wt) and incubated at 4°C for one hour. Approximately 500 mg of biobeads-SM2 adsorbents, previously washed with water and DR buffer, were added, and the reconstitution was rotated at 4°C overnight in a 1.5 ml Eppendorf tube. The following morning, the biobeads were separated, and the supernatant was spun down with a Ti55 rotor at 40,000 rpm for 1 hour at 4°C. The resulting pelleted proteoliposomes were resuspended in 80 μl of DR buffer, snap-frozen in 20 μl aliquots using liquid nitrogen and stored at -80°C for future use. On the day and immediately prior to the electrophysiology recordings, an aliquot was thawed at room temperature and pipetted onto a glass coverslip inside a petri dish. The petri dish was dried overnight in a desiccator at 4°C, followed by rehydration with 20 μl of DR buffer for 2 hours at room temperature. Afterward, 2 μl of the rehydrated proteoliposome were added to a 2 ml bath solution (5 mM HEPES, pH 7.2 (adjusted with KOH), 200 mM KCl, and 40 mM MgCl2) before the recording. A blister was formed immediately and lasted for around 5 hours.

Patch-clamp electrophysiology recordings

Patch pipettes made of thick-walled borosilicate glass capillaries (World Precision Instruments, 1B150F-6) were pulled to resistances of approximately 3–5 MΩ when filled with a pipette solution (5 mM HEPES, pH 7.2 (adjusted with NaOH), 180 mM NaCl, 20 mM KCl). Recordings were acquired using an Axopatch 200B amplifier (Molecular Devices) with an excised inside-out patch configuration. The currents were filtered at 1 kHz and digitized at 10 kHz using a Digidata 1440A (Molecular Devices). Negative pressures were applied through a high-speed pressure clamp (ALA Sciences, HSPC-1). Temperature above ambient was controlled using a perfusion temperature controller (Scientifica).

All recordings were conducted in a 10-fold concentration gradient of K+ and presented in physiological convention: positive currents indicate K+ flux from the high K+ concentration intracellular side (bath solution) to the low K+ concentration extracellular side.

Arachidonic acid (Sigma; 23401) was aliquoted at 50 mg/mL in ethanol and stored under argon at −20 °C until immediately before use. 16:0-18:1 PS (Avanti; 840034C), 18:0-20:4 PI(4,5)P₂ (Avanti; 850165), and L-α-phosphatidylinositol (Sigma; P6636) were aliquoted at 5 mg/mL in DMSO, and 18:1 PE (Sigma; 76548) was aliquoted at 3 mg/mL in DMSO. All lipids were sonicated until dissolved in DMSO prior to use.

Electrospray mass spectrometry lipidomic analysis

The lipids were extracted from ~1.1 mg of TRAAK protein that had been purified as described above using SMA and utilised for spin label experiments (One sample was analyzed). This was achieved by 3 successive vigorous extractions with ethanol (90% v/v)101. The pooled extracts were dried by nitrogen gas in a glass vial and re-extracted using a modified Bligh and Dyer method102. Extracts were dissolved in 15 μL of choloroform:methanol (1:2) and analysed on a Thermo Exploris Orbitrap mass spectrometer by direct infusion in both positive and negative ion modes. Lipid identity based upon accurate mass and MS/MS fragmentation, which was performed using nitrogen as the collision gas with collision energies between 35 and 90 V.

Atomistic molecular dynamics simulations and energy calculations

The PDB 3UM7 and PDB 4WFF were used as initial models for the NS and S states respectively. CHARMM-GUI’s Membrane Builder tool103-105 was used to construct the systems with bilayers containing DOPC, DOPE, and POPS lipid molecules. The NS-state membrane protein complex consisted of 332 lipids in total, whereas the S state contained 329 lipids. All simulations employed the CHARMM36 force field106-108. The systems were solvated using the TIP3P water model and neutralized with potassium and chloride ions at a concentration of 150 mM. The system composition and dimensions are summarized in Supplementary Table 3. Simulations were performed at two temperatures: 19 °C and 40 °C for each state. Energy minimization was carried out for 5,000 steps. Equilibration was conducted in six stages, with positional and dihedral restraints on proteins and lipids gradually released to zero in the final step. Proteins, lipids, and solvent were separately coupled to an external bath using the V-rescale thermostat109 with a coupling constant of 1.0. Semi-isotropic pressure coupling with a compressibility of 4.5×10−5 was applied using the stochastic cell-rescale method (Bernetti and Bussi, 2020). Our simulations were performed by combining the cut-off scheme with Force-switch for the Lenard-Jones, where the interactions are smoothly switched between rvdw switch at 1.0 nm and a cut-off at 1.2 nm. Coulomb interactions were calculated by using the cut-off type PME with a cut-off radius of 1.2 nm. Therefore, the interactions were calculated in real space within the cut-off and in Fourier space using PME after the cutoff. The systems were subjected to production runs of 300–500 ns, using 2 fs time step. All simulations were performed using GROMACS110 versions 2021.5 and 2024.5 at the CSF3 Manchester and the local Pliotas Lab workstations accelerated by GPUs. Energy profiles describing non-bonded interactions between the protein and its environment (membrane and solvent) were computed for each state and temperature. Protein, membrane, and solvent were defined as separate energy groups, and the gmx energy tool in GROMACS was used to extract interaction energies. Non-bonded interactions included both Lennard–Jones (LJ) and Coulomb electrostatic (CE) contributions. The approximate interaction energy for each trajectory was estimated by considering the ideal gas as the reference state, as expressed in Eq. 1:

∆𝐺 = 𝐺(𝑁,𝑝,𝑇)−𝐺 (𝑁,𝑝,𝑇) = 𝑘𝑇𝑙𝑛(⟨𝑒−𝑈⁄𝑘𝑇⟩) (1)
ideal gas

where U is the sum of the LJ and CE interactions energies. PV was calculated by multiplying the reference pressure of 1 bar with the volume giving a contribution of 93 to 96 kJ/mol for all simulations. This contribution is negligible compared to the energies given in Supplementary Table 4 and was therefore omitted. These calculated interactions represent the effective interaction energy of protein–membrane association. Due to long-timescale fluctuations in the interaction energies, strong energy drifts, a block-averaging scheme was applied in Eq. 1 to determine interaction energy values and associated errors. Differences in energies between simulations of the same state (with identical system composition) but at different temperatures were used to estimate the temperature dependence of the interaction energy change (Supplementary Table 4). Analysis was performed for the last half of the trajectories through the python scripts using the MDAnalysis package111,112. This included the inner leaflet resolved 2D density map for the PS lipids headgroups normalized by the mole fraction of the bulk in the membrane plane, the radial distribution function between the PS lipids in the leaflet and the protein, area per lipid molecule in each leaflet and bilayer thickness. Three independent simulation replicates were performed and analysed, and all resulted in consistent outcomes.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In their manuscript entitled “Swapped and non-swapped TRAAK states co-exist in membranes at a ratio influenced by temperature”, the authors demonstrate that these two cap conformations co-exist in the membrane and that their relative proportions are temperature-dependent, with the swapped state being the most prevalent. In addition, the authors present an interesting lipidomic analysis showing that signaling and negatively charged lipids selectively associate with TRAAK, whereas lipids with positively charged head groups, such as phosphatidylcholine, are excluded from the TRAAK membrane microdomain. The topic is of significant interest and will likely attract attention from the community. The authors have supported their experimental findings with structural data to strengthen their claims. However, the manuscript requires revisions before it can be considered for publication.\n\nMajor concerns\n1. Line 152. While Figure S1b is referenced to support the absence of MTSSL labelling in inCysTRAAK (EPR-silent homodimer), the manuscript does not appear to include a CW-EPR spectrum of the spin-labelled heterodimer (e.g., 67R1/144R1) as a positive control. Including such a spectrum would be valuable to demonstrate successful labelling and signal detection under the same conditions, and to allow a direct visual comparison with the negative control in Figure S1b.\n2. While the rainbow colour bars are defined in the figure legends, the manuscript does not discuss how distance-dependent reliability affects the interpretation of the cap state populations. Since the NS state (~3.3 nm) lies within the green “high confidence” zone, and the S state (~5 nm) lies in the less reliable yellow/orange zone, this may influence the precision of population quantification. A brief discussion of this technical limitation would help readers assess the robustness of the S/NS ratio estimation.\n3. In several figures (e.g., Fig. 3c, 3e, S6, and S7b), PDS time-domain traces are presented with varying numbers of curves. While Figures 3c and S6 display four traces, Figure 3e shows only two, and Figure S7b shows a single trace. The manuscript does not clearly explain the rationale behind these differences in the number of traces presented. Does the number of curves shown affect the accuracy or confidence in the resulting distance distributions?\n4. In Supplementary Figure S7d, the experimental distance distribution derived from the 70R1/93R1 spin pair appears to show only a single peak, despite the in silico models predicting distinct distances for the S and NS states. However, the main text states that two distance components are present (lines 288–289). Please clarify why the manuscript states that two populations are present when only a single peak is observed.\n5. A TRAAK concatamer in which Cys67 is introduced in one subunit and Cys144 in the other — that is, placing a single spin label on each subunit at different positions — would serve as a valuable control to distinguish intra-subunit from inter-subunit distance contributions in the PDS measurements.\n\nMinor concerns\n1. Lines 88 to 92: Please consider merging both sentences, as the one beginning with “By utilising a unique approach which combines Å-resolution PDS distance measurements...” appears to be grammatically incomplete and lacks a main clause.\n2. Line 104: Please include “phosphatidylcholine” before “PC,” as this is the first time the abbreviation is mentioned.\n3. Line 115: What do you mean by “species”? Wouldn’t “subfamilies” be more appropriate in this context?\n4. Line 152: You are citing Figure S1b, not the entire Figure S1.\n5. Line 163: Please include “around” before “2 nm,” as the predicted difference between the S and NS states is not exactly 2 nm.\n6. Table S1 must include the legend. Ej: What is DM? D-maltoside? Other abbreviations appear in the manuscript but must appear also in the Table to facilitate interpretation.\n7. The schematic in Figure 2a labels the spin-labelled subunit as “V5_GFP_TRAAK_L144C”, suggesting it contains only a single cysteine mutation. However, the main text and methods describe the use of a double mutant construct (E67C and L144C) for heterologous single-subunit spin labelling. For clarity and consistency, the figure label should be corrected to reflect both mutations (e.g., “V5_GFP_TRAAK_E67C_L144C” or “V5-EL”).\n8. In the legend of Figure 2e, the lipid species \"IPC 36:2;O3\" is listed, but the meaning of the \";O3\" suffix is not explained in the text or figure legend.\n9. In Figure 2e, the lipid species \"PIP3 42:6-10\" is listed, but the meaning of the final \"-10\" is not explained in the text or figure legend.\n10. Line 289: Which part of Figure S7 are you referring to? I assume it is S7d, but this is not clearly stated.\n11. In the main text (lines 350–363), you describe the temperature-induced shift in the TRAAK cap state distribution, but you do not cite the corresponding data shown in Figures 3e and 3f. Since these figures directly illustrate the experimental results at 40 °C that support your conclusions, we suggest explicitly citing them in this section to guide the reader to the relevant data\n12. Lines 407 to 410 list K2P subfamilies, not families — K2P is the family.\n13. From line 364 onward, the text shifts from describing results to interpreting them — discussing the biological relevance of the NS state, potential mechanisms for cap domain transitions, and physiological implications. However, this transition to the discussion section is not explicitly marked. For clarity, especially for readers scanning the manuscript for structure, we suggest including a subheading or transition sentence to clearly delineate the beginning of the discussion.\n14. From around line 455 onward, the manuscript revisits several points that were already discussed earlier in the Discussion section — including the energetic considerations for cap transitions, structural flexibility, and the potential regulatory role of the cap domain. These repetitions reduce the impact of the argument and may affect the overall clarity. We suggest revising this section to avoid redundancy and ensure a more concise and focused conclusion.\n15. Please check the formatting of “K2P” throughout the document — the “2” should be a subscript.", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "Two-pore domain potassium (K2P) channels generate the leak or background potassium current that sets the resting membrane potential and controls cell excitability. These channels assemble as dimers and feature a unique extracellular cap structure. Structural studies have revealed that this cap can adopt two distinct conformations: a non-swapped (NS) configuration, in which the outer transmembrane helix (TM1) interacts with the inner helix of the same subunit, and a swapped (S) configuration, where TM1 interacts with the inner helix of the adjacent subunit within the dimer.\n\nIn this study, the authors used pulsed EPR spectroscopy (PSD) to investigate the conformational dynamics of the human TRAAK channel cap. To do so, they first screened several pairs of spin-labelled cysteine residues and identified the 67R1/144R1 pair as providing the optimal resolution to discriminate between the S and NS states. They further developed a dedicated strategy, termed the HSS-SL protocol, to generate a uniform population of heterodimeric inCysTRAAK/inCysTRAAK channels labelled at these positions.\n\nPrior to the EPR measurements, the authors characterized the lipid composition of TRAAK’s immediate membrane environment using electrospray ionization mass spectrometry (ES-MS). They found that TRAAK resides in a lipid microdomain enriched in phosphatidylinositol (PI) and its phosphorylated derivatives (PIP1, PIP2, PIP3), as well as phosphatidylethanolamine (PE) and phosphatidic acid (PA), while phosphatidylcholine (PC) lipids are largely excluded.\n\nUsing PSD, they demonstrated that both the S and NS cap conformations coexist in TRAAK, with the S-state being predominant. Notably, they showed that the equilibrium between these two states is dynamic and can be modulated by temperature, suggesting that such conformational flexibility could have physiological relevance.\n\nWhile the study is technically sound and the methodological approach is innovative the physiological relevance is limited. Key experiments are required to strengthen the physiological relevance of the findings.\n\n1. In Fig. 2b, in order to validate the functionality of the constructs used for PSD measurements, it is essential to directly compare the electrophysiological properties of the mutants (including spin-labelled constructs) with the wild-type TRAAK channel. This control is critical to exclude potential artefacts or functional alterations induced by the mutations or spin labelling. Moreover, this control should actually be ideally carried out on the heteromeric tagged and spin-labelled constructs used for PSD.\n2. Furthermore, the authors should include a current-voltage (I-V) relationship, ideally obtained using a voltage ramp or step protocol, to provide a comprehensive characterization of the channel's biophysical properties. This would confirm the functional integrity of the constructs. Additionally, the authors should make sure the regulation of the mutant channels used is unaltered (sensitivity to arachidonic acid, temperature sensitivity mainly).\n3. While the PSD results clearly demonstrate that both the S and NS cap conformations coexist in TRAAK, and that their ratio can be modulated by temperature, the direct link between cap swapping and channel activity remains unclear. Specifically, it is not demonstrated that temperature-induced changes in cap conformation is related with an increase in current amplitude.\nTo further explore the functional relevance of cap swapping, the authors should consider testing gain-of-function mutations known to affect TRAAK gating as well as other stimuli such as arachidonic acid. If such mutations and stimulus alter the S/NS equilibrium, as determined by PSD, and this correlates with changes in channel activity, it would provide strong evidence supporting the physiological relevance of the cap conformational dynamics.\nMoreover, to further investigate that NS to S conformational transitions are involved in TRAAK thermal sensitivity, it would be interesting to compare the S to NS ratios of TRAAK at different temperatures over the thermal range to which TRAAK is sensitive (from 30°C to 40°C).\n4. Authors carry out an analysis of the nature of lipids surrounding TRAAK. However, they do not clearly expose the aim of this analysis (beyond knowing the lipid partners of TRAAK, which of itself is interesting but constitutes an article on its own) and to what extent it is important within the framework of the article (swapped, non-swapped conformations). Furthermore, authors should provide more in-depth interpretation of their analysis.\n\nMinor points:\n- “The equivalent of IPC in human membranes is sphingomyelin, which also has a choline headgroup, and would likely be undetected in our analysis.”\nWhy would it be undetected? The authors mention that PC lipids have previously been detected in similar experiments.\n- l.359: “The initial and subsequent enhancement of this dominance of the S state within the TRAAK ensemble upon temperature increase, suggests the S state has greater thermodynamic stability than the NS state.” Shouldn’t it be the reverse? Upon temperature increase, there is more energy in the system allowing more instable states to appear?\n- Authors speculate that either TRAAK adopts 2 distinct folding patterns during protein synthesis and/or that there is a conformational transition between the two states. But didn’t they demonstrate that the latter occurs with their experiment on temperature? (Channels analyzed at low and high temperatures were synthetized with the same protocol.) Authors could mention it.\n- l.406: Authors provide an estimate of the swapped to non-swapped conformation transition energy (10 kcal/mol). They should compare it to the energy gain of the system when temperature is increased from 19°C to 40°C in their experiment. Additionally, they could provide a comparison with the energy associated to temperature shifts to which TRAAK is sensitive. This way, the proposition that a mechanism involving cap swapping explains TRAAK thermal sensitivity would be more convincing.\n- The paragraph on heterodimers might be over-speculated at this point, in particular regarding this sentence: “By quantifying the proportional representation of cap states within these ensembles, we can determine each state’s weighted contribution to the channel’s functional output.”\n- In fig2.a, authors should mention the 2 mutations in the vector name: V5_GFP_TRAAK E67C L144C\n\nMistakes or points of clarification in the text:\n- l.156: what do authors mean by “homologous TRAAK sites”?\n- l.254: what does the “+ve” mean?\n- l.271: PIPS to PIPs\n- l.316: contributed to contributing?", "reviewer_4": "This manuscript probing the swapped (S) and non-swapped (NS) TRAAK potassium ion channel states using DEER spectroscopy is excellent. The research presented clearly indicates that both S and NS states coexist in membranes. The researchers noticed a shift in population to the swapped states at a higher temperature from 19 C to 40 C. The work presented in this study represents a clever way to get a complicated membrane protein sample that has a greatly increased DEER sensitivity for the state of the dimer. The DEER data shown in the paper is excellent. While a Tikhonov regularization is better in general for getting a sense of the most likely distance distribution, (there's no reason to assume that any distribution is purely Gaussian), using a dual gaussian fit is the best way to get a reasonably reliable measurement of the relative populations of the NS and S states (it's what everyone uses for this). I think that this is compelling paper for a Nature Communications publication on the TRAAK K+ channel. Also, the methodology could be broadly applied to a lot of biological and membrane systems.", "reviewer_5": "Potassium channels are oligomers of transmembrane subunits. The membrane-spanning domain of a multimeric channel is an assembly of alpha helices that interact with each other and with membrane bilayer lipids. The pore selectivity filter is formed by four reentrant domains which themselves interact with parts of the transmembrane helices. This assembly is highly stable - an essential condition for pore geometry and channel selectivity. Studies to date show that the oligomerization process takes place very early in channel biosynthesis in the endoplasmic reticulum, under the control of chaperone proteins. No studies have shown that the composition or overall conformational organization of the mature complex can be altered at the plasma membrane, probably because this would require too much energy. Under physiological conditions, very minor conformational changes modify the activity or selectivity of the channel and regulate its function.\n\nTwo-pore domain potassium (K2P) channels, including TRAAK, are dimers of subunits. Previous structural studies have shown that mature K2P channels can exist in two conformational states, the so-called non-swapped (NS) and swapped (S) forms. The two forms differ in the positions of the helices forming the extracellular cap and the membrane-spanning domains of each subunit in the dimer. The proportion of these two forms in cells and their functional significance are not known.\n\nIn this manuscript, Ma and colleagues expressed TRAAK in yeast, purified them and used pulse dipolar electron paramagnetic resonance spectrometry to calculate the proportion of N and NS forms. They also show that temperature influences this ratio suggesting that the previously reported effect of temperature on TRAAK could be due to a conversion between the N and NS forms. I’m not familiar with the technique used and the possible artifacts that it may produce, but the conclusions that TRAAK can switch between the N and NS states and that this switch has physiological significance are not supported by the provided data at this stage. Switching between the S and NS forms would require major conformational changes, with the unfolding and refolding of several domains and the modification of a considerable numbers of hydrophilic and hydrophobic interactions. No kinetics of this process are provided in the manuscript. If such rearrangement is possible after biosynthesis, then it is a multi-step process that should be slow. The effect of temperature on TRAAK/TREK1/TREK2 channel activity is rapid, as the effects of other stimuli (stretch, lipids, pH…). Also a 10-fold increase in TRAAK current is measured for a change from 17 to 40°C, while an increase of only 15% in the S form is observed. Nor is there any functional evidence that N and NS differ in their open probability or conductance, which would support the hypothesis of the authors.\n\nIn this work, the authors also took advantage of the way TRAAK is purified by SMA encapsulation to analyze the lipids surrounding the channel. While this is a promising approach, it has no bearing on the main message of the manuscript and its title. Much more work will be needed to show whether what is observed in yeast is relevant in mammalian cells for TRAAK function and/or whether other K2Ps behave differently or similarly to TRAAK with regard to the recruitment of surrounding lipids.\n\nIn conclusion, the authors have used novel and elegant approaches to study the structure of TRAAK and its lipid neighborhood. But this work is still too preliminary to draw any firm conclusions about a functional role of the N and NS states of K2P channels.", "reviewer_6": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response to the comments from the Reviewers\nWe sincerely appreciate the insightful and constructive comments from the Reviewers, which\nhave greatly helped us improve our manuscript. Below is our point-by-point response to the\ncomments. The comments from the Reviewers are shown in blue, our responses are in black,\nand the changes made in the revised manuscript are highlighted in yellow.\nReviewer #1 (Remarks to the Author):\nIn their manuscript entitled “Swapped and non-swapped TRAAK states co-exist in\nmembranes at a ratio influenced by temperature”, the authors demonstrate that these two cap\nconformations co-exist in the membrane and that their relative proportions are temperature-\ndependent, with the swapped state being the most prevalent. In addition, the authors present\nan interesting lipidomic analysis showing that signaling and negatively charged lipids\nselectively associate with TRAAK, whereas lipids with positively charged head groups, such\nas phosphatidylcholine, are excluded from the TRAAK membrane microdomain. The topic is\nof significant interest and will likely attract attention from the community. The authors have\nsupported their experimental findings with structural data to strengthen their claims.\nHowever, the manuscript requires revisions before it can be considered for publication.\nResponse:\nWe sincerely appreciate the positive comments made by Reviewer #1 and we are pleased that\nthe Reviewer believes that the community will find our study of significant interest and that it\nwill attract considerable attention.\nMajor concerns\n1. Line 152. While Figure S1b is referenced to support the absence of MTSSL labelling in\ninCysTRAAK (EPR-silent homodimer), the manuscript does not appear to include a CW-\nEPR spectrum of the spin-labelled heterodimer (e.g., 67R1/144R1) as a positive control.\nIncluding such a spectrum would be valuable to demonstrate successful labelling and signal\ndetection under the same conditions, and to allow a direct visual comparison with the\nnegative control in Figure S1b.\nResponse:\nWe agree with Reviewer 1 that the spectra of the spin-labelled heterodimeric mutants should\nbe also presented alongside the other spin-labelled homodimers. We have now included this\ndata in the new Supplementary Figure 6 in the revised manuscript (reproduced as Figure 1\nbelow). The data further confirm that these hetero-sites can be spin-labelled and also allow for\ndirect comparison between homo- and heteromeric labelled TRAAK mutants (i.e., 67R1,\n144R1 and 67R1/144R1).\nFigure 1: CW-EPR spectra of spin-labelled TRAAK homodimers and heterodimers in\nSMALPs. Continuous-wave (CW) EPR spectra recorded at 4 °C for various spin-labelled\nTRAAK constructs incorporated into SMALPs. Spectra correspond to the labelling sites\nindicated in the figure. R1 denotes the nitroxide side chain formed by the reaction of MTSSL\nwith engineered cysteine residues.\n2. While the rainbow colour bars are defined in the figure legends, the manuscript does not\ndiscuss how distance-dependent reliability affects the interpretation of the cap state\npopulations. Since the NS state (~3.3 nm) lies within the green “high confidence” zone, and\nthe S state (~5 nm) lies in the less reliable yellow/orange zone, this may influence the\nprecision of population quantification. A brief discussion of this technical limitation would\nhelp readers assess the robustness of the S/NS ratio estimation.\nResponse:\nThe Reviewer is correct to mention the technical limitation of the quantification of the cap\nstates. We have now added a brief discussion explaining the limits of our method. This will\nhelp the reader to follow the rationale of fitting the population with a bimodal Gaussian\nmodel.\nChange 1 on “Results”:\nChange 1 on “Methods”:\n3. In several figures (e.g., Fig. 3c, 3e, S6, and S7b), PDS time-domain traces are presented with\nvarying numbers of curves. While Figures 3c and S6 display four traces, Figure 3e shows only\ntwo, and Figure S7b shows a single trace. The manuscript does not clearly explain the rationale\nbehind these differences in the number of traces presented. Does the number of curves shown\naffect the accuracy or confidence in the resulting distance distributions?\nResponse:\nIdeally, what one would want is measuring a single PDS trace at sufficient length for resolving\nall populations (highest reliability range, green colour bar) and excellent signal-to-noise ratio\nfor processing with high confidence. A shorter trace with small time steps can resolve the\ndistance distribution for short distances whereas long times are required for reliable\nbackground extraction and reliable long-distance populations. Measuring a long trace with\nshort time steps and excellent signal-to-noise is experimentally not always feasible (with the\nexception of the sample shown in Figure S8 in the revised manuscript where it was not possible\nto resolve the two populations even though both are in the highest reliability range of the\ndistribution) and this has been demonstrated and discussed in literature (e.g., Rein et al. J.\nMagn. Reson. 2018, 295, 17-26 and Fábregas Ibáñez et al. Magn. Reson. 2020, 1, 209-224).\nWe have measured multiple traces with increasing time windows of most samples to increase\nconfidence that both cap states are indeed present.\nThus, our approach to this limitation was the use of global fitting of the data in DeerLab. The\nactual number of traces is dependent on the data we had available – for the repeat of the\n67R1/144R1 mutant (shown in Fig S7 in revised manuscript) we stayed close to the original\nsample (Fig 4c in revised manuscript). For the heated sample (Fig 4e in revised manuscript) it\nwas more challenging to measure at longer dipolar evolution times, thus we settled with two\ntraces. Generally, we find that two traces suffice but as we had more data, we did not want to\nbias the analysis by selecting which data to exclude from the analysis.\nAnalysing data for the samples with the 4 traces, leaving out one trace at a time, yielded\nessentially the same distributions.\nWe have added clarifying statements to the Results section of the manuscript (Change 1 on\n“Results”) and the Methods section of the SI (Change 1 on “Methods”).\n4. In Supplementary Figure S7d (Fig S8d in revised manuscript), the experimental distance\ndistribution derived from the 70R1/93R1 spin pair appears to show only a single peak,\ndespite the in silico models predicting distinct distances for the S and NS states. However, the\nmain text states that two distance components are present (lines 288–289). Please clarify why\nthe manuscript states that two populations are present when only a single peak is observed.\nResponse:\nWe thank the Reviewer for pointing this out. We have now revised the wording in the main text\naccordingly:\nChange 2 on “Results”:\n5. A TRAAK concatamer in which Cys67 is introduced in one subunit and Cys144 in the\nother — that is, placing a single spin label on each subunit at different positions — would\nserve as a valuable control to distinguish intra-subunit from inter-subunit distance\ncontributions in the PDS measurements.\nResponse:\nWe appreciate this suggestion, and it is indeed a valuable point. However, constructing a new\nconcatemer is not straightforward, as it requires optimisation of the linker between the two\nsubunits. An inappropriate linker may introduce artifacts that could alter the native\nconformation of TRAAK and is out of the scope of this study. Nonetheless, we will consider\nimplementing this approach in a future study.\nMinor concerns\n1. Lines 88 to 92: Please consider merging both sentences, as the one beginning with “By\nutilising a unique approach which combines Å-resolution PDS distance measurements...”\nappears to be grammatically incomplete and lacks a main clause.\nChange 3 on “Results”:\n2. Line 104: Please include “phosphatidylcholine” before “PC,” as this is the first time the\nabbreviation is mentioned.\nCorrected\n3. Line 115: What do you mean by “species”? Wouldn’t “subfamilies” be more appropriate in\nthis context?\nCorrected\n4. Line 152: You are citing Figure S1b, not the entire Figure S1.\nCorrected\n5. Line 163: Please include “around” before “2 nm,” as the predicted difference between the\nS and NS states is not exactly 2 nm.\nCorrected\n6. Table S1 must include the legend. Ej: What is DM? D-maltoside? Other abbreviations\nappear in the manuscript but must appear also in the Table to facilitate interpretation.\nCorrected\n7. The schematic in Figure 2a labels the spin-labelled subunit as\n“V5_GFP_TRAAK_L144C”, suggesting it contains only a single cysteine mutation.\nHowever, the main text and methods describe the use of a double mutant construct (E67C and\nL144C) for heterologous single-subunit spin labelling. For clarity and consistency, the figure\nlabel should be corrected to reflect both mutations (e.g., “V5_GFP_TRAAK_E67C_L144C”\nor “V5-EL”).\nCorrected\n8. In the legend of Figure 2e, the lipid species \"IPC 36:2;O3\" is listed, but the meaning of the\n\";O3\" suffix is not explained in the text or figure legend.\nCorrected\n9. In Figure 2e, the lipid species \"PIP3 42:6-10\" is listed, but the meaning of the final \"-10\" is\nnot explained in the text or figure legend.\nCorrected\n10. Line 289: Which part of Figure S7 are you referring to? I assume it is S7d, but this is not\nclearly stated.\nCorrected\n11. In the main text (lines 350–363), you describe the temperature-induced shift in the\nTRAAK cap state distribution, but you do not cite the corresponding data shown in Figures\n3e and 3f. Since these figures directly illustrate the experimental results at 40 °C that support\nyour conclusions, we suggest explicitly citing them in this section to guide the reader to the\nrelevant data\nCorrected\n12. Lines 407 to 410 list K2P subfamilies, not families — K2P is the family.\nCorrected\n13. From line 364 onward, the text shifts from describing results to interpreting them —\ndiscussing the biological relevance of the NS state, potential mechanisms for cap domain\ntransitions, and physiological implications. However, this transition to the discussion section\nis not explicitly marked. For clarity, especially for readers scanning the manuscript for\nstructure, we suggest including a subheading or transition sentence to clearly delineate the\nbeginning of the discussion.\nCorrected, we added “Discussion” as a subheading.\n14. From around line 455 onward, the manuscript revisits several points that were already\ndiscussed earlier in the Discussion section — including the energetic considerations for cap\ntransitions, structural flexibility, and the potential regulatory role of the cap domain. These\nrepetitions reduce the impact of the argument and may affect the overall clarity. We suggest\nrevising this section to avoid redundancy and ensure a more concise and focused conclusion.\nWe thank the Reviewer for their suggestions. In response, we have removed the section on\nenergetic considerations (now replaced by our more robust MD energy calculations), and\neliminated repetitive text to ensure our main messages remain focused and undiluted.\n15. Please check the formatting of “K2P” throughout the document — the “2” should be a\nsubscript.\nBoth K2P and K P abbreviations are widely accepted in literature, and we use “K2P”\n2\nthroughout our manuscript.\nReviewer 2:\nThis study characterizes the populations of the swapped and nonswapped conformational states\nof the TRAAK channel cap in a near native lipid environment, using cysteine-based spin\nlabeling. Additionally, the authors examine the temperature dependence of these\nconformational states and carry out a lipidomic analysis of lipids in close proximity to TRAAK\nchannels. The research addresses multiple relevant aspects, and it is laudable that the authors\npresent a comprehensive and integrative study, combining both experimental and theoretical\napproaches. There are, however, several points that could be further clarified or expanded upon\nto strengthen the study:\nMajor concerns\n1. Lines 295, 296: It is not clear how it is claimed that the SMA used forms 10 nm lipid disks.\nWhile Postis et al. (2015) cite a maximal diameter of ~15 nm and Lee et al. (2016) provide no\nsize data, both ultimately reference Jamshad et al. (2014) (not directly quoted in your text),\nwhich reports diameters: ~9.8 and 11–16 nm by different techniques. Please mention the source\nof the value mentioned (e.g., your own measurements, Jamshad data or estimates). This point\nis key, because it will provide further evidence for your interpretations of the PDS distances\nfor intermolecular interactions.\nResponse:\nWe thank the Reviewer for their point and agree that the size of the SMA lipid discs is\ninherently variable. Previous studies highlight how the SMA copolymer effectively solubilizes\nmembranes and can create discoidal assemblies of varying sizes, with measurements typically\nranging from 10 nm to 16 nm in diameter, depending on the lipid environment and the presence\nof proteins (Long et al., 2013; Jamshad et al., 2015). This variability underscores the\nadaptability of the SMA discs to different lipid compositions.\nThe SMA-based nanodiscs can support various native lipid-protein interactions, thereby\npreserving their functional state and activity. The adaptability of SMA disks can thus be\nleveraged to optimize conditions for studying such protein-lipid interactions, ensuring the\nphysiological relevance of these environments is maintained (Dörr et al., 2014). Moreover,\neven for the larger SMA diameter reported of 16 nm two TRAAK channels cannot be\naccommodated in a single SMA disc (i.e., TRAAK has a diameter of ~8 nm, Brohawn et al.,\nScience, 2012), thus diminishing any unspecific inter-dimer distance contributions from within\nthe same membrane disc, which could obscure our PDS experiments. In our study, we further\nensured that two TRAAK channels cannot be inserted in a single SMA disc as we selected only\nthe TRAAK fractions consistent with the purified monodisperse peak of a single dimeric\nTRAAK disc size following size exclusion chromatography, and before subjecting to\nsubsequent EPR experiments (i.e., CW and PDS).\nWe now made a change in the revised manuscript to reflect these points.\nChange 4 on “Results”:\nReferences:\nDörr, J., Koorengevel, M., Schäfer, M., Prokofyev, A., Scheidelaar, S., Cruijsen, E., … &\nKillian, J. (2014). Detergent-free isolation, characterization, and functional reconstitution of a\ntetrameric k+ channel: the power of native nanodiscs. Proceedings of the National Academy of\nSciences, 111(52), 18607-18612. https://doi.org/10.1073/pnas.1416205112 [doi.org]\nJamshad M, Grimard V, Idini I, Knowles TJ, Dowle MR, Schofield N, Sridhar P, Lin YP, Finka\nR, Wheatley M, Thomas OR, Palmer RE, Overduin M, Govaerts C, Ruysschaert JM, Edler KJ,\nDafforn TR. Structural analysis of a nanoparticle containing a lipid bilayer used for detergent-\nfree extraction of membrane proteins. Nano Res. 2015 Mar;8(3):774-789. doi:\n10.1007/s12274-014-0560-6. Epub 2014 Oct 23. PMID: 31031888; PMCID: PMC6485620.\nLong, A., O'Brien, C., Malhotra, K., Schwall, C., Albert, A., Watts, A., … & Alder, N. (2013).\nA detergent-free strategy for the reconstitution of active enzyme complexes from native\nbiological membranes into nanoscale discs. BMC Biotechnology,\n13(1). https://doi.org/10.1186/1472-6750-13-41 [doi.org]\n2. It is recommended to include the crystallographic structures used in the in silico analysis\n(e.g. TRAAK 3UM7 for the NS state and 4WFF for S state) to allow replicability. Also, it\nwould be valuable to specify the number of conformations generated (n, sample size) to\nconstruct the distance distributions, and to report descriptive statistics (mean and standard\ndeviation) directly in the text. Also, the distributions presented have a perfect Gaussian curve,\nperhaps you used some additional adjustment that should be mentioned. This would help to\nreinforce the transparency and statistical robustness of the in-silico results presented.\nResponse:\nWe appreciate the suggestions and have now added a descriptive sentence in the Methods\nsection to indicate the crystallographic structures used for the in silico analysis and the number\nof conformations generated. Mean distances of our in silico models are also included in Figure\n1b and c. Both the mean distance and the SD are provided in the main text. Regarding the\ndetails of obtaining the bimodal Gaussian model, we have added clarifying statements to the\nMethods section of the SI.\nChange 1 on “Methods”:\nChange 2 on “Methods”:\nChange 5 on “Results”:\nReviewer 3:\nTwo-pore domain potassium (K2P) channels generate the leak or background potassium\ncurrent that sets the resting membrane potential and controls cell excitability. These channels\nassemble as dimers and feature a unique extracellular cap structure. Structural studies have\nrevealed that this cap can adopt two distinct conformations: a non-swapped (NS) configuration,\nin which the outer transmembrane helix (TM1) interacts with the inner helix of the same\nsubunit, and a swapped (S) configuration, where TM1 interacts with the inner helix of the\nadjacent subunit within the dimer.\nIn this study, the authors used pulsed EPR spectroscopy (PDS) to investigate the\nconformational dynamics of the human TRAAK channel cap. To do so, they first screened\nseveral pairs of spin-labelled cysteine residues and identified the 67R1/144R1 pair as providing\nthe optimal resolution to discriminate between the S and NS states. They further developed a\ndedicated strategy, termed the HSS-SL protocol, to generate a uniform population of\nheterodimeric inCysTRAAK/inCysTRAAK channels labelled at these positions.\nPrior to the EPR measurements, the authors characterized the lipid composition of TRAAK’s\nimmediate membrane environment using electrospray ionization mass spectrometry (ES-MS).\nThey found that TRAAK resides in a lipid microdomain enriched in phosphatidylinositol (PI)\nand its phosphorylated derivatives (PIP1, PIP2, PIP3), as well as phosphatidylethanolamine\n(PE) and phosphatidic acid (PA), while phosphatidylcholine (PC) lipids are largely excluded.\nUsing PDS, they demonstrated that both the S and NS cap conformations coexist in TRAAK,\nwith the S-state being predominant. Notably, they showed that the equilibrium between these\ntwo states is dynamic and can be modulated by temperature, suggesting that such\nconformational flexibility could have physiological relevance.\nWhile the study is technically sound and the methodological approach is innovative the\nphysiological relevance is limited. Key experiments are required to strengthen the\nphysiological relevance of the findings.\nResponse:\nWe sincerely appreciate Reviewer 3 recognising the innovative aspects of our method. We\nhave now performed additional electrophysiology experiments and include in the revised\nmanuscript to better understand channel function and contribute to knowledge towards the\nphysiological relevance of our PDS findings. These new data are consistent with our initial\nconclusions and shed light into the role of native membrane lipids (i.e., identified by our ES-\nMS analysis) into TRAAK’s function. We do hope that these new functional observations\nwill satisfy the Reviewer and we believe these further strengthen our study.\n1. In Fig. 2b, in order to validate the functionality of the constructs used for PDS measurements,\nit is essential to directly compare the electrophysiological properties of the mutants (including\nspin-labelled constructs) with the wild-type TRAAK channel. This control is critical to exclude\npotential artefacts or functional alterations induced by the mutations or spin labelling.\nMoreover, this control should actually be ideally carried out on the heteromeric tagged and\nspin-labelled constructs used for PDS.\n2.Furthermore, the authors should include a current-voltage (I-V) relationship, ideally obtained\nusing a voltage ramp or step protocol, to provide a comprehensive characterization of the\nchannel's biophysical properties. This would confirm the functional integrity of the constructs.\nAdditionally, the authors should make sure the regulation of the mutant channels used is\nunaltered (sensitivity to arachidonic acid, temperature sensitivity mainly).\nResponse to 1 &2:\nWe have now generated a new construct of the wild-type (WT) TRAAK channel using the\nsame backbone as that of the other mutants employed in the PDS experiments. WT TRAAK\nwas measured under the same experimental conditions, as for all mutants described in our\noriginal manuscript. Furthermore, we remeasured all (including new) constructs using the\nvoltage step protocol and assessed their sensitivity to arachidonic acid. The updated results are\npresented in revised Figure S1 (reproduced as Figure 2 below), along with a statistical\ncomparison of channel sensitivity to arachidonic acid and mechanosensitivity including wild-\ntype (WT) TRAAK, InCys TRAAK, and E67C/L144C TRAAK, shown in Figure 2. These data\nsupport that none of the introduced mutations alter channel functionality. Additionally, we\ncompared the currents of E67C/L144C TRAAK before and after MTSSL spin labelling (i.e.,\nE67R1/L144R1) and observed no significant differences, indicating that spin labelling at\npositions 67 and 144 again does not have any effect on ion channel function (panel b-g in the\nrevised Figure 2, reproduced as Figure 3 below).\nRegarding temperature sensitivity, we recently purchased and installed a new temperature\ncontroller along with new high flow rate pumps integrated to our existing electrophysiology\nset up to enable stable and accurate current recordings at distinct temperatures over extended\nperiods of time (i.e., up to one hour). Using our new setup, we tested 3 × replicates of WT\nTRAAK and observed no significant changes in channel activity when the temperature was\nincreased from 24 °C to 41 °C under the inside-out patch clamp configuration. We also\nmonitored channel activity continuously for 1 h at 37 °C (i.e. recordings above 37 °C were only\npossible for shorter time periods due to GUV patches being fragile and rupturing). Channel\ncurrent did not increase with temperature, regardless of the incubation time (revised Figure\nS13, reproduced as Figure 4 below). Our observation is consistent with previous studies,\nwhich showed that TRAAK activation upon heating only occurs in a whole-cell configuration but not in\ninside-out patches, suggesting that cell integrity is required for temperature activation of\nTRAAK in an electrophysiology set up (Kang et al., 2005; Maingret et al., 2000).\nFigure. 2: Comparison of channel activity for wild-type, InCys, and E67C/L144C TRAAK.\na, g, Current responses to incremental negative pressure applied to representative excised\npatches from wild-type (a) and InCys (g) TRAAK GUVs. Pressure steps every 5 s; traces\nvertically offset. Dashed red lines indicate the zero-current baseline. Holding potential (Vh) =\n0 mV, holding pressure (Ph) = 0 mmHg. b, h, Currents recorded from patches of wild-type (b)\nand InCys (h) TRAAK during a voltage-step protocol (Vh=−50mV ; −100 to +100 mV in 10-\nmV increments; displayed every 40 mV). For each voltage step, a −50 mmHg pressure step\nwas applied (lower). Dashed red line marks the current at Vh=−50 mV, Ph=0 mmHg. c, I, I–V\nrelationships from (b, h): average pre-pressure current and peak current during the pressure\nstep vs voltage. d, j, m, Currents from excised patches of wild-type (d), InCys (j), and\nE67C/L144C (m) under a voltage-step protocol (Vh=0 mV; −100 to +100 mV;ΔV=10 mV;\ndisplayed every 40 mV). e, k, n, Recordings from the same patches as (d, j, m) after 50 µM\narachidonic acid (AA) perfusion. f, l, o, I–V relationships corresponding to (d/e), (j/k), and\n(m/n), respectively.p,q, Fold change in current at Vh = 0 mV for wild-type, InCys, and\nE67C/L144C TRAAK in response to MS (−50 mmHg) and AA stimulation. MS increased\ncurrent 1.66 ± 0.15, 1.54 ± 0.10, and 1.60 ± 0.04-fold, respectively; AA increased current 1.47\n± 0.13, 1.69 ± 0.06, and 1.45 ± 0.14-fold, respectively (mean ± SEM). Biological replicate\nnumbers (n) are shown above the bars. r, Fold change in current at Vh = 0 mV of wild-type\nTRAAK in response to lipid perfusion. TRAAK was activated 1.47 ± 0.13, 1.58 ± 0.02, and\n1.51 ± 0.03-fold by AA, PI, and PI(4,5)P2, respectively (mean ± SEM, n = 3). In contrast,\nDOPE inhibited TRAAK by 2.00 ± 0.25-fold (mean ± SEM, n = 3).\nFigure. 3: b, Current response to pressure increments applied to patch excised from TRAAK\nE67C/L144C GUVs. Pressure steps were incremented every 5 sec, and the currents measured\nduring each step are vertically offset for display. Dashed red lines below each current trace\ndenote the zero current baseline. Holding potential (Vh) = 0 mV, holding pressure (Ph) = 0\nmmHg. c, Recorded currents (Upper) obtained from a patch extracted from TRAAK\nE67C/L144C GUVs during a voltage step protocol (Vh = −50 mV, ranging from −100 to +100\nmV, with a ΔV of 10 mV, displayed at intervals of 40 mV). For each voltage step, a pressure\nstep of −50 mmHg was applied through the pipette (Lower). Dashed red line indicates the\ncurrent level at the holding potential of -50 mV and holding pressure of 0 mmHg. d, The\ncurrent-voltage relationship of the data presented in c is shown at 10 mV intervals. The average\ncurrent prior to pressure step and the peak current during the pressure step application are\nplotted against each voltage. e, f, Currents from excised patches of TRAAK E67C/L144C\nGUVs under a voltage step protocol (Vh = 0 mV; −100 to +100 mV in 10 mV increments;\nevery 40 mV step shown), before (e) and after (f) perfusion with 1 μM MTSSL. g, Current–\nvoltage relationship from the data shown in e and f, incremented by 10 mV. The black trace\nrepresents the average current before MTSSL application, and the red trace (+MTSSL)\nrepresents the current after perfusion with 1 μM MTSSL.\nFigure 4: TRAAK is not activated by heating in the inside-out patch configuration. a,\nCurrents from an excised inside-out patch of wild-type TRAAK recorded with a voltage-step\nprotocol at ambient temperature (24 °C; Vh=0 mV; −100 to +100 mV;ΔV=10 mV; displayed\nevery 40 mV). b, c, Recordings from the same patch after the bath was raised to 37 °C (~15\nmin) and after 1 h at 37 °C, respectively. d, I–V relationships from the same patch under the\nthree conditions.\n3. While the PDS results clearly demonstrate that both the S and NS cap conformations coexist\nin TRAAK, and that their ratio can be modulated by temperature, the direct link between cap\nswapping and channel activity remains unclear. Specifically, it is not demonstrated that\ntemperature-induced changes in cap conformation is related with an increase in current\namplitude.\nTo further explore the functional relevance of cap swapping, the authors should consider testing\ngain-of-function mutations known to affect TRAAK gating as well as other stimuli such as\narachidonic acid. If such mutations and stimulus alter the S/NS equilibrium, as determined by\nPDS, and this correlates with changes in channel activity, it would provide strong evidence\nsupporting the physiological relevance of the cap conformational dynamics. Moreover, to\nfurther investigate that NS to S conformational transitions are involved in TRAAK thermal\nsensitivity, it would be interesting to compare the S to NS ratios of TRAAK at different\ntemperatures over the thermal range to which TRAAK is sensitive (from 30°C to 40°C).\nResponse:\nWe agree that establishing a direct link between cap swapping and channel activity is primarily\nlimited by technical constraints, which was also the case for monitoring the entire cap ensemble\nof TRAAK in lipid membranes with high resolution prior to our study. First, as per our response\nabove, activation of TRAAK by heating requires cell integrity (Maingret, F. et al. EMBO J 19,\n2483-2491, 2000; Kang, D. et al. J Physiol 564, 103-116, 2005), which is absent in our current\nsetup using reconstituted liposomes. Second, although PDS is a powerful technique that allows\nus to monitor the conformational landscape of the entire TRAAK population for the first time-\nincluding the minority-populated and so far neglected NS state- we are currently unable to\nseparate the non-swapped (NS) from the swapped (S) TRAAK populations. Even if whole-cell\npatch-clamp recordings were available, we would still lack control over the exact proportion\nof TRAAK molecules present in each cap conformation within an individual recorded patch.\nThus, establishing a direct link between domain swapping and channel activity will require an\nadditional breakthrough that enables separation of the two cap states. To note that even if\ncurrently available state-of-the-art antibodies are used to select for the S state in combination\nwith PDS, still quantifying the entire ensemble will not be possible, while additionally using\nstate-locking antibodies will not allow for unbiased monitoring of the entire ensemble. In this\nstudy, we provide a robust PDS-based method to detect and quantify the conformational states\nof TRAAK in the ensemble, representing an important first step toward investigating domain\nswapping in TRAAK, which could be also generalised for other K2P channels\nmechanosensitive or not (TREK1/2, TWIK1, TASK et al). One potential approach could be\nscreening for a large number of conditions (and out of the scope of this study) using our current\ncombined PDS and HSS-SL approach which may force the entire TRAAK population to a\nsingle species (i.e. ~100% S or NS). We would then be able to study the function (and structure)\nof this species separately and account for its individual contribution to TRAAK’s activity.\nRegarding arachidonic acid (AA) activation: We appreciate the valuable suggestion by the\nReviewer and we have now performed additional PDS measurements in the presence of AA to\nexamine whether there is a change in the S/NS equilibrium. However, the PDS data obtained\nfrom AA-treated spin labelled TRAAK samples were of insufficient quality (i.e. low SNR and\nvery low EPR signal) to allow for reliable conclusions regarding its effect on the S/NS state.\nTo investigate the origin of the poor EPR data quality after AA was added to heterologously\nspin-labelled TRAAK, we directly incubated the spin label MTSSL with AA and monitored the\nMTSSL signal by CW-EPR at room temperature and in solution. Surprisingly, we observed\nthat AA progressively reduces the MTSSL signal over time until it completely diminishes after\nthe course of few minutes (Figure 5).\nTherefore, it seems that the poor quality of the PDS data and the substantially reduced cwEPR\nsignal over AA incubation is due to the chemical properties of AA. Indeed, it has been reported\nthat the bis-allylic groups in AA are prone to oxidation because their electrons are pulled by\nadjacent double bonds, weakening their bond energy (Johnson, D. R.et, al. 2015). As a\nconsequence, the diminishing EPR signal may reflect AA-induced reduction of the spin-label\nfree radicals.\nFigure 5: Time-dependent reduction of CW-EPR signal upon incubation of MTSSL with\narachidonic acid (AA). (a) Continuous monitoring of the maximum intensity amplitude of\nthe central line peak in the CW-EPR spectra during incubation of 17 µM MTSSL with 6 mM\nAA shows a progressive decrease in signal over time. (b) Representative CW-EPR spectra\nfrom panel (a), recorded at three distinct time points, illustrate the decay in signal amplitude\nas incubation time increases.\nReference:\nJohnson, D. R. & Decker, E. A. The Role of Oxygen in Lipid Oxidation Reactions: A Review.\nAnnual review of food science and technology. 6, 171-190, doi:10.1146/annurev-food-022814-\n015532 (2015).\n4. Authors carry out an analysis of the nature of lipids surrounding TRAAK. However, they do\nnot clearly expose the aim of this analysis (beyond knowing the lipid partners of TRAAK,\nwhich of itself is interesting but constitutes an article on its own) and to what extent it is\nimportant within the framework of the article (swapped, non-swapped conformations).\nFurthermore, authors should provide more in-depth interpretation of their analysis.\nResponse:\nWe thank the Reviewer for the thoughtful comment. We now implemented additional\nelectrophysiology experiments and analysis using the native lipids (i.e. identified by our ES-\nMS analysis) to account for their functional role. These data further support our conclusions\nand contribute to its completeness, as per Reviewer’s suggestion to explore the functional role\nof the identified lipids. Our initial aim in analysing the immediate lipid environment of TRAAK\nwas to demonstrate the combined use of ES-MS with SMA detergent-free technology to study\nTRAAK in a native-like lipid environment.\nOur ES-MS data revealed that TRAAK exhibits a preference for specific lipids which could\nhave a functional role, an important aspect we now explore in our revised manuscript. To this\nend, we performed new multiple electrophysiology recordings of GUV-reconstituted TRAAK\nin presence of a) POPS, b) DOPE, c) PI(4,5)P , and d) PI lipids, identified in our previous ES-\nMS analysis to assess the functional effect each of these lipids have on TRAAK function.\nAs shown in panel b-e from revised Figure 3, (reproduced below as Figure 6), we found that\nin our set up both PI(4,5)P and PI can activate TRAAK, while in contrast DOPE exerts an\ninhibitory effect. Finally, POPS does not significantly affect TRAAK’s current amplitude,\nsuggesting that this lipid likely plays a structural rather than regulatory role, possibly\ncontributing to the stability of TRAAK within the lipid bilayer.\nFig. 6: TRAAK associates with specific membrane lipids and is modulated by lipid\ncomposition in reconstituted membranes. b–e, Representative currents from excised patches\nof wild-type TRAAK reconstituted in GUVs during a voltage-step protocol (Vh = 0 mV; steps\nfrom −100 to +100 mV in 10 mV increments; every 40 mV step shown). Left and middle panels\nshow recordings before and after bath perfusion with 4 µM of the indicated lipid: PI (b), PIP\n2\n(c), DOPE (d), and POPS (e). Right panels: I–V relationships from the same patches at 10-mV\nincrements. Black traces indicate average currents prior to lipid application, while red traces\n(+PI, +PIP₂, +DOPE and +POPS) show the response following lipid perfusion.\nMinor points:\n- “The equivalent of IPC in human membranes is sphingomyelin, which also has a choline\nheadgroup, and would likely be undetected in our analysis.”\nWhy would it be undetected? The authors mention that PC lipids have previously been\ndetected in similar experiments.\nResponse:\nWe thank the Reviewer for this observation. We cited several previous studies demonstrating\nthat PC lipids can indeed be detected in SMA-derived lipid particles to clarify that the absence\nof PC in our analysis is not due to any known artifact or selective exclusion by the SMA\npolymer itself. Yeast do not make sphingomyelin, they make IPC instead. Therefore, the lack\nof PC detection in our study reflects a biological exclusion of PC from TRAAK's immediate\nlipid environment. This is particularly notable given that PC is the most abundant lipid species\nin Pichia pastoris membranes, which we used for protein expression. Based on this, we propose\nthat sphingomyelin, another choline-containing lipid and the functional equivalent of IPC in\nhuman membranes, is also likely absent from the immediate environment of TRAAK and thus\nwas not detected in our analysis.\n- l.359: “The initial and subsequent enhancement of this dominance of the S state within the\nTRAAK ensemble upon temperature increase, suggests the S state has greater thermodynamic\nstability than the NS state.” Shouldn’t it be the reverse? Upon temperature increase, there is\nmore energy in the system allowing more instable states to appear?\nResponse:\nWe thank the Reviewer for their comment and agree with their assessment regarding low\nenergy states. This is indeed a quite critical point in our study, which we agree that needs\nclarification and requires further analysis. Because the S state is the dominant state at both\nlow and high temperatures, we concluded that the S state has greater thermodynamic stability\nthan the NS state.\nHowever, to better understand the equilibrium shift caused by the higher temperature and\naddress this specific point raised by the Reviewer, we teamed up with Dr Qaiser Waheed\n(now a new co-author in the revised manuscript and an expert in MD simulations) and\nperformed long fully atomistic MD simulations of both the non-swapped (NS; PDB 3UM7)\nand swapped (S; PDB 4WFF) states at 19 °C and 40 °C. We then compared the non-bonded\ninteraction energies between the protein and its entire environment (lipid bilayer and solvent).\nThe results are shown in the manuscript as:\nThe MD data and energy calculations are shown in revised Figure S14 (reproduced below as\nFigure 7) and revised Table S4 (reproduced below as Table 1).\nFigure 7: Molecular dynamics energy calculations of the S and NS TRAAK states at two\ndifferent temperatures. Time traces of (a) nonbonded interaction energies (Lennard–Jones\nand Coulomb electrostatic contributions) and (b) backbone RMSD between membrane-\nembedded TRAAK and its environment, compared across temperatures and channel states.\nTable 1: Membrane–protein binding interaction energies for the S and NS states at two\ndistinct temperatures. For each state, the energy difference between temperatures is also\nreported.\nStates Temperature (°C) Energy Energy difference\n(kcal mol⁻¹) (kcal mol⁻¹)\nS 19 -37821 ± 500.2 -979 ± 724\n40 -36842 ± 524.3\nNS 19 -42505.2 ± 548.9 -2689 ± 819\n40 -39816.2 ± 609.1\n- Authors speculate that either TRAAK adopts 2 distinct folding patterns during protein\nsynthesis and/or that there is a conformational transition between the two states. But didn’t\nthey demonstrate that the latter occurs with their experiment on temperature? (Channels\nanalyzed at low and high temperatures were synthetized with the same protocol.) Authors\ncould mention it.\nResponse:\nWe thank the Reviewer for their insightful comment and for the opportunity to clarify point.\nWhile the temperature-dependent shift in the NS/S equilibrium suggests the presence of a\nconformational transition between these states, it does not exclude the possibility that TRAAK\ncan adopt two distinct folding patterns during synthesis. The observation that both\nconformations exist prior to temperature treatment supports the idea that alternative folding\npathways may also contribute to TRAAK’s cap conformational example.\n- l.406: Authors provide an estimate of the swapped to non-swapped conformation transition\nenergy (10 kcal/mol). They should compare it to the energy gain of the system when\ntemperature is increased from 19°C to 40°C in their experiment. Additionally, they could\nprovide a comparison with the energy associated to temperature shifts to which TRAAK is\nsensitive. This way, the proposition that a mechanism involving cap swapping explains\nTRAAK thermal sensitivity would be more convincing.\nResponse:\nWe thank the Reviewer for their valuable suggestion. Upon re-evaluating the available data,\nwe realised that the previously reported estimate of ~10 kcal/mol for the cap region was\nobtained from simulations performed at 300 K (26.85 °C). This value reflects the energy\ndifference for a limited number of residues within the cap region and does not represent the\nenergetic profile of the full protein. Moreover, energy differences in protein conformational\nstates are temperature-dependent, making it inappropriate to directly use this value to estimate\nthe energy gain associated with increasing the temperature from 19 °C to 40 °C in our system.\nFurthermore, it will not be fully accurate to directly relate the energy gain of the system to the\nenergy required for the state transition. When the resuspended cell membrane carrying\nexpressed TRAAK is heated in buffer solution, the absorbed energy is distributed (not evenly)\ninto all (or multiple) samples components. Not all of the heat is retained by the system; part of\nit is dissipated, and part is transferred to other membrane proteins and lipids as well. Therefore,\nit is challenging to quantify the exact fraction of energy specifically transferred to TRAAK\nchannels.\nTo better understand the PDS results which indeed show a shift of the equilibrium, we\nperformed long atomistic simulations of both the non-swapped (NS; PDB 3UM7) and swapped\n(S; PDB 4WFF) states at 19 °C and 40 °C and compared the non-boned interaction energy\nbetween the protein and its environment (lipid bilayer and solvent) (revised Figure S14 and\nrevised Table S4)\n(see also previous response). And the results are shown in the manuscript as:\n- The paragraph on heterodimers might be over-speculated at this point, in particular\nregarding this sentence: “By quantifying the proportional representation of cap states within\nthese ensembles, we can determine each state’s weighted contribution to the channel’s\nfunctional output.” Rephrase this sentence\nWe have now removed this sentence.\n- In fig2.a, authors should mention the 2 mutations in the vector name: V5_GFP_TRAAK\nE67C L144C\nWe thank the Reviewer for spotting this mistake. Now corrected.\nMistakes or points of clarification in the text:\n- l.156: what do authors mean by “homologous TRAAK sites”?\nWe rephrase this sentence to:\nFor spin labelling of a single engineered site per subunit in homodimeric TRAAK, the unique\narchitecture of the cap results in highly similar predicted distances for the two states\n- l.254: what does the “+ve” mean?\nIt means “positive ion mode” and it is corrected in the manuscript.\n- l.271: PIPS to PIPs\nCorrected\n- l.316: contributed to contributing?\nCorrected\nReviewer #4 (Remarks to the Author):\nThis manuscript probing the swapped (S) and non-swapped (NS) TRAAK potassium ion\nchannel states using DEER spectroscopy is excellent. The research presented clearly indicates\nthat both S and NS states coexist in membranes. The researchers noticed a shift in population\nto the swapped states at a higher temperature from 19 C to 40 C. The work presented in this\nstudy represents a clever way to get a complicated membrane protein sample that has a\ngreatly increased DEER sensitivity for the state of the dimer. The DEER data shown in the\npaper is excellent. While a Tikhonov regularization is better in general for getting a sense of\nthe most likely distance distribution, (there's no reason to assume that any distribution is\npurely Gaussian), using a dual gaussian fit is the best way to get a reasonably reliable\nmeasurement of the relative populations of the NS and S states (it's what everyone uses for\nthis). I think that this is compelling paper for a Nature Communications publication on the\nTRAAK K+ channel. Also, the methodology could be broadly applied to a lot of biological\nand membrane systems.\nResponse:\nWe sincerely appreciate the Reviewer's encouraging comments. We are pleased our combined\nmethodology is praised by the Reviewer and particularly the fact they recognise the potential\nof a wider applicability of our integrative approach to other membrane protein systems.\nReviewer #5 (Remarks to the Author):\nPotassium channels are oligomers of transmembrane subunits. The membrane-spanning\ndomain of a multimeric channel is an assembly of alpha helices that interact with each other\nand with membrane bilayer lipids. The pore selectivity filter is formed by four reentrant\ndomains which themselves interact with parts of the transmembrane helices. This assembly is\nhighly stable - an essential condition for pore geometry and channel selectivity. Studies to\ndate show that the oligomerization process takes place very early in channel biosynthesis in\nthe endoplasmic reticulum, under the control of chaperone proteins. No studies have shown\nthat the composition or overall conformational organization of the mature complex can be\naltered at the plasma membrane, probably because this would require too much energy. Under\nphysiological conditions, very minor conformational changes modify the activity or\nselectivity of the channel and regulate its function.\nResponse:\nWe appreciate the thoughtful comments by the Reviewer. We agree that potassium channel\nassemblies comprising four pore-forming re-entrant domains are inherently highly stable\nstructures and essential for maintaining optimal selectivity filter geometry. Consequently,\ndomain swapping at the plasma membrane would require breaking strong inter-helical bonds\nwhich would be energetically unfavourable and likely destabilizing to the pore domain.\nHowever, while it is reasonable to question the likelihood of a large-scale conformational\nrearrangement after early biosynthesis in the ER, this does not entirely exclude the possibility\nof such phenomena occurring under certain conditions.\nAn illustrative example of post-folding assembly conformational rearrangement is found in the\nTRPV3 ion channel, which has been shown to transition from a tetrameric to a pentameric state\nin the membrane. Lansky et al., Nature, (2023) used high-speed atomic force microscopy (HS-\nAFM) to observe membrane-embedded TRPV3 and directly detected channels composed of\nfive subunits, coexisting with the canonical tetramers. Importantly, these pentameric\nassemblies were transient and reversible, existing in dynamic equilibrium with tetramers\nthrough subunit exchange, with the entire process occurring within the membrane. HS-AFM\nvideos captured real-time tetramer-to-pentamer and pentamer-to-tetramer transitions, with\npentamers persisting for minutes within the lipid bilayer. The pentameric state was also trapped\nand resolved by CryoEM. This case study provides clear evidence of conformational variability\nin the mature TRPV3 complex, demonstrating that overall human ion channel organization can\nbe altered post biosynthetic assembly. Therefore, it remains possible that human TRAAK\nundergoes a conformational rearrangement at the membrane under certain physiological\nconditions.\nTwo-pore domain potassium (K2P) channels, including TRAAK, are dimers of subunits.\nPrevious structural studies have shown that mature K2P channels can exist in two\nconformational states, the so-called non-swapped (NS) and swapped (S) forms. The two forms\ndiffer in the positions of the helices forming the extracellular cap and the membrane-spanning\ndomains of each subunit in the dimer. The proportion of these two forms in cells and their\nfunctional significance are not known.\nIn this manuscript, Ma and colleagues expressed TRAAK in yeast, purified them and used\npulse dipolar electron paramagnetic resonance spectrometry to calculate the proportion of N\nand NS forms. They also show that temperature influences this ratio suggesting that the\npreviously reported effect of temperature on TRAAK could be due to a conversion between the\nN and NS forms. I’m not familiar with the technique used and the possible artifacts that it may\nproduce, but the conclusions that TRAAK can switch between the N and NS states and that\nthis switch has physiological significance are not supported by the provided data at this stage.\nSwitching between the S and NS forms would require major conformational changes, with the\nunfolding and refolding of several domains and the modification of a considerable numbers of\nhydrophilic and hydrophobic interactions. No kinetics of this process are provided in the\nmanuscript. If such rearrangement is possible after biosynthesis, then it is a multi-step process\nthat should be slow. The effect of temperature on TRAAK/TREK1/TREK2 channel activity is\nrapid, as the effects of other stimuli (stretch, lipids, pH…). Also a 10-fold increase in TRAAK\ncurrent is measured for a change from 17 to 40°C, while an increase of only 15% in the S form\nis observed. Nor is there any functional evidence that N and NS differ in their open probability\nor conductance, which would support the hypothesis of the authors.\nResponse:\nWe thank the Reviewer for their comment. We have now removed text which directly links\nTRAAK’s activation upon temperature increase to domain swapping and state that heating\nalters NS/S equilibrium ratio. However, whether changes in the NS/S ratio contribute to the\nincrease in current amplitude remains unknown. It is possible that two distinct mechanisms are\ninvolved.\nIn this work, the authors also took advantage of the way TRAAK is purified by SMA\nencapsulation to analyze the lipids surrounding the channel. While this is a promising approach,\nit has no bearing on the main message of the manuscript and its title. Much more work will be\nneeded to show whether what is observed in yeast is relevant in mammalian cells for TRAAK\nfunction and/or whether other K2Ps behave differently or similarly to TRAAK with regard to\nthe recruitment of surrounding lipids.\nIn conclusion, the authors have used novel and elegant approaches to study the structure of\nTRAAK and its lipid neighbourhood. But this work is still too preliminary to draw any firm\nconclusions about a functional role of the N and NS states of K2P channels.\nWe are pleased that the Reviewer finds our combined approach novel and innovative, but we\ndisagree with their assessment that our study is preliminary. We have here successfully\nmanaged to unbiasedly monitor the entire conformational ensemble of TRAAK’s cap domain\nin lipid membranes. However, we agree that what we observed in a yeast expression system\nwill need to be tested in mammalian cell lines for TRAAK and other K2P channels. Such\nextensive experiments will require substantial time and resources, and we intend to explore in\nthe future, but are well out of the scope of this study.\nNevertheless, we agree with the Reviewer that showing functional relevance of human-like\n(yeast) lipids identified in our ES-MS analysis will add to our current story. To this end we\nperformed multiple additional electrophysiology experiments on TRAAK in presence of the\nequivalent (to yeast) human plasma cell membrane lipids identified in our ES-MS analysis.\nIn brief (see also previous response to Reviewer 3), we performed additional electrophysiology\nrecordings of GUV-reconstituted TRAAK in presence of a) POPS, b) DOPE, c) PI(4,5)P , and\n2\nd) PI lipids, identified in our previous ES-MS analysis to assess the functional effect each of\nthese lipids have on TRAAK function. We now show in panel b-e from revised Figure. 3\n(reproduced as Figure 6), that PI(4,5)P and PI can activate TRAAK, while in contrast DOPE\n2\nexerts an inhibitory effect. Moreover, we tested POPS, a lipid also found in our ES-MS native\nmembrane analysis, which does not significantly affect TRAAK’s current amplitude. These\nnew electrophysiology data suggest that PS likely plays a structural rather than regulatory role\nfor TRAAK, possibly contributing to the stability of the channel within the lipid bilayer.\nFig. 6: TRAAK associates with specific membrane lipids and is modulated by lipid\ncomposition in reconstituted membranes. b–e, Representative currents from excised\npatches of wild-type TRAAK reconstituted in GUVs during a voltage-step protocol (Vh =\n0 mV; steps from −100 to +100 mV in 10 mV increments; every 40 mV step shown). Left and\nmiddle panels show recordings before and after bath perfusion with 4 µM of the indicated\nlipid: PI (b), PIP (c), DOPE (d), and POPS (e). Right panels: I–V relationships from the\n2\nsame patches at 10-mV increments. Black traces indicate average currents prior to lipid\napplication, while red traces (+PI, +PIP₂, +DOPE and +POPS) show the response following\nlipid perfusion.\nResponse to comments\nReviewer #1 (Remarks to the Author):\nI do not have any further comments for the authors. All my concerns have been\naddressed.\nReply: We thank the Reviewer for their constructive comments and are glad that all\ntheir concerns have now been addressed.\nReviewer #2 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed\nreports. This is part of the Nature Communications initiative to facilitate training in\npeer review and to provide appropriate recognition for Early Career Researchers\nwho co-review manuscripts.\nReviewer #3 (Remarks to the Author):\nIn their revised manuscript and their answers to the points we raised, Ma et al. have\nprovided extensive supplementary data, additional details and justifications that\nstrengthen their study on the temperature-dependent existence of the swapped and\nnon-swapped conformations of the K2P TRAAK channel.\nMore specifically, authors successfully demonstrated through patch-clamp\nrecordings that the mutated version of TRAAK used for their pulse dipolar EPR\nspectroscopy experiments is functional and retains the main regulatory properties of\nthe wild type channel. We are now convinced of the scientific soundness of their\nexperimental approach.\nRegarding the part where authors study the lipids surrounding TRAAK, the authors\nclarified their analysis and answered our questions satisfactorily.\nFinally, the authors successfully addressed our concerns regarding energetic\nconsiderations. The details they added in the manuscript as well as their molecular\ndynamics study largely improve the understanding of these energetic aspects.\nHowever, even if authors clearly explain why it is technically challenging to assess\nthe physiological relevance of the coexistence of the two TRAAK conformations\n(swapped and non-swapped), they failed to provide convincing evidence for the\nphysiological significance of their findings.\nOverall, their results represent a clear breakthrough in the field of K2P channel\nstructural biology, and their approach is technically innovative. However, the limited\nphysiological relevance of their findings may constrain the potential readership and\noverall impact of the study.\nReply: We thank the reviewer for recognizing the innovation and strengths of our\nstudy. We acknowledge that deep understanding of the physiological role of the co-\nexistence of two cap conformations in TRAAK (and potentially other K2P channels)\nin lipid membranes requires further investigation, which is out of the scope of our\ncurrent study. Although this aspect is technically challenging to investigate at\npresent, it is an important direction that we plan to explore in our future work.\nReviewer #5 (Remarks to the Author):\nWe thank the authors for their response. As they agreed that \"domain swapping at\nthe plasma membrane would require breaking strong inter-helical bonds which would\nbe energetically unfavourable and likely destabilizing to the pore domain\" and that it\nis therefore reasonable to \"question the likelihood of a large-scale conformational\nrearrangement after early biosynthesis in the ER\", it is important to mention this point\nin the manuscript as something to keep in mind for future studies.\nReply: We thank the Reviewer for this helpful suggestion. We have now\nincorporated this point in our revised manuscript.\nReviewer #6 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed\nreports. This is part of the Nature Communications initiative to facilitate training in\npeer review and to provide appropriate recognition for Early Career Researchers\nwho co-review manuscripts.\nReviewer #7 (Remarks to the Author):\nThis manuscript shows that swapped (S) and non-swapped (NS) TRAAK states co-\nexist in membranes and that their population shifts with temperature, supported by\ncareful PDS/DEER measurements. The MD simulations aim to rationalize the\ntemperature dependence by estimating “free energies” from non-bonded interaction\nenergies. However, the current MD analysis has methodological and interpretational\nissues that require major revision.\nMajor Comments:\nThe statement that Coulombic interactions were “shifted to zero between 0–12 Å”\nimplies that long-range electrostatics were truncated rather than treated with Ewald\nsummation. For a membrane-protein system, this is not acceptable because\nelectrostatics strongly govern protein–lipid and protein–solvent interactions. All\nsimulations must be rerun with Particle Mesh Ewald (PME) to appropriately treat\nlong-range interactions.\nWe thank the Reviewer for pointing this out to us. Our simulations were performed\nby combining the cut-off scheme with Force-switch for the Lenard-Jones, where the\ninteractions are smoothly switched between rvdw switch at 1.0 nm and a cut-off at\n1.2 nm. Coulomb interactions were calculated by using the cut-off type PME with a\ncut-off radius of 1.2 nm. Therefore, the interactions were calculated in real space\nwithin the cut-off, and in Fourier space using PME after the cutoff. We now include\nthis information in the Methods section and clarify this point.\nThe “approximate free energy” defined from total non-bonded interaction energy\nusing an ideal-gas reference (Eq. 1) is unconventional and thermodynamically\ninconsistent. Entropic and PV terms are neglected, yet the quantity is interpreted as\na free energy. Either rename this metric as an interaction-energy descriptor and\navoid free-energy language, or replace it with a recognized method (alchemical\nFEP/TI, or a PMF along a physically motivated coordinate). If PV is omitted, provide\nquantitative justification.\nWe agree with the Reviewer, and we have now changed the terminology (i.e. to\ninteraction energy) in the revised manuscript. We have further calculated the PV\nterm and found that this is negligible compared to the high energetics contribution\nfrom the non-bonded interactions. This quantification analysis has now been\nincluded in the revised manuscript.\nThe molecular origin of the temperature-dependent changes in non-bonded\ninteractions remains unclear. Do these shifts primarily reflect membrane mechanics\n(e.g., changes in bilayer thickness, area per lipid, or leaflet asymmetry), or\ndifferences in the stability of the membrane-exposed cap? What is the pressure\nprofile along the membrane normal? Please quantify and discuss how these\nproperties vary with temperature for both S and NS.\nWe thank the Reviewer for their thoughtful suggestions. We have now calculated these\nmembrane properties, including membrane thickness and the area per lipid for each\nmonolayer. These are now shown in Figure 1 below and the new Supplementary\nFigure S16 in the revised manuscript. Our new analysis reveals relatively small\ndifferences in membrane-thickness fluctuations.\nFor the NS state, the bilayer exhibits larger thickness fluctuations at 19 °C. These\nfluctuations reflect dynamic local deformations of the membrane around the protein.\nBy allowing the bilayer to adapt more closely to the surface of NS-state TRAAK,\nthese\ndeformations lower the overall protein–membrane interaction energy, making the NS\nstate more favourable at 19 °C. At 40 °C the membrane-thickness fluctuations are\nreduced, so the bilayer adapts less to the protein, leading to less favourable\ninteractions and a higher interaction energy for the NS state. In contrast, for the S state\nthe fluctuations in membrane thickness are highly similar for the two temperatures.\nThese observations are consistent with our previous protein–environment interaction\nanalysis, which showed that the temperature-dependent energy difference is larger for\nthe NS state than the S state.\nThe number(s) of lipid molecules for the two leaflets show small asymmetries in PC\n(revised Supplementary Table S3). The NS state has four additional PC molecules in\nthe outer leaflet. This explains the difference in area per lipid between the inner\n(cytoplasmic) and outer leaflets for the NS state, while the S state has one additional\nPC lipid molecule in the outer leaflet. The number(s) of PS and PE lipids are the same\nin each leaflet for the respective states, and the total number of lipids is constant\nacross all trajectories.\nFigure 1: Time evolution of membrane thickness and area per lipid for NS and S\nTRAAK simulations at different temperatures. In each row, the left panel shows the\ninstantaneous bilayer thickness as a function of time; the red dashed lines and red numbers\nindicate the time-averaged thickness. The right panels show the corresponding area per lipid\nfor the outer (gold) and inner (blue) leaflet of the bilayer, with horizontal dashed lines and\ncoloured numbers indicating the respective time-averaged values.\nThe simulated bilayers contain DOPC:DOPE:POPS (72:14:14), i.e., an explicit fraction\nof phosphatidylserine (PS). Because PS is negatively charged and typically enriched\nin the inner leaflet, its spatial organization relative to TRAAK is crucial for interpreting\nthe reported protein–membrane interaction energies. The manuscript should show\nwhere PS resides relative to the protein and whether it engages specific basic patches\non the channel surface. Please provide leaflet-resolved 2D density maps of PS\nheadgroups in the membrane plane, normalized by the bulk PS mole fraction, for both\nS and NS (with an inset radial profile g(r) to quantify enrichment). Comparing these\nmetrics between S and NS would clarify whether the reported energetic differences\nreflect PS-mediated contacts or generic bilayer mechanics.\nIn response to the reviewer’s request, we have computed leaflet-resolved two-\ndimensional density maps of PS headgroups in the inner leaflet, normalized by the\nbulk PS mole fraction, together with the corresponding radial distribution functions\n𝑔(𝑟)around the TRAAK channel (Figure 2; new Supplementary Figure S17 in the\nrevised manuscript).\nFor the NS state, the temperature dependence of the PS enrichment is more\npronounced. At 19 °C, the 2D density map displays extended PS-rich patches around\nthe protein, and the radial profile shows a clear maximum with 𝑔(𝑟) being significantly\nabove a value of 2 at short distances away from the protein. At 40 °C, PS remains\nenriched relative to the bulk bilayer, but PS hotspots become weaker and more\ndiffused, while the maximum of 𝑔(𝑟)is reduced. This indicates that PS molecules are\nmore strongly and persistently associated with the NS state at 19 °C than at 40 °C.\nFor the S state, PS is also enriched near the channel, but the changes between 19 °C\nand 40 °C are comparatively modest: the 𝑔(𝑟) curves are more similar for the two\ntemperatures, indicating that the PS environment of the S state is less temperature\ndependent.\nTo further address which specific regions of TRAAK interact with PS, we inspected the\ntrajectories from all four conditions. In all cases, PS binding sites are mainly clustered\nin the channel regions between TM2–TM3 and TM1–TM4 (Figure 3). Because these\ninteraction patterns are broadly similar across all conditions and do not highlight clear\nstate- or temperature-specific differences, we did not include additional structural\nsnapshots in the revised manuscript.\nImportantly, all these new analyses and observations are consistent with our previous\nprotein–environment interaction analysis. For the S state, the temperature\ndependence of the interaction energy was relatively small (ΔΔG ≈ −979 ± 724 kcal\nmol⁻¹), whereas the NS state showed a significantly larger change (ΔΔG ≈ −2689 ±\n819 kcal mol⁻¹). Thus, the stronger temperature sensitivity of the NS state could arise\nfrom a combination of temperature-dependent PS (re)organisation around the channel\nand changes in local bilayer mechanics (e.g., membrane-thickness fluctuations),\namong other factors. A full dissection of all contributing terms would require a new set\nof additional extensive simulations and analysis, which is beyond the scope of our\npresent study.\nTaken together, our new PS-density data are fully consistent with our previous\nanalysis and further support the main conclusions of our original manuscript.\nThe change in the revised manuscript:\nFigure 2: 2D density maps of PS lipid headgroups and radial distribution functions for\nthe inner leaflet of the NS and S TRAAK cap states at different temperatures. The left\npanels show the local PS headgroup mole fraction in the membrane plane (X–Y), normalised\nby the bulk PS mole fraction (colour bar; values > 1 indicate enrichment, values < 1 indicate\ndepletion). The red contour indicates the projected outline of the protein complex. The right\npanels show the corresponding radial distribution function g(r) of PS around the protein, with\nthe dashed line marking g(r) = 1 (bulk density), quantifying PS enrichment or depletion in the\ninner leaflet.\nFigure 3. Snapshots from each of the four simulation conditions, viewed from the\ncytoplasmic (inner leaflet) side. PS Lipids surrounding the protein (wheat-coloured sticks).\nIn the NS state, the binding asymmetry is mostly confined to the structurally unresolved\nregion between TM2 and TM3 of one of the two TRAAK subunits (pink coloured cartoon).\nMinor Comments:\nTRAAK gating is tension-sensitive. make the effective tension depend on leaflet lipid\nnumbers and box-area fluctuations. Please clarify inner/outer leaflet lipid counts.\nWe now provide the total number of lipids in both leaflets in the revised\nSupplementary Table S3.\nA representative snapshot showing the protein within the simulation box and the\nbilayer dimensions would help readers understand boundary conditions and system\npacking.\nWe now include a new Supplementary Figure S15 as follows:\nFig. S15: Representative snapshots showing NS and S TRAAK cap states within the\nbilayer of the simulation boxes. The water molecules are not shown for clarity. Grey\nspheres represent K⁺ ions, and olive green spheres represent Cl⁻ ions. DOPC lipids are shown\nin wheat, POPS lipids in pink, and POPE lipids in cyan colours. Side views (top two panels)\nand cytoplasmic views (bottom two panels) are shown for clarity."}, {"round": 2, "reviewer_comments": {"reviewer_1": "I do not have any further comments for the authors. All my concerns have been addressed.", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "In their revised manuscript and their answers to the points we raised, Ma et al. have provided extensive supplementary data, additional details and justifications that strengthen their study on the temperature-dependent existence of the swapped and non-swapped conformations of the K2P TRAAK channel.\n\nMore specifically, authors successfully demonstrated through patch-clamp recordings that the mutated version of TRAAK used for their pulse dipolar EPR spectroscopy experiments is functional and retains the main regulatory properties of the wild type channel. We are now convinced of the scientific soundness of their experimental approach.\n\nRegarding the part where authors study the lipids surrounding TRAAK, the authors clarified their analysis and answered our questions satisfactorily.\n\nFinally, the authors successfully addressed our concerns regarding energetic considerations. The details they added in the manuscript as well as their molecular dynamics study largely improve the understanding of these energetic aspects.\n\nHowever, even if authors clearly explain why it is technically challenging to assess the physiological relevance of the coexistence of the two TRAAK conformations (swapped and non-swapped), they failed to provide convincing evidence for the physiological significance of their findings.\n\nOverall, their results represent a clear breakthrough in the field of K2P channel structural biology, and their approach is technically innovative. However, the limited physiological relevance of their findings may constrain the potential readership and overall impact of the study.", "reviewer_5": "We thank the authors for their response. As they agreed that \"domain swapping at the plasma membrane would require breaking strong inter-helical bonds which would be energetically unfavourable and likely destabilizing to the pore domain\" and that it is therefore reasonable to \"question the likelihood of a large-scale conformational rearrangement after early biosynthesis in the ER\", it is important to mention this point in the manuscript as something to keep in mind for future studies.", "reviewer_6": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_7": "This manuscript shows that swapped (S) and non-swapped (NS) TRAAK states co-exist in membranes and that their population shifts with temperature, supported by careful PDS/DEER measurements. The MD simulations aim to rationalize the temperature dependence by estimating “free energies” from non-bonded interaction energies. However, the current MD analysis has methodological and interpretational issues that require major revision.\n\nMajor Comments:\nThe statement that Coulombic interactions were “shifted to zero between 0–12 Å” implies that long-range electrostatics were truncated rather than treated with Ewald summation. For a membrane-protein system, this is not acceptable because electrostatics strongly govern protein–lipid and protein–solvent interactions. All simulations must be rerun with Particle Mesh Ewald (PME) to appropriately treat long-range interactions.\n\nThe “approximate free energy” defined from total non-bonded interaction energy using an ideal-gas reference (Eq. 1) is unconventional and thermodynamically inconsistent. Entropic and PV terms are neglected, yet the quantity is interpreted as a free energy. Either rename this metric as an interaction-energy descriptor and avoid free-energy language, or replace it with a recognized method (alchemical FEP/TI, or a PMF along a physically motivated coordinate). If PV is omitted, provide quantitative justification.\n\nThe molecular origin of the temperature-dependent changes in non-bonded interactions remains unclear. Do these shifts primarily reflect membrane mechanics (e.g., changes in bilayer thickness, area per lipid, or leaflet asymmetry), or differences in the stability of the membrane-exposed cap? What is the pressure profile along the membrane normal? Please quantify and discuss how these properties vary with temperature for both S and NS.\n\nThe simulated bilayers contain DOPC:DOPE:POPS (72:14:14), i.e., an explicit fraction of phosphatidylserine (PS). Because PS is negatively charged and typically enriched in the inner leaflet, its spatial organization relative to TRAAK is crucial for interpreting the reported protein–membrane interaction energies. The manuscript should show where PS resides relative to the protein and whether it engages specific basic patches on the channel surface. Please provide leaflet-resolved 2D density maps of PS headgroups in the membrane plane, normalized by the bulk PS mole fraction, for both S and NS (with an inset radial profile g(r) to quantify enrichment). Comparing these metrics between S and NS would clarify whether the reported energetic differences reflect PS-mediated contacts or generic bilayer mechanics.\n\nMinor Comments:\nTRAAK gating is tension-sensitive. make the effective tension depend on leaflet lipid numbers and box-area fluctuations. Please clarify inner/outer leaflet lipid counts.\n\nA representative snapshot showing the protein within the simulation box and the bilayer dimensions would help readers understand boundary conditions and system packing."}, "author_response": ""}, {"round": 3, "reviewer_comments": {"reviewer_7": "I do not have any further comments for the authors. All my concerns have been addressed."}, "author_response": ""}]

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