To assess the involvement of PLD2 mechanosensitivity to a mechanically evoked current from TREK-1 channels, we conducted pressure current measurements in HEK293T cells both with and without the expression of a catalytically inactive K758R PLD2 mutant (xPLD2) (Toschi et al., 2009). We selected HEK293T cells for this study due to their minimal endogenous potassium currents, which allowed us to attribute the recorded currents specifically to TREK-1. The expression of TREK-1 was tracked using an EGFP tag attached to the channels C-terminus, and we confirmed successful expression of all constructs, observing their presence on the cell surface (Figure 2—figure supplement 1A and B).

In intact HEK293T cells, the overexpression of xPLD2 substantially inhibited nearly all mechanically evoked TREK-1 currents (Figure 1A–C). When currents were measured in the inside out configuration under negative pressure conditions (from 0 to –60 mmHg), they exhibited a reduction of over 80% from 0.181 ± 0.04–0.035 ± 0.013 pA/µm², despite an overall increase in the total levels of TREK-1 (Figure 2—figure supplement 1B and C). This observation suggests that if the effect is through a specific PLD2-TREK-1 interaction, then removal of the PLD2 binding domain should also inhibit PLD2-evoked currents.

To investigate further, we eliminated the putative PLD2 binding site on TREK-1 by truncating the C-terminus at residue 322 (TREK trunc) (see Figure 1—figure supplement 2A). The pressure-induced current in TREK trunc decreased by more than 85%, measuring 0.025 ± 0.013 pA/µm² (Figure 1C, Figure 1—figure supplement 2B). Remarkably, this reduction was nearly identical to the results obtained when full-length TREK-1 (TREK FL) was expressed in conjunction with xPLD2. Importantly, previous studies have confirmed the functionality and mechanosensitivity of this similar truncated TREK-1 when reconstituted into crude soy PC lipids (Brohawn et al., 2014a).

Conversely, the overexpression of TREK-1 with wild-type (WT) mouse PLD2 (mPLD2) resulted in a substantial augmentation of TREK-1 pressure-dependent currents within HEK293T cells (Figure 1A, see Figure 1—figure supplement 2B for raw traces). The TREK-1-evoked currents were markedly significant, both with and without overexpression of mPLD2, when compared to xPLD2 (p<0.002 and 0.007, respectively; Figure 1C). HEK293T cells, like all cells, have endogenous PLD2 (enPLD2). The xPLD2 expression is >10× the endogenous PLD2 (enPLD2) that we previously showed outcompetes the binding of enPLD2 to TREK-1 under similar conditions (Pavel et al., 2020).

The threshold for PLD2-dependent TREK-1 activation proved to be quite sensitive. TREK-1 exhibited responsiveness to negative pressure (ranging from 0 to –60 mmHg, in a non-saturating regime) with a half maximal pressure requirement of approximately 32 mmHg. This pressure led to the generation of up to 200 pA of TREK-1 current (Figure 1A, Figure 1—figure supplement 2C), aligning with findings from prior investigations (Brohawn et al., 2014b; Patel et al., 1998).

As previously mentioned, previous studies demonstrated TREK-1 direct sensitivity to mechanical force in purified lipids, independent of PLD2 (Berrier et al., 2013; Brohawn et al., 2014a). To delineate a PLD2-independent (i.e., direct) component of TREK-1 mechanotransduction within a cellular membrane, we conducted a comparative analysis between cells expressing TREK trunc and mock transfected HEK293T cells (lacking TREK-1 expression). Our results demonstrated that TREK trunc exhibited a modest increase in current, measuring 0.046 ± 0.023 pA/µm² (p=0.08, Figure 1C inset), when compared to the condition with no TREK-1. Similarly, in cells expressing xPLD2 alongside full-length TREK-1, the channel exhibited a small increase in current measuring 0.060 ± 0.035 pA/µm² (p=0.11). This PLD2 independent current aligns with direct mechanical activation of TREK-1 observed in liposomes. However, it is important to note that we cannot rule out the possibility of other contributing mechanisms.

Due to the minimal pressure-activated TREK-1 current observed in the presence of xPLD2, we conducted control experiments to validate the presence of functional TREK-1 on the plasma membrane—channels capable of conducting current. TREK-1 is a potassium leak channel with inherent basal currents in cultured cells (Comoglio et al., 2014). It is reasonable to assume that the basal current arises from the presence of basal levels of anionic lipids in the cellular membrane.

As anticipated, we observed TREK-1 basal current both in the presence (16.9 ± 4.3 pA/pF) and absence (24.0 ± 4.2 pA/pF) of xPLD2 (Comoglio et al., 2014; Figure 1D). These basal resting currents were recorded at 0 mV in HEK293T cells overexpressing TREK-1 FL with and without xPLD2. Notably, the current in the presence of xPLD2 was significantly higher than that observed in a nonfunctional control TREK-1 (xTREK), which we previously determined to exhibit no measurable current in unrelated studies. These controls affirm that TREK-1 FL is expressed and functionally active in the presence of xPLD2. Moreover, they highlight the essential role of PLD2 in pressure-activated currents (Figure 1C), contrasting with the basal leak currents (Figure 1D; TREK FL, gray bar vs. +xPLD2, red bar), which did not necessitate PLD2.

As previously reported in a prior publication, we proposed that fluid shear induces the disassociation of PLD2 with cholesterol-dependent GM1 clusters and the association with PIP₂ clusters, leading to a change in spatial organization and activation of the enzyme (Figure 1—figure supplement 1; Petersen et al., 2016). Given that TREK-1 binds to PLD2, we hypothesized the channel could undergo a similar mechanically induced spatial reorganization between GM1 and PIP₂ clusters. To visualize this spatial patterning of TREK-1 in response to shear forces, we developed a method to chemically fix membranes during shear (Figure 2A). In our experiments, we employed cultured HEK293T cells expressing both TREK-1 and mPLD, which matched the conditions used in our electrophysiology experiments. These cells were subjected to shear forces of 3 dynes/cm², a physiologically relevant force to cells (Petersen et al., 2016; Schneck, 1992), using a rotary shear approach. Subsequently, the cells were fixed and labeled for two- or three-color dSTORM. TREK-1 proteins were tagged with EGFP, leveraging its inherent self-blinking properties for detection (Call et al., 2023). Lipids were labeled using Alexa 647 (A647) anti-PIP₂ antibodies or A647-labeled cholera toxin B (CTxB), which selectively bind to PIP₂ and GM1 clusters, respectively (Petersen et al., 2016). By concurrently labeling both lipids and proteins, we were able to monitor the dynamic changes in spatial organization within nanoscopic lipid domains (Yuan and Hansen, 2023). To access the PIP₂ domains on the inner leaflet, cells were permeabilized (see ‘Materials and methods’). The effectiveness of cellular staining was confirmed through confocal microscopy (see Figure 2—figure supplement 1A–G).

Before the application of shear forces, TREK-1 and GM1 lipids displayed a robust correlation (Figure 2B and C), suggesting a close association between TREK-1 and GM1 lipids. However, after the application of shear forces, this correlation significantly diminished (p<0.01 at 50 nm). A similar experiment using a cy3b-labeled anti-TREK-1 antibody yielded nearly identical results, confirming the validity of both our EGFP-dSTORM method (Call et al., 2023) and the specificity of the TREK-1 antibody (Figure 1—figure supplements 1A and 2A).

Following the release of TREK-1 from GM1 lipids, we anticipated that it would undergo a nanoscale repositioning toward PIP₂ clusters. PIP₂ forms distinct nanodomains that are separate from GM1 clusters (Petersen et al., 2016; van den Bogaart et al., 2011; Wang and Richards, 2012) by an average distance of approximately 133 nm in HEK293T cells (Yuan et al., 2022). Cells were subjected to shear forces of 3 dynes/cm², permeabilized, fixed, and subsequently labeled with an anti-PIP₂ antibody. As predicted, the correlation between TREK-1 and PIP₂ was initially low prior to shear application, but it significantly increased after shear forces were applied (Figure 2D), in contrast to the observations made with GM1 (Figure 2C). Therefore, shear forces induce TREK-1 to disassociate from GM1 clusters and associate with PIP₂ clusters. Statistical analysis using a Student’s t-test confirmed the significance of TREK-1 clustering with PIP₂ at both a single radius and multiple point comparisons along the curve.

Subsequently, we investigated the rearrangement of TREK-1 induced by shear forces in the presence of xPLD2. Like mPLD2, we overexpressed xPLD2 in HEK293T cells, mirroring the conditions employed in our electrophysiology experiments in Figure 1B. Furthermore, as with mPLD2, HEK293T cells were subjected to shear forces, permeabilized, labeled, and imaged with two-color dSTORM.

In the presence of xPLD2, shear caused TREK-1 to leave GM1 domains (Figure 2E). However, unlike the response observed with mPLD2, the association of TREK-1 with PIP₂ clusters remained relatively weak following shear forces (Figure 2F), despite an overall increase in TREK-1 and PIP₂ levels in the membrane prior to shear (Figure 2—figure supplement 1C and D). Figure 2G shows a model of shear-induced movement of TREK-1 from GM1 to PIP₂ clusters.

It is presumed that TREK-1 translocates between nanodomains as a complex with PLD2 (Comoglio et al., 2014). Lacking its own palmitoylation, TREK-1 interacts with PLD2 through its unstructured C-terminus. In earlier research, we demonstrated that cholesterol drives PLD2 associates with GM1 lipids. The application of mechanical force activated PLD2, presumably through the release of PLD2 from GM1 lipids. However, this PLD2 patterning under mechanical shear has not been directly demonstrated with dSTORM.

To validate our proposed mechanism regarding the shear-induced movement of PLD2 with dSTORM, we employed calibrated shear chambers (ibidi µ-Slide I^0.4 parallel-plate) with cultured C2C12 muscle cells (mouse myocytes) and N2a mouse neuroblastoma cells, which naturally express TREK-1 (see Figure 2—figure supplement 1E). Utilizing endogenous expression helped circumvent potential artifacts that may arise from artificially saturating GM1 clusters through protein overexpression.

Our shear experiments were initiated by perfusing phosphate-buffered saline (PBS) through the calibrated shear chambers, applying a precisely controlled force of 3 dynes/cm². To maintain a constant temperature of 37℃, we employed a digitally controlled inline heater. Immediately following the application of shear forces (within <10 s), we introduced fixative agents into the shear buffer, facilitating the rapid fixation of cells in their mechanically stimulated state. This approach paralleled the methodology used in our experiments involving rotary shear (see Figure 3A, Figure 3—figure supplement 1A), but with tighter control of temperature.

Utilizing two-color dSTORM and analysis by the pair correlation function, we observed that shear forces introduce the mobilization of PLD2 within the cell membrane, and this effect was independent of temperature fluctuations. Prior to the application of shear forces, PLD2 exhibited a strong association with GM1 clusters (Figure 3B). However, after the application of shear forces, the correlation of PLD2 with GM1 clusters decreased significantly, while it robustly increased in association with PIP₂ clusters (Figure 3C). Notably, the release of PLD2 from GM1 domains induced by shear forces was more pronounced than the disruption caused by anesthetic agents, as previously reported (Pavel et al., 2020). The stability in temperature, maintained within a range of ±0.1℃, suggests the mechanism is unlikely to be attributed to the thermal melting of ordered lipids near a phase transition state.

Conducting cluster analysis of GM1 particles subjected to shear revealed a notable reduction in the apparent size of GM1 clusters, decreasing from 167 ± 3 to 131 ± 3 nm (approximately 20%, Figure 3D) in C2C12 cells. A similar effect on the size of GM1 clusters was also observed in N2a cells (Figure 3E). PIP₂ clusters also remained largely intact with a reduction in size (from 154 ± 1 to 139 ± 1) after shear (Figure 3—figure supplement 1B).

The observed reduction in cluster size closely resembled the outcomes of prior experiments in which we depleted cholesterol with methyl-beta-cyclodextrin (MBCD) (Pavel et al., 2020; Petersen et al., 2016). Given that MBCD is a known cholesterol removal agent, we conducted assays on N2a cells subjected to orbital fluid shear (3 dynes/cm²) to examine changes in cholesterol levels. The cells were subjected to 10 min of shear at 37℃, followed by fixation.

Our results revealed a 25% reduction in free cholesterol levels in N2a cells exposed to orbital shear (p<0.001) (Figure 3F), which coincided with the activation of PLD (Figure 3G). Importantly, this decrease in cholesterol was statistically significant p<0.0001 and reversible. Allowing the cells to recover for approximately 30 s prior to fixation resulted in the restoration of cholesterol levels to those observed in non-sheared cells (Figure 3F).

In C2C12 cells, TREK-1 and PLD2 exhibited a strong correlation both before (Figure 3—figure supplement 1C, gray trace) and after shear (Figure 3—figure supplement 1C, red trace), providing further evidence that they form a complex at least intermittently, in both shear and unsheared states. Following shear, a minor fraction of TREK-1 was found to associate with PIP₂ (Figure 3—figure supplement 1D). Interestingly, in contrast to HEK293T cells, C2C12 cells displayed minimal TREK-1/GM1 correlation (Figure 3—figure supplement 1E, gray trace). Notably, PLD2 regulation is known to be influenced by cholesterol (Petersen et al., 2016). In various cell types, we have observed that cultured cells tend to have lower cholesterol levels than human tissues (Wang et al., 2023; Wang et al., 2021; Yuan et al., 2022). These observations lead us to consider the possibility of introducing more physiological levels of cholesterol in our experimental setup for cultured cells.

Cholesterol levels, especially in the brain, can be notably high (Hansen, 2023; Zhang and Liu, 2015). We hypothesized that cholesterol might influence the endogenous TREK-1 to associate with PLD2 in GM1 clusters. To test this hypothesis, we introduced cholesterol into C2C12 cells using apolipoprotein E (apoE) lipidated with 10% serum as apoE is a naturally occurring cholesterol transport protein (Wang et al., 2023; Wang et al., 2021; Yuan et al., 2022; see Figure 4A). Remarkably, lipidated apoE induced a substantial clustering of TREK-1 with GM1 lipids in membranes of C2C12 cells (Figure 4B). This effect was also observed with TREK-1 in N2a cells. Importantly, the application of fluid shear (3 dynes/cm²) completely reversed the effect of cholesterol (Figure 4C).

We anticipated that the cholesterol-induced association of TREK-1 with GM1 lipids would lead to a reduction in TREK-1 currents as PLD2 would be inhibited due to a lack of substrate. To directly assess the activity of TREK-1 under conditions of elevated cellular cholesterol, we overexpressed TREK-1 in HEK293T cells and quantified the current density in whole-cell patch-clamp mode, both with and without cholesterol uptake (Figure 4D). Consistent with our proposed model, TREK-1 current density decreased nearly 2.5-fold in cholesterol-loaded cells, and this reduction in current was statistically significant (p<0.05).

In our earlier investigation, we established that astrocytes play a pivotal role in regulating the clustering of proteins in neurons. This regulation is achieved through the release of apoE-containing particles, which are primarily laden with cholesterol. Remarkably, our studies revealed that cholesterol amplifies the clustering of proteins within GM1 domains. Furthermore, we demonstrated that disrupting the cholesterol synthesis in astrocytes leads to the reversal of protein clustering, as previously reported (Wang et al., 2021). It stands to reason that the same astrocytic cholesterol dynamics could govern the localization of TREK-1 within GM1 lipids in the brain of a mouse.

To investigate the in vivo regulation of TREK-1 clustering by astrocytes, we stained 50 micron brain slices obtained from both control mice and mice specifically engineered to lack cholesterol synthesis in astrocytes. The depletion of cholesterol from astrocytes was achieved by targeting sterol regulatory element-binding protein 2 (SREBP2), a crucial regulator of cholesterol synthesis (Ferris et al., 2017; Wang et al., 2021). Before slicing, the brains underwent fixation via whole-body perfusion. The resulting free-floating brain slices were mounted onto circular cover slips using a fiberglass filter paper, which facilitated dSTORM buffer access to the tissue with minimal background interference (see Figure 2—figure supplement 2E). These brain slices, taken from coronal sections near the hippocampus, were subjected to staining with cy3b-anti-TREK-1 antibody and A647-CTxB. The imaging process encompassed multiple unspecified locations (approximately 10 regions), and subsequent analysis was performed employing the pair correlation function.

In flox control mice (wild-type SREBP2), TREK-1/GM1 correlation exhibited a distinct pattern, with TREK-1 clustering near GM1 (Figure 4E), aligning with the GM1 correlation seen in N2a cells with elevated levels of cholesterol (Figure 4B). As anticipated, in the brains of mice with astrocyte-specific SREBP2 deficiency, TREK-1 displayed a reduced association with GM1 lipids (p<0.0001). While highly variable, these findings serve as evidence that, in the brain of an animal, astrocytic cholesterol serves as a critical regulator of TREK-1’s affinity for inhibitory GM1 clusters.

Figure 4F presents a comprehensive model detailing the regulation of TREK-1 by astrocyte-derived cholesterol, using the insights derived from both cultured cells and ex vivo studies. In this model, cholesterol originating from astrocytes is transported to neurons via apoE. Under conditions of elevated membrane cholesterol, TREK-1 forms associations with inhibitory GM1 lipids, a scenario in which PLD2 has limited substrate availability. Conversely, when membrane cholesterol levels are reduced, TREK-1 relocates away from GM1 lipids toward activating PIP₂ lipids. In this context, PLD2 gains improved access to its substrate phosphatidylcholine (PC), leading to the production of lipid agonists such as PA, which ultimately evoke activation of the TREK-1 channel (see also Figure 4—figure supplement 1).

Osmotic stretch is a well-documented activator of TREK-1 (Patel et al., 1998). To investigate whether osmotic stretch also activates PLD2, we conducted experiments to monitor PLD product release in live cells subjected to hypotonic stretch. Our findings revealed that membrane stretch by a 70 mOsm swell (representing a high degree of stress) led to approximately 50% increase in PLD2 activity in N2a cells (Figure 5A). It is worth noting that this level of activation, although significant, was less pronounced compared to shear stress in N2a cells (Figure 3E), which increased approximately threefold in activity.

Subsequently, we conducted dSTORM experiments to investigate whether stretch had the capacity to release PLD2 from GM1 lipids within the membranes of N2a cells. In these experiments, cells were exposed to either a hypotonic solution with an osmolality of 70 mOsm (indicating a state of swelling) or an isotonic solution with an osmolality of 310 mOsm (representing a control condition) for 15 min at 37℃. Following this treatment, the cells were fixed, labeled, and subjected to imaging using the same procedures employed in the shear stress experiments. Our results, consistent with the outcomes of shear stress experiments and PLD2 assay, clearly demonstrated a discernible shift in the spatial distribution of PLD2, transitioning from ordered GM1 clusters to the PIP₂ clusters after osmotic stretch (Figure 5B–D).

In order to gain deeper insights into the in vivo role of PLD2 in mechanotransduction, we conducted investigations into mechano-thresholds and pain perception in Drosophila, employing a PLD-knockout model (Thakur et al., 2016). It is worth noting that, while the specific downstream effectors of PLD in fruit flies remain unidentified, and they possess only distantly related TREK-1 orthologs, (as discussed in a later section), the utilization of fruit flies as a model organism is advantageous because they possess a single PLD gene (dPLD).

Our analysis revealed the presence of GM1 domains in dissected fly brains, and we observed that dPLD exhibited a robust response to shear stress of 3 dynes/cm² when cultured in neuronally derived fly cell line, BG2-c2 (see Figure 6A and B). The activity of PLD increased by nearly fourfold in the fly cell line (Figure 6B), a result consistent with the response of PLD to mechanical force observed in HEK293T cells (Figure 3G).

Subsequently, we preceded to examine the in vivo role for PA in mechanosensation employing single-animal measurements of arousal threshold (Murphy et al., 2017; Murphy et al., 2016; see Figure 6—figure supplement 1A). The arousal assay quantifies the amount of mechanical stimulation required to elicit movement from a resting fly. Over a period of 24 hr, the flies were exposed to a series of incremental vibrational stimuli every 30 min. For each series, the threshold of stimulation necessary to induce movement in the fly, as indicated by observable motion, was recorded using automated machine vision tracking. These recorded measurements were then compared to genetically matched control groups.

PLD^null flies exhibited a notably lower arousal threshold compared to their control counterparts (Figure 6C). This lower arousal implies an increased sensitivity to mechanical force. Furthermore, we employed a neuronal-specific driver, Nsyb GAL4, in conjunction with a PLD RNAi line (PLD-KD) to investigate the role of PLD in the central nervous system. The neuronal knockdown of PLD yielded a similar increase in mechanosensitivity, providing clear evidence that this phenotype is specific to neuronal functions (Figure 6D).

Furthermore, we conducted assessments to determine the role of PLD in fly shock avoidance, serving as a measure of an adverse electrical stimulus (Figure 6—figure supplement 1B). To assess their responses, PLD-KD flies were positioned at the choicepoint of a T maze, where they were given the option to select between a chamber inducing a noxious shock or a non-shock control chamber (Drago and Davis, 2016). Flies were subjected to incrementally increasing voltages of electrical shock. Our findings revealed that PLD^null flies exhibited heightened sensitivity to electric shock compared to control groups (Figure 6E and F). In Figure 6G, we present a potential model illustrating how PA may contribute to mechanosensitivity in a fly.

HEK293T cells (ATCC Cat# CRL-3216, RRID:CVCL_0063), C2C12 cells (ATCC Cat# CRL-1772), and N2a (ATCC Cat# CCL-131) were cultured in DMEM (Corning cellgro) with 10% FBS, 100 units/ml penicillin, and 100 µg/ml streptomycin. Mycoplasma testing was performed by PCR and found to be negative. Cells were plated on poly-d-lysine-coated 12 mm microscope cover glass at approximately 12 hr, 36 hr, or 60 hr before transfection with genes for target proteins. Transfections were performed using X-tremeGENE 9 DNA transfection agent (Roche Diagnostics). Full-length human TREK-1(TREK-1 FL) with C-terminus GFP tag in pCEH vector was obtained from Dr. Stephen Long. Mouse PLD2 constructs (mPLD2) and inactive mutant (K758R single mutation, xPLD2) without GFP tag in pCGN vector were provided by Dr. Michael Frohman. TREK-1 constructs were co-transfected with mPLD2 or xPLD2 at a 1:4 ratio (0.5 g of TREK-1 and 2 g of PLD DNA) (Comoglio et al., 2014). All the salts for internal/external solutions were purchased from either Sigma or Fisher Scientific.

The transfected HEK293T cells were used in 24–36 hr after transfection for standard excised inside-out patch-clamp recordings of TREK-1 (Brohawn et al., 2014a; Comoglio et al., 2014). Currents were recorded blinded at room temperature using an Axopatch 200B amplifier and Digidata 1440A (Molecular Devices). Borosilicate glass electrode pipettes (B150-86-10, Sutter Instrument) were pulled with the Flaming/Brown micropipette puller (Model P-1000, Sutter Instrument), resulting in 3–6 MΩ resistances with the pipette solution (in mM): 140 NaCl, 5 KCl, 1 CaCl₂, 3 MgCl₂, 10 TEA-Cl, 10 HEPES, pH 7.4 (adjusted with NaOH). Bath solution consists of (in mM) 140 KCl, 3 MgCl₂, 5 EGTA, 10 TEA-Cl, 10 HEPES, pH 7.2 (adjusted with KOH). Low concentration of TEA (10 mM) was added into both pipette/bath solutions to block the endogenous potassium channels in HEK293T cells. Patch electrodes were wrapped with parafilm to reduce capacitance. Currents measured using Clampex 10.3 (Molecular Devices) were filtered at 1 kHz, sampled at 20 kHz, and stored on a hard disk for later analysis.

Pressure clamping on the patch was performed using high-speed pressure clamping system (ALA Scientific) through the Clampex control. Data was analyzed offline by a homemade procedure using IgorPro 6.34A (WaveMetrics).

TREK-1 current, either co-expressed with mPLD2 or xPLD2, was activated by negative pressure steps from 60 to 0 mmHg in 10 mmHg decrements at +30 mV membrane potential, and five traces for each case were recorded and averaged for the analysis. Patch size was estimated using the current density (I_density; pA/µm²). Then, a Boltzmann equation, I_density = base + {max/[1+exp((P50-P)/slope)]} was used to fit the data with a constraint of base = 1 due to poor saturation of the current at high pressure. P is the applied pressure, P50 is the pressure that activates 50% of maximum current measured (non-saturating), and slope shows the sensitivity of current activation by pressure. In some experiments with hTREK-1+xPLD2 co-expression where the activated currents were too small to fit to the Boltzmann equation, the current amplitude at P=-30 mmHg (I_m30) was compared with its 5× standard deviation(I_5×SD). If I_m30 < I_5×SD, the experiment was excluded from the Boltzmann equation fitting and corresponding P50-slope analysis. This empirical rule (we call it 5×SD rule) can discern four out of five wild-type (TREK-1 FL) cell-attached recording cases as null experiments, suggesting that it could be a usable/useful empirical criterion for our experiment. Then, the current density at –60 mmHg and P50-slope data were used for statistical analysis. Mann–Whitney test was done to assess statistical significance using Prism6 (GraphPad Software), and outliers were eliminated using a built-in function in Prism with Q = 1%. The values represented are mean ± SEM.

Housing, animal care, and experimental procedures were consistent with the Guide for the Care and Use of Laboratory Animals and approved by the Institutional Animal Care and Use Committee of the University of Virginia. The tissue from SREBP2 mice are from strains 030826 and 004600 from the Jackson Laboratories as described in Wang et al., 2021. No new animals were used for this study.

Cells (C2C12, HEK293T, and N2a) were grown to 80% confluence (C2C12 were allowed to differentiate overnight in serum free media). Cells were rinsed, treated as needed, and then fixed with 3% paraformaldehyde and 0.1% glutaraldehyde for 10 min to fix both protein and lipids. Glutaraldehyde was reduced with 0.1% NaBH₄ for 7 min followed by three 10 min washes with PBS. Cells were permeabilized for 15 min with 0.2% Triton X-100, blocked with 10% BSA/0.05% Triton/PBS at room temperature for 90 min. Primary antibody (PLD2, Cell Signaling #13891; TREK-1, Santa Cruz #sc-50412; PIP₂, Echelon Biosciences #z-P045) was added to a solution of 5% BSA/0.05% Triton/PBS for 60 min at rt at a concentration of 1:100 followed by five washes with 1% BSA/0.05% Triton/PBS for 15 min each. Secondary antibody (Life Technologies #A21244 and A21235; cy3B antibodies were produced as described previously; Petersen et al., 2016) was added in the same buffer as primary for 30 min at room temperature followed by five washes as above. A single 5 min wash with PBS was followed by a post-fix with fixing mixture, as above, for 10 min w/o shaking. This was followed by three 5 min washes with PBS and two 3 min washes with dH₂O. Cells only receiving CTxB treatment were not permeabilized.

Brain slices from a hGFAP-Cre driving the SB2 knockout mouse and its Flox control were from the same animals that were previously characterized (Wang et al., 2021). The protocol is the same as cells except the permeabilized samples were treated with anti-TREK cy3b+CTxB-647 for 3 d (4℃) and then washed five times (room temperature) for 1 hr prior to post fixing.

The dual-fixation protocol is used to minimize any effects from post-fixation aberrations. While always good practice for super-resolution in general, this dual fixation also ensures that the movement of any molecule of interest which may not have been immobilized by the initial fixation can be fully immobilized after labeling since the antibodies or toxins used for labeling will be efficiently cross-linked during this post-labeling fixation step. While some have proposed that this problem should be solved by adding the label before the initial fixation (Tanaka et al., 2010), we believe that in the absence of easily attainable monomeric labeling molecules it would have likely led to clustering artifacts due to the (often) multimeric nature of the labeling proteins.

For cells loaded with cholesterol, 4 ug/ml apolipoprotein E3 (BioLegend, USA) was mixed with fresh 10% FBS and applied to the cells for 1 hr prior to shear and/or fixing.

Shear force was applied to cells in ibidi µ-Slide I^0.4 Luer chambers with a flow rate calibrated to apply 3.0 dynes/cm². Fixation media (see above) was applied to cells using a syringe pump (Harvard Apparatus PHD ULTRA) and kept at 37°C using an in-line heater (Warner SH-27B).

TREK-1 in HEK293T cells was labeled with a EGFP concatenated to the C-Terminus of full-length human TREK-1 (Figure 1—figure supplement 2B) or by applying anti-TREK-1 antibody (sc-398449, Santa Cruz) conjugated to cy3b (Petersen et al., 2016). Anti-PIP₂ antibody was directly conjugate Alexa 647 using the same protocol.

Images were recorded with a Vutara 352 and VXL super-resolution microscopes (Bruker Nano Surfaces, Salt Lake City, UT), which is based on the 3D Biplane approach. Super-resolution images were captured using a Hamamatsu ORCA Flash4.0 sCMOS camera and a ×60 water objective with numerical aperture 1.2. Biological replicates (6–12) are individual cells from at least two independent experiments. Data were analyzed using the Vutara SRX software (version 5.21.13 for the data in Figures 3—5 and version 7.0.07 for the data in Figure 2). Single molecules were identified by their brightness frame by frame after removing the background. Identified particles were then localized in three dimensions by fitting the raw data in a customizable region of interest (typically 16 × 16 pixels) centered on each particle in each plane with a 3D model function that was obtained from recorded bead datasets (10,000–750,000 localization per biological replicate). Fit results were stored as data lists for further analysis.

Fixed samples were imaged using a 647 nm and 561 nm excitation lasers, respectively, and 405 nm activation laser in photo switching buffer comprising of 20 mM cysteamine (Sigma, #30070), 1% betamercaptoethanol (BME) (Sigma, #63689), and oxygen scavengers (glucose oxidase, GLOX) (Sigma #G2133) and catalase (Sigma #C40) in 50 mM Tris (Affymetrix, #22638100) + 10 mM NaCl (Sigma, #S7653) + 10% glucose (Sigma, #G8270) at pH 8.0 at 50 Hz and maximal powers of 647 nm, 561 nm and 405 lasers set to 8, 10, and 0.05 kW cm^-2, respectively.

C2C12 cells were grown in DMEM with 10% FBS until 16 hr before use in which they were switched into serum-free DMEM. On the day of the experiment, DMEM in live cells was replaced with DMEM w/o phenol red. Fixed cells were rinsed once with PBS and then put into a mixture of PBS with 3% PFA and 0.1% glutaraldehyde for 20 min at 37°C. Fixed cells were then rinsed with PBS 5 × 5 min and placed back into phenol-free DMEM. CTxB (Thermo Fisher C34778, 100 ug/ml) was then applied 1:200 into each plate and allowed to incubate for >30 min before imaging. Imaging and data collection was performed on a Leica SP8 confocal microscope with the Application Suite X v.1.1.0.12420. Five images were taken as baseline after which a selection of one or more ROI was bleached at 100% laser power for 6–8 frames. Recovery was measured out to 5 min, and fluorescence of the ROI(s) was quantified. The fluorescence before bleaching was normalized to 1 and after the bleaching step was normalized to 0.

N2a cells were cultured in 48-well plates with 200 ul media in each well and then changed to 200 ul PBS for the shear treatment. The shear plate was incubated with PBS on an orbital rotator at 3 dynes/cm² for 10 min in a 37°C incubator. The control plate was incubated with PBS for 10 min in the same incubator with no shear. Then the shear plate was incubated with 200 ul 4%PFA + 0.1% glutaraldehyde in PBS for 10 min with 3 dynes/cm² shear and 10 min without shear. The control plate was fixed for 20 min with no shear. Fixed cells were lysed in RIPA buffer (Thermo) and assay in 100 µl PBS with, 4 U/ml cholesterol oxidase, 100 μM amplex red, and 2 U/ml horseradish peroxidase (HRP). The data shown in the figure are from two biological replicates (independent experiments) with 5–10 technical replicates.

In vitro cellular PLD2 activity was measured in cultured HEK293T cells by an enzyme-coupled product release assay (Petersen et al., 2016) using amplex red reagent. Cells were seeded into 96-well plates (~5 × 10⁴ cells per well) and incubated at 37°C overnight to reach confluency. The cells were starved with serum-free DMEM for a day and washed once with PBS. The PLD reaction was initiated by adding 100 μl of reaction buffer (100 μM amplex red, 2 U/ml HRP, 0.2 U/ml choline oxidase, and 60 μM C8-PC in PBS). The assay reaction was performed for 2–4 hr at 37°C, and the activity was kinetically measured with a fluorescence microplate reader (Tecan Infinite 200 Pro) at excitation and emission wavelengths of 530 nm and 585 nm, respectively. The PLD2 activity was calculated by subtracting the background activity (reaction buffer, but no cells). For the bar graphs, samples were normalized to the control activity at the 120 min time point.

For behavior experiments, 1- to 5-day-old flies were collected in vials containing ~50 flies at least 12 hr before the experiment. Flies were allowed to acclimate to behavior room conditions for >30 min (dim red light, ~75% humidity) before each assay. Shock avoidance was tested by placing flies in a T-maze where they could choose between an arm shocking at the indicated voltage every 2 s and an arm without shock. Flies were given 2 min to choose which arm, after which flies were collected and counted to determine the shock avoidance index for each voltage and genotype. Control and knockout flies were alternated to avoid any preference, and the arm used for shock was also alternated to control for any non-shock preference in the T-maze itself. Shock avoidance index (AI) was calculated as AI = (# flies shock arm-# flies control arm)/# flies total. Plotting the inverse of this metric, we obtain a pain sensitivity curve in which we observe a right-shift when the pld gene was knocked down (Figure 3F). Flies were averaged from 2 to 3 biological replicates with ~10 groups per replicate.

Arousal threshold protocol has been described in detail previously (Murphy et al., 2016). Briefly, animals were exposed hourly to a series of vibrations of increasing intensity ranging from 0.8 to 3.2 g, in steps of 0.6 g. Stimuli trains were composed of 200 ms vibration with 800 ms inter-vibration interval and 15 s inter-stimuli train interval. Stimulation intensity and timing were controlled using pulse-width modulation via an Arduino UNO and shaftless vibrating motors (Precision Microdrives, model 312-110). Arousal to a given stimulus was assigned when an animal (1) was inactive at the time of the stimulus, (2) satisfied a given inactivity criteria at the time of the stimulus, and (3) moved within the inter-stimuli train period (15 s) of that stimulus. Statistics are from two biological replicates.

All statistical calculations were performed using a Student’s t-test or Mann–Whitney test in Prism software (v9) unless otherwise noted. For statistics of more than one point along a pair correlation curve, a nested Student’s t-test was used. Significance is noted as follows: ns, p>0.05; *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.