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. Author manuscript; available in PMC: 2022 Apr 23.
Published in final edited form as: Methods Enzymol. 2021 Mar 4;652:49–79. doi: 10.1016/bs.mie.2021.01.025

Methods to study phosphoinositide regulation of ion channels

Yevgen Yudin 1, Luyu Liu 1, Janhavi Nagwekar 1, Tibor Rohacs 1,1
PMCID: PMC9034691  NIHMSID: NIHMS1797112  PMID: 34059290

Abstract

Ion channel are embedded in the lipid bilayers of biological membranes. Membrane phospholipids constitute a barrier to ion movement, and they have been considered for a long time as a passive environment for channel proteins. Membrane phospholipids, however, do not only serve as a passive amphipathic environment, but they also modulate channel activity by direct specific lipid-protein interactions. Phosphoinositides are quantitatively minor components of biological membranes, and they play roles in many cellular functions, including membrane traffic, cellular signaling and cytoskeletal organization. Phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2] is mainly found in the inner leaflet of the plasma membrane. Its role as a potential ion channel regulator was first appreciated over two decades ago and by now this lipid is a well-established cofactor or regulator of many different ion channels. The past two decades witnessed the steady development of techniques to study ion channel regulation by phosphoinositides with progress culminating in recent cryoEM structures that allowed visualization of how PI(4,5)P2 opens some ion channels. This chapter will provide an overview of the methods to study regulation by phosphoinositides, focusing on plasma membrane ion channels and PI(4,5)P2.

Keywords: Phosphoinositides, lipids, PIP2, phospholipids, phospholipase C, voltage sensitive phosphatase, excised patch

1. Introduction

Phosphoinositides are quantitatively minor membrane lipids, typically constituting less than 1% of the lipid content in any given biological membrane. They are formed by phosphorylation of phosphatidylinositol (PI) at the 3’, 4’ or 5’ hydroxyl group, resulting in seven different phosphoinositide species: PI(4)P, PI(5)P, PI(3)P, PI(4,5)P2, PI(3,4)P2, PI(3,5)P2 and PI(3,4,5)P3 (Balla, 2013). Different subcellular membranes have different patterns of phosphoinositide composition. Given their high charge densities, phosphoinositides can form strong and specific interactions with proteins and thus, play important roles in defining molecular identities of various membrane compartments. PI, the precursor, is not considered a phosphoinositide and it is relatively abundant in cellular membranes (10–20%) (Balla, 2013; Dickson and Hille, 2019). Recent data indicate that PI in the plasma membrane is barely detectable with newly developed fluorescent probes, and much of this lipid is found in intracellular membrane compartments such as the Golgi, mitochondria and the endoplasmic reticulum. (Pemberton et al., 2020; Zewe et al., 2020).

PI(4,5)P2 (Fig. 1) is mainly found in the inner leaflet of the plasma membrane, constituting ~1% of phospholipids under resting conditions. Estimates range from 0.2 – 3.2 % (Hilgemann, 2007; Falkenburger et al., 2010; Balla, 2013; Dickson and Hille, 2019). PI(4,5)P2 is best known for being hydrolyzed by phospholipase C (PLC) enzymes to form the two classical second messengers: inositol 1,4,5 trisphosphate (IP3), which releases Ca2+ from the endoplasmic reticulum, and diacylglycerol (DAG), which activates protein kinase C (PKC). PI(4,5)P2 regulates the activity of a large number of ion channels (Suh and Hille, 2008; Logothetis et al., 2015). For the vast majority of them this lipid is a positive regulator or cofactor. For constitutively active PI(4,5)P2 dependent channels, such as most inwardly rectifying K+ (Kir) channels, this lipid by itself is sufficient to keep the channel open (Zhang et al., 1999). Other PI(4,5)P2 dependent channels require additional stimuli to open, such as Gβγ for G-protein activated Inwardly Rectifying K+ (GIRK, Kir3.x) channels (Sui et al., 1998), or menthol or cold for Transient Receptor Potential Melastatin 8 (TRPM8) channels (Rohacs et al., 2005).

Figure 1. Phosphoinositide metabolism.

Figure 1

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A: Chemical formula of PI(4,5)P2, with arachidonyl and stearyl (AASt) acyl chains, the most common acyl chain combination in biological membranes. B: PI(4,5)P2 is generated by phosphorylation of PI to PI(4)P by phosphatidylinositol-4-Kinases (PI4K) followed by phosphorylation of PI(4)P by phosphatidylinositol 4-phosphate 5-kinases (PIP5K). Phospholipase C (PLC) enzymes hydrolyze PI(4,5)P2 into diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (IP3). PI(4,5)P2 can be further phosphorylated by PI3K enzymes to generate PI(3,4,5)P3.

PI(4)P is the precursor of PI(4,5)P2. This lipid is found in the Golgi as well as in the plasma membrane where its levels are comparable to those of PI(4,5)P2 (Balla, 2013). PI(4)P also affects the activity of some PI(4,5)P2 sensitive ion channels (Nilius et al., 2006), but its effects are usually weaker than those of PI(4,5)P2. In some cases, PI(4)P was proposed to play roles distinct from PI(4,5)P2 in regulating ion channels (Hammond et al., 2012; Lukacs et al., 2013b).

PI(3,4)P2 and PI(3,4,5)P3 are products of the activation of phosphoinositide 3-kinase (PI3K) enzymes. They are largely found in the plasma membrane and play important roles in signaling by receptor tyrosine kinases activated by growth factors. While PI(3,4)P2 and PI(3,4,5)P3 activate some PI(4,5)P2 dependent ion channels (Rohacs et al., 2003; Nilius et al., 2006), their concentrations in the plasma membrane even in stimulated cells are much lower than that of PI(4,5)P2 (Balla, 2013). Thus, in most cases PI(4,5)P2 is likely to override their effects.

PI(3)P and PI(3,5)P2 are thought to be predominantly in membranes of intracellular organelles, such as endosomes and lysosomes. PI(3,5)P2 has been show to activate endolysosomal Transient Receptor Potential Mucolipin (TRPML) (Dong et al., 2010) and Two Pore Calcium (TPC) channels (Wang et al., 2012) with high affinity and specificity.

This review will provide an overview of the techniques to study phosphoinositide regulation of ion channels, focusing on the methods to assess the effects of PI(4,5)P2 on plasma membrane ion channels. Our goal here is to be comprehensive, which will go to the expense of technical detail. There have been several earlier reviews on the topic; the reader is also directed to those (Rohacs et al., 2002; Suh and Hille, 2008; Gamper and Rohacs, 2012; Rohacs, 2014; Robinson et al., 2019), as well as cited literature for more detail.

2. Applying phosphoinositides directly in excised patches or intact cells

PI(4,5)P2 is found in the inner leaflet of the plasma membrane. PI(4,5)P2 is high charged, therefore it is unlikely to cross biological membranes. Thus, for exogenously applied PI(4,5)P2 to reach the inner leaflet of the plasma membrane, it needs to be applied either to excised inside-out patches, or to be included in the patch pipette (intracellular) solution in whole cell patch clamp experiments.

2.1. Excised inside-out patches

Dependence of ion channel activity on PI(4,5)P2 was discovered by studying the mechanism of current rundown in excised inside out patches (Hilgemann, 1997). Rundown is a gradual loss of channel activity after patch excision (Fig. 2A,B) caused by the loss of PI(4,5)P2 from the patch membrane. Accordingly, rundown can be prevented by inhibition of lipid phosphatases, but not by inhibition of protein phosphatases (Huang et al., 1998; Sui et al., 1998). Rundown can be accelerated by applying compounds that bind to PI(4,5)P2, such as poly-Lysine (Lopes et al., 2002), see also Figure 2C, or an anti-PI(4,5)P2 antibody (Huang et al., 1998). Poly-Lys is available in a variety of chain lengths, and while it reliably inhibits PI(4,5)P2 dependent channels, its effect is not specific. This reagent is expected to bind all negatively charged lipids, not only PI(4,5)P2. The anti-PI(4,5)P2 antibody, by contrast, is thought to be specific but it is less reliable than Poly-Lys (Rohacs et al., 2002; Rohacs and Nilius, 2007). For example, it was reported to be inefficient on ion channels that were shown to be PI(4,5)P2 sensitive by other methods (Estacion et al., 2001; Liu and Liman, 2003).

Figure 2: Application of phosphoinositides and MgATP in excised inside out patches.

Figure 2:

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For all panels TRPV6 channels were expressed in Xenopus leavis oocytes, top traces show measurements at 100 mV, bottom traces at −100 mV, dashed lines show zero current. The beginning of the traces shows cell attached configuration (CA), the arrows show the establishment of the inside out (i/o) configuration, which is followed by current rundown. A: Channel activity after rundown was evoked by repeated alternating applications of 25 μM diC8 PI(4,5)P2 and 2 mM MgATP as indicated by the horizontal lines. The effect of diC8 PI(4,5)P2 develops quickly, and is readily reversible. The effect of MgATP develops slowly, and reverses slowly. The current increase after removal of MgATP is caused by relief from Mg2+ block; from (Zakharian et al., 2011). B: Channel activity is robustly stimulated by 10 μM AASt PI(4,5)P2, but 10 μM AASt PI(4)P had only minimal effect; from (Zakharian et al., 2011). C: Channel activity was evoked by various diC8 phosphoinositides (50 μM each), as indicated by the horizontal lines, numbers denote the positions phosphorylated in the inositol head group. At the beginning of the experiment 30 μg/ml Poly Lysine was applied to accelerate rundown; from (Thyagarajan et al., 2008).

After rundown, channel activity can be restored by applying MgATP (Fig. 2A) (Hilgemann and Ball, 1996; Sui et al., 1998), and the effect of MgATP can be prevented by inhibition of phosphoinositide 4-kinases (Xie et al., 1999b; Zakharian et al., 2011; Badheka, 2015). Application of MgATP is a simple and convenient way of regenerating endogenous PI(4,5)P2 by stimulating endogenous phosphoinositide kinases. This approach avoids perturbing the physicochemical properties of the membrane, which presumably takes place in the case of applying excess exogenous lipids.

Experiments with MgATP, however, provide only indirect proof that a channel is regulated by PI(4,5)P2. Direct proof requires applying phosphoinositides to the patch membrane. The two often used variants are synthetic, water-soluble dioctanoyl (diC8) and natural PI(4,5)P2. Early articles on the field used natural PI(4,5)P2 (Huang et al., 1998; Sui et al., 1998) purified from animal tissues, such a bovine, or porcine brain. Natural PI(4,5)P2 has a mixture of various acyl chains. Arachidonyl stearyl (AASt) is the most common, but the exact acyl chain combination shows some variability between different tissues (Traynor-Kaplan et al., 2017). The advantage of AASt PI(4,5)P2 is that it is the natural form. The disadvantage is its low solubility in water. As a consequence, this reagent needs to be sonicated before use (Rohacs et al., 2002) and forms micelles, the fusion of which to the patch membrane may induce non-specific effects. Given the slow incorporation in the membrane, AASt PI(4,5)P2 application usually activates channels slowly (Fig. 2B). Further, its effective concentration is difficult to control and can reach well above physiological concentrations. Finally, the effect of AASt PI(4,5)P2 is not, or only very slowly reversible. Despite these disadvantages, we recommend that the effect of natural PI(4,5)P2 is established during the characterization of a new, potentially PI(4,5)P2 sensitive ion channel for two reasons. One, in some rare cases, only the natural lipid, but not the short acyl chain analogues have an effect (Borbiro et al., 2015). Two, if only the short acyl chain analogues are used, one can argue that these synthetic lipids do not exist in nature, thus their effect does not reflect a physiologically occurring situation. Once the effect of both the natural and the short chain PI(4,5)P2 is established, the latter is far more practical to use for detailed characterization, as described below.

The synthetic diC8 PI(4,5)P2 and other diC8 phosphoinositides act quickly, and their effects are readily reversible on most channels (Fig. 2A,C). A major advantage of these lipids is that they can be applied to the same patch several times. Thus, concentration response relationships, as well as head group specificity profiles can be conveniently obtained. DiC8 phosphoinositides probably quickly equilibrate between the aqueous and lipid phase, and generally activate channels almost instantaneously (Fig. 2A,C). However, for some channels slower kinetics of activation (Rohacs et al., 2002) and deactivation (Lukacs et al., 2007) have also been reported. Additionally, several other synthetic versions of phosphoinositides are available from Cayman or Echelon, these include DiC4, DiC6 and DiC16, but these are less widely used than diC8 or AASt. In our hand, DiC4 PI(4,5)P2 did not activate inwardly rectifying K+ channels, and diC16 PI(4,5)P2 had similar effects to AASt PI(4,5)P2 (Rohacs et al., 1999; Rohacs et al., 2002).

Technical notes: Our lab routinely uses the Xenopus oocyte expression system to study the effects of phosphoinositides directly applied to excised patches. Given the large size of the oocyte (~1 mm diameter), large patches can be used to measure macroscopic currents. Technical details of this method have been described earlier (Rohacs, 2013). Mammalian cells, such as HEK293 cells have also been used by several laboratories (Ufret-Vincenty et al., 2011; Poblete et al., 2014). Divalent cations can aggregate PI(4,5)P2 into larger vesicles (Flanagan et al., 1997). Thus, leaving out Mg2+ from the lipid solutions is advisable. Additional details on handling lipids can be found in our earlier review (Rohacs et al., 2002).

2.2. Applying phosphoinositides in whole cell patch clamp through the patch pipette

Another way to apply phosphoinositides is to supplement the whole cell patch pipette solution with the lipid. After establishing the whole cell configuration, the pipette solution replaces much of the cytoplasm in a couple of minutes, and the added phosphoinositide presumably incorporates into cellular membranes, including the plasma membrane. This experimental setting is most useful when a stimulus is applied that decreases cellular PI(4,5)P2 levels, such as PLC activation, and the experimenter would like to test if the change in ion channel activity was caused by the decrease in PI(4,5)P2 levels. Figure 3 shows an example of such an experiment in peripheral sensory neurons of the dorsal root ganglia (DRG) (Liu et al., 2019). Endogenous cold- and menthol-sensitive TRPM8 channels were activated by consecutive pulses of the specific TRPM8 agonist WS12, before and during the third pulse, a cocktail of inflammatory mediators largely signaling through PLC was applied, inducing a ~50% inhibition in control neurons. In the group where diC8 PI(4,5)P2 was included in the patch pipette, the inflammatory cocktail did not induce any inhibition, supporting the idea that inhibition was induced by decreased PI(4,5)P2 levels (Fig. 3).

Figure 3. Applying PI(4,5)P2 through the whole cell patch pipette.

Figure 3

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Current traces measured at −60 mV in DRG neurons from mice expressing GFP under the promoter of TRPM8, from (Liu et al., 2019). Inward currents were evoked by repeated applications of the TRPM8 agonist WS12 (10 μM). To stimulate PLC and induce PI(4,5)P2 breakdown, a cocktail of agonists for Gq coupled cell surface receptors (100 μM ADP, 100 μM UTP, 500 nM bradykinin, 100 μM histamine, 100 μM serotonin, 10 μM PGE2, and 10 μM PGI2) was applied. A: control experiment, B: the whole cell patch pipette was supplemented with 100 μM diC8 PI(4,5)P2.

Most published experiments applying phosphoinositides through the whole cell patch pipette used the water soluble diC8 analogue (Trebak et al., 2009; Tang et al., 2010; Lukacs et al., 2013b), but AASt PI(4,5)P2 has also been used (Borbiro et al., 2015).

Including or excluding MgATP from the whole cell patch pipette is another way of influencing endogenous PI(4,5)P2 levels, as type III PI4K enzymes have low affinity for ATP (Km ~400–700 μM) (Balla and Balla, 2006). This is in sharp contrast to most protein kinases, which have high affinity for ATP, therefore they can function even when ATP is not provided in the whole cell patch pipette, presumably because the cell is capable of generating / buffering enough ATP for high affinity enzymes (Hilgemann, 1997). Leaving out MgATP from the patch pipette has been used to inhibit recovery from PLC mediated inhibition (Suh and Hille, 2002), or to potentiate PLC mediated inhibition (Borbiro et al., 2015), by preventing PI(4,5)P2 re-synthesis.

3. Manipulating endogenous phosphoinositides in intact cells

Activation of PLC enzymes is the best documented mechanism by which PI(4,5)P2 decreases in intact cells. Whether a physiological decrease in PI(4,5)P2 inhibits the activity of a given PI(4,5)P2 dependent ion channel depends on the apparent affinity of PI(4,5)P2 to the channel (Gamper and Shapiro, 2007). Channels with low PI(4,5)P2 affinity, such as GIRK channels can be readily inhibited by decreasing PI(4,5)P2 levels (Kobrinsky et al., 2000). Channels with high apparent affinity, such as Kir2.1, are less likely to be inhibited by physiological decreases in PI(4,5)P2 (Kobrinsky et al., 2000). Stimulation of channel activity by agonists may alter PI(4,5)P2 affinity by allosteric mechanisms, for example it was shown for TRPM8 channels that both cold and menthol increases the apparent affinity for PI(4,5)P2 (Rohacs et al., 2005).

3.1. PLC activation

PLC enzymes catalyze the hydrolysis of PI(4,5)P2 to IP3 and DAG (Fig. 1). The PLC family consists of 16 enzymes in mammals (Katan and Cockcroft, 2020). The three best characterized subfamilies are PLCβ-s, PLCγ-s and PLCδ-s.

Activation of cell surface receptors that couple to Gq, such as M1 muscarinic and bradykinin B2 receptors, activate PLCβ-beta (PLCβ1–4) isoforms. Stimulation of overexpressed Gq-coupled receptors induces a robust decrease in PI(4,5)P2 levels as monitored by fluorescent lipid sensors (Horowitz et al., 2005) or by mass spectrometry (Traynor-Kaplan et al., 2017). In response to activating endogenous receptors in native cells, the decrease in PI(4,5)P2 is generally smaller than that induced by overexpressed receptors. Whether PI(4,5)P2 decreases in response to stimulating endogenous receptors, and to what extent, is cell type and agonist specific (van der Wal et al., 2001; Winks et al., 2005). In peripheral sensory DRG neurons, for example, activation of endogenous bradykinin receptors was shown to induce either no decrease (Liu et al., 2010) or a small decrease (Lukacs et al., 2007) as monitored by a fluorescent PI(4,5)P2 sensor fusion protein, but application of a cocktail of different receptor agonists that stimulate PLC resulted in a more robust decrease in PI(4,5)P2 (Liu et al., 2019). When performing whole cell patch clamp measurements and stimulating GPCR-s, most publication include GTP (0.1 – 0.5 mM) in the patch pipette (Xie et al., 1999a; Suh and Hille, 2002; Horowitz et al., 2005), as GTP is required for G-protein activation. However, inclusion of GTP may not be essential, as there are also reports describing GPCR effects on ion channels without added GTP in the patch pipette (Liu and Qin, 2005; Quallo et al., 2017). The likely reason for this is the very high (nanomolar) affinity of Gα for GDP and GTP (Clapham, 1994). It is also possible that ATP in the patch pipette allows for some GTP regeneration

PLCγ-s (PLCγ1–2) are activated by receptor tyrosine kinases (RTK). Activation of overexpressed RTK-s has been used to inhibit PI(4,5)P2 dependent ion channels (Wu et al., 2002; Liu and Qin, 2005; Rohacs et al., 2005; Tong and Stockand, 2005; Badheka, 2015), but we are not aware of reports of endogenous RTK-s regulating PI(4,5)P2 dependent ion channels.

PLCδ-s (PLCδ1,3,4) are not activated by cell surface receptors per se. While most PLC isoforms require Ca2+ for their activity, PLCδ-s are activated by increased cytoplasmic Ca2+ alone (Allen et al., 1997; Yudin et al., 2016). We and others showed that activation of Ca2+ influx through the cold- and menthol-sensitive TRPM8 activates a PLCδ isoform leading to a decrease in PI(4,5)P2 levels and channel desensitization (Rohacs et al., 2005; Daniels et al., 2009). Similarly, the activation of the heat and capsaicin activated TRPV1 channels leads to a robust decrease in both PI(4,5)P2 and its precursor PI(4)P in both heterologous systems (Borbiro et al., 2015) and in native DRG neurons (Lukacs et al., 2013b). In both cases this decrease is more robust than that induced by activation of Gq-coupled receptors.

The compound m-3M3fbs was shown to activate PLC in intact cells as well as purified PLCβ2, β3, γ1, γ2 and δ1 (Bae et al., 2003). M-3M3fbs was shown to inhibit the activity of PI(4,5)P2 dependent TRPM8 (Daniels et al., 2009) and KCNQ channels (Horowitz et al., 2005). The most widely used inhibitor of PLC is U73122 (Balla, 2001). This compound is not very specific and has a number of side effects, including PI(4,5)P2 hydrolysis at higher concentrations. An excellent description of its proper use with PI(4,5)P2 dependent ion channels, including length of application, concentration and careful use of its inactive analogue U73343 can be found in (Horowitz et al., 2005). Edelfosine is another PLC inhibitor, but it also inhibits PKC (Horowitz et al., 2005).

3.2. Chemically inducible phosphatases

While PLC activation is the most likely physiological cause to decrease PI(4,5)P2 levels, the ensuing formation of second messengers and Ca2+ signal makes it difficult to attribute an effect on ion channels to decreased PI(4,5)P2 levels. What makes studying the effects phosphoinositides more straightforward than studying most other lipids, is the availability highly specific inducible phosphatases (and kinases) that can rapidly change PI(4,5)P2 levels without generating second messengers.

The most versatile and often used rapidly inducible phosphatases are based on the rapamycin-induced heterodimerization of the FKBP protein and the FRB fragment of mTOR (Fig. 4A–C). Two laboratories developed the prototype of this system independently. FRB is targeted to the plasma membrane and FKBP is fused to a PI(4,5)P2 5’-phosphatase. Upon the application of rapamycin (or its analogue) the FKBP-5’ phosphatase fusion protein translocates to the plasma membrane where it dephosphorylates PI(4,5)P2 converting it to PI(4)P. This approach provided a direct proof for the dependence of KCNQ channels (Suh et al., 2006) and TRPM8 channels (Varnai et al., 2006) on PI(4,5)P2 in intact cells. The introduction of this approach gave a substantial boost to the acceptance of PI(4,5)P2 as an ion channel regulator, and it has been used ever since to demonstrate PI(4,5)P2 regulation of a number of other ion channels, see Figure 4A–C for an example on TRPV6.

Figure 4: Inhibiting channel activity using inducible phosphoinositide phosphatases.

Figure 4:

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A: HEK293 cells were co-transfected with TRPV6 and the components of the rapamycin-inducible 5-phosphatase system. Inward monovalent currents at −60 mV were evoked by removing extracellular Ca2+; the translocation of the 5-phosphatase to the plasma membrane was initiated with the application of 100 nM rapamycin. B: control experiment in a cell transfected with TRPV6 and all components, except the 5-phosphatase domain. Figure redrawn from (Thyagarajan et al., 2008) C: cartoon of the rapamycin-inducible 5-phosphatase system. D-E: HEK293 cells were co-transfected with TRPV6 and ciVSP, inward currents at −60 mV were evoked by a Ca2+ free solution containing 1 mM EGTA. ciVSP was activated by voltage steps to 100 mV for 0.1, 0.3, 1 and 5 seconds, panel E shows the 0.3 s voltage step on a shorter time scale, redrawn from (Velisetty et al., 2016). F: Cartoon explaining ciVSP. The exact manner of conformational change that induces phosphatase activity is not known. For illustrative purposes the cartoon shows a simple movement towards the plasma membrane.

A further improvement to this approach was the development of pseudojanin, a rapamycin-inducible dual phosphatase that removes both the 5’ and 4’ phosphate from PI(4,5)P2 converting it to PI (Hammond et al., 2012). The pseudojanin clone is also available with single 4’ or 5’ phosphatase activities, as well as a negative control where both phosphatase activities were inactivated. A rapamycin-inducible PI(4)P-5 kinase (PIP5K) that increases PI(4,5)P2 concentration in the plasma membrane was also described, and was used to potentiate the activity of KCNQ channels (Ueno et al., 2011).

Technical notes: Application of rapamycin in these inducible phosphatase systems induces movement of the phosphatase from the cytoplasm to the plasma membrane, increasing its local concentration there. The cytoplasmic enzyme is also active, however, and if is expressed at high levels, it may also dephosphorylate PI(4,5)P2 to some extent even without the application of rapamycin. For this reason, in patch clamp experiments it is advisable to select cells that do not have very high expression (fluorescence) of the FKBP-phosphatase-RFP construct. It is also worth noting that the system developed by Tamas Balla used a different 5’-phosphatase from that developed by the Meyer lab, which may cause some differences between the results obtained with one or the other phosphatase. Even though the 5’ phosphatase in the dual phosphatase pseudojanin (Hammond et al., 2012) is the same as the one used by the Balla lab (Varnai et al., 2006), the 5’ phosphatase activity of pseudojanin is somewhat less than that of the originally described rapamycin-inducible 5’phosphatase (Badheka, 2015), most likely because pseudojanin is a larger construct. The inducible phosphatase system developed by Tobias Meyer originally used an analogue of rapamycin (iRap) but subsequent publications also successfully used it with the more readily available rapamycin (Klein et al., 2008; Daniels et al., 2009).

3.3. Voltage activated phosphatases (VSPs)

An alternative approach to chemically inducible phosphatases is voltage sensitive phosphoinositide 5’ phosphatases that convert PI(4,5)P2 into PI(4)P (Iwasaki et al., 2008). Two commonly used naturally occurring voltage sensitive phosphatase proteins are from the sea squirt ciona intestinalis (ciVSP) (Murata et al., 2005) and from zebrafish (drVSP) (Hossain et al., 2008). drVSP requires larger depolarizations than ciVSP (Okamura et al., 2009), and shows negligible activity at resting membrane potentials. Thus, reduced PI(4,5)P2 levels at baseline conditions are less of a concern with drVSP. Both VSPs have been used to demonstrate dependence of ion channel activity of PI(4,5)P2, (Falkenburger et al., 2010), see also example in Figure 4D,E. There are two major advantages of VSP-s over chemically inducible phosphatases. First, their effect is quickly reversible in 10–20 seconds (Falkenburger et al., 2010), which allows performing complex protocols in the same cell (Lukacs et al., 2013b). Second, they can be used in a graded fashion by increasing either the length of the depolarizing pulses or the extent of depolarization, see Figure 4D–F for TRPV6 and (Liu et al., 2019) for TRPM8 channels. The disadvantage of VSP-s is that they only remove the 5’ phosphate, depleting PI(4,5)P2, but generating excess PI(4)P and thus they may not inhibit very efficiently channels that have substantial activation by PI(4)P (Lukacs et al., 2013b).

The phosphatase domain of VSP proteins is homologous to PTEN, which removes the 3’ phosphate from PI(3,4,5)P3. ciVSP on the other hand dephosphorylates both PI(4,5)P2 and PI(3,4,5)P3, and it was shown that it removes the 5’ from PI(4,5)P2 (Iwasaki et al., 2008) as well as from PI(3,4,5)P3 (Kurokawa et al., 2012). At stronger depolarization, ciVSP also removes the 3’ from PI(3,4)P2 (Kurokawa et al., 2012). This complex specificity profile however rarely gains importance in studying ion channel activity, as in resting conditions PI(3,4)P2 and PI(3,4,5)P3 are barely detectable, and PI(4,5)P2 is the dominant phosphoinositide, and it can be safely assumed that ciVSP converts this lipid into PI(4)P upon depolarization.

Technical notes: As mentioned earlier, basal activity of ciVSP in cells with depolarized membrane potentials, e.g. HEK293 cells can cause problems such as decreased basal current amplitudes (Zhelay et al., 2018) and long term effects of decreased PI(4,5)P2. This can be circumvented either by using drVSP, which requires larger depolarizations to activate (Falkenburger et al., 2010), and/or by co-expressing K+ channels with constitutive activity that will shift the resting membrane potential to negative values (Lukacs et al., 2013b). Even though several labs reported use of VSP-s in the Xenopus oocyte expression system (Iwasaki et al., 2008; Tang et al., 2015), in our hands the oocytes became extremely leaky after expressing ciVSP, which essentially prevented us from using this system in oocytes (unpublished). There is no single established voltage protocol to activate VSPs. Figure 4D shows our often used protocol with 4 pulses of increasing length that worked well for studying TRP channels (Velisetty et al., 2016; Liu et al., 2019). Other laboratories used a variety of different protocols such as keeping the cell at depolarized holding potentials followed by fast voltage ramps to measure channel activity (Toth et al., 2015), or episodic voltage steps, or voltage ramps (Tang et al., 2015). When designing a voltage protocol for studying an ion channel, a number of factors need to be taken into consideration, including the relatively fast (10–20 s) resynthesis of PI(4,5)P2, the voltage sensitivity of the given VSP used, and the optimal voltage to measure channel activity (Falkenburger et al., 2010). Recently, an enhanced version of drVSP has been engineered, with increased phosphatase activity and improved plasma membrane localization (Kawanabe et al., 2020).

3.4. Light-induced phosphatases

An additional rapidly inducible phosphatase system has also been developed based on the light-induced dimerization between cryptochrome 2 (CRY2) and the transcription factor CIBN, two plant proteins (Idevall-Hagren et al., 2012). Similar to the rapamycin inducible phosphatase systems, here the 5’-phosphatase domain of OCRL translocates to the plasma membrane in response to blue light where it dephosphorylates PI(4,5)P2 and PI(3,4,5)P3. Activation of this system has been shown to inhibit PI(4,5)P2 dependent KCNQ channels (Idevall-Hagren et al., 2012), TRPC4 and TRPC5 channels (Ko et al., 2019) and ENaC channels (Archer et al., 2020).

3.5. Pharmacological tools to inhibit phosphoinositide kinases

Before the development of the rapidly inducible phosphatases, lipid kinase inhibitors were used in addition to, and in combination with PLC activation to decrease cellular PI(4,5)P2 levels. PI(4,5)P2 is generated by sequential phosphorylation of PI by PI4K and PIP5K enzymes (Fig. 1). For a long time, PI4K had no specific inhibitors, but two PI3K inhibitors, wortmannin and LY294002 were shown to inhibit type III PI4 Kinases (Balla, 2001). Prolonged incubation with wortmannin (> 10 μM for 1–2 hours) have been used to inhibit PI(4,5)P2 dependent ion channels (Zhang et al., 2003; Badheka, 2015). High concentration of wortmannin have also been used to inhibit recovery from PLC mediated inhibition of ion channels, to support the claim that channel inhibition was mediated by PI(4,5)P2 depletion (Xie et al., 1999a; Suh and Hille, 2002; Liu et al., 2005; Lopes et al., 2005). Both wortmannin and LY294002, as well as the PI3K inhibitor PIK93 that inhibits PI4KIIIβ, and the selective PI4K-IIIα inhibitor A1, were also used to demonstrate that MgATP-induced recovery from rundown in excised patches is mediated by PI(4,5)P2 synthesis (Xie et al., 1999b; Zakharian et al., 2011; Badheka, 2015).

New selective PI4K-IIα inhibitors (Li et al., 2017; Sengupta et al., 2019), and PIP5K inhibitors ISA-2011B (Semenas et al., 2014) and UNC3230 (Wright et al., 2015) have also been reported, but their effects on ion channel function have not yet been reported.

4. Monitoring phosphoinositides

Traditionally, phosphoinositides were measured in cells incubated with radioactively labeled inositol, and the lipids were separated on thin layer chromatography, or other separation methods (Gonzalez-Sastre and Folch-Pi, 1968; Rana and Hokin, 1990; Singh and Jiang, 1995). These techniques went out of favor, due to several reasons including the lack of subcellular information as well as the need for long labeling times to reach equilibrium. Here, we will focus on detecting phosphoinositides in live cell imaging, using fluorescent protein probes, and also briefly describe other techniques.

4.1. Genetically encoded fluorescent phosphoinositide probes

The prototype of this approach was described independently in 1998 by the laboratories of Tamas Balla (Varnai and Balla, 1998) and Tobias Meyer (Stauffer et al., 1998). Both labs fused GFP to the Pleckstrin Homology domain of PLCδ1, which binds PI(4,5)P2 with high specificity and affinity. Because it binds PI(4,5)P2, this fluorescent protein is located at the plasma membrane under resting conditions. PLC activation by either Gq-coupled cell surface receptors, or by large Ca2+ signals lead to the translocation of the probe to the cytoplasm due to the breakdown of PI(4,5)P2, which can be followed by confocal microscopy and other techniques, see last five paragraphs of this section. These publications were followed by the development of a variety of similar probes to monitor distribution of various phosphoinositides; a comprehensive list of these can be found in recent reviews (Varnai et al., 2017; Wills et al., 2018). Here, we will briefly describe some of the phosphoinositide probes that are relevant to the study of plasma membrane ion channels.

The PH domain of PLCδ1 binds not only PI(4,5)P2, but also the head group of the lipid, IP3, raising the possibility that its translocation after PLC activation is induced not only by decreasing PI(4,5)P2 levels but also by the increased IP3 that would compete with PI(4,5)P2 for binding to the probe (Xu et al., 2003). While this claim has been debated (Szentpetery et al., 2009), there are other PI(4,5)P2 probes that do not have this uncertainty. The most frequently used alternative PI(4,5)P2 probe that is devoid of this problem is based on the PI(4,5)P2 biding domain of the tubby protein (Santagata et al., 2001), which does not bind IP3 (Quinn et al., 2008). Since this probe binds PI(4,5)P2 with very high affinity, most publications used its mutant version (R332H) that has lower affinity for PI(4,5)P2 and translocates more readily in response to receptor stimulation (Quinn et al., 2008). An alternative option is a probe based on the PH domain of PLCδ4, which also shows negligible binding to IP3. This probe has a lower affinity than the PLCδ1-based probe and thus, shows a somewhat weaker plasma membrane localization (Lee et al., 2004). It has been used to monitor PI(4,5)P2 changes together with ion channel activity (Yudin et al., 2011).

DAG and IP3 are breakdown products of the hydrolysis of PI(4,5)P2 by PLC. A number of fluorescent protein-based probes have been developed for both these second messengers. The most frequently used DAG probe is the GFP tagged C1 domain of PKCγ (Oancea et al., 1998; Horowitz et al., 2005). Other sensors are based on a circularly permutated GFP placed between the C1 domain and pseudosubstrate domain of PKCδ. Different versions of this probe respond to DAG with increased or decreased fluorescence (Tewson et al., 2012), and they are commercially available from Montana Molecular. A probe with higher affinity for DAG based on the tandem C1 domain of PKD fused to GFP was also developed (Kim et al., 2011) and was used to study ion channel regulation by PKC (Liu et al., 2020).

PI(4)P is the precursor of PI(4,5)P2. It is also a substrate for most PLC isoforms. Strong PLC stimulation usually results in a robust decrease in PI(4,5)P2, and a smaller or equal decrease in PI(4)P, while weaker PLC stimulation leads to a selective decrease in PI(4,5)P2 (Lukacs et al., 2013b). Developing specific probes for PI(4)P has been challenging, as very few proteins bind to PI(4)P but not to PI(4,5)P2. The tandem PH domain of OSH2 (Balla et al., 2008) has been used in several publications to assess changes in plasma membrane PI(4)P levels (Lukacs et al., 2007; Hammond et al., 2012; Lukacs et al., 2013b). This probe, however binds to both PI(4,5)P2 and PI(4)P and depletion of both lipids is required for its translocation. When used together with a specific PI(4,5)P2 probe, one can deduce PI(4)P changes upon receptor stimulation by comparing these two signals (Hammond et al., 2012; Lukacs et al., 2013b). More recently, a specific probe for PI(4)P has been developed based on the PI(4)P binding domain of the SidM from Legionella pneumophila (Hammond et al., 2014).

PI is the precursor of PI(4)P, and for a long time no specific probes were available to detect its subcellular localization. Recent reports describe a new probe based on the bacterial PI-PLC from B. Cereus and concluded that the plasma membrane contains minimal amounts of PI, and most of this lipid is located in the Golgi, the mitochondria, and to some extent in the ER (Pemberton et al., 2020; Zewe et al., 2020).

Phosphoinositides containing 3’ phosphate are generated by various PI3 Kinase enzymes. PI(3,4,5)P3, and PI(3,4)P2 are mainly found in the plasma membrane. Even though they activate some PI(4,5)P2 dependent ion channels (Rohacs et al., 2003; Nilius et al., 2006), the much higher concentration of PI(4,5)P2 makes it unlikely that those effects have physiological relevance. We are not aware of specific activation of ion channels by these lipids.

PI(3,5)P2 and PI(3)P are mainly found in intracellular membranes, and PI(3,5)P2 is known to be a specific activator of several ion channels located in intracellular membranes such as endolysosomal TPC channels (Wang et al., 2012) and TRPML channels (Dong et al., 2010). Fluorescent probes have also been developed for most of these 3’ phosphorylated phosphoinositides. We refer to two recent reviews for details (Varnai et al., 2017; Wills et al., 2018).

Most of the probes described so far bind specifically to phosphoinositides in the plasma membrane, and a decrease in their level can be deduced from the translocation of the probe to the cytoplasm. There are multiple techniques that can detect this translocation. Here, we will briefly describe them and discuss their advantages and disadvantages.

Confocal microscopy is still the gold standard to look at translocation (Fig. 5A–C) as its focal plane is a lot thinner than for a traditional fluorescence microscope, allowing a better assessment of plasma membrane and cytoplasmic localization. The disadvantage of using a confocal microscope is the price of the equipment and the relatively slow speed of most confocal microscopes. While combining confocal microscopy with patch clamping is feasible, it requires a dedicated rig, and it is usually not a realistic option for equipment located in core facilities. A robust translocation from the plasma membrane to the cytoplasm can be detected with a regular epifluorescence microscope, but due to the cross talk from out of focus fields, smaller signals are difficult to detect.

Figure 5. Monitoring PI(4,5)P2 levels with fluorescent fusion protein sensors.

Figure 5

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A-C detecting translocation of the Tubby-YFP PI(4,5)P2 sensor from the plasma membrane to the cytoplasm using confocal microscopy in HEK293 cells expressing the tubby-YFP PI(4,5)P2 sensor and M1 muscarinic receptors, redrawn from (Liu et al., 2019). A: confocal images before and after the application of the M1 agonist carbachol (100 μM). B: Fluorescence intensity before and after carbachol plotted at the cross section of the cell as indicated by the white lines in panel A. C: time course of the change in fluorescence at the plasma membrane and the cytoplasm. D: Cartoon explaining FRET-based monitoring of the translocation of the PLCδ-PH domain tagged with CFP and YFP, see details in main text. E: Combined emission spectra of YFP and CFP in the absence (cyan) and presence (orange) of FRET. F: Measurement in a cell expressing the CFP and YFP tagged PI(4,5)P2 sensor and M1 muscarinic receptors, fluorescence traces at 480 nm (cyan) and 535 nm (orange) emission wavelengths are shown, the black trace shows the ratio of the two traces. G: Representative TIRF images before and after the application of 100 μM carbachol in a cell expressing the tubby-YFP PI(4,5)P2 sensor and M1 muscarinic receptors, redrawn from (Liu et al., 2019). I: cartoon explaining TIRF; see main text for details; H: time course of the TIRF fluorescence intensity from panel A.

PLCδ1-based PI(4,5)P2 sensors that can be readily detected with a standard fluorescence microscope, or even a plate reader, have also been developed (Tewson et al., 2013) and are commercially available from Montana Molecular. These probes are based on dimerization-dependent green and red fluorescent proteins, and they respond with a decrease in fluorescence once translocated from the plasma membrane to the cytoplasm. The commercial versions of these probes are supplied as baculovirus particles that can be used to transduce cells that are difficult to transfect with other techniques. We have used this system to detect changes in PI(4,5)P2 in DRG neurons (Liu et al., 2019).

Fluorescence Resonance Energy Transfer (FRET) is also often used as a readout of translocation-based (Fig. 5D–F) and conformation change-based fluorescent probes. FRET is a non-radiative energy transfer that takes place when the emission spectra of a donor fluorophore overlaps with the excitation spectra of an acceptor fluorophore. If the two fluorescent molecules are in molecular proximity, and in the proper orientation, the excited electron of the donor transfers its energy to the acceptor, which emits a photon at its characteristic emission wavelength (Siegel et al., 2000; Lakowicz, 2006). Changes in FRET can be monitored by exciting the donor, and measuring the emission of the acceptor and the donor simultaneously, which change in opposite directions when FRET changes (Fig. 5F). These ratiometric measurements offer the advantage of correcting for non-specific changes in the individual wavelengths (Fig. 5F). The first report of a FRET-based probe to monitor phosphoinositide changes used the PLCδ1 PH domain tagged with CFP and with YFP (van der Wal et al., 2001). In cells transfected with PLCδ1-PH-CFP and PLCδ1-PH-YFP, these fluorescent fusion proteins localize to the plasma membrane, and at sufficiently high expression levels they undergo FRET, presumably because they are located close enough in the quasi 2 dimensional plane of the plasma membrane. Upon PLC activation, the probes translocate to the cytoplasm, their distance increases, and FRET decreases. CFP/YFP FRET-based PI(4,5)P2 probes using either the PLCδ4 PH domain (Yudin et al., 2011) or tubby (Borbiro et al., 2015) have been used to study ion channel regulation. To detect dynamic changes in FRET in real time, a fluorescence setup is required that is capable of measuring two different emission wavelengths simultaneously. This can be achieved by using two photomultipler tubes (PMT) and a dichroic beam splitter. These are relatively inexpensive to add to an existing patch clamp setup on an epifluorescence microscope. The analogue signal from the PMT-s can be connected to the A/D converter of the patch clamp rig, providing simultaneous detection of FRET with ion currents with the same time resolution (Yudin et al., 2011). The disadvantage of the PMT-based system is that it does not provide spatial resolution, and only one cell can be studied in one experiment. Camera based solutions offer the possibility of studying several cells on the coverslip at once. Splitting the emission wavelength can be achieved either with a dichroic mirror and an image splitter, or a small filter wheel in front of the camera. FRET-based sensors are also available to measure IP3 levels, these include IRIS (Matsu-ura et al., 2006; Yudin et al., 2011) various versions of LIBRA (Tanimura et al., 2009) and several other improved probes (Gulyas et al., 2015).

Total Internal Reflection Fluorescence (TIRF) microscopy is another alternative to monitor fluorescent probe translocation from the plasma membrane (Fig. 5G–I). In this method, the excitation light hits the cover glass-cell interface at a critical angle so that it is completely reflected. The generated evanescent wave only penetrates the bottom of the cell to less than 100 nm, predominantly exciting the plasma membrane and a narrow sub plasmalemmal region. Translocation of a plasma membrane bound fluorescent probe to the cytoplasm will manifest as a drop in fluorescence. This method has also been used to monitor PI(4,5)P2 levels (Yao and Qin, 2009). One common concern with TIRF, especially when using water soluble agonists, is that the agonist has to reach the narrow space between the bottom of the cell and the cover glass to stimulate receptors at the right place. In our hands stimulation of M1 muscarinic receptors with the water soluble carbachol induced a rapid translocation of the YFP-tagged tubby PI(4,5)P2 sensor in HEK293 cells (Fig. 5G,I) (Liu et al., 2019). TIRF microscopes are expensive, and the technique is very sensitive to small drifts in focus thus a focus stabilizing mechanism is advisable, which adds to the overall equipment cost.

Bioluminescence Resonance Energy transfer (BRET) is another alternative that can be used to detect phosphoinositide changes in cell populations using a plate reader. In this assay, the donor is the chemiluminescent protein luciferase, which is fused to the lipid binding domain, and the acceptor is the fluorescent protein Venus, which is targeted to the plasma membrane, (Varnai et al., 2017). BRET does not use an excitation light, as the donor energy is created by the luciferase enzyme acting on its substrate, coelenterazine. The emitted light intensity from a single cell is too weak to be detected, thus this technique can only be used in cell populations. Its overall signal to noise ratio however exceeds that of the single cell measurements described before, but is time resolution is a lot lower than that of FRET. As it can be used in a multi well plate reader, it is also a higher throughput technique than the previously described ones. See Varnai et al 2017 and references therein for more details.

4.2. Other techniques to detect phosphoinositide changes

Hilgemann and colleagues developed a non-radioactive approach to measure phosphoinositide levels in intact tissues. This method is based on deacylating the lipids to glycero-head groups, separating them by anion-exchange chromatography and detecting them by suppressed conductivity measurements (Nasuhoglu et al., 2002). The advantage of this method is that the cells are not perturbed by introducing exogenous probes. The disadvantage is that it does not give information on subcellular localization.

Mass spectrometry has recently been used to measure phosphoinositide levels (Robinson et al., 2019). An advantage of this technique is that it can differentiate between various combinations of acyl chains, (Traynor-Kaplan et al., 2017), and the disadvantage is that it does not offer subcellular resolution and does not differentiate between phosphoinositides with the same number of phosphates, e.g. PI(4,5)P2 and PI(3,4)P2.

5. Techniques using purified proteins

The techniques described so far studied ion channels in a cellular environment. To demonstrate direct effect of a phosphoinositide on the channel, techniques with purified proteins provide more conclusive evidence.

5.1. Binding experiments:

In the early days of the field many binding techniques were developed mostly using isolated cytoplasmic fragments to demonstrate direct binding of PI(4,5)P2 to ion channels, as well as to attempt identifying PI(4,5)P2 binding sites (Suh and Hille, 2008). PI(4,5)P2, due to its high charge density, tends to bind to essentially any positively charged surface (McLaughlin and Murray, 2005). Binding to an isolated peptide fragment, therefore, does not necessarily mean that the lipid binds to the same place in the context of the full length channel. For example, if the peptide fragment is not close to the plasma membrane, where PI(4,5)P2 is located, physiological binding of this lipid is unlikely. More recently soluble binding assays with fully assembled ion channels have also been developed allowing determination of binding affinities for phosphoinositides (Robinson et al., 2019). Mass spectrometry can also be used to identify lipids bound to ion transmembrane proteins (Robinson et al., 2019). This approach has not yet been applied to ion channels and phosphoinositides, but PI(4,5)P2 was recently identified to be bound to G-protein coupled receptors (Yen et al., 2018), pointing to potential feasibility of this approach.

5.2. Planar lipid bilayers & lipid vesicles

Demonstrating direct binding of the lipid to a channel protein does not prove that the binding is biologically relevant. Excised patches contain a number of proteins other than the expressed channel, including cytoskeletal elements, and lipid kinases and phosphatases. Therefore, an effect of PI(4,5)P2 in excised patches can be mediated by other proteins. For these reasons showing activation of a purified reconstituted channel may serve as an ultimate proof for direct activation of a channel by phosphoinositides. This approach has been applied to TRPM8 (Zakharian et al., 2010), TRPV6 (Zakharian et al., 2011) and GIRK channels (Leal-Pinto et al., 2010) using the planar lipid bilayer technique (Zakharian, 2019), as well as to Kir2.1 and Kir2.2 (Cheng et al., 2011) and TRPV1 channels (Cao et al., 2013) using lipid vesicles. While this approach is ideal to demonstrate the direct effect of PI(4,5)P2 on an ion channel, it is not free from disadvantages. The most important of these is that the lipid composition of the plasma membrane is difficult to mimic, especially given the asymmetrical distribution between the inner and outer leaflets, plus not all lipid combinations form stable bilayers. In the case of TRPV1, two labs using purified channel protein but two different techniques (planar lipid bilayer vs. patching lipid vesicles) and different lipid compositions, arrived at opposite conclusions on whether PI(4,5)P2 activates (Lukacs et al., 2013a) or inhibits (Cao et al., 2013) the TRPV1 channel; see (Rohacs, 2015) for discussion.

5.3. Structure determination:

Before the recent cryoEM structural revolution, a lot of effort has been dedicated to locate PI(4,5)P2 binding sites in various ion channels using site directed mutagenesis and functional assays (Suh and Hille, 2008). The consensus from these studies was that the negatively charged head group of PI(4,5)P2 binds to positively charged residues in the cytoplasmic domains of the channel. When the structure of some of these ion channels were determined together with PI(4,5)P2, this key conclusion was confirmed. Not all details matched, however, as some of the residues identified with mutagenesis were not in direct contact with PI(4,5)P2. Thus, their mutations likely affected PI(4,5)P2 activation via indirect effects. In the structural era, the ideal approach to locate the binding site for PI(4,5)P2 is to determine the structure of the protein together with the lipid. In the absence of a co-structure, molecular docking to a known structure or a model based on a close homolog, combined with mutational analysis is a potentially useful approach. If the cytoplasm plasma membrane interface is not too large, computational approaches may successfully identify a PI(4,5)P2 binding site, as for example for TRPV6 (Hughes et al., 2018).

The first co-structures of PI(4,5)P2 were determined for Kir2.2 and Kir3.2 (GIRK2) channels using crystallography (Hansen et al., 2011; Whorton and MacKinnon, 2013). The first cryoEM structure of an ion channel with a phosphoinositide was TRPV1, where PI was identified in the vanilloid binding site (Gao et al., 2016), but the functional relevance of this binding site is quite complex (Yazici, 2019). The first cryoEM structure where PI(4,5)P2 appeared on the structure and induced a conformation change compatible with channel opening was TRPV5 (Hughes et al., 2018), followed by KCNQ1 (Sun and MacKinnon, 2020). The structure of TRPM8 with PI(4,5)P2 is also available (Yin et al., 2019), as well as the structures of the intracellular TPC1 (She et al., 2018) and TRPML1 (Fine et al., 2018) with PI(3,5)P2. Unexpectedly, the structure of GABAA receptors contained an endogenous PI(4,5)P2 molecule (Laverty et al., 2019), but the functional relevance of this lipid interaction is not clear.

6. Summary

Here, we provided an overview of the techniques to study phosphoinositide regulation of ion channels. Convergence of evidence from multiple approaches always provides a more convincing case than relying on one technique. This is especially true for demonstrating the PI(4,5)P2 dependence of an ion channel. In many cases, such as inwardly rectifying K+ channels, KCNQ channels, TRPM8, and TRPV5/6, we have fully congruent cases for PI(4,5)P2 dependence, based on the use of the majority of techniques described here, including co-structures with PI(4,5)P2. For some channels, such as TRPV1, however, the picture is more elusive, despite the use of essentially all these techniques described here, highlighting the inherent complexity of the biology of phosphoinositides.

Acknowledgements

The work in the Rohacs lab is supported by NIH Grants NS055159, GM093290, and GM131048

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