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Jan 8, 2013 - and Makielski, 1997; Kim and Bang, 1999; Hilgemann et al., 2001 ...... in lipid bilayer properties (Andersen and Koeppe, 2007;. Lundbæk et al.
Research Article

Phosphoinositides alter lipid bilayer properties Radda Rusinova,1 E. Ashley Hobart,1 Roger E. Koeppe II,2 and Olaf S. Andersen1 1

Department of Physiology and Biophysics, Weill Cornell Medical College, New York, NY 10065 Department of Chemistry and Biochemistry, University of Arkansas, Fayetteville, AR 72701

The Journal of General Physiology

2

Phosphatidylinositol-4,5-bisphosphate (PIP2), which constitutes 1% of the plasma membrane phospholipid, plays a key role in membrane-delimited signaling. PIP2 regulates structurally and functionally diverse membrane proteins, including voltage- and ligand-gated ion channels, inwardly rectifying ion channels, transporters, and receptors. In some cases, the regulation is known to involve specific lipid–protein interactions, but the mechanisms by which PIP2 regulates many of its various targets remain to be fully elucidated. Because many PIP2 targets are membrane-spanning proteins, we explored whether the phosphoinositides might alter bilayer physical properties such as curvature and elasticity, which would alter the equilibrium between membrane protein conformational states—and thereby protein function. Taking advantage of the gramicidin A (gA) channels’ sensitivity to changes in lipid bilayer properties, we used gA-based fluorescence quenching and single-channel assays to examine the effects of long-chain PIP2s (brain PIP2, which is predominantly 1-stearyl-2-arachidonyl-PIP2, and dioleoyl-PIP2) on bilayer properties. When premixed with dioleoyl-phosphocholine at 2 mol %, both long-chain PIP2s produced similar changes in gA channel function (bilayer properties); when applied through the aqueous solution, however, brain PIP2 was a more potent modifier than dioleoyl-PIP2. Given the widespread use of short-chain dioctanoylphosphoinositides, we also examined the effects of diC8-phosphoinositol (PI), PI(4,5)P2, PI(3,5)P2, PI(3,4)P2, and PI(3,4,5)P3. The diC8 phosphoinositides, except for PI(3,5)P2, altered bilayer properties with potencies that decreased with increasing head group charge. Nonphosphoinositide diC8 phospholipids generally were more potent bilayer modifiers than the polyphosphoinositides. These results show that physiological increases or decreases in plasma membrane PIP2 levels, as a result of activation of PI kinases or phosphatases, are likely to alter lipid bilayer properties, in addition to any other effects they may have. The results further show that exogenous PIP2, as well as structural analogues that differ in acyl chain length or phosphorylation state, alters lipid bilayer properties at the concentrations used in many cell physiological experiments . INTRODUCTION

Phosphoinositides regulate the function of structurally and functionally distinct membrane proteins (Hilgemann et al., 2001; McLaughlin et al., 2002; Suh and Hille, 2008; Logothetis et al., 2010). This regulation may result from phosphoinositide binding to the target protein, and several specific phosphoinositide-binding domains have been identified. Specific binding mainly (but not exclusively; Rosenhouse-Dantsker and Logothetis, 2007) involves the interaction of positively charged residues with the negatively charged phosphatidylinositol-4,5-bisphosphate (PIP2) phosphates (McLaughlin and Murray, 2005; Hurley, 2006; Suh and Hille, 2008; Hansen et al., 2011; Leonard and Hurley, 2011; Whorton and MacKinnon, 2011). In many cases, the regulation seems to be more general, however, in the sense that other anionic phospholipids can substitute for the phosphoinositides (Fan and Makielski, 1997; Kim and Bang, 1999; Hilgemann et al., 2001; Zhang et al., 2003; Vaithianathan et al., 2008), suggesting that the regulation involves less specific electrostatic interactions with a positively charged region—but

leaving open the question of how the phosphoinositides alter function. The unifying feature among the diverse class of membrane proteins that are regulated by PIP2 is that many are embedded in a lipid bilayer, and changes in bilayer properties are known to alter membrane protein function (Bienvenüe and Marie, 1994; Lee, 2004; Andersen and Koeppe, 2007; Lundbæk et al., 2010a). In this context, it is important that PIP2 has 50% polyunsaturated fatty acids (Marsh, 2012b), which would tend to reduce the bilayer bending modulus (Rawicz et al., 2000; Bruno et al., 2013). PIP2 constitutes 1% of total cell lipid, varying among different cell types (Ferrell and Huestis, 1984; Nasuhoglu et al., 2002; Wenk et al., 2003; Xu et al., 2003; Horowitz et al., 2005; Li et al., 2005; Hilgemann, 2007). Because the total cell lipid profile includes intracellular membranes, and PIP2 is localized predominantly to the inner leaflet of the plasma membrane (Balla and Várnai, 2002), the area-averaged mole percentage of PIP2 in the plasma membrane is likely to be >1% (Hilgemann,

Correspondence to Radda Rusinova: r­a­r­2­0­2­1­@­m­e­d­.­c­o­r­n­e­l­l­.­e­d­u­ Abbreviations used in this paper: ANTS, 8-aminonaphthalene-1,3,6trisulfonate; CMC, critical micelle concentration; gA, gramicidin A; LUV, large unilamellar vesicle.

© 2013 Rusinova et al.  This article is distributed under the terms of an Attribution– Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.rupress.org/terms). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).

The Rockefeller University Press  $30.00 J. Gen. Physiol. Vol. 141 No. 6  673–690 www.jgp.org/cgi/doi/10.1085/jgp.201310960

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ROUGH GALLEY PROOF

percentage of PIP2 in the plasma membrane is likely to conformational equilibrium of membrane proteins. (It be >1% (Hilgemann, 2007)—and local PIP2 concentrais given that Gdef will change, but the changes may or tions may be even higher (Wang and Richards, 2012). not be sufficient to cause measurable changes in out 2007)—and local PIP2 concentrations may be even higher mayanalogues alter bilayer properties and, if so, to parse Given the lack of a clear mechanism for many of the protein function.) We therefore set out to determine (Wang and Richards, 2012). Given the lack a clear which bilayer properties are affected. PIPmechanism it thus important to exPIPdifferent structural analogues alter 2-mediated effects, 2 and its probes for many of thebecomes PIP2-mediated effects, it thus whether Many have been used to bilayer characterplore whether changes in extant, or exogenously approperties and, if so, to parse out which bilayer becomes important to explore whether changes in exize the changes in bilayer properties that arepropercaused by plied, at relevant are affected. 2s alter bilayer tant,PIP or exogenously applied,properties PIP2s alter bilayer properties ties drugs and other amphiphiles (Seddon et al., 2009). For concentrations, in which case some of PIP may different probes used toAcharacterize 2’s effects at relevant concentrations, in which case some of PIP2’s Many the present studies, wehave usedbeen gramicidin (gA) channels be caused by nonspecific bilayer-mediated mechanisms, the changes in bilayer properties that are caused by effects may be caused by nonspecific bilayer-mediated as probes that serve as prototypical bilayer-spanning as recently was proposed in the case of invertebrate drugs and other amphiphiles (Seddon et al., 2009). For mechanisms, as recently was proposed in the case of inverproteins, which experience minimal interference from phototransduction (Hardie and Franze, 2012). the present studies, we used gramicidin A (gA) chantebrate phototransduction (Hardie and Franze, 2012). specific lipid–protein interactions. gA channels form by Bilayer-spanning proteins are are energetically coupled as transmembrane probes that serve as prototypical bilayer-span-moBilayer-spanning proteins energetically coupled nelsthe dimerization of nonconducting to the host lipid bilayer through hydrophobic interacning proteins, which experience minimal interference to the host lipid bilayer through hydrophobic interacnomeric subunits residing in apposing bilayer leaflets tions between the domains and andthe thelipid from specific lipid–protein interactions. gA channels tions between thetransmembrane transmembrane domains (O’Connell et al., 1990). Channel formation is associated lipidbilayer bilayerhydrophobic hydrophobiccore, core,which whichcan caninfluence influencethe thelipid form by the transmembrane dimerization of et nonconwith a local bilayer compression (Elliott al., 1983; lipidpacking packing around protein (Killian et al., 1989, monomeric subunits in which apposing bi- gA around thethe protein (Killian et al., 1989, 1996) ducting Huang, 1986; Harroun et residing al., 1999), makes 1996) or the protein conformation (Keller et al., 1993; layer leaflets (O’Connell et al., 1990). Channel or the protein conformation (Keller et al., 1993; Brown, channels sensitive to changes in bilayer properties Brown, Greathouse al., 1994). Thisreorganization lipid reorformation is associated withLundbæk a local bilayer 1994;1994; Greathouse et al.,et1994). This lipid (Andersen et al., 2007; et al.,compression 2010a,b). ganization (bilayer deformation) has an associated en(Elliott et al., 1983; Huang, 1986; Harroun et al., 1999), the Using gA channels as probes, we have explored (bilayer deformation) has an associated energetic cost, ergetic cost, the bilayer deformation energy (G ). If which makes gA channels sensitive to changes in bilayer def effects of phosphoinositides on lipid bilayers while varythe bilayer deformation energy (Gdef). If protein con(Andersen et al.,membrane 2007; Lundbæk al., protein conformational changes (e.g., state II) I toalter properties ing the method by which PIP2 wasetmanipuformational changes (e.g., from statefrom I to state 2010a,b). statethe II)local alterlipid the bilayer local lipid bilayer deformations, the deformations, the resulting differlated, the acyl chain composition, and the number and I → II gA channels as probes, wethe have explored the 1). resulting difference in the bilayer deformation energy, ence in the bilayer deformation energy, ∆G , be- Using posi­tions of phosphate groups on inositol ring (Fig. bilayer I → II ∆G comes , becomes the bilayer contribution to the total effects of phosphoinositides on lipid bilayers while varythe bilayer contribution to the total free energy For comparison, we also examined the bilayer-perturbing bilayer I → II (∆G I → II ) freeofenergy of the conformational the method by related which membrane 2 was maniputhe conformational transitiontransition of other short-chainPIP phospholipids. (∆G total ) (Nielsen total and ing effects (Nielsen and Andersen, 2000; Andersen and Koeppe, lated, the acyl chain composition, and the and the Andersen, 2000; Andersen and Koeppe, 2007; Lundbæk Our results show that increasing or number decreasing 2007; Lundbæk et al., 2010a): positions of phosphate groups on the inositol ring (Fig. et al., 2010a): mole percentage of long-chain PIP2 in the plasma mem1). brane For comparison, we also lipid examined bilayer-perresults in altered bilayerthe properties. Both I∆ II I → II ⎫ →G  turbing effects of other related short-chain n II n II =I →KII I → II = exp total  ∆⎧G  longand short-chain phosphoinositides, as well as nontotal − = =K = exp − ⎨ =⎬ phospholipids. phosphoinositide short-chain phospholipids, are bilay­ernI nI  ⎩kBT kBT  ⎭ (1) (1) Our results agents. show that increasing or decreasing the modifying II I → II ⎫ I∆ II I → II + I∆ →G →G  ∆⎪⎧G protein  ⎪ + ∆G bilayer protein bilayer mole percentage of long-chain PIP2 in the plasma memexpexp − ⎨− , ⎬, k T brane results in altered lipid bilayer properties. Both kBT B  ⎩⎪  ⎭⎪ M and A T Eshort-chain RIALS AN D M E T H O D S as well as nonlongphosphoinositides, where nI and nII denote surface densities of the where nI and nII denote the the surface densities of the pro-pro- phosphoinositide short-chain phospholipids, are biMaterials I → II I → II I → II I → II agents. ∆G bilayer in states II,protein denote ∆G protein teintein in states I andI and II, ∆G andand denote the the layer-modifying ∆G bilayer All phospholipids were purchased from Avanti Polar Lipids, Inc., exI → II protein bilayer contributionsto to ∆G total , and protein andand bilayer contributions andkBkTBTconcept for diC8PI (1,2-dioctanoyl-sn-glycero-3-phosphoinositol), which sists of Boltzmann’s constant constant and and the the temperature temperature in M Awas consists T Epurchased R I A L S from A N Echelon D M E Biosciences T H O D S (Fig. 1). The diC8 phospholipid stock solutions were 5 mM in water, except for the diC8PE Kelvin. Changes in properties, which in Kelvin. inlipid lipidbilayer bilayerphysical physical properties, stock solution, which was 5 mM in ethanol (Pharmco-AAPER). I → II I → II ∆G,bilayer are are expressed as changes in ∆G thus, will which expressed as changes inbilayer thusalter will the Materials The long-chain PIP2s were 1,2-dioleoyl-sn-glycero-3-phosphoconformational equilibrium and function. All phospholipids were purchased from and Avanti Lipids, alterprotein the protein conformational equilibrium and (1-myo-inositol-4,5-bisphosphate) thePolar porcine brainInc., extract for , diC 8PI (1,2-dioctanoyl-sn-glycero-3-phosphoinositol), Amphiphiles modulate lipid bilayer properties by ad- except function. of PIP 2 which is 80% C18/20:4PIP2. To make aqueous “solutions” purchased from Biosciences (Fig. 1). The diC8 of was diC18:1 sorbing to the bilayer–solution interface (Lundbæk et al., which and brain PIPEchelon Amphiphiles modulate lipid bilayer properties by ad2, the powder was suspended in H2O. The phospholipid stock solutions were 5 ultrasonic mM in water, except for (model the suspension was sonicated in an homogenizer 2010a; et al., 2011; Bruno et al., 2013), thereby sorbing toRusinova the bilayer–solution interface (Lundbæk et diC8300 PE stock solution, which was 5 mM in ethanol (PharmcoV/T; Biologics Inc.) for 30 min on ice at 90% power and 90% the bilayeretresponse to aBruno deformation al., altering 2010a; Rusinova al., 2011; et al., (changing 2013), AAPER). The using long-chain PIP2sThe were 1,2-dioleoyl-sn-glycero-3duty cycle 1-s pulses. resuspended long-chain phosGdefaltering and thusthe Gbilayer bilayertoproperties that may phospho-(1-myo-inositol-4,5-bisphosphate) and the inporcine thereby response a deformation bilayer). Key phoinositides were flash frozen and stored at 40°C a glass vial. PIP2PC , which is 80% C18/20:4PIP2. To make aquebe affected byand amphiphile adsorption includeproperthe intrin- brainToextract (changing Gdef thus Gbilayer ). Key bilayer make of DC (1,2-dioleoyl-sn-glycero-3-phosphocholine)/ 18:1 ous PIP “solutions” of diC18:1 and brain PIP2, the powder was suscurvature, as a resultbyofamphiphile the altered profile of intermoties sic that may be affected adsorption in2 lipid mixtures, the long-chain PIP2 was dissolved in chloropended in(Honeywell H2O. The Burdick suspension was sonicated in an ultrasonic form Jackson) and mixed with DC18:1PC lecular interactions along the normal (Seddon, homogenizer (model 300 V/T;and clude the intrinsic curvature, as abilayer result of the altered Biologics Inc.) for 30 Chloroform min on ice was in chloroform at the appropriate mole percentage. 1990; 1997; Andersen and Koeppe, 2007; Marsh, at 90% power and 90% duty cycle using 1-s pulses. The resusprofile ofCantor, intermolecular interactions along the bilayer evaporated under N2, and the mixture was suspended in n-decane 2007), bilayer elasticity (Evans1997; et al.,Andersen 1995), lipid disorder, pended long-chain phosphoinositides were flash frozen (ANTS) and normal (Seddon, 1990; Cantor, and Ko(Sigma-Aldrich). 8-Aminonaphthalene-1,3,6-trisulfonate stored 40°C in afrom glass Invitrogen. vial. To make DCthe (1,2-dioleoyl-sn18:1PC and possibly lipid demixing Heerklotz, eppe, 2007; Marsh, 2007), bilayer(Zhelev, elasticity1998; (Evans et al., wasatpurchased For single-channel experilipid mixtures, the long-chain 1 ments, we used a 15–amino 2acid analogue, [Ala 2002). amphiphile-mediated changes in bilayer glycero-3-phosphocholine)/PIP ]gA (AgA(15)), a 1995), lipidThese disorder, and possibly lipid demixing (ZhePIP was dissolved in 13–amino chloroformacid (Honeywell Burdick and Jack-helix gA analogue of opposite in turn may shiftThese the conformational equilib- son)2chain-shortened lev, properties 1998; Heerklotz, 2002). amphiphile-mediand mixed with DC  PC in  chloroform at the appropriate 18:1 sense, des-d-Val-Gly-gA (gA (13)), and a 15–amino acid analogue of membrane given that Gthe atedrium changes in bilayer proteins. properties(Itinisturn may shift def will mole percentage. Chloroform was evaporated under N2, and the in which the NH groups on the four Trp residues are methylated, change, but the changes may or may not be sufficient to [Val1,N-Me-Trp9,11,13,15]gA (N-Me-gA(15)); they were synthesized cause measurable changes in protein function.) We thereand purified as described previously (Greathouse et al., 1999). For 2 of 22 alter lipid bilayer properties the fluorescence assay, we used the natural mixture of gramicidins fore setPhosphoinositides out to determine whether PIP2 and its structural 674

Phosphoinositides alter lipid bilayer properties

Figure 1.  Structures of the phospholipids used in this study; brain PIP2 is predominantly 1-C18:0-2-C20:4-PI(4,5)P2, which is shown in

the figure.

from Bacillus brevis (Sigma-Aldrich), which is 80% [Val1]gA (AboRiziq et al., 2006); the mixture is denoted as gD (for R. Dubos, who discovered the gramicidins; Dubos, 1939). Electrophysiology Planar lipid bilayers were formed across a 1.5-mm–diameter aperture in a Teflon partition using DC18:1PC or DC18:1PC–long-chain PIP2 suspensions in n-decane. The total phospholipid concentration was the same (15 mg/ml) in the pure and mixed lipid suspensions. The aqueous solution was a 1 M NaCl and 10 mM HEPES, pH 7.0, solution; the temperature was 25 ± 1°C. Single-channel measurements were performed at 200 mV using a patch clamp amplifier (model 3900A; Dagan Corp.) and the bilayer punch method (Andersen, 1983). The current signal was filtered at 5 kHz, sampled at 20 kHz, and digitally filtered at 500 Hz. After the bilayer was formed, gA analogues of different lengths and helix senses were added to both sides of the bilayer, and the solutions were stirred for 5 min (in a few experiments, only N-Me-gA(15) was added). After 30 min, to allow for gA incorporation, control current traces were recorded. In the experiments in which the lipids were added through the aqueous phase, short- or long-chain phospholipids were added to both sides of the bilayer. The solutions were stirred for 1 min and equilibrated for 10 min before recording. Single-channel current transitions were identified using the algorithm described previously (Andersen, 1983), implemented in software written in Visual Basic (Microsoft Corp.). Single-channel lifetimes () were determined by fitting single exponential distributions to survivor histograms of the lifetime distributions using Origin 6.1 (OriginLab). At least 300 channels were collected in each experiment. Changes in appearance frequency ( f ), the number of channels (opening followed by closing events) per unit of time, were determined only in experiments in which the bilayer remained intact after adding the phospholipid. The results were reported as means ± SD; in case there were only two observations, the results were reported as means ± range. Relative changes in appearance frequency were determined by normalizing the appearance frequency in the first two recordings right after the 10-min incubation 

with the compound to the appearance frequency from a recording right before compound addition. Only recordings lasting ≥300 s were used to evaluate changes in relative channel appearance frequencies. Energetics of gA channel formation gA in lipid bilayers exists in equilibrium between nonconducting monomers and conducting dimeric channels: k1 ⎯⎯→ M + M ←⎯⎯ D,



k−1

where k1 and k1 are the association and dissociation rate constants, respectively. The dimer dissociation rate constant can be estimated as 1/, and changes in the association rate constant can be estimated from changes in the channel appearance frequency because k1 = f/[M]2. The channel’s hydrophobic length (l), 2.2 nm for a 15–amino acid analogue (Elliott et al., 1983; Huang, 1986), is less than the lipid bilayer’s unperturbed hydrophobic thickness (d0), 3–4 nm (Benz et al., 1975; Lewis and Engelman, 1983; Lundbæk et al., 2010b), and gA channel formation is associated with a local bilayer compression with an associated deformation energy, Gdef. Using the theory of elastic bilayer deformations (Huang, 1986; Helfrich and Jakobsson, 1990; Dan et al., 1994; Nielsen et al., 1998; Nielsen and Andersen, 2000; Partenskii and Jordan, 2002; Rusinova et al., 2011), Gdef can be expressed as

2 ∆G def = H B × (l − d 0 ) + H X × (l − d 0 ) × c 0 + H C × c 02 ,

(2)

where c0 is the intrinsic monolayer curvature and HB, HX, and HC are phenomenological elastic coefficients determined by the bilayer elastic moduli, the channel radius, and d0. The bilayer responds to the compression by imposing a disjoining force (Fdis) on the channel,

Fdis = −

∂∆G def = 2 × H B × (d 0 − l ) − H X × c 0 , ∂ (l − d 0 ) Rusinova et al.

(3)

675

Figure 2.  Effects of increasing the mole percentage

of long-chain PIP2 on gA channel activity in mixed DC18:1PC/long-chain PIP2 bilayers. Single-channel current traces recorded in DC18:1PC planar lipid bilayers containing 0 (control), 0.2, 0.6, and 2 mol % brain PIP2 or 2 mol % diC18:1PIP2. acting to dissociate the dimer. Changes in bilayer physical properties will alter the phenomenological elastic coefficients and thus Gdef, f, and  (Lundbæk et al., 2010b; Greisen et al., 2011), which make the gA channels suitable as probes of changes in bilayer properties. The energetics of the gA monomer↔dimer equilibrium is described by

(4) M→D M→D M→D ⎧ ⎫ G G ∆ + ∆ ⎧ ⎫ [D] G ∆ ⎪ ⎪ prot bil tot = K M → D = exp ⎨− ⎬ = exp ⎨− ⎬, kBT ⎭ kBT [M]2 ⎩ ⎩⎪ ⎭⎪

where [M] and [D] denote the surface concentrations of gA monoM→ D mers and dimers, ∆G bil is the change in Gdef upon dimer forM→ D mation, and ∆G tot is the free energy contribution from protein interactions. We are interested in the amphiphile-induced changes M→ D in bilayer properties, and the associated ∆∆G bil is determined by the changes in the time-averaged gA activity in the presence relative to that in the absence of amphiphiles. From Eq. 4: M→D ⎪⎧ K amph ⎪⎫ M→ D ∆∆G tot = −kBT ln ⎨ M → D ⎬ , ⎩⎪ K cntrl ⎭⎪



(5)

where the subscripts denote the presence or absence of amphiM → D can be estimated as phile and ∆∆G tot M→D

∆∆G tot

(6)

⎧[D]amph ⎫ ⎧ f amph × τ amph ⎫ M→D ≈ −k BT × ln ⎨ ⎬ = −kBT × ln ⎨ ⎬ ≈ ∆∆G bil . ⎩ [D]cntrl ⎭ ⎩ f cntrl × τ cntrl ⎭

The first approximation holds as long as [M] is in excess and does not vary after the amphiphiles have been added—as long as [M]amph  [M]cntrl/[M]cntrl ≈ 0. The second approximation holds when the subunit–subunit interactions do not vary—as long as M→D ∆∆G prot ≈ 0. 676

Phosphoinositides alter lipid bilayer properties

Additional information can be obtained through simultaneous measurements on two gA channels of different lengths (with different hydrophobic mismatch and helix sense, to prevent the formation of heterodimers), which allows for distinguishing between effects on bilayer curvature and elasticity. This is possible because, according to Eqs. 2 and 3, if the bilayer elasticity (expressed as changes in HB) is altered, the effects will be hydrophobic mismatch dependent, with greater changes in the shorter channel’s appearance frequency and lifetime (Lundbæk et al., 2005; Rusinova et al., 2011). If the relative changes in lifetime are similar for the longer and shorter channels, curvature effects dominate (Artigas et al., 2006; Ashrafuzzaman et al., 2006). Fluorescence-based gA assay To assay for the phosphoinositides’ bilayer-modifying effects, we used a gramicidin-based fluorescence quench assay (Ingólfsson and Andersen, 2010; Ingólfsson et al., 2010; Rusinova et al., 2011) in which fluorophore-loaded lipid vesicles were mixed with a gramicidin-permeable fluorescence quencher (Tl+) such that changes in gramicidin channel activity (the number of channels in the large unilamellar vesicle [LUV] membrane) were monitored as changes in the rate of fluorescence quenching. In brief, after solvent evaporation, DC22:1PC (1,2-dierucoyl-sn-glycero-3phosphocholine) was rehydrated in 100 mM NaNO3, 25 mM ANTS, and 10 mM HEPES, pH 7. After vesicle extrusion and desalting to remove excess ANTS, the ANTS-loaded LUVs were prepared for fluorescent experiments by diluting 1:20 in 140 mM NaNO3 and 10 mM HEPES, pH 7. ANTS-loaded LUVs were doped with gD; after incubation for 24 h at 12.5°C to allow for bilayer incorporation of gD, the LUVs were incubated at 25°C with or without (control) the test phospholipid for 10 min (unless otherwise noted). The LUVs and the Tl+-containing quench buffer were mixed in a stopped-flow spectrometer (SX.20; Applied Photophysics) with a dead time of 1 nm to reach the transition state for channel formation, whereas the two subunits need to move apart by only a small distance to reach the transition state for channel dissociation (Greisen et al., 2011). M→ D The changes in ∆G bilayer are modest (Fig. 7 C) but of sufficient magnitude to alter gA channel function. We M→ D also evaluated the changes in ∆G bilayer for N-Me-gA(15) M→ D channels, where ∆∆G bilayer was 2.3 kJ/mol at 50 µM diC8PI(4,5)P2 (1,2-dioctanoyl-sn-glycero-3-phospho-(1myo-inositol-4,5-bisphosphate; Fig. S1). As was observed previously with other amphiphiles (Lundbæk et al., 2010b), the changes in the gA–(13) single-channel lifetimes (expressed as ln{ τ13 / τ13,cntrl }) vary as a linear function of the changes in the AgA(15) single-channel lifetimes (expressed as ln{ τ15 / τ15,cntrl }; Fig. 7 D). The slope is 1.30 ± 0.08 (R = 0.996) and not significantly different from a slope of 1. Thus, though Rusinova et al.

679

Figure 7.  Effects of long- and short-chain PIP2s on

gA single-channel function. (A and B) Normalized changes in  (A) and f (B) of gA(13) (red bars) and AgA(15) (blue bars) channels produced by diC8PIP2, brain PIP2, and diC18:1PIP2. (A) †, in the presence of 30 µM diC8 or 50 µM brain PIP2, /cntrl is significantly >50 µM diC18:1PIP2 (P ≤ 0.05) for gA(13) as well as gA(15); ‡, gA(13) /cntrl in the presence of 10 µM diC8PIP2 is significantly >10 µM brain PIP2. (B) †, gA(13) f/fcntrl in the presence of 10 µM diC8 is signifiM→ D cantly >10 µM brain PIP2. (C) The resulting ∆∆G bil values are calculated from the changes in  and f as described in Materials and methods (Eq. 6). We could not determine the changes in f at 50 µM brain PIP2 because of bilayer instability. (A–C) All values are plotted as means ± SD or range; n = 2–6. *, means are significantly different from 1 (P ≤ 0.05). Dotted lines indicate the level where there is no change relative to control (y = 1). (D) Plot of the natural logarithm of /cntrl of gA(13) as a function of /cntrl of gA(15) channels in the presence of different concentrations of diC8 and brain PIP2 (square). The 95% confidence intervals (green lines) indicate that the slope is not significantly different from the slope of 1 (dashed line), suggesting that both short- and long-chain PIP2s produce larger increases in the lifetimes of gA(13) channels than of gA(15) channels.

the relative changes in single-channel appearance frequencies, f, and lifetimes, , elicited by either PIP2 species were not significantly different for the longer AgA(15) and the shorter gA(13) channels, which would suggest that they primarily alter intrinsic curvature, the aggregate results suggest that they also increase bilayer elasticity (Lundbæk et al., 2010b; Rusinova et al., 2011). Fluorescence quench studies. We also explored the longchain PIP2s’ and diC8PIP2’s bilayer-modifying effects using a gramicidin-based fluorescence assay that takes advantage of the gramicidin channels’ permeability to Tl+, a heavy-ion quencher of the water-soluble ANTS fluorophore. In this assay, changes in the fluorescence 680

Phosphoinositides alter lipid bilayer properties

quench rate reflect changes in the Tl+ influx and channel activity (Ingólfsson and Andersen, 2010; Ingólfsson et al., 2010). The gA dimerization constant varies with changes in lipid bilayer properties (Andersen and Koeppe, 2007; Lundbæk et al., 2010a), meaning that one can attribute the changes in quench rate after amphiphile addition to modulation of bilayer properties, which in turn govern the gramicidin monomer↔dimer equilibrium and thus the number of channels in the LUV membrane (Ingólfsson and Andersen, 2010; Rusinova et al., 2011). Brain PIP2 and diC8PIP2 increased the fluorescence quench rates, whereas diC18:1PIP2 did not alter the quench rates at the tested concentrations (Fig. 8). The results in Fig. 8 are consistent with the electrophysiological

ROUGH GALLEY PROOF ROUGH GALLEY PROOF Figure 8. Testing for bilayer-modifying effects long-chain PIP2s and diC8PIP2 Figure 8. Testing for of bilayer-modifying usingfor a bilayer-modifying gramicidin-based fluorescence effects PIP Figure of 8. long-chain Testing 2s and diC8PIP2

assay. (A) of2 Tl+ quenching using effectsaofgramicidin-based long-chain PIPTime andcourse diC8PIP 2s fluorescence + the ANTS assay. Timeof course of Tlfluorescence quenching in the absence using(A) a gramicidin-based + (top trace) and presence of the ANTS fluorescence the absence (bottom three assay. (A) Time course of in Tl quenching traces) of gD inthree the absence or pres(top trace) presence (bottom of the ANTSand fluorescence inand the absence ence of absence different traces) of gDand and in the orPIP pres(top trace) presence (bottom three 2s. The trace for each condition shows data points from ence ofofdifferent PIP The trace for traces) gD and in the or pres2s.absence all mixing (gray points) and the each shows datatrials points ence condition of different PIP trace forfrom each 2s. The corresponding (black lines, conall mixing trials (gray and the condition shows data points) points mean from all trols; cyan line, 50 diC18:1PIP2; navy corresponding mean (black lines, conmixing trials (gray points) and theµM corblue line,diC 50 µM trols; cyan line, 50 (black µM PIP ; navyPI(4,5)P2). (B) responding mean lines, controls; 18:1 2brain Fluorescence traces from a sinblue brain (B) cyan line, line, 50 50 µM µM diC18:1PI(4,5)P PIP2quench ; navy 2). blue glePI(4,5)P mixing for each Fluorescence quench traces from a sin-condition in A. line, 50 µM brain ). (B) Fluores2trial The points colored gle mixing trialtraces for each in A. like the correcence quench fromcondition a are single mixing sponding mean in A; the solid The colored in like thetraces corretrial points for eachare condition A. The points denote fitsthe of amean stretched exponensponding traces in A; solid are coloredmean likelines the corresponding tial to denote the data, lines denote ofsolid a stretched exponentraces in A; fits the lines fits β tial to exponential the data, of a stretched to the data, F (t ) = F (∞) + ( F (0) − F (∞)) × exp {−(t / τ ) } ,

{ is

} fluorescence

F (t ) = F (∞) + ( F (0) − F (∞)) × exp −(t / τ 0 ) , β

0

where F(t) intensity at time t,  is a intensity measure where at where F(t) F(t) isis fluorescence fluorescence intensity of at sample dispersity (0