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Electrostatic Sequestration of PIP₂ on Phospholipid Membranes by Basic/Aromatic Regions of Proteins

Alok Gambhir ^* , Gyöngyi Hangyás-Mihályné , Irina Zaitseva , David S. Cafiso , Jiyao Wang , Diana Murray , Srinivas N. Pentyala ^¶, Steven O. Smith ^|| and Stuart McLaughlin 

^* Department of Physics and Astronomy, Department of Physiology and Biophysics, ^¶ Department of Anesthesiology, and ^|| Department of Biochemistry and Cell Biology, SUNY Stony Brook, Stony Brook, New York 11794; Department of Chemistry and Biophysics Program, University of Virginia, Charlottesville, Virginia 22904; and Department of Microbiology and Immunology, Weill Medical College of Cornell University, New York, New York 10021

Correspondence: Address reprint requests to Stuart McLaughlin, Dept. of Physiology and Biophysics, HSC BST, SUNY Stony Brook, Stony Brook, NY 11794-8661. Tel.: 631-444-3615; Fax: 631-444-3432; E-mail: SMCL{at}epo.som.sunysb.edu.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS SUMMARY DISCUSSION APPENDIX 1: PHOSPHOLIPID... APPENDIX 2: Effects of... ACKNOWLEDGEMENTS REFERENCES

 
The basic effector domain of myristoylated alanine-rich C kinase substrate (MARCKS), a major protein kinase C substrate, binds electrostatically to acidic lipids on the inner leaflet of the plasma membrane; interaction with Ca²⁺/calmodulin or protein kinase C phosphorylation reverses this binding. Our working hypothesis is that the effector domain of MARCKS reversibly sequesters a significant fraction of the L--phosphatidyl-D-myo-inositol 4,5-bisphosphate (PIP₂) on the plasma membrane. To test this, we utilize three techniques that measure the ability of a peptide corresponding to its effector domain, MARCKS(151–175), to sequester PIP₂ in model membranes containing physiologically relevant fractions (15–30%) of the monovalent acidic lipid phosphatidylserine. First, we measure fluorescence resonance energy transfer from Bodipy-TMR-PIP₂ to Texas Red MARCKS(151–175) adsorbed to large unilamellar vesicles. Second, we detect quenching of Bodipy-TMR-PIP₂ in large unilamellar vesicles when unlabeled MARCKS(151–175) binds to vesicles. Third, we identify line broadening in the electron paramagnetic resonance spectra of spin-labeled PIP₂ as unlabeled MARCKS(151–175) adsorbs to vesicles. Theoretical calculations (applying the Poisson-Boltzmann relation to atomic models of the peptide and bilayer) and experimental results (fluorescence resonance energy transfer and quenching at different salt concentrations) suggest that nonspecific electrostatic interactions produce this sequestration. Finally, we show that the PLC-₁-catalyzed hydrolysis of PIP₂, but not binding of its PH domain to PIP₂, decreases markedly as MARCKS(151–175) sequesters most of the PIP₂.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS SUMMARY DISCUSSION APPENDIX 1: PHOSPHOLIPID... APPENDIX 2: Effects of... ACKNOWLEDGEMENTS REFERENCES

 
Although PIP₂ comprises only 1% of the phospholipids in a plasma membrane from a typical animal cell, it plays a key role in many cell-signaling pathways. Specifically, it is the source of three important second messengers (Berridge and Irvine, 1984; Payrastre et al., 2001; Cantley 2002; Vanhaesebroeck et al., 2001) and is involved in membrane trafficking (Czech, 2003; Cremona and De Camilli, 2001; Martin, 2001; Osborne et al., 2001; Simonsen et al., 2001); it also modulates several ion channels (Hilgemann et al., 2001; Runnels et al., 2002; Prescott and Julius, 2003), mediates cytoskeleton-membrane interaction (Yin and Janmey, 2003; Raucher et al., 2000), and regulates the activity of a variety of molecules (e.g., PLD (Sciorra et al., 2000); N-WASP (Lim, 2002). How does one lipid play so many different roles? We and others have hypothesized that proteins bind a significant fraction of the PIP₂ in the plasma membrane, then release it locally in response to specific signals (Caroni, 2001; McLaughlin et al., 2002). Putative PIP₂-sequestering proteins should satisfy three criteria: they must be present at concentrations comparable to PIP₂, bind PIP₂ with high affinity, and release it locally in response to specific stimuli such as an increase in the local calcium ion concentration.

One such candidate is the myristoylated alanine-rich C kinase substrate (MARCKS) protein (Aderem, 1992; Blackshear, 1993; McLaughlin and Aderem, 1995; Arbuzova et al., 1998, 2002). Other viable candidates include GAP43 and CAP23 in neuronal tissue (Laux et al., 2000). As illustrated in Fig. 1 A, MARCKS is a natively unfolded protein that binds to the plasma membrane through hydrophobic insertion of its N-terminal myristate (shown in yellow) and electrostatic interaction of its basic effector domain with acidic lipids (shown in the box). The available evidence suggests that MARCKS may satisfy the three criteria listed above. First, MARCKS is present at high concentrations (10 µM) in neuronal and other tissues (Blackshear, 1993; Albert et al., 1987); all measurements to date indicate this is in the same range as the PIP₂ concentration. PIP₂ comprises 0.3–1.5% of the phospholipid in the plasma membrane of mammalian erythrocytes (Ferrell and Huestis, 1984; Hagelberg and Allan, 1990), lymphocytes (Mitchell et al., 1986), and hepatocytes (Tran et al., 1993); for a 10-µm cell, this corresponds to an effective concentration of PIP₂ of 5–30 µM. Bunce et al. (1993) determined that the total concentration of PIP₂ in human myeloid cells is 30 µM. Studies using GFP-PH domain constructs also suggest the effective concentration of PIP₂ in cells is 2 < [PIP₂] < 30 µM, as discussed elsewhere (McLaughlin et al., 2002). Second, a peptide corresponding to the effector domain, MARCKS(151–175), binds with high affinity to phospholipid vesicles containing a mixture of the zwitterionic lipid phosphatidylcholine (PC) and PIP₂ (Arbuzova et al., 2000b; Wang et al., 2001, 2002; Rauch et al., 2002). As illustrated in Fig. 1 A, the effector domain interacts with approximately three PIP₂ to form an electroneutral complex. This report provides evidence that MARCKS(151–175) can laterally sequester PIP₂ in the presence of a significant excess of monovalent acidic lipids. Third, the sequestration of PIP₂ can be reversed by protein kinase C (PKC) phosphorylation or by interaction with calcium/calmodulin (Ca²⁺/CaM). Introduction of three negatively charged phosphates by PKC or binding of Ca²⁺/CaM causes translocation of both the MARCKS protein (Kim et al., 1994) and MARCKS(151–175) (Arbuzova et al., 1998) from the membrane to solution by diminishing its electrostatic binding to acidic phospholipids. The translocation of the native MARCKS protein from the plasma membrane due to PKC phosphorylation or Ca²⁺/CaM binding has been observed in vivo (Swierczynski and Blackshear, 1995; Ohmori et al., 2000; J. Sable and M. P. Sheetz, Columbia University, personal communications).

View larger version (79K): [in this window] [in a new window]   FIGURE 1  (A) Cartoon of the "natively unfolded" MARCKS protein, shown as a black line, interacting with the inner leaflet of the plasma membrane. The N-terminal myristate, shown in yellow, inserts hydrophobically into the bilayer. The MARCKS effector domain (residues 151–175 of bovine MARCKS, shown in the box) interacts electrostatically with acidic lipids (3 PIP2 shown in red) through its 13 basic residues (in blue with + signs) and hydrophobically through its five aromatic residues (in green). (B) Molecular model of a peptide corresponding to the MARCKS effector domain overlaid on a molecular model of a bilayer membrane. Experimental evidence (see text) shows that the peptide is located at the polar headgroup region in an extended conformation; the five aromatic phenylalanine residues (colored green) penetrate to the level of the acyl chains, and the highly charged N-terminal region (basic residues colored blue) is in the aqueous phase. The lipids are shown in white with the carbonyl oxygen atoms colored red to illustrate the interface between the headgroup region and the hydrocarbon interior of the membrane. The extended peptide is 9 nm in length.

Arbuzova et al. (1998) discuss the evidence that the MARCKS(151–175) peptide is a good model for studying the interaction of MARCKS with membranes. For example, the peptide binds Ca²⁺/CaM with the same high (4 nM) affinity as the intact protein and contains the three serine residues phosphorylated by PKC; either binding of Ca²⁺/CaM or phosphorylation by PKC produce translocation of both protein and peptide from membrane to solution. There is strong evidence that the MARCKS(151–175) peptide binds with high affinity to PIP₂/PC vesicles by electrostatically sequestering approximately three PIP₂ (Wang et al., 2002; Rauch et al., 2002). There is only indirect evidence, however, that the effector domain can sequester PIP₂ when the membrane contains a physiological mol fraction of monovalent acidic lipids (i.e., 15–30% phosphatidylserine (PS) (White, 1973; Yorek, 1993)); specifically, both MARCKS and MARCKS(151–175) inhibit the PLC-catalyzed hydrolysis of PIP₂ in vesicles or monolayers containing both PS and PIP₂ (Murray et al., 2002; Wang et al., 2002). We have used three different techniques, fluorescence resonance energy transfer (FRET), quenching, and EPR, to obtain more direct evidence that the effector domain of MARCKS can sequester PIP₂ when PS is present.

We first measured FRET between labeled MARCKS(151–175) and PIP₂ using PC/PS/PIP₂ vesicles containing 0.1% PIP₂ and up to 30% PS. We eliminated the possibility that sequestration of PIP₂ is due to probe-probe interactions by measuring the effect of unlabeled MARCKS(151–175) on the quenching of fluorescent PIP₂ and on the EPR spectra of spin-labeled PIP₂ in these PC/PS/PIP₂ vesicles.

Calculations using atomic-level models of bilayers and adsorbed basic peptides show that the peptides produce a local positive electrostatic potential, even when the membrane contains 30% monovalent acidic lipid; this positive potential should electrostatically sequester the multivalent acidic lipid, PIP₂. To test that the sequestration is due to electrostatics, we measured the effect of increasing the salt concentration on lateral sequestration.

Aromatic Phe residues embedded within a basic peptide drag the adjacent basic residues into the polar headgroup region (Fig. 1 B). Our electrostatic model predicts that this should increase the positive potential produced by the peptide when it adsorbs to the bilayer. Hence, aromatic residues should enhance PIP₂ sequestration. To test this prediction, we compare the PIP₂ sequestration produced by MARCKS(151–175) versus FA-MARCKS(151–175), a peptide synthesized with alanine residues substituted for phenylalanine (Table 1).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS SUMMARY DISCUSSION APPENDIX 1: PHOSPHOLIPID... APPENDIX 2: Effects of... ACKNOWLEDGEMENTS REFERENCES

 
Materials
Fig. 2 shows the structures of the fluorescent and spin-labeled PIP₂ lipids that we used in these studies. Bodipy-TMR-PIP₂ (Fig. 2 A) was purchased from Echelon (Salt Lake City, Utah) as a triethylammonium salt. 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine (PC), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylserine, and the ammonium salt of PIP₂ were purchased from Avanti Polar Lipids (Alabaster, AL). Radioactively labeled [dioleoyl-1-¹⁴C]-L--dioleoyl-phosphatidylcholine (¹⁴C-DOPC) and [inositol-2-³H(N)]-L--phosphatidyl-D-myo-inositol 4,5-bisphosphate (³H-PIP₂) were from Perkin Elmer Life Sciences (Boston, MA). Texas Red C₅ bromoacetamide, Bodipy-507 iodoacetamide, Oregon Green 488 maleimide, fluorescein-5-iso-thiocyanate (FITC), and N-(6- tetramethylrhodaminethiocarbamoyl)-1,2-dihexadecanoyl-sn-glycero-3- phosphoethanolamine (TRITC-PE) were from Molecular Probes (Eugene, OR). The Molecular Probes catalog illustrates the structures of these fluorescent probes. FITC-labeled neomycin (Arbuzova et al., 2000a) was obtained from Glenn Prestwich (Echelon). Recombinant human PLC-₁ was purified from Escherichia coli as described elsewhere (Garcia et al., 1995). The EGFP-PLC-₁ PH domain construct (EGFP-PH domain) was prepared in E. coli as described elsewhere (Pentyala et al., 2003; Tall et al., 2000).

View larger version (14K): [in this window] [in a new window]   FIGURE 2  Molecular structures of (A) Bodipy-TMR-PIP2 and (B) proxyl-PIP2 .

Peptide labeling
We used a protocol modified from "Conjugation with Thiol-Reactive Probes" (Molecular Probes) to label peptides with the thiol-reactive fluorescent probes. In brief, we mixed 1 ml of 1 mM peptide in 10 mM K₂HPO₄/KH₂PO₄, pH 7.0 with the probe dissolved in N,N'-dimethylformamide (probe-to-peptide molar ratio of 1:1) for 1 h. We purified the labeled peptide using high-performance liquid chromatography and checked that it has the correct molecular weight using MALDI-TOF mass spectrometer (Proteomics Center, SUNY at Stony Brook, Stony Brook, NY).

Vesicle preparations
We used 100-nm diameter large unilamellar vesicles (LUVs) for the FRET, self-quenching, PLC-₁ hydrolysis, and centrifugation binding experiments, as described in detail elsewhere (Wang et al., 2002). Briefly, we added solutions of PIP₂ (or Bodipy-TMR-PIP₂), PS, and PC in chloroform to a 50-ml round-bottom flask, which was then well immersed in a 30–35°C water bath and attached to a rotary evaporator. The flask was rotated without vacuum for 5 min to warm the flask and solution. We then rapidly evaporated most of the solvent by applying the maximum vacuum that does not boil the chloroform. (Rapid evaporation is required to produce a uniform mixture of PC, PS, and PIP₂ because PIP₂ is less soluble in chloroform than either PC or PS.) The flask was kept under full vacuum for 30 min to remove all traces of chloroform. A solution containing 100 mM KCl, 1 mM MOPS, pH 7.0 was added for most of our experiments. Subsequently, we rapidly froze and thawed the multilamellar vesicles five times. Finally, we formed LUVs by extrusion of the multilamellar vesicles through 100-nm diameter polycarbonate filters. (A solution containing 176 mM sucrose, 1 mM MOPS, pH 7.0 was added to make sucrose-loaded LUVs for centrifugation binding measurements. The solution bathing the LUVs was then exchanged for 100 mM KCl, 1 mM MOPS, pH 7.)

Preparation of lipid vesicles for EPR spectroscopy
LUVs (100 nm) were prepared as described previously by mixing appropriate volumes of stock solutions of POPC and POPS in chloroform, drying the lipid mixtures under vacuum, hydration of the lipid in 100 mM KCl, 10 mM MOPS, pH 7.0 followed by extrusion through 100-nm pore size polycarbonate filters (Rauch et al., 2002). The spin-labeled proxyl-PIP₂ (shown in Fig. 2 B; provided by G. Prestwich and C. Ferguson, Echelon, Salt Lake City, UT) was incorporated into the outer leaflet of the vesicles by adding the lipid vesicle suspension to a dried film of labeled lipid. As discussed previously, this procedure resulted in incorporation of the proxyl-PIP₂ into the outer leaflet of the vesicle bilayer (Rauch et al., 2002).

FRET experiments
FRET and other fluorescence experiments were performed on an SML-AMINCO spectrofluorometer. FRET experiments used LUVs prepared with 0.1% Bodipy-TMR-PIP₂ (shown in Fig. 2 A) incorporating varying mol fractions of POPS and POPC. We excited the donor fluorophore and collected the complete emission spectra. When Bodipy-TMR-PIP₂ was the donor and Texas Red MARCKS(151–175) the acceptor, we excited at 547 nm and collected emission spectra from 560 to 660 nm. Direct binding measurements using the centrifugation technique (not shown) allowed us to choose a sufficiently high lipid concentration so that essentially all the peptide we added was bound to the LUVs; we monitored FRET as peptide was added to the solution containing the LUVs. We deconvoluted the data by performing a four-parameter two-peak Lorentzian fit keeping the peak wavelengths fixed at 571 nm for the Bodipy-TMR-PIP₂ and at 607 nm for Texas Red MARCKS(151–175). After checking the quality of fit, we used the peak amplitudes and full width at half-maximum amplitude for each peak to calculate the fluorescence intensity. The energy transfer efficiency, % transfer, was analyzed by calculating the quenching of the fluorescence intensity of the donor,

(1)

where I_da is the donor fluorescence intensity determined at the given excitation wavelength, , in the presence of the acceptor, and I_d is the corresponding intensity in the absence of the acceptor. The % transfer was also determined by calculating the fluorescence intensity of the acceptor,

(2)

where _ad is the absorbance of the acceptor when donor is present; _da is the absorbance of the donor when acceptor is present; ₁ is the peak absorbance wavelength of the donor; I_ad is the intensity determined at the acceptor wavelength, ₂, in the presence of the donor; and I_a is the corresponding intensity in the absence of the donor (Berney and Danuser, 2003; Lakowicz 1999; Van Der Meer et al., 1994).

We performed two control measurements to estimate effects due to probe-probe interactions and other artifacts. First, we replaced fluorescent PIP₂ with TRITC-PE, a zwitterionic lipid that should not be sequestered by fluorescent MARCKS(151–175) and should not exhibit FRET. TRITC is a rhodamine analog that has excitation and emission spectra comparable to the Bodipy-TMR. Second, we added 1–5 nM of PLC-₁ to hydrolyze the Bodipy-TMR-PIP₂ on the same vesicles used to perform FRET experiments. The amount of FRET was remeasured on these hydrolyzed vesicles.

We performed FRET experiments with vesicles containing 0.1% or 1% fluorescent PIP₂. We note, however, that random interactions produced significant energy transfer when Texas-Red-labeled peptide bound to vesicles containing 1% fluorescent PE (an electrostatically neutral lipid that should not bind to the basic peptides) instead of fluorescent PIP₂. This is expected because, for a vesicle containing 1% fluorescent lipid, the average distance between a fluorophore on the lipid and a fluorophore on the peptide is 50 Å, which is approximately the R_o value for the fluorophores we used (Fluorescence Resonance Energy Transfer in Molecular Probes Handbook). For a vesicle containing 0.1% fluorescent lipid, the average distance between the fluorophores on the peptide and lipid is 160 Å; thus, FRET measurements of lateral interaction are more informative at the lower PIP₂ concentration. Moreover, measurements using 0.1% PIP₂ are a more stringent test of our hypothesis that much of the PIP₂ is sequestered under physiological conditions.

Quenching experiments
Bodipy-TMR-PIP₂ was excited at 547 nm and emission spectra were recorded from 560 to 660 nm in the presence of different concentrations of unlabeled peptide. We calculated the % quenching as described above using Eq. 1 (Berney and Danuser, 2003; Lakowicz 1999; Van Der Meer et al., 1994). The vesicle concentration was sufficiently high that essentially all the peptide added to the solution bound to the membrane. We excluded quenching artifacts by performing control experiments on vesicles in which the negatively charged Bodipy-TMR-PIP₂ was replaced with zwitterionic or neutral fluorescent lipids (TRITC-PE or Bodipy-TMR-DAG), as discussed in the FRET section above.

The average distance between PIP₂ molecules is 100 Å and 300 Å in vesicles containing 1% and 0.1% PIP₂, respectively; the R_o value of the Bodipy-TMR probe is 50 Å, so self-quenching should be negligible in the absence of peptide. Indeed, for 1% Bodipy-TMR-PIP₂, fluorescence increases linearly with the % labeled PIP₂ in the vesicles. We observed qualitatively similar results in experiments performed with vesicles containing either 1% or 0.1% Bodipy-TMR-PIP₂, but present data only for the vesicles with 1% Bodipy-TMR-PIP₂, where the effect was more pronounced.

EPR spectroscopy
EPR spectra were recorded at X-band from 5 µL of sample using a Varian E-line Century series spectrometer fitted with a MITEQ microwave amplifier (Hauppauge, NY) and a two-loop one-gap resonator (Medical Advances, Milwaukee, WI). Spectra were recorded using a modulation of 1 Gauss peak-to-peak and microwave power of 2 mW.

EPR spectra with titration of MARCKS(151–175) were performed by adding concentrated solutions of the peptide to 100 µL of a 7–20 mM lipid vesicle suspension with 0.25–0.85 mol% proxyl-PIP₂ incorporated into the outer leaflet. Spectra were recorded by withdrawing a small sample of the vesicle suspension into the loop-gap resonator as described previously (Rauch et al., 2002). The binding of MARCKS(151–175) to proxyl-PIP₂ was analyzed by recording the amplitude of the first derivative peak-to-peak EPR spectrum.

Measurement of PIP₂ Hydrolysis
We previously reported that MARCKS and a peptide corresponding to its effector domain inhibit PLC-catalyzed PIP₂ hydrolysis (Wang et al., 2001, 2002; Murray et al., 2002). In those experiments, however, the vesicles or monolayers contained 1% PIP₂ and a high concentration of protein or peptide was needed to inhibit the hydrolysis. The experiments reported here used vesicles containing only 0.1% PIP₂, allowing us to use lower concentrations of peptide and minimize effects other than PIP₂ sequestration (for example, effects due to the insertion of Phe residues into the acyl chain region of the bilayer or to the charges on the peptide decreasing the net negative charge of the PC/PS/PIP₂ bilayer).

We added PLC-₁ to vesicles containing ³H-PIP₂ and terminated hydrolysis at different times by adding 375 µl ice-cold 10% trichloroacetic acid and 50 µl 10% Triton X-100 to 75 µl of the reaction mixture, then incubating the samples on ice until a white precipitate formed. The samples were then centrifuged at 14,000 x g for 5 min and the supernatant was mixed with 1 ml of chloroform/methanol (2:1 volume ratio); subsequently, the upper phase containing the ³H-IP₃ products was transferred to a scintillation vial for counting.

Data were analyzed by first plotting the % PIP₂ hydrolyzed versus time (Fig. 13 A). We obtained the rate of hydrolysis or PLC-₁ activity by calculating the initial (first 2–3 min) slopes from the time curves of individual experiments. Because PLC-₁ activity varies from one day to the next, we normalized the activity to controls where no peptide was present; the data in Fig. 13 B are averages of normalized activity from individual experiments.

View larger version (23K): [in this window] [in a new window]   FIGURE 13  Inhibition of PLC-1-catalyzed hydrolysis of PIP2 by peptides that sequester PIP2. (A) The % accessible PIP2 hydrolyzed versus time after addition of PLC-1. These data are for zero peptide (•) and 0.5 µM MARCKS(151–175) (). A control with no PLC shows no hydrolysis (), as expected. The results illustrate the average of six separate experiments. (B) Bar graph of relative PLC-1 activity, calculated from the initial slopes of hydrolysis versus time curves similar to those shown in A. These bars represent an average of six experiments (±SD) for each peptide. LUVs were composed of PC/PS/3H-PIP2 (83:17:0.15); 0.2 mM total lipid concentration. Solution contains 100 mM KCl, 25 mM HEPES, 100 µM EGTA, 102 µM CaCl2 (5 µM free Ca2+), 2 mM DTT, 10 nM PLC-1, pH 7.0.

Electrostatic calculations
Electrostatic potentials and free energies are obtained from a modified version of the DelPhi program (Gallagher and Sharp, 1998) that solves the nonlinear Poisson Boltzmann (PB) equation for protein/membrane systems (Ben-Tal et al., 1996). DelPhi produces finite difference solutions to the PB equation (the FDPB method) for a system where the solvent is described in terms of a bulk dielectric constant and concentrations of mobile ions, whereas solutes (here, basic peptides, the PLC-₁ PH domain, PIP₂, and phospholipid membranes) are described in terms of the coordinates of the individual atoms as well as their atomic radii and partial charges (Brooks et al., 1983; Peitzsch et al., 1995).

The application to the PIP₂/peptide/membrane systems considered here is described in more detail in our companion computational paper (Wang et al., 2003). PIP₂ is assumed to have a valence of -4 and the basic peptides are assumed to be in their minimum free-energy orientations as determined for the interaction with PC/PS bilayers in 0.1 M KCl in the absence of PIP₂; i.e., it is assumed that the interaction with PIP₂ does not affect the orientation of the basic peptides at the membrane surface. The structure of the PH domain from PLC-₁ (Ferguson et al., 1995) was used in the calculation of the electrostatic potentials illustrated in Fig. 15. The electrostatic potentials depicted in Figs. 12 and 15 were calculated by solving the nonlinear PB equation (Gallagher and Sharp, 1998) and visualized in GRASP (Nicholls et al., 1991).

View larger version (68K): [in this window] [in a new window]   FIGURE 15  MARCKS(151–175) laterally sequesters the PIP2-bound PH domain of PLC-1 as well as PIP2. (A) See text for description of cartoon. Equilibrium 1: membrane-bound MARCKS(151–175) (blue ovals represent the 13 basic residues; green ovals the five phenylalanine residues) laterally sequesters PIP2 (red). Equilibrium 2: The PLC-1 PH domain (light green) binds to PIP2 (red) with high specificity; PIP2 forms multiple hydrogen bonds with positively charged residues (blue + signs) in the binding pocket. Equilibrium 3: We propose that the PIP2-bound PH domain, which contains a patch of acidic residues (red - signs) on its surface, is (like PIP2) sequestered electrostatically by the membrane-bound MARCKS(151–175). Panel B shows the potential produced by PIP2 (yellow) on a PC membrane. Panel C shows the potential produced by PIP2-bound PH domain (green) on a PC membrane as viewed from the side. Panel D illustrates the view from the top of the membrane: -25 mV and +25 mV potential profiles are shown in red and blue; salt concentration = 100 mM.

View larger version (52K): [in this window] [in a new window]   FIGURE 12  Electrostatic potentials produced by basic peptides FA-MARCKS(151–175) or Lys-13. (A) FA-MARCKS(151–175) (colored green) binds to a 5:1 PC/PS membrane; a single PIP2 (colored yellow) is visible far from the peptide in the upper right-hand side of the bilayer. The blue and red meshes show +25 and -25 mV equipotential profiles. (B) FA-MARCKS(151–175) binds to a 5:1 PC/PS membrane and sequesters a PIP2. (C) FA-MARCKS(151–175) in 100 mM KCl solution. The blue line shows a two-dimensional representation of the +25 mV equipotential profile. In panels D, E, and F the peptide is adsorbed to a 5:1 PC/PS bilayer and viewed from above. For clarity, we do not show the membrane and -25 mV potential profile. (D) FA-MARCKS(151–175) binds to a 5:1 PC/PS membrane; (E) membrane-bound FA-MARCKS(151–175) sequesters a PIP2; (F) membrane-bound Lys-13.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS SUMMARY DISCUSSION APPENDIX 1: PHOSPHOLIPID... APPENDIX 2: Effects of... ACKNOWLEDGEMENTS REFERENCES

 
FRET shows membrane-bound MARCKS(151–175) sequesters PIP₂, even when the vesicles contain a 300-fold excess of PS. Fig. 3 A shows energy transfer from Bodipy-TMR-PIP₂ to membrane-bound Texas Red MARCKS(151–175) as the peptide concentration increases from 0 to 200 nM. Fig. 3, B and C, show the deconvoluted, emission spectra of the two fluorophores. As the peptide concentration increases, the donor (Bodipy-TMR-PIP₂) fluorescence decreases (Fig. 3 B) and the acceptor fluorescence increases (Fig. 3 C). Fig. 4 shows the % energy transfer calculated using both the quenched Bodipy-TMR fluorescence (•) and the energy transferred Texas Red fluorescence () data. The two calculations agree, as they should. These experiments illustrate that the Bodipy-TMR-PIP₂ transfers energy effectively to membrane-bound Texas Red MARCKS(151–175) on LUVs containing 30% monovalent acidic lipids, suggesting that the basic peptide sequesters the multivalent acidic lipid PIP₂ under physiological conditions (i.e., 300-fold excess PS).

View larger version (24K): [in this window] [in a new window]   FIGURE 3  FRET from Bodipy-TMR-PIP2 to membrane-bound Texas Red MARCKS(151–175). The LUVs are formed from PC/PS/Bodipy-TMR-PIP2 (70:30:0.1). The total lipid concentration is 0.75 mM; approximately all the peptide is bound to the LUVs. Bodipy-TMR-PIP2 is excited at 547 nm. (A) Representative corrected emission spectra for the peptide concentrations indicated. Spectra are deconvoluted to two peaks: (B) the quenching of Bodipy-TMR-PIP2, centered at 571 nm, and (C) the energy transfer to Texas Red MARCKS(151–175), centered at 607 nm. The solutions in these experiments, and in those shown in the following figures (except Figs. 9 and 13), contain 100 mM KCl, 1 mM MOPS, pH 7.0.

View larger version (17K): [in this window] [in a new window]   FIGURE 4  The % energy transfer versus concentration of Texas Red MARCKS(151–175). We calculate the % energy transfer from the quenching data (•) in Fig. 3 B and similar experiments, or from the energy transferred data () in Fig. 3 C and similar experiments. All subsequent energy transfer results are calculated using the quenching spectra. Each point shown in the graph is an average of 7 separate experiments (±SD).

View larger version (19K): [in this window] [in a new window]   FIGURE 9  Increasing the salt concentration decreases both energy transfer and quenching. (A) Energy transfer between Bodipy-TMR-PIP2 and Texas Red MARCKS(151–175) (•) or Texas Red FA-MARCKS(151–175) () in solutions containing 100 mM, 200 mM, or 300 mM KCl. Lipid composition of LUVs is PC/PS/Bodipy-TMR-PIP2 (70:30:0.1). (B) Quenching of Bodipy-TMR-PIP2 due to MARCKS(151–175) (•), FA-MARCKS(151–175) (), or Lys-13 () in solutions containing 100 mM, 200 mM, or 300 mM KCl. Lipid composition of LUVs is PC/PS/Bodipy-TMR-PIP2 (69:30:1).

View larger version (14K): [in this window] [in a new window]   FIGURE 5  The % energy transfer for PC/PS/Bodipy-TMR-PIP2 (70:30:0.1) LUVs plotted against the molar ratio of Texas Red MARCKS(151–175) to Bodipy-TMR-PIP2. The total lipid concentrations are 0.75 mM (), 0.5 mM (), and 0.1 mM (•).

We also studied FRET to Bodipy-TMR-PIP₂ using peptides labeled with donor fluorophores (Bodipy-507 or Oregon Green) and obtained results similar to those shown for Texas Red MARCKS(151–175) (data not shown). Specifically, 200–300 nM of either Bodipy-507-MARCKS(151–175) or Oregon Green MARCKS(151–175) energy transferred 50% to Bodipy-TMR-PIP₂ on PC/PS/Bodipy-TMR-PIP₂ (70:30:0.1) LUVs, as observed in Fig. 4 with the Texas Red label. The energy transfer with these probes also approached 100% quenching for high (>5) peptide:PIP₂ ratios. MARCKS(151–175) labeled with these hydrophobic probes also rapidly permeates vesicles (see Appendix 1).

The data shown in Figs. 3, 4, and 5 were obtained with vesicles containing 30% PS. We also performed experiments with vesicles containing 0%, 10%, and 17% PS (data not shown). The 0% PS data are consistent with spin label (Rauch et al., 2002), kinetic (Wang et al., 2002), competition, and -potential (Wang et al., 2001) measurements that show each membrane-bound MARCKS(151–175) sequesters approximately three PIP₂ on a PC/PIP₂ membrane. As expected, the vesicles with 10% and 17% PS show stronger energy transfer than those with 30% PS. Because biological membranes contain PE and cholesterol as well as PC, PS, and PIP₂ (White, 1973; Yorek, 1993), we measured FRET with 1:1:1:1 POPC:POPS:POPE:cholesterol + 0.1% Bodipy-TMR-PIP₂ LUVs; the energy transfer was comparable to that illustrated in Fig. 4 (data not shown). Appendix 2 considers in more detail the role of cholesterol in the binding of MARCKS effector domain to membranes.

Effect of aromatics
Fig. 6 shows the effect of replacing the five aromatic (Phe) residues in Texas Red MARCKS(151–175) with Ala: Texas Red FA-MARCKS(151–175) showed approximately twofold weaker energy transfer than Texas Red MARCKS(151–175), indicating that the aromatics increase the sequestration of PIP₂. The lipid concentration was 0.75 mM, sufficient to bind essentially all of the peptide.

View larger version (18K): [in this window] [in a new window]   FIGURE 6  The % energy transfer from Bodipy-TMR-PIP2 to Texas-Red-labeled peptides: Texas Red MARCKS(151–175) (•),Texas Red FA-MARCKS(151–175) (), Texas Red Lys-13(), and Texas Red Lys-7(). The total lipid concentration is 0.75 mM in PC/PS/Bodipy-TMR-PIP2 (70:30:0.1) LUVs.

One potential complication with these experiments is that direct interactions may occur between the fluorophores on the peptide and on PIP₂. We performed control FRET experiments using Bodipy-TMR-DAG, which is produced by hydrolyzing Bodipy-TMR-PIP₂ with PLC-₁; the same vesicles used in the original FRET measurements were hydrolyzed. Because Bodipy-TMR-DAG is electrically neutral, we expect no electrostatic sequestration and no energy transfer between Bodipy-TMR-DAG and Texas Red MARCKS(151–175). The energy transfer (not shown) is significantly less than illustrated in Fig. 4 (maximum 10% transfer rather than 50%), and similar results were obtained using TRITC-PE instead of Bodipy-TMR-PIP₂. Thus, most of the FRET seen in Figs. 4–6 is due not to probe-probe interaction but to lateral interaction of MARCKS(151–175) with PIP₂. We used two techniques, quenching and EPR, to perform experiments with unlabeled peptide, which allowed us to eliminate any probe-probe interactions and to test the possibility that the probe on the peptide is enhancing the lateral sequestration of PIP₂ by some other mechanism (e.g., enhanced image charge effect).

MARCKS(151–175) produces self-quenching of fluorescent PIP₂
Because one MARCKS(151–175) peptide can bind approximately three PIP₂ on a PIP₂/PC membrane (Rauch et al., 2002; Wang et al., 2002), we postulated that the peptide should induce local clustering, and thus self-quenching of fluorescent PIP₂. Fig. 7 A illustrates that unlabeled MARCKS(151–175) decreases the fluorescence of Bodipy-TMR-PIP₂ in PC/Bodipy-TMR-PIP₂ (99:1) vesicles. Fig. 7 B plots the % quenching of Bodipy-TMR-PIP₂ as a function of MARCKS(151–175) concentration (•); data were from Fig. 7 A and similar experiments. These results are consistent with previous experiments (spin-label, kinetic measurements, etc.) showing that MARCKS(151–175) can sequester approximately three PIP₂ lipids on a PC/PIP₂ membrane. Our control experiments examined the ability of MARCKS(151–175) to quench electrostatically neutral TRITC-PE or Bodipy-TMR-DAG: as expected, no significant quenching was observed (data not shown).

View larger version (21K): [in this window] [in a new window]   FIGURE 7  Quenching of Bodipy-TMR-PIP2 due to membrane-bound unlabeled MARCKS(151–175). Total lipid concentration is 0.1 mM. Bodipy-TMR-PIP2 is excited at 547 nm. (A) Representative experiments showing the raw emission spectra for Bodipy-TMR-PIP2 at different concentrations of MARCKS(151–175); vesicles are PC/Bodipy-TMR-PIP2 (99:1) LUVs. (B) The % quenching of Bodipy-TMR-PIP2 as a function of MARCKS(151–175) concentration on vesicles comprised of PC, 1% Bodipy-TMR-PIP2, and either 0 (•), 17% (), or 30% () PS. Each point on the plot is an average of 7 independent experiments.

Fig. 8 shows similar quenching measurements using either unlabeled FA-MARCKS(151–175) or Lys-13; in contrast to MARCKS(151–175), these peptides bind outside the envelope of the polar headgroup region. These measurements allow us to investigate how aromatics and the linear charge density affect the electrostatic sequestration of PIP₂. MARCKS(151–175) (•) produces stronger quenching of Bodipy-TMR-PIP₂ than FA-MARCKS(151–175) (); thus, aromatics enhance the electrostatic sequestration of PIP₂. Lys-13 () produces stronger quenching than FA-MARCKS(151–175) (), even though both peptides contain 13 basic residues and no aromatics; the simplest, but not the only, explanation is that a higher linear density of basic residues also increases the sequestration. (We note in passing that if the quenching shown in Figs. 7 B and 8 were due only to the proximity of the PIP₂ bound to the peptide, it would have exhibited a maximum for 100 < [peptide] < 1000 nM, where the molar ratio of bound peptide to PIP₂ decreases to a value <1. No maxima are observed, which indicates that other factors contribute to the quenching produced by the peptide-lipid interaction.)

View larger version (16K): [in this window] [in a new window]   FIGURE 8  Quenching of Bodipy-TMR-PIP2 due to MARCKS(151–175) (•), Lys-13 (), and FA-MARCKS(151–175) (). PC/PS/Bodipy-TMR-PIP2 (69:30:1) LUVs at a lipid concentration of 0.5 mM are sufficient to bind essentially all of the peptide.

Increasing the salt concentration decreases the lateral sequestration of PIP₂
If the sequestration of PIP₂ by a membrane-bound peptide is due mainly to electrostatics, theory predicts that sequestration should decrease as the salt concentration is increased. FRET measurements with labeled peptides (Fig. 9 A) and quenching measurements with unlabeled peptides (Fig. 9 B) show the lateral sequestration of PIP₂ indeed decreases as the salt concentration increases.

EPR experiments show MARCKS(151–175) can sequester proxyl-PIP₂ in the presence of monovalent acidic lipids
Fig. 10 shows EPR spectra of the proxyl-PIP₂ spin label incorporated into LUVs formed from PC or mixtures of PC and PS in the absence and presence of MARCKS(151–175). In each lipid mixture, addition of MARCKS(151–175) produced a substantial reduction in the EPR resonance amplitude, reflecting a broadening of the normalized EPR spectrum. Previous studies showed addition of molecules known to interact with PIP₂, such as neomycin and the PH domain of PLC-₁, decrease the amplitude of the EPR spectra (Rauch et al., 2002). For MARCKS(151–175), the decrease in spectral amplitude of proxyl-PIP₂ reflects an 30% reduction in the rotational correlation time. Under certain conditions (for example the PC sample in Fig. 10 A), dipolar interactions may produce additional broadening when more than one proxyl-PIP₂ lipid is bound to MARCKS(151–175) (Rauch et al., 2002). The changes seen upon MARCKS(151–175) addition are reversed when the peptide is removed from the membrane interface, e.g., by the addition of Ca²⁺/calmodulin (data not shown).

View larger version (21K): [in this window] [in a new window]   FIGURE 10  EPR spectra of proxyl-PIP2 in bilayers composed of (A) PC, (B) PC/PS (9:1), and (C) PC/PS (7:3) in the absence (black line) and presence (gray line) of MARCKS(151–175). The peptide was added to concentrations of 50, 80, and 120 µM in A, B, and C, respectively. The spin label was present at 0.5 mol%; total lipid concentration is 20 mM; solution contains 100 mM KCl, 10 mM MOPS, pH 7.0. The amplitudes of the EPR spectra have been normalized against the total spin concentration.

View larger version (17K): [in this window] [in a new window]   FIGURE 11  Titration of the central (mI = 0) nitroxide EPR resonance, A(0), as a function of the concentration of added MARCKS(151–175). The titration is shown in vesicle suspensions of PC (), PC/PS (9:1, ), PC/PS (7:3, •) at a total lipid concentration of 7 mM; proxyl-PIP2 is present at 0.85 mol%.

	SUMMARY

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS SUMMARY DISCUSSION APPENDIX 1: PHOSPHOLIPID... APPENDIX 2: Effects of... ACKNOWLEDGEMENTS REFERENCES

Calculations of the sequestration of PIP₂ by FA-MARCKS(151–175) provide support for a nonspecific electrostatic mechanism
We used the Finite Difference Poisson-Boltzmann (FDPB) method to examine two hypotheses: i), lateral sequestration of PIP₂ by membrane-adsorbed basic peptides is due to nonspecific electrostatic interactions, and ii), electrostatic sequestration is significant, even when the membrane contains an appreciable mol fraction of PS. The calculations incorporate molecular models of PIP₂, the peptide, and the PC/PS membrane. Fig. 12 A illustrates the predicted electrostatic properties of a 5:1 PC/PS membrane under a variety of conditions. The front portion of the figure represents the membrane far from either peptide or PIP₂: the -25 mV equipotential profile above this region of "bulk" membrane undulates gently, with hills corresponding to the location of the charged PS lipids. The middle portion of Fig. 12 A illustrates how the adsorption of the basic peptide FA-MARCKS(151–175) to the membrane surface produces a highly localized region of positive potential in its vicinity. The multivalent anionic PIP₂ (shown in yellow and visible in the upper right portion of Fig. 12 A) produces a localized enhancement of the negative potential of the 5:1 PC/PS membrane. The membrane-adsorbed FA-MARCKS(151–175) electrostatically attracts the highly charged PIP₂; as discussed in Wang et al. (2003), the calculated change in electrostatic free energy when PIP₂ partitions from bulk membrane to a position adjacent to the peptide is -2.2 kcal/mol (Fig. 12 B). Therefore, FA-MARCKS(151–175) adsorbed to the surface of a membrane containing 17 mol % PS provides a strong basin of attraction for the electrostatic sequestration of PIP₂. Additional quantitative results, as well as the calculated dependence of PIP₂ partitioning on the ionic strength of the solution, the mol percent acidic lipid in the membrane, and the residue composition of the peptide, are provided in our companion computational paper (Wang et al., 2003).

Fig. 12, C–E, provide an alternative representation of the electrostatic interactions that occur first between FA-MARCKS(151–175) and bulk membrane, then between the membrane-adsorbed peptide and PIP₂. As depicted in Fig. 12 C, FA-MARCKS(151–175) in solution ([KCl] = 0.1 M) has a strong positive electrostatic profile. Fig. 12 D shows the decrease in this positive contour when the peptide binds to the surface of a 5:1 PC/PS membrane (the view is from above, looking down on the membrane surface). FDPB calculations indicate this decrease is due to the favorable electrostatic interaction between the basic peptide and acidic lipids in the membrane; details of the membrane and its electrostatic potential have been removed for clarity. Fig. 12 E illustrates how interaction with PIP₂ further decreases the positive contour of the N-terminal portion of the membrane-adsorbed peptide; this is the same interaction depicted in Fig. 12 B.

Fig. 12, D and F, contrast the positive potential contours surrounding FA-MARCKS(151–175) and Lys-13, respectively, adsorbed to the surface of a 5:1 PC:PS membrane. Although both peptides have a net charge of +13, Lys-13 has a significantly higher linear charge density. Consequently, FDPB calculations predict that the positive equipotential contour of membrane-adsorbed Lys-13 is significantly larger than that of membrane-adsorbed FA-MARCKS(151–175), which consists of 25 amino acid residues. This suggests that Lys-13 interacts with PIP₂ more strongly than FA-MARCKS(151–175) because of its higher linear charge density.

One limitation of our atomic model is obvious: PS is not at electrochemical equilibrium, but is assumed to be spatially fixed. Only PIP₂ redistributes (is laterally sequestered) when the peptide binds to the membrane. Experiments are in progress to address the possibility that PS is also sequestered by basic peptides (May et al., 2000) and that when PIP₂ is sequestered from a membrane containing both PS and PIP₂, there is an exchange of several sequestered PS for PIP₂. Simplified models of peptides and membranes allow one to take into account the electrochemical equilibrium of both PS (May et al., 2000; Groves et al., 2000) and PIP₂ (Haleva et al., 2003) as the peptide adsorbs. If the peptide is assumed to be a disk larger than the Debye length, the charge density on the membrane beneath the adsorbed disk is approximately "matched" to the charge density on the disk (Haleva et al., 2003). In this case there is a release of approximately three PS as one PIP₂ is sequestered. The major conclusion of this disk model agrees with our atomic model calculations: basic peptides can laterally sequester multivalent anionic lipids such as PIP₂ by an electrostatic mechanism.

Simple electrostatics predicts increased sequestration due to aromatics
Figs. 6, 8, and 13 B show that aromatic residues embedded in a cluster of basic residues enhance the lateral sequestration of PIP₂, but how do aromatic residues exert this effect? Fig. 1 B depicts a model based on the results of several different experiments showing that the aromatics insert into the acyl chain region and consequently pull the adjacent basic residues into the polar headgroup region. Moving a charge close to or within the polar headgroup region increases the electrostatic potential it produces—and thus the sequestration of PIP₂—for two reasons. First, when a charge moves close to a region of low dielectric constant, the "image charge" effect enhances the potential, as discussed in standard texts (Dill and Bromberg, 2003; see Chapter 21) and reviews (McLaughlin, 1989; see Fig. 1). In the simplest case (no salt in aqueous, a, or membrane, m, phases) moving a single point charge from a medium of dielectric 80 to the interface with a homogeneous phase of dielectric 2 increases the potential by a factor 2_a/(_m + _a) 2 at any distance from the charge (see Fig. 4.4 in Jackson, 1975). Second, if the aqueous phase contains salt, and the membrane phase does not, moving a point charge q to the interface perturbs the ion atmosphere around the charge. Even if image charge effects are absent (e.g., dielectric constant of the membrane phase is identical to that of the aqueous phase), the potential increases at distances comparable to the Debye length. For example, in a 100-mM salt solution (where 1/, the Debye length, is 1 nm) the potential a distance r = 2 nm from the charge q increases approximately twofold at the membrane surface due to the perturbation of the double layer (Mathias et al., 1992, see Fig. 4 and Eq. 10 for the complicated expression that describes the potential).

A surprisingly simple expression for the potential emerges when the membrane phase has both a low dielectric and excludes ions. For a charge q far from the interface in the aqueous phase, the monovalent ions in solution "screen" the charge, and the potential is described by Debye-Hückel theory: _a = q exp(-r) / (4_oar) where r is the distance from the charge. If the charge q is moved to the interface with a low dielectric membrane phase (_a / _m >> 1) the potential in the aqueous phase is simply 2 times the Debye-Hückel expression at all distances r from the charge (Mathias et al., 1992, see Fig. 3; Stillinger, 1961).

Of course, only the acyl chain region of a phospholipid bilayer has a dielectric 2: the polar headgroup region contains 50% water by volume, and its average dielectric constant is significantly higher. Nevertheless, the two factors discussed above should approximately double the potential produced by a charge on a peptide (at least for r > 1/ 1 nm, z = distance from surface = 0) when the charge is located within the polar headgroup region. These electrostatic phenomena provide a simple explanation for the ability of aromatic residues to enhance the sequestration of PIP₂ by clusters of basic residues observed in Figs. 6, 8, and 13 B.

The energy required to increase the electrostatic potential adjacent to the basic residues comes from the hydrophobic energy gained by insertion of the Phe residues into the acyl chain region of the membrane (Fig. 1 B). Engelman et al. (1986) estimate this energy is 4 kcal/mol per Phe residue, sufficient to account for the observed effects.

More detailed, realistic theoretical calculations are highly model dependent, and are not presented here. We merely note that J. Wang (unpublished) has shown that the potential produced by a model basic peptide does increase in the expected manner as the charges on the peptide approach the polar headgroup interface of an atomic model of a bilayer similar to that shown in Fig. 12.

Biological implications
MARCKS(151–175) sequestration of PIP₂ on LUVs inhibits PLC-catalyzed hydrolysis of PIP₂
Fig. 13 A plots the % PIP₂ hydrolyzed in PC/PS/PIP₂ LUVs as a function of time. In the absence of PLC-₁ (), we observed no significant hydrolysis over 15 min, whereas addition of enzyme produced significant hydrolysis of PIP₂ within the first 3 min (•). Adding 0.5 µM MARCKS(151–175) with the PLC () decreased the initial rate of hydrolysis by 50%. Fig. 13 B shows the effect of peptide on the initial rate of hydrolysis seen in Fig. 13 A, along with the results of similar experiments done with different concentrations of MARCKS