Area Expansion and Permeation of Phospholipid Membrane Bilayers

of Pediatrics Martin Luther King Jr./Drew University Medical Center and Perinatal ... the membrane area expansion resulting from the insertion of fusi...
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Langmuir 1998, 14, 2385-2395

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Area Expansion and Permeation of Phospholipid Membrane Bilayers by Influenza Fusion Peptides and Melittin Marjorie L. Longo,†,‡ Alan J. Waring,§ Larry M. Gordon,§ and Daniel A. Hammer*,‡,| School of Chemical Engineering, Cornell University, Ithaca, New York 14853, and Department of Pediatrics Martin Luther King Jr./Drew University Medical Center and Perinatal Laboratories, Harbor-UCLA, Los Angeles, California 90059 Received August 18, 1997. In Final Form: December 22, 1997 The fusion of membrane bilayers is an ubiquitous cellular process. Exocytosis, organelle formation, cellular trafficking, cell division, fertilization, and numerous other cellular activities all involve membrane fusion. Fusion can be rapid, occurring on the order of seconds or minutes. In viral infection, specific glycoproteins mediate the fusion of the viral lipid envelope with a cellular membrane. Viral fusion glycoproteins typically contain a segment which embeds into the target cellular membrane, referred to as the fusion peptide. The mechanism of action of viral fusion proteins is still not certain; in particular, the extent and rate of insertion of the fusion peptide are not well quantified. In this report, we use micropipet aspiration and video microscopy of large unilamellar phosphatidylcholine vesicles to determine the membrane area expansion resulting from the insertion of fusion peptides into the lipid bilayer. The fusion peptide of the viral fusion protein, influenza hemagglutinin, inserts into phosphatidylcholine bilayers, resulting in an increase in the membrane area on a time scale (i.e., seconds) similar to that of viral fusion. Following peptide insertion, porous defects form in minutes. We show that chemical changes in the Nand C-termini of this peptide can either eliminate, decrease, or enhance surface activity of the peptide; particularly the propensity to form pores can be diminished. In control studies, the well-studied lytic peptide, melittin, similarly increases the membrane area and forms pores in the membrane. The observation that the wild-type influenza fusion peptide and melittin each form pores below the areal expansion limit of 5% suggest that membrane disrupting proteins act through specific and localized perturbation.

Introduction For many membrane-enveloped animal viruses, infection involves the fusion of the viral lipid bilayer envelope with the lipid bilayer membrane of the target cell, either at the cell surface or inside the cell. Fusion is mediated by specific membrane bilayer-associated viral glycoproteins (for reviews see refs 1-3).1-3 Of these, the best characterized is the hemagglutinin protein (referred to as HA) of influenza virus. HA consists of two disulfidebonded subunits, HA1 and HA2 formed from a proteolitic cleavage of the entire HA protein.4-7 As is typical of fusion proteins, HA contains a highly conserved, moderately hydrophobic sequence of approximately 20 amino acids located at the amino terminus (N-terminus) of HA2, referred to as the fusion peptide, 8 which embeds into the neighboring cellular membrane prior to and during fusion.9-12 † School of Chemical Engineering, Cornell University, Ithaca, NY 14853. ‡ Present address: Department of Chemical Engineering and Materials Science, University of California, Davis, CA, 95616. § King-Drew University Medical Center and Perinatal Laboratories-UCLA, Los Angeles, CA 90059. | Corresponding author. Current address: Department of Chemical Engineering, University of Pennsylvania, 220 S. 33rd St., Philadelphia, PA 19104.

(1) Hoekstra, D. J. Bioenerg. Biomembr. 1990, 22, 121-153. (2) White, J. M. Annu. Rev. Physiol. 1990, 52, 675-697. (3) Stegmann, T.; Doms, R. W.; Helenius, A. Annu. Rev. Biophys. Biophys. Chem. 1989, 18, 187-211. (4) Gething, M. J.; Doms, R. W.; York, D.; White, J. M. J. Cell Biol. 1986, 102, 11-23. (5) Steinhauer, D.; Wharton, S.; Skehel, J.; Wiley, D. C. J. Virol. 1995, 69, 6643-6651. (6) Garten, W.; Bosch, F. X.; Linder, D.; Rott, R.; Klenk, H.Virology 1981, 115, 361-374. (7) Melikyan, G. B.; Niles, W. D.; Peeples, M. E.; Cohen, F. S. J. Gen. Physiol. 1993, 102, 1131-1149.

To initiate infection, influenza virus attaches to sialic acid residues on the target cell membrane via a receptor site (located on HA1).9 This attachment triggers the cell to take in the virus through endocytosis. Thereafter, the virus resides inside the cell in a lipid membrane capsule, referred to as the endosome.13 In the endosome, the pH is initially approximately 7, but through progressive pumping of protons into the endosome, the pH decreases to approximately 5. At a threshold pH between 5 and 6 there is a conformational change in HA,3,4,11,14,15 such that the fusion peptide is exposed to insert in the adjacent endosomal membrane. Insertion is followed by membrane fusion.9-12 As a result of fusion, the nucleocapsid of the virus is deposited in the cell cytoplasm to ultimately enter the nucleus for viral replication. Often, the structure of viral proteins involved in membrane disruption is R-helical. In the presence of lipid membrane, approximately 50% of the amino acid sequence of the synthesized fusion peptide is R-helical.16 If the HA fusion peptide assumes a helical conformation, it will form (8) Lamb, R. A. In Genetics of Influenza Viruses; Palese, P., Kingsbury, D. W., Eds.; Springer-Verlag: Berlin, 1983; pp 21-69. (9) Wiley, D. C.; Skehel, J. J. Annu. Rev. Biochem. 1987, 56, 365369. (10) Stegmann, T.; Delfino, J. M.; Richards, F. M.; Helenius, A. J. Biol. Chem. 1991, 266, 18404-18410. (11) Harter, C. J. Biol. Chem. 1989, 264, 6459-6464. (12) Weber, T.; Paesold, G.; Galli, C.; Mischler, R.; Semenza, G.; Brunner, J. J. Biol. Chem. 1994, 269, 18353-18358. (13) Matlin, K. S.; Reggion, J.; Helenius, A.; Simons, K. J. Cell Biol. 1981, 91, 601-613. (14) Ruigrok, R. W. H.; Martin, S. R.; Wharton, S. A.; Skehel, J. J.; Bayley, P. M.; Wiley, D. C. Virology 1986, 155, 484-487. (15) White, J. M.; Wilson, I. A. J. Cell Biol. 1987, 105, 2887-2896. (16) Gray, C.; Tatullian, S. A.; Wharton, S. A.; Tamm, L. K. Biophys. J. 1996, 70, 2275-2286.

S0743-7463(97)00932-3 CCC: $15.00 © 1998 American Chemical Society Published on Web 04/08/1998

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a “sided helix” or amphipathic helix17 with most of the bulkier, more hydrophobic residues found on one hemiface of the helix and most of the smaller apolar amino acids (e.g., alanine and glycine) residing on the other side of the helix.2,11 A possible role of this “sidedness” is to promote interaction of a hydrophobic peptide face with the hydrophobic chains of the membrane lipids during and prior to fusion. Fusion peptides from influenza virus HA have been studied intensely.16,18-25 The activity of synthetic HA fusion peptide has been measured by its ability to fuse and/or lyse (cause leakage of) model lipid vesicles or red blood cell membranes.18-20,23,25 The lytic activity of HA fusion peptides correlates well with the in vivo fusion activity of the full HA molecule. For example, the lytic activity of fusion peptides found in native influenza viruses (so-called “wild-type” peptides) increases greatly as the pH decreases to near 5.0 (endosomal pH).19,20,23,25 Furthermore, fusion peptides derived from altered forms of influenza viruses that are unable to infect cells induce very little lysis of lipid-containing membranes.19,23 The exact nature of the membrane perturbation that results in lysis is unknown. There has been much focus on the particular amino acids responsible for fusion activity and the attenuation or elimination of fusion activity caused by changing amino acid residues or deleting residues of the HA fusion peptide.4-6,16,19,23,26 A primary goal of this work is the design of pharmaceuticals that might block the active site of the viral fusion molecule, thus preventing viral entry. It has become clear that slight mutations of the three HA2 N-terminal amino acids (NH2-glycine-leucine-phenylalanine, one letter amino acid sequence ) GLF) can lead to a significant decrease or elimination of fusion (or lytic) activity. Concomitantly, these sequence changes result in a significant decrease in the R-helical content of the peptide in comparison to the native sequence when lipid is present.16 Wharton et al.19 demonstrated that deletion of the N-terminal glycine of a synthetic 20 amino acid HA fusion peptide (∆G1 peptide) greatly decreased fusion activity at pH 5.0 and 7.0 and virtually eliminated the peptide’s hemolytic (lysis of red blood cells) activity at pH 5.0 in comparison to the wild type peptide. This lack of fusion and lytic activity in the synthetic ∆G1 fusion peptide corresponds to an absence of fusion in cells expressing HA in which the N-terminal glycine of the fusion peptide was chemically removed.5 Additionally, it has been demonstrated that substitution of the N-terminal apolar amino acid glycine with many other amino acid residues results in a significant reduction in fusion activity (substitution with alanine is the only exception found thus far).5,19 Steinhauer et al.5 addressed the role of positional specificity of the N-terminal glycine by producing two HA fusion peptides, 16 amino acids in length, containing alanine (A) inserted at two different locations at the (17) Eisenberg, D. Annu. Rev. Biochem. 1984, 53, 595-623. (18) Lear, J. D. J. Biol. Chem. 1987, 262, 6500-6505. (19) Wharton, S. A.; Martin, S. R.; Ruigrok, R. W. H.; Skehel, J. J.; Wiley, D. C. J. Gen. Virol. 1988, 69, 1847-1857. (20) Du¨zgu¨nes, N.; Shavnin, S. A. J. Membr. Biol. 1992, 128, 71-80. (21) Du¨zgu¨nes, N., Gambale, F. FEBS. Lett. 1988, 227, 110-114. (22) Burger, K. N.; Wharton, S. A.; Demel, R. A.; Verkleij, A. J. Biochemistry 1991, 30, 11173-11180. (23) Rafalski, M.; Oritz, A.; Rockwell, A.; van Ginkel, L. C.; Lear, J. D.; DeGrado, W. F.; Wilschut, J. Biochemistry 1991, 30, 10211-10220. (24) Ishiguro, R.; Kimura, N.; Takahashi, S. Biochemistry 1993, 32, 9792-9797. (25) Wagner, E.; Plank, C.; Zatloukal, K.; Cotten, M.; Birnstiel. M. L. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 7934-7938. (26) Daniels, R. S., Downie, J. C., Hay, A. J., Knossow, M., Skehel, J. J., Wang, M. L., Wiley, D. C. Cell 1985, 40, 431-439.

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N-terminus (GALF and AGLF). AGLF showed slightly lowered hemolytic activity in comparison to the wild-type peptide, whereas GALF demonstrated significantly lower activity. Although mutations of the N-terminus can detrimentally affect fusion capability and hemolytic activity of HA fusion peptides, this effect does not necessarily result from a lack of binding to a lipid membrane. Fusion peptides containing substitution or deletion of the N-terminal glycine, in addition to the AGLF and GALF mutants, can bind liposomes.5,16,19 Clearly, the ability of a peptide to bind a membrane is not a direct indicator of its ability to perturb or insert in the membrane. Also, the ability of a peptide to lyse a membrane (an all-or-none phenomenon) is not a sensitive indicator of the kinetics or extent of protein insertion in the membrane. Only in the work of Gray et al.16 was some measure of the extent of insertion of peptide or perturbation of the liposomes included. This recent work,16 employing Fourier transform infrared (FTIR) spectroscopy, suggests that the nonfusogenic fusion peptides of HA interact more strongly with lipid headgroups compared to the fusogenic peptides. However, to our knowledge, with the exception of our recent paper,27 no measurements of the extent of binding or insertion of fusion peptides have been performed on the time scales over which fusion occurs (i.e., seconds to minutes7,10,18-20,23,24,28-37). Recently, we investigated the use of micropipet aspiration to quantify peptide insertion by measuring bilayer area expansion.27 Similar studies have been performed by Needham and Zhelev38 to quantify lysolipid insertion. We focused on the native fusion peptide of influenza virus (wt-20) of the X31 strain. We used micropipet aspiration to measure wt-20 insertion and desorption, wt-20 induced bilayer defect formation, and pH triggering of wt-20 insertion and desorption. It was necessary to find a control which was unlikely to insert in the lipid bilayer. Therefore, the ∆G1 fusion-diminished influenza fusion peptide was used. Micropipet aspiration was used to directly expose a single-membraned vesicle to a peptide-containing solution while holding the membrane at a low, constant tension (0.1-0.2 dyn/cm). Solution conditions were readily altered, such as pH and concentration, to match those that correspond to fusion. The vesicles used for micropipet aspiration were sufficiently large (approximately 25 µm in diameter) that changes in membrane area were quantified through optical microscopy observations of changes in the vesicle projection into a micropipet (inner (27) Longo, M. L.; Waring, A. J.; Hammer, D. A. Biophys. J. 1997, 73, 1430-1439. (28) Slepushkin, V. A.; Melikyan, G. B.; Sidorova, M. S.; Chumakov, V. M.; Andreev, S. M.; Manulyan, R. A.; Karamov, E. V. Biochem. Biophys. Res. Commun. 1990, 172, 952-957. (29) Mobley, P. W; Lee, H. F.; Curtain, C. C.; Kirkpatrick, A.; Waring, A. J.; Gordon, L. M. Biochim. Biophys. Acta 1995, 1271, 304-314. (30) Glushakova, S. E.; Omelyaneko, V. G.; Lukashevitch, I. S.; Bogdanov, A. A., Jr.; Moshnikova, A. B.; Kozytch, A. T.; Torchilin, V. P. Biochim. Biophys. Acta 1992, 1110, 202-208. (31) Stegmann, T. Biochemistry 1985, 24, 3107-3113. (32) Hoekstra, D. Biochemistry 1985, 24, 4739-4745. (33) Zimmerberg, J. J. Cell Biol. 1994, 127, 1885-1894. (34) Hug, P., Sleight, R. G. J. Biol Chem. 1994, 269, 4050-4056. (35) Struck, D. K.; Hoekstra, D.; Pagano, R. E. Biochemistry 1981, 20, 4093-4099. (36) Fonteijn, T. A. A.; Engberts, J. B. F. N.; Nir, S.; Hoekstra, D. Biochim. Biophys. Acta 1992, 1110, 185-192. (37) Bagai, S.; Puri, A.; Blumenthal, R.; Sarkar, D. P. J. Virol. 1993, 67, 3312-3318. (38) Needham, D.; Zhelev, D. V. Annu. Biomed. Eng. 1995, 23, 287298.

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diameter approximately 8 µm).27,38,39 Therefore, substantial and rapid area changes accompanying wt-20 insertion and desorption were assessed by optical microscopy.27 By use of solutes of different size within and outside vesicles, membrane permeation was detected by the swelling or collapse resulting from unidirectional flux of solute and cotransport of water.27,38 In addition, alterations in mechanical properties and tensile strength of vesicles that result from peptide insertion were measured.27,40,41 In our control study, we found that ∆G1 did not expand the bilayer significantly in comparison to wt-20.27 Here, we built on our previous work, by investigating two new structural variants of HA fusion peptide by micropipet aspiration of large unilamellar vesicles. Additionally, for comparison to HA fusion peptides, we investigate the insertion of the potent lytic (pore forming) peptide of bee venom, melittin. We also performed experiments with wt-20 and ∆G1 for comparison and investigation of N-terminus composition effects. In the first new peptide (FITC-wt-20), we appended the wt-20 peptide by attaching fluoroscein-5-isothiocyanate to the N-terminal glycine. With this new peptide, we were able to observe several phenomena. First, by a combination of micropipet aspiration and fluorescence microscopy, we were able to detect fluorescence associated with inserted peptides and loss of fluorescence associated with desorption. Second, we were able to investigate the consequences of N-terminus labeling on insertion and permeation. This situation is similar to previous lytic experiments by Steinhauer et al.,5 discussed above, in which a single amino acid is added to the wild type N-terminus. Our results were compared to the ∆G1 peptide in which the Nterminus has been removed. Third, because it was found that FITC-wt-20 does not rapidly permeate vesicles, but does expand the area to a significant extent, we were able to look at the effect of flow on area expansion and thus the role of mass transfer in these experiments. In the second new peptide, the carboxyl (C-terminus) was capped by carboxyamidation. The rationale behind this peptide structure was that the NH of the CONH2 at the C-terminus can hydrogen bond to the fourth residue from the C-terminus, thus encouraging R-helical formation at the C-terminus. Theoretically, the peptide may behave as if it is part of the helical structure of a longer protein. In addition, this peptide was lengthened to include 23 residues of the N-terminal protein domain contained in the HA protein. This peptide is uncharged at the C-terminus, further mimicking the pH dependent charge of the native protein at that location in the amino acid sequence. With this peptide, we were able to investigate a shift in the active pH to a higher value. The conformation of this peptide in phospholipid bilayers was compared to that of wt-20 using CD spectroscopy. The lack of pH and sequence-dependent conformational change, similar to results reported by Gray et al.16 for wt-23 (no carboxyamidation), results in our conclusion that peptide charge is likely to be the most important determinant of lipid bilayer insertion for pH-activated HA fusion peptides. Finally, as a comparison to the HA fusion peptide, we performed area expansion and permeation experiments using another well-studied lytic peptide, melittin.42 Melittin is similar in structure to the native HA fusion peptide, containing a large percentage of apolar amino acids in the (39) Simon, S. A.; Disalvo, E. A.; Gawrisch, K.; Borovyagin, V.; Toone, E.; Schiffman, S. S.; Needham, D.; McIntosh, T. J. Biophys. J. 1994, 66, 1943-1958. (40) Evans, E.; Needham, D. J. Phys. Chem. 1987, 91, 4219-4228. (41) Evans, E.; Rawicz, W. Phys. Rev. Lett. 1990, 64, 2094-2097. (42) Dempsey, C. E. Biochim. Biophys. Acta 1990, 1031, 143-161.

Langmuir, Vol. 14, No. 9, 1998 2387 Table 1. Amino Acid Sequences of Peptides Derived from the N-Terminal Segment of Hemagglutinin and the Lytic Peptide Melittina wt-20 FITC-wt-20 wt-23-CONH2 ∆G1 melittin

NH2-GLFGAIAGFIENGWEGMIDG-COOH FITC-GLFGAIAGFIENGWEGMIDG-COOH NH2-GLFGAIAGFIENGWEGMIDGWYG-CONH2 NH2-LFGAIAGFIENGWEGMIDG-COOH NH2-GIGAVLKVLTTGLPALISWIKRKRQQ-CONH2

a Residues are represented by the one-letter amino acid code. NH2 indicates the amino terminal, COOH represents the carboxy terminus, and CONH2 represents the carboxyamidation of the C-terminal residue.

26 amino acid sequence. Also like the HA fusion peptide, melittin folds into a helical rod of amphiphathic character when in the presence of lipid. Materials and Methods Materials. 1-Stearoyl-2-oleoylphosphatidylcholine (SOPC) and 1-palmitoyl-2-oleoylphosphatidylserine (POPS) were purchased from Avanti Polar Lipids (Alabaster, AL) and used without further purification. Chloroform and methanol, used to form lipid films for vesicle production, were obtained from Fisher (Fairlawn, NJ) and were HPLC grade. All solutes used were from Sigma (St. Louis, MO) and were ultra grade. Water was obtained from a Milli-Q UV Plus system (Millipore, Bedford, MA). The four peptides synthesized (wt-20, wt-23-CONH2, FITCwt-20, and ∆G1) were based on the HA fusion-active (wt-20, wt-23) and fusion-diminished (∆G1) peptides previously synthesized by Wharton et al.19 The sequences of these peptides and melittin are shown in Table 1. For micropipet aspiration experiments, peptides were stored frozen in DMSO (Fisher, HPLC grade) at 2 mM concentration, except melittin which was combined with the glucose buffer (described below) at a concentration of 20 µM and remained unfrozen. Peptides were mixed with buffers solution immediately before use. Melittin was purchased from Calbiochem-Novabiochem (La Jolla, CA) and is 98% in purity. The remainder of the peptides were synthesized as follows. Peptide synthesis reagents including solvents and Fmoc amino acid derivatives were from Perkin-Elmer Applied Biosystems (Foster City, CA). The peptides were synthesized on a 0.25 mmol scale with an ABI model 431A peptide synthesizer using FastMoc chemistry.43 A prederivatized Wang p-benzyloxybenzyl alcohol resin (Applied Biosystems, Foster City, CA) with the C-terminal glycine was used for the stepwise synthesis of the peptides. All residues were double coupled to optimize the yield of the fusion peptide sequence. After cleavage of the peptides from the resin, the crude product was purified by reverse-phase high-performance liquid chromatography (HPLC) employing a Vydac C4-column (Vydac, Hesperia, CA). The material was chromatographed using a 60min linear gradient with the starting phase as water and the eluting phase as 100% acetonitrile containing 0.1% TFA as an ion pairing agent. After concentration of the peptides by vacuum centrifugation, the peptides were freeze-dried from acetonitrile: 10 mM HCl, 1:1 (v:v) to remove any counterions from the chromatography solvent. The molecular masses of the peptides were then confirmed by fast atom bombardment mass spectroscopy (Center for Molecular and Medical Sciences Mass Spectrometry, University of California, Los Angeles). The 23-residue hemagglutinin peptide (wt-23-CONH2) was synthesized with a Rink amide MBHA resin (AnaSpec, Inc., San Jose, CA) and was double coupled and purified as described above. The crude peptide was purified by reverse-phase HPLC with a Vydac C4-column (Vydac, Hesperia, CA) as described above. The molecular mass of the peptide was confirmed by fast atom bombardment mass spectroscopy as described above. The N-terminal fluorescein derivative of the hemagglutinin fusogenic peptide (FITC-wt-20) was prepared by reaction of the (43) Fields, C. G.; Lloyd, D. H.; Macdonald, R. L.; Ottenson, K. M.; Noble, M. L. Peptide Res. 1991, 4, 95-101.

2388 Langmuir, Vol. 14, No. 9, 1998 peptide with fluorescein-5-isothiocyanate (Molecular Probes, Eugene, OR). The peptide was suspended in 1,1,1,3,3,3hexafluoro-2-propanol (Aldrich Chemical Co., Milwaukee, WI): 100 mM bicarbonate buffer of pH 9, (7:3, v:v) and a 4-fold excess of the fluorescent label was added to the reaction mixture. The reaction was allowed to proceed for 1 h before removing the unreacted label from the derivative by gel filtration with a Sephadex G-10 column. Unlabeled peptide was then separated from the covalently labeled hemagglutinin peptide by reversephase HPLC as described for the purification of the crude peptide. Preparation of Vesicles for Micropipet Aspiration. Vesicles containing 100 mM sucrose were formed of 99.5% SOPC and 0.5% POPS (mol:mol) by an electric field technique developed by Angelova et al.,44 with minor modifications. We included 0.5 % POPS to prevent vesicle aggregation and adhesion to chamber surfaces. Briefly, 0.25 mg of lipid was dried onto two electrodes contained in a chamber from a chloroform/methanol solution. The chamber was filled with 100 mM sucrose solution. A 3-V sine wave was applied across the electrodes in the following sequence: 10 Hz, 1/2 h; 3 Hz, 15 min; 1 Hz, 7 min; 0.5 Hz, 7 min. The vesicles that formed on the electrodes were harvested by flowing sucrose solution through the chamber. This technique yields mainly unilamellar large vesicles. Vesicles were no longer used 3 days after production. The complete preparation is described in ref 27. Micropipet Aspiration Chamber. The chamber used in these experiments (described in ref 27) contains two sections separated by a 1 mm thick Teflon sheet. A 0.9-mm hole was bored in the Teflon sheet to allow transfer of vesicles via a micropipet from one section (section A), containing vesicles, to another section (section B), containing peptide. The hole remained blocked by capillary tubing. For vesicle transfer, from section A to section B, the capillary tubing was removed from the hole. Water-saturated nitrogen was delivered to the air-solution interface of both sections to minimize evaporation. Micropipet Aspiration. One-half hour before aspiration began, section A of the chamber was filled with a solution containing 100 mM glucose, 1 mM Na citrate, 1 mM HEPES, 1 mM MES, and 0.04 wt % bovine serum albumin (fraction V, low heavy metals; Calbiochem-Novabiochem, La Jolla, CA) pH adjusted with 1 N and 0.1 N NaOH or HCl solutions. Section B was filled with the same solution, containing peptide (which had been dispersed from a 2 mM solution in DMSO). The maximum concentration of DMSO was 1% for a 20 µM peptide solution. Pretreating the chamber for 1/2 h with the albumin present in the test solutions results in the dissipation of charge on the glass microscope slides used in the chamber.45 After 1/2 h, approximately 6 µL of vesicle solution in 100 mM sucrose was transferred to section A of the chamber. The same amount of 100 mM sucrose was also added to chamber section B to balance the osmotic strengths in the two chamber sections as closely as possible. The difference in refractive indices between the sucrose solution inside of the vesicles and glucose solution in the surrounding media allows for high contrast between the vesicles and their surroundings. During all stages of the experiment, the chamber was held between 20 and 23 °C (determined by a Minco platinum RTD temperature probe; Minco, Minneapolis, MN) by adjusting the room temperature. Aspiration of vesicles was performed by the use of a twochambered manometer with attached Validyne (Northridge, CA) pressure transducers and monitor. A micropipet was attached to the manometer (for preparation and calibration of micropipets, see ref 27) and manipulated with a micromanipulator (Technical Products International, St. Louis, MO). Initially, a test vesicle was aspirated by micropipet suction. Subsequently, the suction was increased until the vesicle lysed. This test was performed to obtain an average lysis tension (τlyse, see below for calculation of τ) for the populations of vesicles used in these experiments (for good references of materials properties measurements using micropipet aspiration see refs 40 and 41). Values of τlyse were compared to literature values for the same lipid composition to (44) Angelova, M. I.; Sole´au, S.; Me´le´ard, Ph.; Faucon, J. F.; Bothorel, P. Prog. Colloid Polym. Sci. 1992, 89, 127-131. (45) Zhelev, D. V.; Needham, D. Biochim. Biophys. Acta 1993, 1147, 89-104.

Longo et al. verify unilamellarity of the vesicles. These τlyse measurements were compared to τlyse measurements from vesicles which had been exposed to peptides. Immediately following the test of lysis tension, a vesicle was chosen for the insertion experiment. Initially, we determined the area compressibility (K) of the vesicle. From all the area compressibility measurements taken at the beginning of the experiment, we obtained an average K for the population of vesicles used in these experiments. This compressibility modulus, K, could then be compared to K for vesicles that had interacted with peptide and literature values of K for vesicles of the same composition. Area compressibility data were obtained by initially increasing the suction pressure to approximately 5 dyn/cm and then decreasing the suction pressure on the vesicle in a stepwise manner (approximately 4 cm of water per step). From the suction pressure changes (∆P) we calculated the isotropic tension, τ (τ ) ∆PRp/2{1 - Rp/Ro}),41 where Rp and Ro are the pipet and exterior vesicle segment radii, respectively. From the accompanying changes in the projection length (∆L), we calculated the areal strain, R ) ∆A/Ao where Ao is the initial area of the vesicle and ∆A is the change in area. R can be closely approximated by 1/ {(R /R )2 - (R /R )3}∆L/R .41 K was determined by the slope 2 p o p o p of the isotropic tension, τ, versus areal strain, R. Subsequently, the aspiration pressure was lowered and maintained throughout the rest of the experiments to give a constant membrane tension between 0.1 and 0.2 dyn/cm. The vesicle was positioned approximately 150 µm from the capillary tubing blocking the hole connecting the two sections of the chamber. To transfer a vesicle from one chamber to another (A to B), we held the pipet steady and manipulated the lateral position of the microscope stage. The transfer also resulted in the removal of the capillary tubing blocking the hole between the two chamber sections. Insertion was monitored by measuring the change in projection (∆L) of the vesicle in the pipet as the vesicle membrane area increased. When vesicles were subject to flow, a flow pipet was used. The inner diameter of the flow pipets were approximately 100 µm. A flow pipet, filled with peptide solution, was attached to a manometer. Seconds before the experiment began, the pressure in the manometer was adjusted to give a constant flow of approximately 150 µm/s (the speed of a particle flowing out of the flow pipet, at the entrance). The entrance of the flow pipet was moved to the vesicle very quickly with the vesicle centered several micrometers from the flow pipet opening. Area expansion, area compressibility, and lysis tension data were recorded using a Nikon inverted Diaphot-TMD microscope equipped with Hoffman optics (Modulation Optics; Greenvale, NY) and epifluorescence illumination. Events were recorded with a Videoscope International CCD-200 video camera (Washington, DC), and images contained video overlay (suction pressure and time data are overlaid frame by frame on all images) with a video encoder (courtesy of R. Waugh, University of Rochester, NY) and stored on videotape using a Sony SVO-9500 VTR. Image analysis software and hardware (Inovisions Corp., Durham, NC) were used to obtain vesicle dimensions, projection versus time data, and the images shown here. An intensifier attachment (Videoscope International grade A image intensifier) was used to monitor fluorescence associated with the peptide, FITC-wt20. Peptide Solutions Conditions. To compare the bilayer interaction of various structural motifs of the fusion peptide of HA with the native HA fusion peptide (wt-20), we chose a concentration (10 µM) corresponding to the concentration used in hemolytic and fusion experiments previously performed by Wharton et al.19 We performed our tests at a pH of 7.0 (corresponding approximately the pH encountered by the peptide outside a target cell) and pH 5.0 (corresponding to the active, endosomal pH). Preparation of Liposomes for Circular Dichroism (CD) Spectroscopy. Unilamellar vesicles used for studies of peptide conformation in lipid bilayers were prepared by extrusion through polycarbonate membrane filters.46 Briefly, a dry SOPC lipid film was hydrated with 10 mM Na citrate buffer, pH 5.0, or 10 mM (46) Waring, A. J.; Harwig, S. S. L.; Lehrer, R. L. Protein Peptide Lett. 1996, 3, 177-184.

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Figure 1. Sequential videomicrographs of a large unilamellar vesicle held under constant, low membrane tension (0.17 dyn/cm) exposed to 10 µM wt-20 (influenza hemagglutinin fusion peptide) at pH 5.0: (A) at the moment of exposure; (B, C) as the vesicle projection increases in length from the insertion of wt-20; (D, E) as porous defects form in the membrane and the permeation of glucose and water into the vesicle causes the vesicle to swell, resulting in a decreased projection length; (F) as the vesicle lyses and is aspirated down the pipet bore because of the mild suction in the pipet. HEPS buffer, pH 7.0, at a concentration of 500 nmol total lipid per mL of buffer. The suspension was vortexed to form multilamellar vesicles, then freeze thawed five times, and finally extruded through 100 nm pore size polycarbonate filters (Nuclepore Corp., Pleasanton, CA) seven times using a LipoFast device (Avestin Inc., Ottawa, ON). The peptide in hexafluoro-2propanol, HFI (0.4 mM) was added to the vesicle dispersion at a lipid to peptide ratio of 50:1, mole:mole and allowed to incubate with the liposomes for 1 h before measurements by CD. HFI was used rather than DMSO since DMSO absorbs strongly in the ultraviolet range used in CD spectroscopy whereas HFI is UV transparent. The size distribution of extruded SOPC vesicles was measured by dynamic light scattering with a Microtrac 9230 UPA ultrafine particle analyzer (Leeds and Northrup, St. Petersburg, FL). Extrusion through 100 nm pore filters yielded a single population of SOPC vesicles with mean diameter of 974 nm and a standard deviation of 110 nm. CD Spectroscopy. CD measurements were made with an AVIV 62DS spectropolarimeter (AVIV Associates, Lakewood, NJ). The spectropolarimeter was fitted with a cell holder that mounted the sample cuvette close to the photomultiplyer to minimize light scattering artifacts,47 and the temperature of the sample was maintained at 25 °C using a thermoelectric temperature controller. Lipid-peptide dispersions in 0.01-0.05 cm light path demountable cells were scanned from 250 to 195 nm at a rate of 10 nm/min and a sample interval of 0.2 nm. The instrument (47) Bruni, R.; Taeusch, H. W.; Waring, A. J. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 7451-7455.

was routinely calibrated with (+)-10-camphorsulfonic acid (1 mg/ mL) and a 1-mm path length cell,48 and the ellipticity expressed as the mean residue ellipticity, [θ]MRE (deg cm2 dmol-1). Peptide sample concentrations were determined from the UV absorbance at 280 nm using an extinction coefficient based on aromatic amino acid residues49 and by quantitative amino acid analysis (UCLA Microsequencing Facility, Los Angeles, CA). The percentage of R-helix conformation in the peptide was estimated using the formalism of Chen et al.50 This approach assumes the maximum theoretical ellipticity for a given peptide or protein at 222 nm may be derived from the number of amino acid residues n, and the ellipticity at 222 nm of a helix of infinite length described by: % R-helix ) [Θ]MRE222/[-39500(1 - (2.57/n))] deg cm2 dmol-1.

Results Area Expansion Resulting from Insertion of WildType Influenza Hemagglutinin Fusion Peptide. For the purpose of comparison, we have provided insertion data of the wild-type fusion peptide (wt-20). When we exposed a vesicle at pH 5.0 to a solution containing 10 µM wt-20, the projection rapidly increased due to the insertion (48) Johnson, W. C., Jr. Proteins Struct. Funct. Genet. 1990, 7, 205214. (49) Gill, S. C.; von Hippel, P. H. Anal. Biochem. 1989, 182, 319326. (50) Chen, Y. H.; Yang, J. T.; Chau, K. H. Biochemistry 1974, 13, 3350-3359.

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Table 2. Area Expansion and Permeation Time Values for Vesicles Exposed to 10 µM of Influenza HA Fusion Peptide (Unless Otherwise Stated) peptide (pH) wt-20 (5.0) wt-20 (7.0) control (5.0)a control (7.0)a wt-23-CONH2 (5.0) wt-23-CONH2 (7.0) FITC-wt-20 (5.0) FITC-wt-20 (5.0)c FITC-wt-20 (7.0) FITC-wt-20 (5.0)d ∆G1 (pH 5.0) ∆G1 (pH 7.0) melittin, 0.1 µM (pH 5.5) melittin, 0.5 µM (pH 5.5)

permeation ∆A/Ao,maxb start time (s) (maximum ∆A/Ao increase) 197 403 35 35

51

29

0.086 0.0062 0.0043 0.0066 0.067 0.037 0.026 0.026 0.010 0.085 0.015 0.0059 0.012 0.0099

a

These measurements should be viewed as baseline measurements of ∆A/Ao,max as a result of projection increases caused by increases in osmotic strength. b Maximum uptake before plateau (if any). c 20 µM peptide. d Using a flow pipet, flow of approximately 150 µm/s.

of wt-20 into the membrane bilayer (Figure 1 A-C). Subsequently, the vesicle swelled due to permeation of the relatively smaller sugar species, glucose, through porous defects in the lipid bilayer and cotransport of water. This volume increase caused the projection to decrease (Figure 1D,E). Eventually, swelling of the vesicle increased the tension in the plane of the membrane and led to mechanical failure (lysis) of the vesicle membrane. When lysis occurred, the vesicle was slowly aspirated into the pipet bore (Figure 1F). Increases in projection (∆L) (Figure 1) were converted into membrane area changes, normalized by the initial area of the vesicle (∆A/Ao), plotted versus time (Figure 2); ∆A ) 2πRp(1 - Rp/Ro)∆L.40 Normalization by the vesicle initial area allows for the comparison of measurements taken from various-sized vesicles. Swelling of the vesicles was monitored by the magnitude of the decrease in the projection length. These measurements were converted into volume changes normalized by the volume of the vesicle at the onset of swelling (∆V/Vo). The calculation of ∆V/Vo was based on the volume change indicated by the decrease in projection length. The total change in volume was found to be consistent with measurements taken of the vesicle volume outside of the pipet prior to rupture. In order to compare the insertion behavior of the five peptides, investigated in this work, we have tabulated (Table 2) the maximum area expansion (∆A/Ao,max) before either a plateau in area expansion or permeation and the time to the start of the permeation of the typical plots shown here. At a pH of 5.0, wt-20 insertion resulted in a ∆A/Ao,max of approximately 0.086 in roughly 3 min. In contrast, at a pH of 7.0, no significant area expansion occurs. Lysis occurred more slowly for a vesicle held at pH 7.0 compared to a vesicle held at pH 5.0 (approximately 14.5 and 5 min, respectively). DMSO Control. To demonstrate that the small amount of DMSO used to solubilize the peptides caused no significant changes in vesicle strength, we calculated the lysis tension of vesicles transferred to pH 5.0 and 7.0 glucose buffer solutions containing 0.5% DMSO. We then compared those lysis tensions (τlyse) to τlyse of control vesicles that were not exposed to DMSO and found them to be similar. The average τlyse of control vesicles was found to be 9.5 ( 2.0 dyn/cm. The τlyse of pH 5.0 and pH 7.0 vesicles exposed approximately 12 min to 0.5% DMSO was 9.0 and 8.3 dyn/cm, respectively.

Figure 2. Fractional area change of vesicles (∆A/Ao, axis on left) versus time (points connected by smooth solid curves). Fractional volume (∆V/Vo, axis on right) versus time (points connected by dashed curves). A vesicle exposed to 10 µM wt-20, pH 5.0 (filled squares) initially expands in area (a, b) due to the insertion of the peptide. Then the vesicle swells in volume (c, d) due to the permeation of sucrose and cotransport of water into the vesicle. A vesicle exposed to 10 µM wt-20, pH 7.0 (filled circles). When a vesicles is exposed to the buffer solution containing the same concentration of DMSO used to solubilize the peptide, the vesicle volume decreases slightly due to a small increase in osmotic strength: pH 5.0 (open squares). Therefore a baseline ∆A/Ao in all experiments can be estimated by the apparent area expansion (solid line, open squares).

We found that transfer of vesicles to 0.5% DMSO in buffer solution resulted in small increases in projection length relative to vesicles in wt-20 at pH 5.0. This baseline change in projection was due to a difference in osmotic strength between the two sections of the chamber and some evaporation that occurs during the experiment. Transferring a vesicle to this slightly hypertonic solution caused the vesicle volume to shrink, thus balancing the osmotic strength inside and outside of the vesicle. The projection length readjusted to accommodate the decrease in volume. The volume decrease, normalized by the initial volume of the vesicle (∆V/Vo ), versus time is shown if Figure 2. This change in osmotic strength was present in all experiments shown here; thus this apparent contribution to ∆A/Ao versus time is shown in Figure 2. Thus, the values of ∆A/Ao,max for all measurements should be compared to DMSO control values. The change in volume causes an overcalculation of ∆A/Ao,max of approximately 0.005 (average of pH 5.0 and pH 7.0 measurements). Often, the extent of area changes was approximately an order of magnitude greater than these errors. Effect of Carboxyamidation of the C-Terminus. It is possible that a small peptide, such a wt-20, based on a larger protein, may not fully capture the conformation of the N-terminal section of the complete fusion protein. In particular, in a small peptide the C-terminus is not able to hydrogen bond to other amino acid residues. Therefore, the last few amino acids of the C-terminus of the peptide may be prevented from forming an R-helical conformation, which may be present in the larger protein. Additionally, the complete amino acid sequence which interacts with the target membrane may not be properly represented in a small peptide. Therefore, we performed area expansion experiments with a peptide in which the C-terminus was carboxyamidated and lengthened by three amino acids of the native sequence. The rational behind this peptide structure (referred to as wt-23-CONH2) is that the NH of the CONH2 at the C-terminus will hydrogen

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B

Figure 3. (A) ∆A/Ao (solid curves) and ∆V/Vo (dashed curve) versus time for vesicles exposed to 10 µM of wt-23-CONH2 containing the 23 N-terminal amino acids of the native HA protein and a carboxyamidated C-terminus (designed to “lock in” a C-terminal R-helix): pH 5.0 (squares) and pH 7.0 (circles). (B) wt-23-CONH2 causes the membrane to become permeable to both glucose and sucrose at pH 7.0: (left) immediately prior to permeation of sucrose, there is good contrast between the vesicle and surrounding solutions due to the refractive index differences of sucrose (inside vesicle) and glucose (outside vesicle); (right) permeation to sucrose leads to an immediate change in contrast between the vesicle and the surrounding solution.

Figure 4. (A) ∆A/Ao versus time for vesicles exposed to FITCwt-20 which contains the fluorescent probe fluorescein-5isothiocyanate appended to the N-terminal glycine: 10 µM, pH 5.0 (squares); 10 µM pH 7.0 (circles); 20 µM, pH 5.0 (diamonds, corresponding to the fluorescent image (Figure 4B)). A vesicle exposed to 10 µM FITC-wt-20, pH 5.0, at a flow rate of approximately 150 µm/s (triangles) expands in area (solid curve) and subsequently swells (dashed curve). (B) Images, taken seconds after transfer of vesicle exposed to 20 µM FITC-wt-20 to a solution free of peptide: (i) Hoffman optics image of vesicle (image by intensified CCD camera accounts for the graininess of the image); (ii) corresponding fluorescence image showing diffuse fluorescence associated with the membrane of the vesicle.

bond to the fourth residue from the C-terminus, thus minimizing conformational fraying. Thus, theoretically, the peptide behaves like it is part of the helical structure of a longer protein. For example, the native lytic peptide of bee venom, melittin, contains a carboxyamidated C-terminus, presumably to ensure the helical conformation of the C-terminus is maintained. The major feature of the area expansion behavior of wt-23-CONH2 (Figure 3A) is that ∆A/Ao,max at pH 7.0 (≈0.037) is significant and much closer to ∆A/Ao,max at pH 5.0 (≈0.067) for the same peptide concentration (10 µM). Additionally, the cycles of insertion, permeation, and lysis at both pH 7.0 and pH 5.0 for the amidated peptide occurred on similar time scales (both approximately 45 s). Another interesting feature of the amidated peptide is that its insertion into the membrane at pH 7.0 caused the vesicle to permeate to both glucose and sucrose shortly before rupture. This permeation was seen by the disappearance of optical contrast between the vesicle and the surrounding solution (Figure 3B). We repeated this experiment four times; each time, peptide insertion resulted in a loss of optical contrast. For all the other peptides studied, permeation did not result in a loss of optical contrast. Effect of FITC Labeling of the N-Terminus (Detection of Insertion and Insertion in Flow). In order to confirm that association of wt-20 peptide with the lipid bilayer was responsible for the observed area expansion of the vesicle membrane, we produced a wt-20 peptide in which the N-terminal glycine was fluorescently labeled with fluorescein-5-isothiocyanate. This peptide is referred

to as FITC-wt-20. Insertion of this peptide at pH 5.0 caused area expansion of the membrane bilayer (Figure 4A). Area expansion did not result in permeation of the vesicle (over 12 min). However, the incorporation of the peptide into the membrane at both 20 and 10 µM weakens the membrane significantly. The lysis tension, τlyse, of vesicles exposed to 10 and 20 µM of FITC-wt-20 was 3.6 and 2.2 dyn/cm, respectively. These values of τlyse are significantly lower than the mean lysis tension of 9.5 dyn/ cm for normal vesicles. There was no significant change in area compressibility modulus (K), 173 dyn/cm, for the vesicle exposed to 10 µM FITC-wt-20 in comparison to 195 ( 25 dyn/cm determined for normal vesicles. Since the vesicle exposed to 20 µM FITC-wt-20 lysed at a very low tension, we were not able to obtain enough data points to accurately determine K. We used fluorescence microscopy to visualize the fluorescence associated with bound FITC-wt-20. We exposed a vesicle to 20 µM of FITC-wt-20 for approximately 2 min. Subsequently, we transferred the vesicle to chamber section A which contained no FITC-wt-20. This allowed us to visualize a background free of fluorescent dye. Although, FITC-wt-20 desorbs quite rapidly from the surface of the vesicle (detected by a decrease in the projection length, data not shown), the fluorescence associated with the vesicle is bright enough to be detected for approximately 10 s after transfer (Figure 4B). It has been previously shown38 that the area expansion caused by rapidly inserting amphiphilic species can be enhanced if the membrane is exposed to a flow (using a

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Figure 5. ∆A/Ao versus time for vesicles exposed to 10 µM of ∆G1 mutant: pH 5.0 (squares); pH 7.0 (circles). Unlike wt-20, this peptide causes little area expansion of the membrane area at pH 5.0 and no pore formation (on the time scale of these experiments).

Figure 6. ∆A/Ao (solid curves) and ∆V/Vo (dashed curve) versus time for vesicles exposed to the bee venom peptide melittin at pH 5.5: 0.1 µM (diamonds); 0.5 µM (triangles). At 0.5 µM melittin, uptake results in rapid permeation of the vesicle to glucose.

flow pipet) containing amphiphilic species at a concentration above the critical micelle concentration. This flowenhanced insertion demonstrates that the overall rate of insertion of amphiphile aggregates is strongly influenced by transport of the aggregate to the membrane. This suggests that the overall rate of insertion can be enhanced if the amphiphile is delivered to the vesicle surface more quickly. We found that exposure of a vesicle to a flow (approximately 150 µm/s) of 10 µM FITC-wt-20 (Figure 4A) led to enhanced insertion (initial rate of area change ≈0.0028 s-1) in comparison to stagnant conditions (initial rate of area change ≈0.00063 s-1). Additionally, area expansion was greatly enhanced (∆A/Ao,max ≈0.085) in comparison to stagnant conditions (∆A/Ao,max ≈0.026). Under flow conditions, insertion of this peptide resulted in permeation of the vesicle to glucose in 51 s, which was similar to our results for wt-20 and wt-23-CONH2 under stagnant conditions. Fusion and Lytic Diminished HA Peptide. At a pH of 7.0, exposure of vesicles to a 10 µM solution of the ∆G1 “less active” peptide resulted in a change of ∆A/Ao indistinguishable from the apparent ∆A/Ao of the control experiment containing DMSO alone (Table 1, Figure 5). This result suggests that the change in projection length was simply due to an increase in osmotic strength that resulted in a vesicle internal volume decrease. At a pH of 5.0, ∆A/Ao,max for ∆G1 is significantly lower (≈0.014) than ∆A/Ao,max for wt-20 (≈0.086) at pH 5.0. In addition, τlyse of these vesicles was tested after approximately 12 min of exposure to ∆G1 and was found to be only slightly decreased (5.5 dyn/cm at pH 5.0 and 6.9 dyn/cm at pH 7.0) in comparison with the lysis tension of control vesicles (mean lysis tension 9.5 dyn/cm). Melittin. As a comparison to the fusion peptides of HA, we investigated the area expansion of the membranes of vesicles resulting from the uptake of bee venom melittin (Figure 6). We found melittin to permeate vesicles instantaneously at a concentration of 10 µM and pH 5.5. Therefore, data could not be obtained at the same concentrations as used for the HA peptides. Instead, at a concentration of 0.5 µM melittin, the cycle of insertion, permeation, and lysis occurred on a similar time scale (≈3 min) in comparison to wt-20 (≈5 min). At a concentration of 0.1 µM melittin, insertion of melittin did not result in permeation in the times scales of this experiment; however, the vesicle was significantly weakened (τlyse )

1.5 dyn/cm after 14 min of exposure) in comparison to normal vesicles (τlyse 9.5 ) dyn/cm). r-Helical Content of wt-20 and wt-23-CONH2 Influenza Fusion Peptides in SOPC Vesicles. Circular dichroic measurements of influenza peptides in SOPC vesicles provided insights into the structure of the peptides as the peptides associated with lipid bilayers. At pH 5.0, both wt-20 and wt-23-CONH2 showed a definitive dichroic minimum at 222 nm typical of R-helical folding conformations (Figure 7A,B). Measurements of the lipidpeptide dispersions at pH 7.0 also had a minimum at 222 nm with an amplitude similar to that at pH 5.0 (Figure 7A,B). Using the amount ellipticity at 222 nm and the formalism of Chen et al.,50 the helical conformations of wt-20 and wt-23-CONH2 in SOPC vesicles at both pH 5 and pH 7 were estimated at 46% and 38%, respectively. Below 222 nm there was a shift to more positive ellipticities suggestive of β-sheet structure. However, the lipidpeptide dispersion light scattering at these wavelengths precluded quantitative analysis of the complete spectral range for sheet and coil conformational contributions. Discussion In this report, we have shown that simple structural changes in the native sequence of the HA fusion peptide can alter how the peptide inserts into, expands, and disrupts the membrane. These effects were demonstrated with the use of micropipet aspiration of large vesicles. This technique, can be used unambiguously to determine area changes in lipid membrane bilayers.27,38-41 In our experiments, we can measure the area expansion of the membrane on time scales relevant to fusion (seconds to minutes). We show here that alterations in the native sequence of wt-20, by deletions of the N-terminal glycine or appendage of a fluorescent probe, resulted in a diminished ability to expand the area of the membrane through insertion and less propensity to form membrane pores. In contrast, the lytic peptide, melittin, and fusogenic peptide, wt-20, expand the membrane area and cause rapid membrane permeability. Carboxyamidation of the C-terminus designed to “lock in” helical structure in the C-terminus produced an enhanced ability to expand the membrane and form pores at pH 7.0. We show here and in previous work (Longo et al.27) that insertion of a synthesized wild-type fusion peptide of HA (wt-20) expands the membrane area rapidly and to a larger

Fusion of Membrane Bilayers

A

B

Figure 7. Circular dichroism spectra of influenza fusion peptides (A) wt-20 and (B) wt-23-CONH2 in SOPC unilamellar vesicles at pH 5 and pH 7. The lipid to peptide mole ratio was 50:1.

extent at a pH of 5.0 (endosomal pH) in comparison to pH 7.0. The pH activity observed here correlates with the well documented observation10,12,26 that the influenza fusion virus significantly fuses with or lyses lipid membranes at a pH of 5 (corresponding to endosomal pH), and not at 7. At pH 5.0, membrane expansion results in the permeation of the membrane to glucose (diameter ≈ 0.5 nm), but not sucrose, and rapid swelling of the vesicle. However, at a pH of 7.0, permeation occurs, but over a longer time scale. Interestingly, we find no significant difference in the R-helical content in the bilayer of this peptide in SOPC vesicles at a pH of 5.0 in comparison to pH 7.0 by use of CD spectroscopy. These results are in contrast with the work of Wharton et al.19 that investigated the pH dependent R-helical content of wt-20 and found an increase in R-helical content as the pH is lowered from 7 to 5. However, our CD work is in agreement with the work of Gray et al.16 that found that there were not significant differences in the R-helical content of the 23 amino acid version of the native fusion peptide at pH 5 and 7. In fact, they found that the helical content was slightly decreased in a pH 5 environment. Therefore, we postulate that the differences in surface activity at pH 5

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compared to pH 7.0 are the result of a charge difference on the peptide as the three negatively charged amino acids of wt-20 are neutralized. Since the pKa values of the negatively charged amino acid residues are approximately 4, one might expect that any charge-derived change in surface activity should occur at a pH of 4, and not near 5 as we previously observed27 and observe here. It is possible that the presence of bulky hydrophobic amino acids (wt-20 contains two phenylalanines and one tryptophan) in the peptide act to shift the pKa of the charged amino acids to higher values. Such pKa rises, through the placement of phenylalanine groups, have been thoroughly documented in the work of Urry et al.51,52,53 on pH sensitive hydrophobic elastin proteins. In Urry’s work, it has been possible to fine tune the pKa values of negatively charged groups by adjusting the positions of phenylalanines with respect to negatively charged residues in the beta spiral conformation of elastin.51 In the case of wt-20, the unique protein conformation and relative position of amino acids may exist to fix the pKa values of the negatively charged amino acids near 5 (the pH of the endosome). Our observation of some surface activity at pH 7 is consistent with the neutralization hypothesis. At a pH of 7, there will be a statistically small number of peptides in which the negatively charged amino acids are neutralized or partially neutralized and thus exhibit pH 5 type of insertion. Thus it is difficult to produce a pH-triggered fusion peptide that is completely inactive at pH 7 and completely activated at pH 5. Here, in an attempt to more closely mimic the native state of the fusion peptide (in comparison to wt-20) in vivo, we produced a peptide containing the 23 native amino acids of the fusion peptide and carboxyamidation of the C-terminus (wt-23-CONH2). In principle, carboxyamidation prevents the loss of the R-helical conformation of the C-terminus of small peptides. In contrast to our work with wt-20, we observed that there was little difference in percent of area change or permeation time when working at pH 5.0 or pH 7.0 when vesicles were exposed to 10 µM of wt-23-CONH2. Additionally, as with wt-20, our CD data show no significant difference in R-helical content at pH 5 or pH 7. Once again, a shift in the pH of neutralization may be responsible for the observed area expansion behavior. In the case of wt-23-CONH2, there has been a shift in the pH of neutralization of the entire peptide resulting from carboxyamidation of the C-terminus; the normally present negatively charged terminal carboxylic acid has been replaced by a carboxyamide group which is charge neutral at a pH of 5 and 7. Therefore, the pH for neutralization of the whole peptide will increase, possibly allowing significant insertion at higher pH as observed here. We have confirmed that area expansion of the membrane is associated with the binding of peptide to the membrane by imaging the fluorescence at the vesicle surface resulting from the insertion of FITC labeled wt20 peptide (FITC-wt-20). As has been demonstrated earlier (AGLF peptide5,16), we see that appending a molecule of low molecular weight to the N-terminus of the wild type peptide (i.e., FITC, mw ) 389) alters, but does not eliminate, the surface activity of this peptide. Gray et al.16 showed that the lowered surface activity corresponded with a slightly reduced R-helical content and (51) Urry, D. W.; Gowda, D. C.; Peng; S. Q.; Parker, T. M.; Jing, N.; Harris, R. D. Biopolymers 1994, 34, 889-896. (52) Urry, D. W.; Peng, S. Q.; Parker, T. M. Biopolymers 1992, 32, 373-379. (53) Urry, D. W.; Peng, S. Q.; Xu, J.; McPherson, D. T. J. Am. Chem. Soc. 1997, 119, 1161-1162.

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higher β-structure content in comparison to the wild-type peptide. Unlike wt-20, FITC-wt-20 does not cause pores to form in the membrane (on the time scale of this experiment), despite the fact that it is taken up significantly in the membrane (area expansion plateaus at approximately 4%). It should be mentioned here that it has been shown by Zheelev and Needham38 that true equilibrium has not been reached at the plateau. Instead, a slow “flip-flop” process transports lipid (and maybe peptide in this case) from the outer monolayer of the bilayer to the inner monolayer; thus new molecules are slowly and continually added to the outer monolayer. This results in a much slower rate of area expansion until final equilibrium is reached. The slow uptake is evident by the sloping nature of the plateaus observed here. By using flow, we could roughly double the area expansion of the membrane resulting from the uptake of FITC-wt-20 and enhance the area expansion rate. These results indicate that the area expansion rates that we obtain, without flow, are affected by diffusion of peptide to the surface of the vesicle. Therefore, by using flow, a better approximation of the kinetic rate of expansion can be measured.38 This phenomenon occurs because in flow, the concentration local to the membrane can be enhanced in comparison to the stagnant conditions, since peptide is delivered more quickly as it is depleted. The great increase in maximum area expansion under flow observed here probably cannot be explained by enhanced mass transfer rates and may be related to unexplored structural changes that occur to the peptide or vesicle under flow that enhance the equilibrium membrane concentration. It is possible that flow may change the aggregation state of the peptide and possibly the conformational state. For example, it has been shown that flow can dramatically change the aggregation state of wormlike aggregates.54,55 It is not known if FITC-wt-20 forms wormlike structures. However, Wharton et al.19 observed that wt-23 forms thin wormlike aggregates greater than 1 µm in length. This flow experiment demonstrates that it is important to scrutinize the history of the peptide, which may affect insertional behavior. We show here as a control, and in our previous work,27 that a simple change in the native fusion peptide sequence (deletion of the N-terminal glycine) can almost completely eliminate the ability of the peptide to expand the membrane area. In comparison to the ∆G1 peptide at pH 5.0, FITC-wt-20, wt-23-CONH2, and wt-20 expand the membrane area to a larger extent at the same concentration and pH (approximately 4%, 6%, and 8% respectively). Wharton et al.19 and Gray et al.16 have shown that the ∆G1 peptide has a much lowered level of R-helix content in comparison to the wild-type peptide at pH 5.0. The conformational change could affect the propensity of the peptide to insert into the bilayer. Additionally, the pKa values of the amino acids could be less affected by the bulky hydrophobic residues in this conformation in comparison to the more helical state of wt-20. In an attempt to determine if area expansion and vesicle permeation were typical of other lytic peptides, we have exposed vesicles to the well-studied lytic peptide, melittin.42 We found that melittin is able to form pores in the membrane at very low concentrations (0.5 µM). This result is in agreement with Rex56 who determined (using similarly low concentrations of melittin and phosphati(54) Botenhagen, P.; Hu, Y. T.; Matthys, E. F.; Pine, D. J. Europhys. Lett. 1997, 38, 389-394. (55) Wheeler, E. K.; Izu, P.; Fuller, G. G. Rheol. Acta 1996, 35, 139149. (56) Rex, S. Biophys. Chem. 1996, 58, 75-85.

Longo et al.

dylcholine large unilamellar vesicles) that approximately 0.5 nm radii pores were formed containing about six melittin molecules. As a comparison, wt-20 loses its ability to form pores at approximately 1 µM on the time scales of these experiments (Longo et al.27). At pH 5.5, 0.5 µM, melittin forms pores in approximately the same exposure time as 10 µM of wt-20 at pH 5.0 (approximately 4 min and 2 min, respectively). Similar to wt-20, the uptake of melittin in the membrane results in measurable area expansion of the membrane (approximately 1%), even at 0.1 µM. It appears that melittin, like wt-20, has a propensity to form membrane pores, even when the amount of area expansion is small (less than 1%). A membrane bilayer can only be stretched by approximately 5% before mechanical failure occurs.40,41 It has been demonstrated previously that the rate of insertion of an amphiphile to the outer monolayer of the bilayer can be much faster than flip-flop processes from the outer monolayer to the inner monolayer.38 Therefore, a membrane should mechanically fail when the outer monolayer area has been expanded (by insertion) more than approximately 5%. Needham and Zhelev38 showed that, rather than lysing, the membrane fails by the formation of porous defects when the outer monolayer is stretched relative to the inner monolayer. We show here that the peptides wt-20 and melittin cause the membrane to become porous even when their uptake results in area expansion in the range of 1% or less. These pores do not widen significantly during the experiment, as is evidenced by a lack of vesicle permeation to sucrose, the larger sugar species. This work is in concert with that of Zimmerberg et al.,33 which indicates that fusion of cells by influenza virus may proceed through a number of small pores. For wt-20, membrane perturbation may result from an oblique angle of insertions57 or through aggregation of peptide in the membrane bilayer. Both possibilities may stress the membrane more effectively than simple area expansion (or localized expansion) of the outer monolayer with respect to the inner monolayer. Most likely, in the case of melittin, pores are formed by the aggregation of melittin monomers58 allowing the membrane to become porous at area expansions around 1% seen here. The observation that slight chemical changes to the native fusion peptide can result in a lack of porous defect formation even when area expansion is relatively large (as much as 4%) may indicate that pore formation by peptides is a useful indicator of the mode of activity of complete fusion molecules. However, how pore formation by these small fusion peptides relates to the mechanism of activity of the full HA fusion protein remains to be determined. Our work shown here demonstrates that HA fusion peptides containing simple changes in the chemical structure of the N- and C-terminus interact with lipid bilayers and have various consequences regarding insertion and permeation. Attachment of a small fluorescent molecule (FITC) to the N-terminus yielded a peptide with rapid area expansion of the bilayer at pH 5.0, a property of the wild-type peptide. Yet, unlike the wildtype HA peptide, the fluorescent peptide did not cause the membrane to permeate on the time scales of these experiments (approximately 12 min). Deletion of the N-terminal glycine eliminates most of the insertion of this peptide into a lipid bilayer and eliminates its ability to form porous defects in the lipid membrane. These results correlate with the work of Steinhauer et al.5 which (57) Brasseur, R.; Vandenbranden, M.; Cornet, B.; Burny, B.; Ruysschaert, J. M. Biochim. Biophys. Acta 1990, 1029, 267-273. (58) Fattal, E.; Nir, S.; Parente, R. A.; Szoka, F. C., Jr. Biochemistry 1994, 33, 6721-6731.

Fusion of Membrane Bilayers

showed that small chemical changes to the N-terminus of the fusion peptide of HA can lead to decreased fusogenicity. Moreover, this work extends the work of Gray et al.16 which showed that these same changes were associated with a decreased R-helical content and enhanced β-content of the peptides and correlated with an increase in interaction with lipid head groups. We have demonstrated, using CD, that no significant change in R-helical content of wildtype peptide (wt-20) accompanies the change in insertion that occurs between pH 5 and 7. These results suggest that further investigations are required to ascertain the cause of the dramatic difference in insertion in a lipid bilayer observed. Charge neutralization, coupled with an increase in the pKa of the negative amino acids through interactions with hydrophobic amino acids, and not a change in conformation, is a more likely explanation for the observed results. Activation of the wild-type peptide at pH 7.0 through carboxyamidation (wt-23-CONH2) observed here may also be caused by a shift in the pH of neutralization of the peptide. Additionally, our results shown here suggest that studies of the exact mechanism of lipid membrane pore formation by fusion peptides and characterization of these pores on short time scales relevant to fusion may be critical to discerning the complete

Langmuir, Vol. 14, No. 9, 1998 2395

mechanism of lipid disruption in viral fusion. Finally, it would be useful for further investigations to be performed which probe the possible presence of pKa changes due to positioning of hydrophobic and negatively charged residues. Acknowledgment. We offer special thanks to the members of Dr. Richard Waugh’s laboratory (University of Rochester, NY) for their helpful guidance and expert advice regarding the micropipet aspiration technique. We thank Professor Charles Knobler for use of the dynamic light scattering device (Department of Chemistry, UCLA) and Professor James Bowie for access to the AVIV 62DS spectropolarimeter (Department of Chemistry and Biochemistry, UCLA). Acknowledgment is made to the National Institutes of Health HL18208 for support of this research. M. L. Longo is grateful for the support of the National Institutes of Health NRSA Postdoctoral Fellowship GM 16506-02. The ABI 431A peptide synthesizer was acquired by an NIH small equipment grant GM 50483 to A. Waring and L. Gordon. Alan Waring and Larry Gordon were supported by NIH Grant GM08140. LA970932P