A Novel Mechanism of Pore Formation

MEMBRANE PENETRATION BY THE N-TERMINAL AMPHIPATHIC REGION OF EQUINATOXIN*

Petra Malovrh {ddagger} § , Gabriella Viero  ||, Mauro Dalla Serra ||, Zdravko Podlesek {ddagger}, Jeremy H. Lakey **, Peter Macek {ddagger}, Gianfranco Menestrina || and Gregor Anderluh {ddagger} {ddagger}{ddagger}

From the {ddagger}Department of Biology, Biotechnical Faculty, University of Ljubljana, Vecna pot 111, 1000 Ljubljana, Slovenia, the ||CNR-ITC, Istituto di Biofisica, Sezione di Trento, Via Sommarive 18, 38050 Povo (TN), Italy, and the **School of Cell and Molecular Biosciences, University of Newcastle upon Tyne, Framlington Place, Newcastle upon Tyne, NE2 4HH, United Kingdom

Received for publication, January 20, 2003 , and in revised form, March 19, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Equinatoxin II is a representative of actinoporins, eukaryotic pore-forming toxins from sea anemones. It creates pores in natural and artificial lipid membranes by an association of three or four monomers. Cysteine-scanning mutagenesis was used to study the structure of the N terminus, which is proposed to be crucial in transmembrane pore formation. We provide data for two steps of pore formation: a lipid-bound monomeric intermediate state and a final oligomeric pore. Results show that residues 10–28 are organized as an {alpha}-helix in both steps. In the first step, the whole region is transferred to a lipid-water interface, laying flat on the membrane. In the pore-forming state, the hydrophilic side of the amphipathic helix lines the pore lumen. The pore has a restriction around Asp-10, according to the permeabilization ratio of ions flowing through pores formed by chemically modified mutants. A general model was introduced to derive the tilt angle of the helix from the ion current data. This study reveals that actinoporins use a unique single helix insertion mechanism for pore formation.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Pore-forming toxins (PFT)1 are considered as a link between soluble and membrane proteins (1). These proteins, produced in a soluble form, act upon target cell membranes by forming transmembrane channels. During this toxic action they adapt their structure to expose the hydrophobic parts needed for membrane association and pore formation. PFT can be divided according to the transmembrane structural elements of the final inserted state. Some bacterial PFT form stable oligomeric {beta}-barrels (2). Examples include {alpha}-toxin from Staphylococcus aureus (3), a family of cholesterol-dependent cytolysins from Gram-positive bacteria (4), anthrax protective antigen (5), and others. PFT pores lined by {alpha}-helices (similar to colicins) (6) are fewer and less well understood, largely because of the inherent instability of the resulting pores.

One interesting family of PFT is that of the actinoporins of sea anemones (7). They are closely related cysteine-less proteins that form pores in natural and artificial lipid membranes. Their peculiar three-dimensional structure and biochemical properties reveal them to form an entirely novel class of PFT (8, 9). They are one-domain proteins; smaller than bacterial PFT, they lyse erythrocytes at picomolar concentrations and exhibit sphingomyelin dependence. The most studied representative is equinatoxin II (EqtII), an actinoporin from Actinia equina. EqtII is composed of a tightly folded {beta}-sandwich flanked on two sides by {alpha}-helices (Fig. 1A) (8, 9). The first 30 N-terminal residues, encompassing the N-terminal {alpha}-helix (residues 16–26 from the crystal and NMR structures), is the only part that can be dislocated from the body of the molecule without disrupting the {beta}-sandwich (Fig. 1B). Pore formation by EqtII is a multistep process (Fig. 1C). It was shown recently that the toxin binds to the lipid bilayer with the aromatic amino acid cluster located on a broad loop at the bottom of the molecule and on the C-terminal {alpha}-helix (10, 11). In the next step an N-terminal segment translocates to the lipid-water interface (10, 12), and, finally, a transmembrane pore is formed by helices from three or four monomers as proposed by Belmonte et al. (13). The final pore is not a rigid structure, like the stable {beta}-barrels of bacterial PFT that can be studied by x-ray crystallography or electron microscopy (3, 5). Actinoporin pores are not resistant to SDS and have not yet been directly visualized. The number of monomers in the pore was deduced by cross-linking and kinetic experiments (13, 14).



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FIG. 1.
The three-dimensional model of EqtII. A, three-dimensional model of EqtII. The N-terminal amphipathic region 10–28 encompassing helix A is shown in green. B, helical wheel conservation analysis of 10–28 region. The size of the circle corresponds to the degree of conservation of that particular amino acid in all known actinoporin sequences. The residues are colored according to physical properties (black, hydrophobic; yellow, polar; red, negatively charged; blue, positively charged). C, the current hypothesis of EqtII pore formation. The pore formation involves at least four different conformational states of the toxin: a soluble form, a membrane-bound form attached to the membrane with the aromatic cluster (M1-state), a membrane-bound form with a dislocated helix (M2-state), and oligomeric form (P-state), when helix is a part of the conductive pathway. The lipid-water interface is shaded gray.

 

According to helical wheel analysis, the whole region of residues 10–28, including the N-terminal {alpha}-helix, is amphipathic and could be involved in forming the walls of the functional pore (Fig. 1B) (15). This region is highly conserved in all known actinoporins. In particular, the hydrophobic face is almost completely conserved, while on the polar face the most conserved are negatively charged residues (i.e. Asp-10, Asp-17, and Asp-24), which could account for the cation selectivity of actinoporins. We decided to study the topology of this important region in its membrane environment by employing cysteine-scanning mutagenesis. This approach proved to be useful in obtaining detailed structural information of membrane regions of some PFT (1618). A desired amino acid is changed to a cysteine by site-directed mutagenesis in a protein that normally does not possess any surface-exposed, and therefore reactive, cysteines. Extrinsic labels can then be covalently attached to this unique thiol group; thus giving the site-specific information about their immediate environment (19). We have already performed low resolution cysteine-scanning mutagenesis, where regions of EqtII that interact with the lipid membranes were determined (12). In the current work we performed a detailed scanning of the region from Asp-10 to Asn-28. Mutants were modified with thiol reactive reagents and assayed by fluorescence spectroscopy and electrophysiological methods to determine the structural arrangement of this N-terminal region at each stage of pore formation. Results indicate that the whole region is transferred to the lipid-water interface during the initial binding and that the same region forms the walls of the final pore in an {alpha}-helical arrangement.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cloning, Expression, and Isolation of Cysteine Mutants—Single cysteine mutants were produced by substituting the corresponding wild-type (WT) amino acid residue with cysteine as described (12). Mutants were expressed from a T7-based expression vector in Escherichia coli BLR(DE3) strain and recombinant proteins purified from bacterial supernatants as described (20). S13C, D17C, and E24C were prepared as fusion proteins using His-tagged third domain of TolA (TolAIII, a bacterial periplasmic protein) as a fusion partner. They were expressed in E. coli using the pTolT plasmid and purified from bacterial supernatants by Ni-chelate chromatography (21). All mutants were purified to homogeneity as observed on SDS-PAGE gels.

Characterization of Mutants—Mutants were tested for hemolytic activity using a microplate reader (MRX; Dynex Technologies, Dekendorf, Germany). The percentage of hemolysis was determined as described (10). In short, bovine red blood cells (100 µl, OD630 = 0.5) were added to 2-fold serially diluted mutants in 100 µl of 0.13 M NaCl, 20 mM Tris-HCl, pH 7.4, and hemolysis was monitored by measuring absorbance at 630 nm for 20 min at room temperature. The ability of mutants to bind to bovine red blood cells and small unilamellar vesicles (SUV), with or without 1-palmitoyl-2-stearoyl-(7-doxyl)-sn-glycero-3-phosphocholine (7-NO-PC), at a lipid/toxin (L/T) ratio of 1:1000 was determined from the residual hemolytic activity as described (10). In this test, unbound protein is detected by hemolysis assay by adding fresh erythrocytes at the end of the incubation. Hemolytic activity is observed only, when unbound protein is present in the sample (10). The amount of unbound toxin was determined from the calibration curves obtained using purified mutants. For the bovine erythrocytes assay toxins, at a final 76 nM concentration, were preincubated with erythrocytes for 10 min. Thereafter, a fresh suspension of erythrocytes was added, and hemolysis measured as described above. To assay binding to SUV, 100 µl of sample after fluorescence assays (see below) was mixed with 100 µl of erythrocytes, and activity was measured as described above.

IANBD Labeling—Typically, 0.5–1 mg of protein was incubated in 50 mM Tris-HCl, pH 7.1, with dithiothreitol (DTT) (toxin/DTT molar ratio 1:1) for 30 min at room temperature. Thereafter, N-((2-(iodoacetoxy)ethyl)-N-methyl)amino-7-nitrobenz-2-oxa-1,3-diazole (IANBD) (Molecular Probes, Eugene, OR), in 5x molar excess over DTT and toxin, was added. The mixture was incubated on a magnetic stirrer at 4 °C overnight. Labeled proteins were separated from unlabeled by using a hydrophobic interaction chromatography (HIC) column (Brownlee, Aquapore) as described. The extent of protein modification was calculated from the relative areas of the two peaks that appeared at 280 nm. The first one corresponded to the unmodified and the second one to the labeled mutant. S13C, D17C, and E24C were "on-column" labeled as TolAIII fusion proteins. 0.5 ml of Ni-NTA gel slurry (Qiagen, Crawley) was placed in the column and washed with 5 ml of 20 mM {beta}-mercaptoethanol, 20 mM NaH2PO4, 300 mM NaCl, pH 7.4. For each mutant a few milligrams of protein were incubated in the above buffer for 1 h at 4 °C. Afterwards, they were applied to the column and washed with 10 ml of the same buffer without {beta}-mercaptoethanol. Bound mutant protein was washed with 4 ml of 5 mM IANBD and incubated in the dark at room temperature for 4 h. After labeling, the column was washed with 15 ml of enterokinase buffer (20 mM Tris-HCl, 50 mM NaCl, 2 mM CaCl2, pH 7.4). Enterokinase (Novagen) was then added (30 units for 1 mg of protein) and incubated at 30 °C for 24 h. Cleaved labeled mutants were washed from the column by 20 mM NaH2PO4, 200 mM NaCl, pH 7.4, and purified using FPLC and a MonoS ion-exchange chromatography column (Amersham Biosciences).

Chemical Modification Using Methanethiosulfonate Derivatives (MTS)—Mutants were chemically modified with MTS reagents to introduce, at the thiol group, either a positive or a negative charge with (2-aminoethyl) MTS hydrobromide (MTSEA) and sodium (2-sulfonatoethyl) MTS (MTSES), respectively (both from Biotium, Inc. Fremont, CA). Mutants were preincubated overnight in a 20 molar excess of DTT. MTS reagents, freshly dissolved in H2O, were then added at 1000 molar excess to 10–50 µM mutants. After a 1-h incubation at room temperature, after which all the excess reagent had hydrolyzed, the modified samples were used for PLM experiments.

Preparation of Lipid Vesicles—All lipids were from Avanti Polar Lipids (Alabaster, AL). Small unilamellar vesicles (SUV) were prepared from brain sphingomyelin (SM) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) at a 1:1 molar ratio. Liposomes used in lipophilic quenching experiments were prepared by using SM and DPPC, except that 20 mol% of DPPC were replaced by 7-NO-PC. Chloroform was removed by a rotary evaporator. Vesicle buffer (140 mM NaCl, 20 mM Tris-HCl, 1 mM EDTA, pH 8.5) was added to the lipid film, and the suspension was vigorously vortex-mixed in the presence of glass beads. The resulting multilamellar vesicles were converted to SUV by sonication (MSE 150W ultrasonic disintegrator) of suspension at room temperature. The SUV suspension was centrifuged at 12,000 x g for 15 min to remove titanium particles released from the probe. Vesicles were stored at 4 °C immediately after the preparation and used the next day.

Fluorescence Spectroscopy—All fluorescence measurements were performed on a Jasco FP-750 spectrofluorimeter (Jasco Corporation, Japan). The sample compartment was equipped with a Peltier thermostatted single-cell holder. All experiments were done at 25 °C at a protein concentration of 250 nM. The excitation and emission slits were set to 5 nm. Tryptophan emission spectra were measured in 50 mM Tris-HCl, pH 8.0. The excitation wavelength was 295 nm, and spectra were recorded from 310–400 nm. NBD-labeled mutants in 140 mM NaCl, 20 mM Tris-HCl, 1 mM EDTA, pH 8.5 were excited at 470 nm, and the emission was scanned from 500–600 nm. Subsequently, the appropriate amount of SUV, with or without 7-NO-PC, at the final L/T ratio of 1000 was added, and the NBD emission was scanned again. The spectra were corrected for the dilution factor, and the background was subtracted using the appropriate blank with SUV. No further correction for wavelength dependence of the photomultiplier tube was done. The {lambda}max was determined from the corrected spectra by Spectra Analysis software (part of Spectra Manager 1.53.00; Jasco Corporation).

Quenching with 7-NO-PC and Iodide—The quenching efficiency, E, for the quenching of NBD emission by 7-NO-PC was calculated as follows in Equation 1,

(Eq. 1)
where FQUENCH and Fsuv represent the normalized fluorescence of IANBD at 530 nm in the presence of SUV with or without 7-NO-PC, respectively.

For quenching with iodide ions, a 2.5 M stock solution of KI in 1 mM Na2S2O3 was freshly prepared prior to each experiment. The aliquots from the stock solution were added to 250 nM final NBD-labeled protein with or without the presence of SUV (L/T ratio 1000). NBD emission spectra were measured as described above. They were corrected for the emission of buffer or buffer with vesicles and for the dilution factor. The data were analyzed according to the Stern-Volmer equation shown in Equation 2,

(Eq. 2)
where F0 is the fluorescence of the probe in the absence of the quencher, F is the observed fluorescence, [Q] is the concentration of iodide ions, and KSV is the collisional quenching constant.

Electrical Recordings of Ion Channel Activity—Electrical properties of unmodified and MTS-modified mutants were studied on planar lipid membrane (PLM) made of 1,2 diphytanoyl-sn-glycerophosphocholine (DPhPC) and 20% (w/w) of SM, both from Avanti Polar Lipids. Mutants were added on one side (cis) to stable preformed bilayers. All experiments were started in symmetrical solutions (10 mM Tris-HCl, 100 mM KCl, pH 8.0). For the selectivity determination a 10-fold KCl gradient was formed, whereby the higher concentration was on the trans side, which was held at virtual ground. Macroscopic currents were recorded by a patch clamp amplifier (Axopatch 200, Axon Instruments). A PC equipped with a DigiData 1200 A/D converter (Axon Instruments) was used for data acquisitions. The current traces were filtered at 0.1 kHz and acquired by the computer using Axoscope 8 software (Axon Instruments). Measurements were performed at room temperature.

Derivation of Helix Orientation Inside the Pore—The selectivity of the pore in MTS modification experiments was analyzed in terms of a model that relates it to the potential generated by the introduced charges. In fact these charges generate a local field at the entrance of the pore that attracts and concentrates counter ions, and therefore the partial conductance Gi of the ion species i through the pore, can be written as in Equation 3 (22),

(Eq. 3)
where ui, zi, and ci are the mobility, valence, and local concentration of the ion i, e is the elementary charge, and K a geometrical constant (in the simplest form K = {pi}r2/l where r and l are radius and length of the pore).

The local concentration ci derives from the bulk concentration cio through Equation 4,

(Eq. 4)
where {Psi}'pore is the reduced potential at the center of the pore entrance (i.e. {Psi}'pore/(kT/e), with k, Boltzmann constant, T, absolute temperature). From this, in the case of 1:1 salt like KCl we used, one can derive the selectivity index in Equation 5,

(Eq. 5)
where P+, P, G+, G, u+, and u are permeability, partial conductance, and mobility of cation and anion, respectively.

Each mutant will have a certain {Psi}'pore, that will be different depending on the residue that was substituted by the cysteine. Upon modification with MTSEA, however, there will be an additional potential generated by the new positive charge introduced at the cysteine, so that we can write Equation 6.

(Eq. 6)

When the same mutant is modified with MTSES, instead, the additional potential will be generated by a negative charge as in Equation 7.

(Eq. 7)

By combining Equations 6 and 7 we obtain Equation 8,

(Eq. 8)
which is a quantity independent from the variable entrance potential of the unmodified mutant {Psi}'pore.

For the potential generated by a fixed charge in solution, at a distance R, we can take the Debye-Hückel expression (23) in Equation 9,

(Eq. 9)
where {kappa} is the Debye-Hückel coefficient, Q is the fixed charge in elementary units, {epsilon} and {epsilon}o the dielectric constants of water (relative) and vacuum (absolute). Because the two modifications carry one, opposite, elementary charge we can write Equation 10.

(Eq. 10)

Finally, if we combine equations 8 and 10, and we take into account that the pore is formed by a tetrameric aggregate of {alpha}-helices, which we assume to be symmetrically distributed in their contribution to the field, we can write Equation 11.

(Eq. 11)

To calculate the distance R of the introduced charge from z, the central axis of the pore, we have to take into account its location around the {alpha}-helix and its position along the {alpha}-helix. In fact, if the helix forms an angle {theta} with the normal to the membrane (assumed to be parallel to the pore axis), the pore would have a funnel shape, as shown in Fig. 2. If Asp-10 is the first residue of the {alpha}-helix, located at the narrowest point of the pore, then R is given by Equation 12,

(Eq. 12)
where d = n{delta} (n is the position of the successive residues along the {alpha}-helix with respect to Asp-10 and {delta} is the pitch of the {alpha}-helix, i.e. 0.15 nm per residue), a is the radius of the {alpha}-helix (taken as 0.5 nm), {alpha} is the angular position of the n residue in the plane perpendicular to the helix axis ({alpha} = 2{pi}n/3.6), and r is the radius of the pore at the narrowest position, where Asp-10 is located (n = 0).



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FIG. 2.
Schematic geometry of one of four {alpha}-helices forming the pore. The membrane-inserted {alpha}-helix forms an angle {theta} with the pore axis (parallel to the membrane normal). Residue Asp-10, indicated by a black dot, is placed at the narrowest point of the pore, where the radius is r. The nth residue is located at a distance d along the {alpha}-helix axis and an angle {alpha} in the plane perpendicular to it (see the enlargement). R is its distance from the pore axis and a is the average radius of the {alpha}-helix. The lipid-water interface is presented as gray.

 

If we substitute Equation 12 into Equation 11, and we give to the constants their corresponding values, we obtain an expression for (P+/P)ES/(P+/P)EA versus n with only two free parameters, r and {theta} as shown in Equations 13 and 14.

(Eq. 13)

(Eq. 14)

This expression was used to fit the experimental results in Fig. 8.



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FIG. 8.
Effects of MTS modifications on the selectivity of channels formed by mutants. The ratio P+/P was calculated from the reversal voltage (Vrev) after modification with either MTSES or MTSEA. The ratio of the values obtained in the two conditions is reported. The solid line is the prediction of Equations 13 and 14. Best fit parameters obtained were r = 1.3 nm and {theta} = 0.36 rad or 21°.

 


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Experimental Design—Pore formation by actinoporins is a multistep process that leads from the monomeric state in solution to a membrane-inserted aggregate composed of three or four monomers (Fig. 1C). This process includes at least three conformationally different membrane-bound states: an initial complex stabilized by the aromatic cluster (M1-state), a monomer-membrane complex with the N terminus inserted into the lipid-water interface (M2-state) (11), and a final functional oligomeric complex (P-state). Calcein release experiments have indicated that the number of pores formed in a single vesicle depends strongly on the L/T lipid to toxin ratio (13, 14). However, even at very high toxin densities (low L/T) the proportion of pores formed was still very low in comparison to the amount of bound monomeric toxin (14). It is, therefore, not reasonable to study the topology of the N-terminal helix in the oligomeric pore by fluorescence techniques, as there will be always a contribution from the monomeric forms also present. For this reason, we decided to study by fluorescence only the topology of the N-terminal region 10–28 of the monomeric membrane-bound toxin. We used high L/T ratios, in order to obtain homogeneous population of the M2-state. At the conditions chosen for all subsequent experiments (SUV DPPC/SM 1:1; 250 nM EqtII; L/T molar ratio 1000) the release of calcein was only 0.4% ± 0.2 (n = 2 ± S.D.), indicating that the amount of functional pores (P-state) was negligible. There was no residual hemolytic activity in the sample after the calcein release experiment; therefore, EqtII was fully bound to the membranes (10). According to the reported equilibrium constants for lipid-bound monomeric states (11) and oligomeric P-state (14) we can reasonably assume prevalence of the M2-state (about 92% of the total EqtII) over all other toxin forms.

PLM experiments were employed to study the topology of the N-terminal region in the final oligomeric pore conformation, since this technique is only sensitive to ion flow through open pores. Any change in ion selectivity observed through the channels should therefore be the result of the side chain alterations introduced by cysteine mutagenesis and by subsequent chemical modifications of the thiol groups.

Production and IANBD Labeling of Cysteine Mutants—Cysteine mutants were prepared by using an E. coli expression system. Intrinsic tryptophan fluorescence was measured in order to check for any conformational changes induced by the mutation. The value of the tryptophan emission maxima ({lambda}max) for all mutants was 339 ± 1 nm, similar to that of the WT; hence we assumed that the conformation of mutants remained unaltered. Mutants were further assayed for hemolytic activity and binding to bovine red blood cells. The majority of the mutants retained >50% of the WT activity. A12C, L26C, G27C, and N28C were exceptions having only 16, 15, 17, and 0.7% of the WT activity, respectively. The reason for the decreased activity of N28C may be a lower conformational stability. As this mutant could be produced only in extremely small amounts and also precipitated over time it was not used in fluorescence studies. The lower activity of A12C and G27C was not due to low binding to the bovine erythrocytes membranes, as they bound fully, but obviously later steps in pore formation were inhibited. Of all mutants only L26C exhibited reduced binding, which resulted in reduced hemolytic activity. However, this mutant bound fully to SUV under the conditions employed for fluorescence studies.

Mutants were labeled with a thiol-specific probe IANBD. G11C, S15C, and F16C were not used in fluorescence experiments due to insufficient expression levels in bacteria. No reaction between IANBD and WT was detected. The extent of labeling was mutant-dependent and ranged from 10% for L26C to 100% for I18C. L19C could not be labeled. The side chain of Leu-19 is oriented from the hydrophobic face of the helix toward the {beta}-sandwich ensuring that it is completely buried and thus inaccessible to IANBD. Most of the labeled mutants were slightly less hemolytically active; however, the majority retained more than 50% of the hemolytic activity prior to labeling. The activity of D17C, E24C, and L26C increased by ~2-fold. All the mutants bound fully to SUV at a L/T ratio of 1000 as no free toxin was detected by residual hemolytic activity measured after the fluorescence measurement.

Changes in NBD Environment Detected by Emission Fluorescence—Emission spectra were obtained for each monomeric NBD-labeled mutant in solution and when bound to SUV at a L/T ratio of 1000 (SM/DPPC 1:1) (Fig. 3). In solution {lambda}max ranged from 535 (L23C) to 543 nm (E24C). Most of the mutants exhibited {lambda}max of 539–540 nm, indicating a relatively solvent-exposed NBD environment. The data correlate with the relative solvent accessibility of individual amino acid residue calculated from the three-dimensional structure (Fig. 3C). Generally, those residues that are less exposed in structure showed blue-shifted {lambda}max, i.e. the least exposed leucines at positions 14, 23, and 26 showed the shortest {lambda}max. The {lambda}max of mutants in positions 20 to 26 followed the {alpha}-helical arrangement (Fig. 3B) in agreement with the recently determined three-dimensional structure. There is less support for an {alpha}-helical model for the region 10–16, which forms an extended region in the three-dimensional model (8, 9).



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FIG. 3.
Fluorescence emission maxima of NBD labeled mutants free in solution and bound to SUV. A, the fluorescence was measured with labeled mutants (250 nM) in 140 mM NaCl, 20 mM Tris-HCl, 1 mM EDTA, pH 8.5 at 25 °C. The excitation wavelength was 470 nm. Spectra are shown for mutants in solution (solid line), in the presence of SM/DPPC 1:1 SUV (dashed line), and SM/DPPC/7-NO-PC SUV 5/4/1 (dotted line) at a L/T ratio of 1000. B, emission maxima were determined from NBD emission spectra for mutants in solution (white squares) and upon addition of SM/DPPC 1:1 SUV (black squares). Each value is the average of 2–6 independent experiments ± S.D. A dashed line represents the {alpha}-helical pattern. C, the correlation between {lambda}max of NBD-labeled mutants in solution and the surface exposure of residue at that position. The surface exposure was determined by the WhatIf program (44).

 

Upon interaction with SUV all mutants exhibited an increase and blue shift in NBD fluorescence indicating the change in the polarity of the NBD environment (Fig. 3A). The fluorescence of free IANBD in the presence of SUV changed only slightly (from 542 to 540 nm) confirming that the observed changes are due to the lipid-bound protein and were not artifacts caused by the association of the probe with vesicles. The comparison of initial and final fluorescence can be misleading as labels on different mutants exhibited different initial intensities in the soluble form. The {lambda}max was used instead as these values reflect more reliably the polarity of the environment (24). The {lambda}max varied for individual mutants upon SUV binding, as represented by the emission spectra of the two extremes; D17C from the hydrophilic and A12C from the hydrophobic face of the helix (Fig. 3A). For the whole region studied, the maxima were shifted to lower wavelengths by ~10 nm. The values now range from 525 (G27C) to 533 nm (D17C). Most of the mutants exhibited values between 528 and 530 nm. Crucially we observed a clear {alpha}-helical pattern for residues 12–26 (Fig. 3B) with maxima at residues 14, 17, 20, 21, and 24.

Determination of the Helix Orientation by Using Hydrophilic and Hydrophobic Collisional Quenchers—To gain information on the orientation of the N-terminal region in the lipid phase we employed an aqueous (iodide) and a lipophilic (7-NO-PC) collisional quencher. When buried in the bilayer, aqueous quenchers should not quench NBD. However, it should be accessible to the nitroxide group of 7-NO-PC, whose position is restricted to the interior of the membrane.

Iodide quenching is shown for NBD-labeled D17C either free in solution or bound to vesicles on Fig. 4A. The Stern-Volmer collisional quenching constant, KSV, was calculated from the slope of the best fitting line. A linear dependence of fluorescence intensity on quencher concentration was obtained for all mutants in soluble and membrane-bound form, indicating that quenching is collisional rather than static. Data for all mutants are summarized in Fig. 4B. The KSV for most of the mutants in solution ranged between 5 and 7 M–1 (Fig. 4B), indicating NBD exposure to the aqueous environment. KSV values correlated well with the hydrophilic/hydrophobic orientation of the residues in the helix, being higher for the mutants on the hydrophilic side, like D17C and E24C, which have KSV of 7.7 M–1 and 7.4 M–1, respectively, and lower for the mutants from the hydrophobic side. From the known structure, L23C has the least exposed side chain of all the mutants studied but gave the highest KSV value (9 M–1). In this mutant NBD appears to be next to the positive charge from Lys-32 that is oriented toward the side chain of Leu-23. This positive charge might specifically enhance the quenching of NBD by increasing the local concentration of I.



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FIG. 4.
Quenching of NBD-labeled mutants with a water soluble quencher. A, quenching of NBD-labeled D17C with iodide. Iodide was added in aliquots to 250 nM mutant in 140 mM NaCl, 20 mM Tris-HCl, 1 mM EDTA, pH 8.5 at 25 °C in solution (open squares) or when bound to SUV at a L/T ratio of 1000 (solid squares). Other conditions were as in the legend to Fig. 3. B, KSV values for all mutants in soluble form (white squares) or when bound to SUV (black squares). KSV values were calculated from the slopes of curves in A after linear fitting. The average of 2–5 independent experiments ± S.D. is shown. C, the change in the environment polarity of NBD is shown as a ratio between KSV after (KSV lip) and prior (KSV sol) to addition of SUV. A dashed line represents the {alpha}-helical pattern.

 

When NBD-labeled mutants were bound to SUV, the KSV for iodide quenching shifted to lower values, largely ranging between 1.3 and 2.5 M–1 (Fig. 4B). Thus, for all mutants, NBD moved to a more hydrophobic environment becoming less accessible for iodide. After binding, and the expected conformational change away from the Lys-32 positive charge, the quenching of L23C was no longer exceptional, and it behaved as one of the least exposed. We summarized these data by comparing the KSV of the lipid-bound form, KSV lipids, with that of the soluble form, KSV sol (Fig. 4C). The lower the ratio KSV lipids/KSV sol the greater was the change in the environment for a given NBD-labeled residue upon vesicle addition. As expected an {alpha}-helical arrangement for residues 10–27 was observed. Higher KSV lipids/KSV sol were observed for the residues on the hydrophilic side, e.g. 13, 14, 21, and 24. To ascertain whether NBD-labeled mutants penetrated the bilayer, the lipid-confined quencher, 7-NO-PC was incorporated into the SUV (SM/DPPC/7-NO-PC 5:4:1, by mole). 7-NO-PC is a short range quencher, and in order to be quenched NBD-labeled residues must be in the proximity of the nitroxide group. Confirming the transfer of the whole region to the lipid phase, all of the mutants were quenched to varying degrees by 7-NO-PC (Figs. 3A and 5). This is nicely represented by the NBD emission spectra of D17C (hydrophilic) and A12C (hydrophobic) (Fig. 3A). Quenching efficiencies were between 20 and 30% for residues from the polar side of the helix (10, 13, 17, 21, and 24) and 30–60% for those from the non-polar side (Fig. 5). This indicates that the hydrophobic side faces more directly the region of the lipid acyl chains, while the residues on the polar side are turned toward the bilayer surface within the lipid-water interface.



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FIG. 5.
Quenching of NBD-labeled mutants with membrane-embedded quencher. Efficiency of quenching, E, was calculated from normalized fluorescence intensities at 530 nm measured in the absence or presence of 7-NO-PC in SUV according to Equation 1 described under "Experimental Procedures." Other experimental conditions were as in the legend to Fig. 3. Average of 2–5 independent measurements ± S.D. is presented. A dashed line represents the {alpha}-helical pattern.

 

Topology of the N-terminal {alpha}-Helix in the Final Pore Assembly—Topological information about the final pore assembly was obtained by assaying cysteine mutants on PLM. Each mutant formed single channels with a rather broad conductance distribution, but mean values were always around 300 pS. Current fluctuations indicated opening and closing of the channels with average lifetime that changed only slightly with the mutation (Fig. 6). Even if present, small differences in conductance could not be easily reconnected to the position of the mutation. The situation was instead quite different when the cation/anion selectivity of the pores was examined (Fig. 7A). Reversal voltages, measured in a 10-fold KCl gradient, were converted, by the standard Goldman-Hodgkin-Katz equation, into a selectivity index giving the ratio of the permeability of cations over the permeability of anions (P+/P). All mutants showed cation selectivity, but the extent was quite different. Four typical levels of selectivity could be identified. Level 1 was that measured for WT, L26C, and some controls, which are not shown (Y156C, K159C, S160C, and F163C). They are non-influential and therefore not linked with the pore lumen. Level 2 is slightly more cation selective than the WT and comprises the majority of the non-charged residues in the helix. This increase is conceivably due to the effect of the mean –0.5 charge introduced by the cysteine at pH 8.0. Level 3 is less cation selective than WT and was obtained with D17C and E24C, corresponding to the fact that the original negative charge of these acidic residues was reduced to the half-negative charge of the cysteine. Finally, level 4, the most cation selective of all, was shown by D10C, T21C, and N28C. These appear to be the most important residues for pore selectivity. These residues are almost perfectly aligned on the polar side of the helix (Fig. 1B), suggesting that this is directly exposed to the pore lumen. This helix orientation is supported by the fact that mutant L26C, with the cysteine on the opposite apolar side, does not change the selectivity with respect to the WT, as it is embedded in the lipid phase.



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FIG. 6.
Current traces of EqtII and some of mutants. Current fluctuations corresponding to the opening of single ion channels with EqtII and two representative mutants (as indicated) are shown. Protein was added to the cis side at a concentration of 10–80 nM, depending on the activity. The applied voltage was in all cases +40 mV.

 


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FIG. 7.
Selectivity of channels formed by mutants in a 10-fold KCl gradient. A, mutants were added at nanomolar concentration to the cis side. Initially both sides (volume of 2 ml) were bathed by symmetrical solution of 10 mM Tris, 100 mM KCl, pH, 8.0. Thereafter the trans side was perfused with 20 ml of 10 mM Tris, 1 M KCl, pH 8.0. In this asymmetrical condition the potential necessary to reach zero current (i.e. the reversal potential (Vrev)) was determined. Vrev were converted into the reported permeability ratio (P+/P) by the Goldman-Hodgkin-Katz equation. Means of 2–5 experiments ± S.D. are presented. B, pH dependence of the reversal voltage observed with the wild type (solid squares), I18C (open circles), and D17C (open squares). Other conditions were as in A.

 

The involvement of the half-negative charge of the cysteine was confirmed by the pH dependence of the effect (Fig. 7B). In fact, the selectivity of D17C (exposed in the lumen of the pore) only reached that of WT at high pH, when a full negative charge was restored at position 17. The selectivity of a control, I18C on the apolar side of the helix, was instead nearly independent of pH.

MTS reagents were used to get further confirmation of the lumen-exposed position of the cysteines for the mutants from the polar side of the helix. MTS are known to modify channel properties when attached to thiol groups exposed in the pore lumen. Two reagents were used, MTSEA and MTSES, introducing either a positive or a negative charge. Thiol modification did not change the hemolytic properties of the mutants, which retained original activity (not shown). Modification was quantitative, according to native PAGE, which demonstrated that MTSEA and MTSES changed the electrophoresis mobility of the mutants, but not of the WT, to indicate the presence of a greater and reduced positive charge respectively (not shown).

MTSEA (positive charge) modification of mutants with a former negative charge, D10C, D17C, and E24C, caused a marked decrease in cation selectivity. In fact D10C became anion-selective, confirming this is as a crucial point of the pore lumen. Gly-11, Leu-14, and Ile-18, all positioned on the border of the hydrophobic face (Fig. 1B), showed a similar trend, suggesting that the added positive charge was able to induce a slight rotation of the helix, shifting these positions into the lumen. To analyze quantitatively these data, we calculated the ratio of the selectivity index obtained after MTSES and after MTSEA modification (Fig. 8). As shown under "Experimental Procedures," this ratio, which reports the change in selectivity when a negative or a positive charge are introduced at a given position, is a very sensitive measure of the level of exposure of that position in the pore lumen. We observed again a clear {alpha}-helical pattern, with maxima at positions 10, 14, 17, 21, 24, and 28. Non-influential positions (ratio 1), indicating no effect on selectivity, were observed with positions on the hydrophobic side. It was also clear that the extent of change decreased as the modification moved away from Asp-10 along the helix, suggesting again that this residue was the most critical.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In this study we have studied the topology of the N-terminal region (residues 10–28) of EqtII in two steps of pore formation, an intermediate membrane-bound state and the final functional pore. Two types of structural organization are likely for the membrane-inserted part of EqtII: {beta}-barrel or {alpha}-helix, as these are the only structural arrangements providing the complete backbone hydrogen bonding required by the transmembrane portion of a protein (25). The pattern that emerges clearly suggests that EqtII region 10–28 is in an {alpha}-helical conformation in both steps. We observed peaks approximately every 3–4 amino acids for each measured property (Figs. 3, 4, 5 and 8) and not the {beta}-sheet pattern that alternates at each consecutive residue. L14C and K20C were notable exceptions to the {alpha}-helical pattern. Both mutants lie on the border between the hydrophilic and hydrophobic face of the helix (Fig. 1B). Mutations of these to cysteine, and their subsequent modification, changed their polarity, thereby slightly modifying the disposition of the whole helix in the membrane. L14C behaved as if it was on the hydrophilic face (Figs. 3 and 8) and K20C as if on the hydrophobic face (Figs. 4 and 5), in agreement with the fact that the cysteine is less hydrophobic than the leucine but more than the lysine. This situation is compatible with an {alpha}-helical structure that can easily rotate around its axis within the membrane, unlike a {beta}-barrel, where each strand is stabilized by extensive hydrogen bonding to neighboring strands. In a {beta}-barrel, changes in the polarity introduced by probe labeling are generally inhibited by the hydrogen bond stabilization of the whole structure, thus providing well defined patterns (1618).

The N-terminal Helix in the Intermediate Membrane-bound Form of EqtII—The step studied by fluorescence represents an intermediate state, where EqtII was bound to the membrane via its tryptophan-rich region (1012), and the helix detached from the rest of the molecule positioned at the lipid-water interface (11, 12). The lipid-water interface is a heterogeneous layer, thick enough to accommodate a helix laying flat on the membrane plane. This was shown by x-ray scattering for melittin (26), an amphipathic peptide from the honey bee venom that shows sequence similarity to the region studied here (15). The spectral properties of NBD attached to EqtII mutants support this proposition. {lambda}max of NBD was shown to be 523–528 for the hydrophobic acyl chain interior of the membrane and 530–535 for the lipid-water interface (2729). Most of values measured here pertain to the second region and derive from residues located on the polar side of the helix. Even D17C and E24C, showing the highest values, were still considerably shifted from the solution values, demonstrating that the whole helix is transferred to the interfacial region. Consistently, mutations on the hydrophobic face of the helix, i.e. I18C, V22C, and L23C, provided the shortest {lambda}max, by accommodating their side chains in the acyl chains of the bilayer.

Quenching data (Figs. 4 and 5) lend further support to the interfacial positioning of the helix. All KSV values for iodide quenching were decreased in the membrane-bound state, indicating a less accessible position of the whole helix (Fig. 4B). Quenching with a lipid-embedded nitroxide moiety was complementary (Fig. 5) and excluded a vertical positioning of the helix for the intermediate M2-state. Similar minima of quenching efficiency were observed for residues on the polar side, away from the quencher. In a vertical arrangement of the helix, some of them should show a higher quenching value, as they should contact the nitroxide label either on one or on the other leaflet of the membrane (18).

The N-terminal Helix in the Pore—Channel properties, in particular ion selectivity, clearly indicated that region 10–28 is also involved in the walls of the pore (Fig. 7). It is arranged as an {alpha}-helix, which is positioned in the bilayer to expose side chains of residues from the polar face toward the interior of the pore. A major portion of the helix faces the lipid phase, in agreement with the helix wheel analysis showing that most of the helix is composed of hydrophobic amino acids (Fig. 1B).

From the analysis of the effects of MTS reagents (Fig. 8), we could conclude that the helices are positioned nearly perpendicular to the plane of the bilayer, with a small tilt angle that gives the pore a funnel shape with a central constriction. Asp-10 is the residue that is closer, or possibly at the constriction. Best fit provided a pore radius of 1.3 nm at the narrowest point and a tilt angle of 21°. Although these parameters should be taken with some caution, it is intriguing that the radius of actinoporin pores estimated by osmotic exclusion experiments (13, 30), permeation of fluorescent molecules (31), and PLM (14) was around 1 nm, in excellent agreement with the present result.

We have assumed that a functional pore is formed by four EqtII molecules, in agreement with cross-linking and kinetic data (13, 14). However, the pore diameter appears to be too large to be explained by a simple cluster of four helices. We have to assume either that the pore is partially formed by membrane lipids or that there are other parts of EqtII involved. The first possibility is plausible, as there is already evidence that actinoporins may trigger the formation of a toroidal lipid pore (32), similar to that formed by short amphipathic peptides like melittin and magainin (33). With regard to the second possibility, only small structural rearrangements have been observed during actinoporin binding and insertion (3436). In the bound form there is a 3% increase in {alpha}-helical content that is consistent with the folding of the unstructured region 10–15 that we observed here. The region 1–10 may form the remaining portion of the pore walls but would require that the whole region 1–28 forms a helical hairpin, a structural motif found in many pores of {alpha}-helical PFT (6, 37, 38). However, this segment is too short to provide a turn and second helix of the hairpin to double back to the surface. Furthermore, it was shown that the first five residues of EqtII could be deleted without impairing its hemolytic activity (39), arguing against an important role of part 1–5 in oligomeric assembly.

EqtII pore formation is clearly different from mechanisms known to be used by bacterial {alpha}-helical PFT and eukaryotic proteins with the same structural scaffold, such as apoptotic regulators of Bcl/Bax family (6, 40, 41). In these proteins the pore-forming domain consists of helices arranged in three layers. The middle layer is composed of a hydrophobic helical hairpin that is inserted into the membrane upon pore formation (37, 38, 42). EqtII instead uses a single amphipathic helix for membrane penetration. Its hydrophobic face touches the {beta}-sandwich core and reorients to contact acyl chains in both the membrane bound intermediate and the final oligomeric assembly. Pore formation and the use of a single helix are reminiscent of the much smaller antimicrobial peptides that use this structural motif for the nonspecific penetration of lipid membranes at higher molar concentrations (33, 43). EqtII is effective at lower concentrations, and this efficiency may be provided by the aromatic cluster on the {beta}-sandwich, which is an increasingly common motif in protein-lipid interactions (11). The molecular mechanism of pore formation by EqtII thus represents a novel paradigm for {alpha}-helical PFT.


    FOOTNOTES
 
* This work was supported by a grant from the Slovenian Ministry of Education, Science and Sport (to P. Mh., Z. P., P. Mk., and G. A.), grants from the Consiglio Nazionale delle Ricerche (CNR) and the Istituto Trentino di Cultura (ITC) (to G. V., M. D. S., and G. M.), and the BBSRC and Wellcome Trust Equipment Grants 56232, 40422, and 55979 (to J. H. L.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Present address: Educell d.o.o., Teslova 30, 1000 Ljubljana, Slovenia. Back

These authors contributed equally to the work presented in this study. Back

{ddagger}{ddagger} To whom correspondence should be addressed. Tel.: 386-1-423-33-88; Fax: 386-1-257-33-90; E-mail: gregor.anderluh{at}uni-lj.si.

1 The abbreviations used are: PFT, pore-forming toxins; DPhPC, 1,2 diphytanoyl-sn-3-glycerophosphocholine; DPPC, 1,2-dipalmitoyl-sn-glycero-3-phosphocholine; DTT, dithiothreitol; EqtII, equinatoxin II; IANBD, N-((2-(iodoacetoxy)ethyl)-N-methyl)amino-7-nitrobenz-2-oxa-1,3-diazole; L/T, lipid/toxin; MTS, methanethiosulfonate; MTSEA, (2-aminoethyl) methanethiosulfonate hydrobromide; MTSES, sodium (2-sulfonatoethyl) methanethiosulfonate; 7-NO-PC, 1-palmitoyl-2-stearoyl-(7-doxyl)-sn-glycero-3-phosphocholine; PLM, planar lipid membrane; SM, sphingomyelin; SUV, small unilamellar vesicles; WT, wild-type. Back



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Gouaux, E. (1997) Curr. Opin. Struct. Biol. 7, 566–573[CrossRef][Medline] [Order article via Infotrieve]
  2. Heuck, A. P., Tweten, R. K., and Johnson, A. E. (2001) Biochemistry 40, 9065–9073[CrossRef][Medline] [Order article via Infotrieve]
  3. Song, L., Hobaugh, M. R., Shustak, C., Cheley, S., Bayley, H., and Gouaux, J. E. (1996) Science 274, 1859–1866[Abstract/Free Full Text]
  4. Rossjohn, J., Feil, S. C., McKinstry, W. J., Tweten, R. K., and Parker, M. W. (1997) Cell 89, 685–692[CrossRef][Medline] [Order article via Infotrieve]
  5. Petosa, C., Collier, R. J., Klimpel, K. R., Leppla, S. H., and Liddington, R. C. (1997) Nature 385, 833–838[CrossRef][Medline] [Order article via Infotrieve]
  6. Lakey, J. H., and Slatin, S. L. (2001) Curr. Top. Microbiol. Immunol. 257, 131–161[Medline] [Order article via Infotrieve]
  7. Anderluh, G., and Macek, P. (2002) Toxicon 40, 111–124[CrossRef][Medline] [Order article via Infotrieve]
  8. Athanasiadis, A., Anderluh, G., Macek, P., and Turk, D. (2001) Structure (Camb.) 9, 341–346[CrossRef][Medline] [Order article via Infotrieve]
  9. Hinds, M. G., Zhang, W., Anderluh, G., Hansen, P. E., and Norton, R. S. (2002) J. Mol. Biol. 315, 1219–1229[CrossRef][Medline] [Order article via Infotrieve]
  10. Malovrh, P., Barlic, A., Podlesek, Z., Macek, P., Menestrina, G., and Anderluh, G. (2000) Biochem. J. 346, 223–232[CrossRef][Medline] [Order article via Infotrieve]
  11. Hong, Q., Gutierrez-Aguirre, I., Barlic, A., Malovrh, P., Kristan, K., Podlesek, Z., Macek, P., Turk, D., González-Mañas, J. M., Lakey, J. H., and Anderluh, G. (2002) J. Biol. Chem. 277, 41916–41924[Abstract/Free Full Text]
  12. Anderluh, G., Barlic, A., Podlesek, Z., Macek, P., Pungercar, J., Gubensek, F., Zecchini, M. L., Dalla, Serra, M., and Menestrina, G. (1999) Eur. J. Biochem. 263, 128–136[Abstract/Free Full Text]
  13. Belmonte, G., Pederzolli, C., Macek, P., and Menestrina, G. (1993) J. Membr. Biol. 131, 11–22[Medline] [Order article via Infotrieve]
  14. Tejuca, M., Dalla Serra, M., Ferreras, M., Lanio, M. E., and Menestrina, G. (1996) Biochemistry 35, 14947–14957[CrossRef][Medline] [Order article via Infotrieve]
  15. Belmonte, G., Menestrina, G., Pederzolli, C., Krizaj, I., Gubensek, F., Turk, T., and Macek, P. (1994) Biochim. Biophys. Acta 1192, 197–204[Medline] [Order article via Infotrieve]
  16. Valeva, A., Walev, I., Pinkernell, M., Walker, B., Bayley, H., Palmer, M., and Bhakdi, S. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 11607–11611[Abstract/Free Full Text]
  17. Shepard, L. A., Heuck, A. P., Hamman, B. D., Rossjohn, J., Parker, M. W., Ryan, K. R., Johnson, A. E., and Tweten, R. K. (1998) Biochemistry 37, 14563–14574[CrossRef][Medline] [Order article via Infotrieve]
  18. Shatursky, O., Heuck, A. P., Shepard, L. A., Rossjohn, J., Parker, M. W., Johnson, A. E., and Tweten, R. K. (1999) Cell 99, 293–299[Medline] [Order article via Infotrieve]
  19. Heuck, A. P., and Johnson, A. E. (2002) Cell Biochem. Biophys. 36, 89–101[Medline] [Order article via Infotrieve]
  20. Anderluh, G., Pungercar, J., Strukelj, B., Macek, P., and Gubensek, F. (1996) Biochem. Biophys. Res. Comm. 220, 437–442[CrossRef][Medline] [Order article via Infotrieve]
  21. Anderluh, G., Gokce, I., and Lakey, J. H. (2002) Prot. Exp. Purif. 28, 173–181
  22. Schultz, S. G. (1980) Basic Principles of Membrane Transport, Cambridge University Press, New York
  23. Bockris, J. O. M., and Reddy, A. K. N. (1970) Modern electrochemistry, Plenum Press, New York
  24. Lakowicz, J. R. (1999) Principles of fluorescence spectroscopy, Kluwer Academic/Plenum Publishers, New York
  25. White, S. H., Ladokhin, A. S., Jayasinghe, S., and Hristova, K. (2001) J. Biol. Chem. 276, 32395–32398[Free Full Text]
  26. Hristova, K., Wimley, W. C., Mishra, V. K., Anantharamiah, G. M., Segrest, J. P., and White, S. H. (1999) J. Mol. Biol. 290, 99–117[CrossRef][Medline] [Order article via Infotrieve]
  27. Rajarathnam, K., Hochman, J., Schindler, M., and Ferguson-Miller, S. (1989) Biochemistry 28, 3168–3176[Medline] [Order article via Infotrieve]
  28. Rapaport, D., and Shai, Y. (1991) J. Biol. Chem. 266, 23769–23775[Abstract/Free Full Text]
  29. Gazit, E., Bach, D., Kerr, I. D., Sansom, M. S. P., Chejanovsky, N., and Shai, Y. (1994) Biochem. J. 304, 895–902[Medline] [Order article via Infotrieve]
  30. Tejuca, M., Dalla Serra, M., Potrich, C., Alvarez, C., and Menestrina, G. (2001) J. Membr. Biol. 183, 125–135[CrossRef][Medline] [Order article via Infotrieve]
  31. De los Rios, V., Mancheño, J. M., Lanio, M. E., Onaderra, M., and Gavilanes, J. G. (1998) Eur. J. Biochem. 252, 284–289[Abstract]
  32. Valcarcel, C. A., Dalla Serra, M., Potrich, C., Bernhart, I., Tejuca, M., Martinez, D., Pazos, F., Lanio, M. E., and Menestrina, G. (2001) Biophys. J. 80, 2761–2774[Abstract/Free Full Text]
  33. Yang, L., Harroun, T. A., Weiss, T. M., Ding, L., and Huang, H. W. (2001) Biophys. J. 81, 1475–1485[Abstract/Free Full Text]
  34. Menestrina, G., Cabiaux, V., and Tejuca, M. (1999) Biochem. Biophys. Res. Comm. 254, 174–180[CrossRef][Medline] [Order article via Infotrieve]
  35. Poklar, N., Fritz, J., Macek, P., Vesnaver, G., and Chalikian, T. V. (1999) Biochemistry 38, 14999–15008[CrossRef][Medline] [Order article via Infotrieve]
  36. Anderluh, G., Barlic, A., Potrich, C., Macek, P., and Menestrina, G. (2000) J. Membr. Biol. 173, 47–55[CrossRef][Medline] [Order article via Infotrieve]
  37. Shin, Y. K., Levinthal, C., Levinthal, F., and Hubbell, W. L. (1993) Science 259, 960–963[Medline] [Order article via Infotrieve]
  38. Oh, K. J., Zhan, H. J., Cui, C., Hideg, K., Collier, R. J., and Hubbell, W. L. (1996) Science 273, 810–812[Abstract]
  39. Anderluh, G., Pungercar, J., Krizaj, I., Strukelj, B., Gubensek, F., and Macek, P. (1997) Protein Eng. 10, 751–755[Abstract]
  40. Parker, M. W., Pattus, F., Tucker, A. D., and Tsernoglou, D. (1989) Nature 337, 93–96[CrossRef][Medline] [Order article via Infotrieve]
  41. Muchmore, S. W., Sattler, M., Liang, H., Meadows, R. P., Harlan, J. E., Yoon, H. S., Nettesheim, D., Chang, B. S., Thompson, C. B., Wong, S. L., Ng, C. S., and Fesik, S. W. (1996) Nature 381, 335–341[CrossRef][Medline] [Order article via Infotrieve]
  42. Engelman, D. M., and Steitz, T. A. (1981) Cell 23, 411–422[Medline] [Order article via Infotrieve]
  43. Oren, Z., and Shai, Y. (1998) Biopolymers 47, 451–463[CrossRef][Medline] [Order article via Infotrieve]
  44. Rodriguez, R., Chinea, G., Lopez, N., Pons, T., and Vriend, G. (1998) Bioinformatics 14, 523–528[Abstract]