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* Department of Biochemistry, Department of Chemical Engineering and Applied Chemistry, Institute of Biomaterials and Biomedical Engineering, University of Toronto, Toronto, Ontario, Canada; Sandia National Laboratories, Biosystems Research, Livermore, California; and Sandia National Laboratories, Biomolecular Interfaces & Systems, Albuquerque, New Mexico
Correspondence: Address reprint requests to Christopher M. Yip, IBBME, Rosebrugh Bldg., University of Toronto, 4 Taddle Creek Rd., Toronto, Ontario, Canada, M5S 3G9. Tel: 416-978-7853; Fax: 416-978-4317; E-mail: christopher.yip{at}utoronto.ca.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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,2). Characterizing the dynamics of their interactions at the membrane interface may therefore provide insights into the development of new treatments, detection strategies, and prophylactic agents. The Clostridial neurotoxins are the most potent toxins known, with TeNT and BoNT responsible for the clinical symptoms of tetanus and botulism, respectively. These toxins target neuronal cells by binding to the trisialoganglioside GT1b in the presynaptic cell membrane of neuromuscular junctions (3,4), and are internalized into the cells where they ultimately inhibit neurotransmitter release (5). However, while BoNTs act primarily in the peripheral nervous system blocking acetylcholine release, TeNT undergoes axonal retrograde transport to the central nervous system where it targets inhibitory motor neurons (6,7). TeNT is synthesized by the bacterium Clostridium tetani as a single 
150 kDa polypeptide chain that must undergo posttranslational proteolytic cleavage to produce the biologically active, disulfide linked two-chain molecule that is composed of an 50-kDa light chain (LC) and an 100-kDa heavy chain (HC) (Fig. 1 a) (8). The binding of tetanus toxin to GT1b in neuronal cell membranes is mediated by the 50-kDa C-terminal region (HC) of the heavy chain, known as the C fragment (Tet C) (Fig. 1 b). The 50-kDa N-terminal region (HN) facilitates the translocation of the LC, a zinc endopeptidase, into the cell cytosol. Once in the cytosol, the LC specifically cleaves the synaptic SNARE (soluble N-ethylmaleimide sensitive-factor attachment protein receptor) protein synaptobrevin, thereby preventing the fusion of neurotransmitter secretory vesicles to the nerve terminal membrane, and thus blocking the release of inhibitory neurotransmitters (9).
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,11). This conformational change facilitates insertion of TeNT into the endosomal membrane with subsequent translocation of the light chain into the cytosol, where it exerts its catalytic activity. Indeed, it has been reported that AnBm-toxins having A1B1-stoichiometry, including the full-length tetanus toxin, form ion channels in planar lipid bilayers at acidic pH (12–18).
To fully understand how toxins exert their action, it has been proposed that "trapping" the toxin molecules at different points along the invasion pathway from the cell surface to the cytosol, would allow for reconstruction of the entire mechanism (19). Although this approach has been applied successfully to characterize colicin Ia activity, it is extremely arduous and time intensive (20). Recently, in situ studies using scanning probe microscopy (SPM), and more specifically, atomic force microscopy (AFM), have provided new insights into the association and assembly of various peptides and proteins, such as filipin (21), amphotericin B (22), and melittin (23), on membrane surfaces, including their role in inducing membrane reorganization and disruption. We have used this technique to study the insertion and subsequent assembly of the amyloid-ß 42 (Aß42) peptide (24), 
-synuclein (25), hemagglutinin (26), Bax protein (27), and NAP22 (28) at membrane interfaces.
For such studies, supported planar lipid bilayers (SPBs) are often used as membrane-mimetic surfaces. Typically prepared in situ by free vesicle fusion onto a freshly cleaved mica surface, SPBs retain many of the properties of free-standing membranes, including lateral fluidity (29–31). Indeed, direct fusion of receptor-containing lipid vesicles onto mica is a particularly facile and attractive route of preparing model membranes for AFM studies of ligand-receptor interactions. We have used this approach in our earlier investigation of the full-length transmembrane insulin receptor (IR) (32).
We report herein an in situ AFM study on the interactions of the tetanus toxin binding domain (Tet C) with biphasic supported planar lipid bilayers containing the ganglioside membrane receptor GT1b. Our results revealed preferential association of Tet C with the surface of the fluid phase regions of the lipid bilayers, and that the affinity was dependent upon the bilayer concentration of GT1b and solution pH. Following an incubation period, the protein-bound regions of the membrane became thicker and circular depressions of 40–80 nm diameters appeared through the action of Tet C upon the membrane. The presence of Tet C in these regions of the bilayer was confirmed using combined atomic force microscopy/total internal reflection fluorescence microscopy (AFM/TIRFM) (33). From these studies, detailed insights into the binding and activity of Tet C and subsequent restructuring of the lipid bilayer were attained at nanoscale resolution.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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-dipalmitoyl-phosphatidylcholine (DPPC; C16:0) were purchased from Sigma-Aldrich (Oakville, ON).
Liposome preparation
Appropriate molar quantities of lipid ((1:1) DPOPC/DPPC) and, when required, the desired amount of GT1b (10 mol % or 1 mol %) were dissolved in chloroform. During preparation, fluorescent liposomes were protected from light to minimize photobleaching. The solvent was removed by evaporation under vacuum (50°C) and the dried lipid films were rehydrated by the addition of 5 mM (N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid)-(2-[N-morpholino] ethanesulfonic acid)-citric acid (HEPES-MES-citric acid) buffer (150 mM NaCl, pH 7.4) to give a final lipid concentration of 2 mM. Unilamellar vesicles were then formed by sonication in a heated water bath (50°C) (Branson 200, Branson Ultrasonics, Danbury, CT) until the solution became clear or only slightly hazy. The resulting liposomes were found to be 145 nm in diameter by dynamic laser light scattering (Brookhaven Instruments BI-200SM equipped with a BI-9000 AT digital correlator and photon counter; Holtsville, NY). The fluorescent liposome solutions were stored in the dark. All liposome solutions were stored at 4°C.
AFM imaging
Solution tapping mode AFM (TMAFM) images were acquired on a Digital Instruments Nanoscope IIIa Multimode SPM (Santa Barbara, CA) equipped with an "E" scanner (13.6 x 13.6 µm maximum lateral scan area) using 120-µm-long oxide-sharpened silicon nitride V-shaped model DNP-S cantilevers installed in a combination contact/tapping mode liquid flow-cell sealed against a freshly cleaved mica substrate. The typical tip oscillating frequency was 7–9 kHz and was chosen to optimize image quality and adjusted for the individual response characteristics of each cantilever. We note that the nominal radius of curvature of the DNP-S tips used in these studies was 15 nm, and that for each experiment, a fresh tip was selected and exposed to ultraviolet irradiation to remove any adventitious organic contaminants prior to imaging. All AFM images were collected as 512 x 512 pixel data sets, at scan rates ranging from 1 to 2 Hz, at a scan angle of 0° to the fast scan axis, and at ambient temperature. For the Tet C studies on mica, 150 µL of a 5 µg/mL Tet C solution (10 mM PBS, 150 mM NaCl, pH 7.4) was applied to the mica surface and the sample sealed in the AFM fluid cell. The fluid cell was flushed through with protein-free buffer after a 30-min incubation period. The sample remained fully hydrated during the incubation period. Supported planar lipid bilayers were formed in situ by injecting 200 µL of the GT1b/DPOPC/DPPC or DPOPC/DPPC liposome solution directly into the fluid cell and allowing it to adsorb to the mica surface for 10 min. The fluid cell was then flushed with 10 mM CaCl2 solution to facilitate liposome fusion. Once the bilayers were formed, calcium ions and excess lipid were removed by flushing the fluid cell with 10 mM ethylenediaminetetraacetic acid (EDTA) solution. Before the introduction of Tet C, reference images of the lipid bilayers were acquired in 5 mM HEPES-MES-citric acid buffer (pH 7.4). For the Tet C-membrane interaction studies, 300 µL of a 10 µg/mL HEPES-MES-citric acid solution (pH 7.4) of Tet C was injected directly into the fluid cell and imaging initiated after 1 h. In the case of experiments carried out at pH 4.0, the Tet C-free bilayers were formed at pH 7.4 and the imaging fluid exchanged with 5 mM HEPES-MES-citric acid buffer (pH 4.0) before the addition of Tet C. For these studies, the 10 µg/mL Tet C solution was also made up in 5 mM HEPES-MES-citric acid buffer (pH 4.0). All AFM imaging experiments were repeated five times.
Combined AFM/TIRFM imaging
Solution tapping mode atomic force microscopy/total internal reflection fluorescence microscopy (TMAFM/TIRFM) images were acquired on a Digital Instruments Nanoscope IIIa Bioscope SPM equipped with an extended z-range "J" scanner (116 x 116 µm maximum lateral scan area) and mounted on an Olympus Fluoview 500 confocal microscope equipped with an Olympus TIRFM accessory (10 mW Ar-ion (488 nm) laser (Melles Griot, CA); Olympus 60x 1.45 NA TIRFM objective). All TIRFM images were acquired simultaneously using a cooled charge-coupled device camera (CoolSnap HQ, Roper Scientific, Tucson, AZ) and the Image Pro Plus software package (Media Cybernetics, Silver Springs, MD). Oxide-sharpened silicon nitride V-shaped cantilevers (120 µm) were installed in a combination contact/tapping mode liquid cell and supported planar lipid bilayers were formed in situ by the introduction of 200 µL of a 10 mol % GT1b/DPOPC/DPPC liposome solution directly onto a freshly cleaved mica surface that was sealed within a custom-built thermostated flow cell. The flow cell was fitted with a motorized syringe pump (Harvard Apparatus, Saint-Laurent, Quebec, Canada) to facilitate exchange of imaging solutions. The liposomes were allowed to adsorb to a mica surface for 10 min and then 200 µL of a 10 mM CaCl2 solution was then added to aid liposome fusion. Once the bilayers had formed, the flow cell was flushed several times sequentially with 10 mM EDTA solution and 5 mM HEPES-MES-citric acid buffer (pH 7.4) to remove calcium ions and any excess lipid. All AFM/TIRFM imaging was conducted in complete darkness at ambient temperature in 5 mM HEPES-MES-citric acid buffer (pH 7.4). For the fluorescently labeled Tet C studies, 200 µL of a 25 µg/mL HEPES-MES-citric acid buffer solution (pH 7.4) of FITC-Tet C was added to the fluid cell. The FITC-Tet C solution contained 5 µg of bound FITC per milligram of Tet C, as stated by the manufacturer. Imaging was initiated after 1 h incubation. Five replicate AFM/TIRFM experiments were performed.
AFM image analysis
Image analyses were conducted using the Digital Instruments Nanoscope software (version 4.42r9). Height images were low-pass filtered and plane-fit in the x-scan direction. Quantitative height measurements were determined by section analysis.
Molecular modeling
Images were prepared using Swiss PDBViewer, Version 3.7b2 (http://www.expasy.org/spdbv/). The Tet C and BoNT B molecular coordinates were obtained from the Protein Data Bank (PDB) using the identification codes 1FV2 and 1SOF, respectively.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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), there have been no reported structural studies of Tet C bound to its native receptor in a membrane or membrane-mimetic environment. To establish a structural baseline, we first characterized Tet C as an adsorbed species on a mica surface. After 30 min incubation in the presence of a 5 µg/mL solution of Tet C, tapping mode AFM (TMAFM) imaging of the mica surface revealed small, randomly distributed particles with a height of 3.5 nm and widths ranging from 15 to 40 nm (Fig. 2 a). Electrostatic potential maps of the Tet C surface reveal a large area of negative charge (red area) opposite to the ganglioside-binding site, which is an area of concentrated positive charge (blue area) (Fig. 2 b). As mica is negatively charged at neutral pH, it is likely that Tet C would adsorb in an orientation such that the area of positive charge, or the ganglioside-binding site, would interact with the mica surface. This orientation would presumably be similar to that with which Tet C interacts with its ganglioside receptor and is consistent with the structures observed on mica by TMAFM. We note that although AFM tip-sample convolution does contribute to significant overestimation of the lateral dimensions of adsorbed molecules (35), image deconvolution of these structures using a nominal tip size of 15 nm was consistent with the adsorption of Tet C, as single molecules, to the mica surface.
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). In this study, solution tapping mode AFM (TMAFM) imaging of the 10 mol % GT1b-containing DPOPC/DPPC lipid bilayers revealed molecularly smooth bilayers that were composed of two types of membrane structures, with tall domains extending 1.5 nm above a shorter continuous lipid phase (Fig. 3). The lipid bilayers themselves were fairly continuous across the AFM imaging window, with only the occasional defect exposing the underlying mica surface. Cross section analysis performed at the edges of these defect areas revealed the shorter continuous phase to be 5.5 nm tall (for example, see Fig. 4 a). These results are consistent with previous AFM studies of binary lipid systems where the 1.5 nm height difference has been attributed to a decrease in the tilt angle of the acyl chains of the gel phase lipids due to tighter packing of the hydrophobic tails (36–40). As such, these data are in agreement with the taller domains being rich in DPPC and the shorter regions of the membrane being rich in DPOPC. We cannot discount the possibility of transient bilayer defects that exist on timescales shorter than that which can be readily resolved by the AFM.
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). Even at high resolution, no features were found that could be attributed to aggregates of GT1b. It is known that gangliosides, as well as other glycosphingolipids, and cholesterol are involved in the formation of "lipid raft" membrane domains. These structures are thought to play a leading role in regulating membrane function and have been implicated in mediating molecular recognition and intracellular signal transduction (41–44). In the past, evidence of such domains has often been obtained by direct imaging of complexes between the ganglioside receptors and their corresponding ligand. In the case of monosialoganglioside GM1, this has been accomplished through the introduction of soluble cholera toxin B-subunits (CTX-B). For instance, Mou et al. performed AFM studies on the CTX-B-GM1 complex to infer that GM1 was homogeneously distributed throughout gel phase dipalmitoylphosphatidylcholine (DPPC) bilayers (45). In yet other AFM studies, Yuan and Johnston used CTX-B as a reporter molecule to confirm clustering of GM1 receptors in DPPC monolayers and bilayers as well as in fluid phase egg-PC lipid bilayers (46,47). To date, there has been very little work done to characterize the distribution of GT1b in bilayers although we note that electron microscopy has been used to visualize GT1b in phosphatidylcholine bilayers using lectin molecules as topographical markers (48).
Although the presence of GT1b was not confirmed by AFM imaging, its presence was verified through binding studies with Tet C. In the absence of GT1b, Tet C did not exhibit any affinity for the DPOPC/DPPC bilayers with no changes observed in the bilayer surface topology or structural morphology after 12 h exposure to a 10 µg/mL solution of Tet C (data not shown). Under the same conditions, in situ TMAFM revealed a structural topology of DPOPC/DPPC bilayers containing 10% GT1b that suggests adsorption of Tet C with selective affinity for the DPOPC-rich areas after 2 h incubation (Fig. 4 a). These regions of the bilayer were populated by numerous protrusions having dimensions that are consistent with bound Tet C molecules. The bound proteins were homogeneously distributed throughout the fluid phase DPOPC extending 2 nm above the bilayer surface. The relatively featureless surface of the areas rich in DPPC suggests little adsorption of Tet C to these gel phase areas, although further confirmation was attained via fluorescence imaging (vide infra).
TMAFM imaging of the same area of the bilayer after 12 h of exposure revealed a dramatic change in the overall bilayer morphology (Fig. 4 b). While the DPPC-rich domains remained relatively unchanged, the previously smooth DPOPC-rich areas now had a granular texture and were populated by numerous small, circular cavities with diameters in the range of 40–80 nm (Fig. 4 c). These changes in the overall structure of the membrane are clearly evident in the three-dimensional image of the bilayer surface shown in Fig. 5. Cross section analysis also revealed that after 12 h the DPOPC-rich lipid phase now exhibited a thickness similar to that of the DPPC domains (7 nm). In addition to the cavity formation and the thickening of the DPOPC lipid phase, the Tet C molecules bound to the surface of the DPOPC-rich regions were also found to have decreased in height (<1 nm in height). Upon closer examination, the small number of Tet C aggregates occasionally found on the surface of the DPPC domains exhibited no change in height nor induced any structure changes on that portion of the membrane after 12 h of incubation (Fig. 4 c). In control studies performed with supported lipid bilayers of the same composition (10 mol % GT1b/DPOPC/DPPC), no change or obvious disruption of the bilayers was observed after 12 h of TMAFM imaging in the absence of Tet C (data not shown).
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25 µg/mL solution of FITC-labeled Tet C fragment (FITC-Tet C). After in situ fusion of the GT1b/DPOPC/DPPC lipid vesicles on the mica surface, TMAFM imaging revealed well-formed phase-separated lipid bilayers with a morphology similar to those previously described. There was no observable auto-fluorescence from these lipid bilayers when TIRFM imaging was performed (data not shown). After 12 h incubation of the bilayers with FITC-Tet C, in situ TMAFM imaging revealed that the DPOPC regions were again granular in texture and populated by numerous circular cavities, whereas the DPPC domains remained relatively unchanged (Fig. 6 a). The corresponding TIRFM images, acquired simultaneously, revealed a bright fluorescence emission from the DPOPC regions, which is consistent with preferential association of the FITC-Tet C with the fluid lipid phase (Fig. 6 b). We note that TIRFM data acquired during intermediate time points was consistent with the association of the FITC-Tet C with the bilayer.
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,50). We also cannot discount the possibility that the dynamics of the lipid bilayer may be better resolved on TIRFM timescales rather than on AFM imaging timescales (33).
The results of our correlated TMAFM/TIRFM study suggest that after continuous incubation of Tet C with the 10 mol % GT1b/DPOPC/DPPC lipid bilayers, the surface-bound Tet C undergo partial insertion into the GT1b-containing DPOPC fluid phase regions of the bilayer, thus explaining both the decrease in height and the apparent decrease in the surface coverage of Tet C in these areas. The absence of Tet C within the gel-phase DPPC domains may be due to the tighter packing of the DPPC molecules, which prevents protein insertion, or due to an absence of GT1b in the DPPC domains. Similar studies using 10 mol % GT1b/DPPC lipid bilayers also revealed no GT1b-induced clustering or domain formation nor any specific association of Tet C molecules with these surfaces after extended incubation periods (data not shown). Although the preferential association with the DPOPC-rich regions observed for Tet C may reflect favorable partitioning of the GT1b receptors into the fluid phase regions of the lipid bilayers, it is also possible that it may simply reflect a greater degree of mobility of the receptors within the fluid DPOPC matrix, and hence greater ease in adopting the correct binding conformation. Although it would be ideal to use combined TMAFM/TIRFM to address the issue as to the presence/absence of GT1b within the DPPC lipid domains, it is becoming increasingly apparent that the behavior of a fluorescently labeled lipid is strongly influenced by the presence of the fluorophore (51–53). The addition of a bulky fluorophore to the acyl chain of a ganglioside or phosphocholine molecule prevents tight packing of the lipid tails, and thus often results in the formation of, or partitioning of the fluorescent probe into, a more disordered phase in the lipid bilayer. Similar behavior has also been observed for headgroup labeled lipids (54,55). As such, under the conditions of our studies, we are unable to quantitatively determine the partitioning of GT1b in either lipid phase.
In situ TMAFM studies were also conducted to examine the effect of varying the GT1b concentration and the pH of the surrounding imaging fluid on Tet C-membrane interactions. In the case of DPOPC/DPPC bilayers containing 1 mol % GT1b, the Tet C molecules were also found to be preferentially associated with the DPOPC-rich areas after an initial 2-h incubation period. In contrast to our studies of the 10 mol % GT1b/DPOPC/DPPC bilayers after exposure to Tet C (vide infra), after 12 h of incubation with Tet C, we saw no evidence of persistent defects in either lipid phase of the 1 mol % GT1b/DPOPC/DPPC bilayers (Fig. 7). Cross section analysis revealed the Tet C molecules associated with the DPOPC-rich regions after 12 h to have a height of 2 nm relative to the surface of the bilayer. The occasional Tet C molecule was also observed on the surface of the DPPC domains with a height of 2 nm.
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,11). These data are consistent with those of Winter et al. who reported that the full-length TeNT preferentially bound to negatively charged lipids in vesicles (56). Based on the results of their studies into the interactions of TeNT with vesicles containing different types of lipid molecules, Winter et al. believed that successful binding of the toxin was determined by lipid headgroup charge and conformation, as well as acyl chain length, with all of these factors being best fulfilled by the ganglioside GT1b. These observations suggest that electrostatic interactions play a large role in the binding of TeNT to the GT1b receptor. As the Tet C fragment is the membrane-binding domain, it is likely that its interactions with GT1b would be sensitive to changes in pH. This result is in contrast to work by others who have examined the formation of pores in lipid membranes by the full-length tetanus toxin at low pH (57–60).
The results of these studies clearly indicate that at physiological pH (7.4) the tetanus toxin C-fragment binds to the fluid phase regions of GT1b-containing membranes and after prolonged exposure, appears to organize into structures or assemblies that generate fairly uniform and relatively large cavity depressions within these regions. The gel phase domains, however, were relatively resistant to Tet C binding and activity. Our preliminary studies also suggest a threshold GT1b concentration is necessary to initiate Tet C association and cavity formation, although at this time we cannot confirm whether the distribution of GT1b is uniform throughout the model membranes. Studies by Winter et al. of the full-length tetanus neurotoxin (TeNT) also found that a critical concentration of GT1b was necessary for pore formation in phosphatidylcholine lipid vesicles (61). Calorimetric measurements indicated that each TeNT molecule required 20 bound GT1b molecules to adopt the correct conformation for stable pore formation.
Cross section analysis performed on the larger cavities formed in the DPOPC-rich areas revealed that they extend 1.5 nm into the bilayer and do not span the entire SPB to expose the underlying mica surface (Fig. 6 c). Although the cavities themselves do not have the characteristics of transmembrane pores, they may instead be precursors toward the formation of the smaller membrane pores (estimated at 2 nm diameter for tetanus neurotoxin) (15) or perhaps the initial stages of endosome formation. Neuronal endosomes are known to form within the size range of these observed cavities (62). The presence of a fluorescence signal in the areas of the defects in the correlated TIRFM images indicates that Tet C was present within the cavities, which likely contributed to their stability over extended periods of AFM imaging (greater than hours) (Fig. 6 b). Choi and Dimitriadis performed in situ AFM studies of the binding of cytochrome c to anionic lipid bilayers (61). At higher concentrations, cytochrome c was found to insert into the fluid phase bilayers and over time, formed circular defects in the bilayers. The authors believed that the defects were nanodomains of pure protein. Studies by others have suggested that the Tet C fragment itself does not form pores in lipid bilayers (18,57–59,63). This model is in reasonable agreement with our studies where Tet C appears to affect only the upper leaflet of the bilayer. In the context of this study, we cannot discount the possibility that this effect is specific to the use of a supported planar lipid bilayer model of a membrane.
We note that the larger size of the cavities observed in our combined TMAFM/TIRFM studies may be attributed to the higher concentration of FITC-labeled Tet C used. Sharpe and London found that the size of the pores formed by the A1B1-diphtheria toxin in lipid vesicles increased with increasing toxin concentration (64). The authors believed that the change in pore size was due to the oligomerization of the diphtheria molecules, with the number of toxin subunits determining the size of the pore formed in the membrane. In our AFM studies, we did not observe the formation of any supramolecular assemblies of Tet C on the surface of the lipid bilayer before the appearance of the cavity structures.
The insertion of Tet C into the fluid phase regions of the lipid bilayer and concomitant cavity formation may explain the observed increase in the thickness of these areas after the 12-h incubation period. This process would likely cause tighter packing of the DPOPC molecules, and hence a decrease in the tilt angle of their acyl chains and subsequent increase in the height of these regions (36–38). As cell membranes are dynamic in nature, this change in the morphology of the bilayer may reflect an actual physiological cellular response. Cantor proposed a mechanism whereby changes in the lateral pressure of a cell membrane can affect the behavior of membrane proteins, thereby influencing cell surface recognition and initiating intracellular signaling events (65,66).
In addition to the circular cavities found throughout the fluid phase of the 10 mol % GT1b/DPOPC/DPPC lipid bilayers, prominent defects were found at the border between the gel phase DPPC domains and the fluid phase DPOPC lipid matrix (Figs. 4, b and c, and 5). These defects exist between the intact DPPC and granular, cavity-filled DPOPC domains, and were found on both the leading and trailing edges, as defined by the scanning AFM tip, of various domains, suggesting that these structures are not imaging artifacts. Indeed, Barlic et al. found that the pore-forming equinatoxin-II preferentially binds at the boundaries between the ordered and disordered phases in biphasic membranes (67). They proposed that lipid packing defects and differences in the membrane thickness occurring at these interfaces may facilitate favorable interactions with the toxin. As Tet C inserts into the lipid bilayer, we are unable to detect by AFM, any accumulation of Tet C at the interface between the DPOPC and DPPC lipid phases. Ideally, we would be able to characterize, using TIRFM, any accumulation of Tet C at these interfaces as an increase in the intensity of the fluorescence emission in these regions. Unlike AFM, however, TIRFM is a diffraction-limited technique. With such a large amount of fluorescently labeled Tet C present throughout the DPOPC fluid phase of the bilayer contributing to the overall fluorescence emission, it was difficult to resolve whether the intensity within the DPOPC regions differs from that at the DPOPC-DPPC interfaces. Nevertheless, our observations of the membrane-disruptive activity of the C-fragment of the A1B1-tetanus toxin may have physiological relevance as it suggests that Tet C may actually be involved or aid in the formation of a "true" membrane pore by the full-length TeNT.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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,59). Furthermore, we observed preferential association of Tet C with the GT1b-containing fluid phase regions in gel-fluid biphasic membranes. Over time, these interactions facilitated a dramatic change in the morphology of the supported lipid bilayer that resulted in the formation of 40–80 nm cavities with a concomitant increase in the thickness of the fluid phase regions of the bilayer upon insertion of bound Tet C. We also found that the membrane activity of Tet C was dependent upon both the ganglioside concentration in the membrane and solution pH. Although TMAFM studies were unable to resolve Tet C on the surface of the lipid bilayer after the structural change in the fluid phase regions had occurred, combined TMAFM/TIRFM studies were able to confirm that Tet C is indeed present in the bilayer. This demonstrates the power of this coupled approach for performing functional imaging of lipid membranes and specifically ligand/protein-membrane interactions. The results of these studies now provide us with the framework necessary to perform in situ AFM studies of the membrane interactions and subsequent pore formation by the full-length tetanus toxin.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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Submitted on December 27, 2005; accepted for publication August 22, 2006.
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REFERENCES |
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