Biophysical Journal Volume 105 September 2013 1397–1405

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Real-Time Visualization of Assembling of a Sphingomyelin-Specific Toxin
on Planar Lipid Membranes
Neval Yilmaz,† Taro Yamada,† Peter Greimel,† Takayuki Uchihashi,‡§ Toshio Ando,‡§ and Toshihide Kobayashi†{*†

Lipid Biology Laboratory, RIKEN, Wako, Saitama, Japan; ‡Department of Physics and §Bio-AFM Frontier Research Center,
Kanazawa University, Kanazawa, Ishikawa, Japan; and {Institut National de la Santé et de la Recherche Médicale U1060-Université Lyon 1,
Villeurbanne, France

ABSTRACT Pore-forming toxins (PFTs) are soluble proteins that can oligomerize on the cell membrane and induce cell deathby membrane insertion. PFT oligomers sometimes form hexagonal close-packed (hcp) structures on the membrane. Here, weshow the assembling of the sphingomyelin (SM)-binding PFT, lysenin, into an hcp structure after oligomerization on SM/
cholesterol membrane. This process was monitored by high-speed atomic force microscopy. Hcp assembly was driven byreorganization of lysenin oligomers such as association/dissociation and rapid diffusion along the membrane. Besides rapid
association/dissociation of oligomers, the height change for some oligomers, possibly resulting from conformational changes
in lysenin, could also be visualized. After the entire membrane surface was covered with a well-ordered oligomer lattice, thelysenin molecules were firmly bound on the membrane and the oligomers neither dissociated nor diffused. Our results reveal
the dynamic nature of the oligomers of a lipid-binding toxin during the formation of an hcp structure. Visualization of this dynamic
process is essential for the elucidation of the assembling mechanism of some PFTs that can form ordered structures on themembrane.

INTRODUCTION
Pore-forming toxins (PFTs) are soluble proteins found in awide spectrum of organisms, including bacteria, fungi, andanimals, and they exert toxic effects by binding the target
membranes, where they assemble into oligomers (1). PFT
oligomers are sometimes organized into two-dimensional
hexagonal close-packed (hcp) structures that have been
resolved by electron and atomic force microscopy in model
membranes (2–4). The mechanism of the formation of hcp
structures on lipid membranes is of interest, because oligomer formation can occur randomly on the membrane.
In this study, we focused on the assembly of the earthworm-derived 297-aa-long pore-forming toxin, lysenin(5,6). Lysenin is known for its specific interaction withsphingomyelin (SM) (7,8). Recently, the crystal structureof lysenin revealed the presence of multiple SM bindingsites (9). SM is one of the major lipids in the outer leafletof the plasma membrane of mammalian cells. SM is a reservoir of the signaling lipid, ceramide (10). It also forms lipid
domains with cholesterol (chol) known as ‘‘lipid rafts’’ (11).
The specific interaction between SM and lysenin enables
the detection of SM, and hence, allows the examinationof the lipid distribution within the membrane (12,13). By
transmission electron microscopy, it was revealed thatlysenin oligomerizes and forms hcp structures on liposomesof SM and chol (8). Although lysenin oligomerizes onlyin the presence of SM, chol is essential for the formationof an hcp structure, as it facilitates the oligomerization oflysenin (14).

Submitted April 2, 2013, and accepted for publication July 22, 2013.
*Correspondence: kobayasi@riken.jp
Editor: Simon Scheuring.
Ó 2013 by the Biophysical Society
0006-3495/13/09/1397/9 $2.00

Here, we report the spatiotemporal dynamics of lysenin
oligomers during their assembling on the SM/chol bilayer.
To monitor this process, we utilized high-speed atomic force
microscopy (HS-AFM). HS-AFM, which was recently
developed (15), has enabled the real-time imaging of the
dynamics of some membrane proteins at subsecond resolution (16–18). By using this state of the art technique for apore-forming toxin, we revealed (to our knowledge, forthe first time) the mechanism of the formation of an hcp
structure, which involves cooperative association/dissociation (i.e., the simultaneous association or dissociation ofseveral oligomers) and lateral diffusion of the toxin along
the membrane surface in milliseconds.

MATERIALS AND METHODS
Materials
Lysenin from earthworm, Eisenia foetida, was obtained from Peptide Institute (Osaka, Japan). Brain SM and chol (R99%) were purchased from
Avanti Polar Lipids (Alabama, AL) and Sigma (St. Louis, MO), respectively. Multilamellar vesicles (MLVs) of SM and SM/chol (1:1) were prepared from a homogeneous lipid mixture in chloroform. The thin lipid
film formed after evaporation of chloroform was hydrated with a solutionof phosphate-buffered saline (PBS, 10 mM, pH ¼ 7.5; Sigma) to a total
lipid concentration of 1 mM by heating to 55 C and vortex mixing. The
lipid suspensions were additionally sonicated in a bath sonicator at 38
kHz. The sonication duration was 10 min for SM suspension and 5 min
for SM/chol suspension. The obtained MLVs were frozen at 80 C andthawed at 55 C twice. Unilamellar vesicles were prepared by sonicationof MLVs at 20 kHz with an ultrasonic homogenizer, UH-50 from SMT(Tokyo, Japan), for 10 min. The vesicle size was measured by dynamic lightscattering using a Zetasizer Nano ZS (Malvern, Worcestershire, UK). The
size of the vesicles was found to be ~120 nm (see Fig. S1 in the Supporting
Material). Lysenin was dissolved in PBS to a final concentration of 15 mM.
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Visualization of lysenin oligomers on lipid bilayer
High-speed atomic force microscopy (HS-AFM) measurements to visualize
lysenin binding and assembling on SM-containing bilayer were performed
at Kanazawa University and also in part at Research Institute of Biomolecule Metrology (RIBM, Tsukuba, Japan) using the Nanoexplorer and Nano
Live Vision systems (RIBM). A sample stage made of quartz was placed on
the Z-scanner. A mica disk with a diameter of 1.5 mm was glued onto thesample stage. A detailed report by Uchihashi et al. (19) on HS-AFM setup isavailable. To prepare lipid bilayers, 1.5 mL of multilamellar or unilamellar
vesicle suspension was dropped onto the mica disk and incubated for 5 min
at 55 C. Mica surface was rinsed with pure water and placed onto the scanner. The surface was imaged in 70-mL PBS solution with cantilevers that
had a resonant frequency of 500 kHz in water and a spring constant of
0.1 N/m. For in-situ measurements, AFM imaging was started just after
introduction of lysenin diluted with a PBS solution to a concentration of
15 mM. At half-concentration, lysenin did not assemble into an hcp
structure. This could be due to its slow diffusion from bulk solution to
the vicinity of the membrane. The adsorption of lysenin on mica wasvisualized in situ at the same lysenin concentration, used for the observationof lysenin assembling on lipid bilayer. For preincubation experiments, 2 mLof diluted lysenin was dropped onto a mica-supported SM/chol bilayer
and incubated for 15 min at room temperature. HS-AFM imaging was
performed at room temperature.

Calculation of the average neighboring distance
We took the following processes to quantify the randomness of oligomer
positions: To begin, we generated a table of X-Y coordinates of all oligomer
centers in every AFM image, using a spot-readout software (GRAPHCEL,
http://www9.plala.or.jp/kobo333/). Then, the two-dimensional plots of theoligomer positions were subjected to two-dimensional numerical Fouriertransformation. The Fourier-transformed images are composed as the reciprocal-space lattice. This reciprocal lattice was converted to the real lattice.
The real lattice exhibits hexagonal symmetry, and the real hexagonal lattice
constant is hereafter designated as the average neighboring distance. The
randomness (i.e., the average displacement of oligomers from the lattice
points) was quantified by using the autocorrelation plots. The autocorrelation plots were obtained by calculating the two-dimensional displacement
vectors of all the pairs of oligomers that appear in the image of oligomer
X-Y coordinates. The autocorrelation plots, according to this definition,
are significant only within the first nearest plots from the origin. Overlaying
the two-dimensional real lattice (generated by the above-mentioned Fouriertransformation) onto the autocorrelation plots within the first-nearest range,
we can see that the autocorrelation plots are concentrated near the reallattice points. We calculated the distance between each autocorrelation
plot and the real lattice point nearest to it, and the obtained distanceswere averaged over one frame. This average distance is the standard deviation, which indicates the randomness of oligomer positions in the frame.

Molecular modeling
The model of lysenin was based on Protein Data Bank entry PDB:3ZX7(lysenin in complex with choline phosphate). The secondary structure of
N-terminal residues 1–10 was predicted by the homology modeling andthreading server I-TASSER without specified restraints (20–22), superimposed on N-terminal residues 5–10 of the x-ray structure and the four
missing N-terminal residues 1–4 were manually connected to the Lyseninmodel. The protonation state of the resulting model was predicted utilizing
the Hþþ server (23). The resulting model was parameterized utilizing theCHARMM C36 force field (24). The lysenin oligomers were assembled
either utilizing pore-forming proteins featuring a central b-barrel structureof varying size as templates (e.g., PDB:1UYN, PDB:1PHO, PDB:3SZD) or
manually. Subsequently, the oligomer assemblies were equilibrated for 1 ns
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Yilmaz et al.
at 310 K utilizing the NAMD 2.8 software package (25), before superimposition with AFM topology data utilizing custom-made VMD scripts (26).

Radial distribution function
The radial height distribution of stable hcp oligomers was established by
mapping the relative brightness in the vicinity of lysenin oligomers. The
ideal center of lysenin oligomers in the AFM images was determined via
a custom-made computer script. Usual computational approaches basedon particle identification procedures produced poor results due to thecentral cavity of lysenin oligomers. In total, the relative radial brightness
distribution in the vicinity of ~900 lysenin oligomers was analyzed andaveraged.

RESULTS AND DISCUSSION
Assembly of lysenin oligomers on SM/cholbilayerPreviously, using negative staining electron microscopy werevealed that lysenin oligomers assemble into an hcp structure when incubated with SM/chol liposomes (8). In thisstudy, we preincubated SM/chol bilayer with lysenin (seeMaterials and Methods) before imaging by HS-AFM.
Fig. 1, A and B, shows two- and three-dimensional imagesof the hcp arrangement of lysenin oligomers on the lipid
bilayer. In Fig. 1 A, the hexagon that appears in open representation indicates a single hcp unit. The average neighboring distance in the hcp structure (i.e., the distance
between the centers of two neighboring oligomers), estimated by two-dimensional Fourier transformation, was13.1 5 1.7 nm. The oligomers had an outer diameter of
10 nm with an inner pore of 5 nm, which was calculated using the radial distribution function (see Fig. S2). These
values are in good agreement with the diameters of thelysenin oligomers and the pore-like structures on SM/cholliposomes, visualized by electron microscopy (8). In a
recent study, the oligomeric assembly of lysenin was revealed by two-dimensional electron crystallography andthe oligomer diameter was estimated to be 11.2 nm withan inner pore of 5.7 nm (9). The oligomeric assembly wasconsidered to be a trimer or a hexamer. A hexamer can
provide a sufficient number of b-strands to form membranepores. The high similarity of our oligomer and pore diameters with De Colibus et al. (9) suggests the presence ofcomparable oligomerization state.
Whereas the oligomers of lysenin exhibit a uniform
lateral size, the height profile revealed the presence of two
distinct populations (Fig. 1 B). Due to the complete coverageof the observed membrane patches with lysenin oligomers,
unambiguous determination of the oligomer height proofed
to be challenging. To quantify the oligomer height, we
assumed that the darkest areas represent the lipid bilayer.
Based on this assumption, the height distribution in Fig. 1
C yielded two maxima: at 2.6 nm for the (darker) lower
and 5 nm for the (brighter) taller oligomers, representing
the majority of the population. The presence of lower-heightAssembling of a Toxin on Lipid Membrane

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FIGURE 1 AFM images of hexagonal closepacked (hcp) assembly of lysenin oligomers on
SM/chol (1:1) bilayer. Lysenin was preincubatedwith SM/chol (1:1) bilayer for 15 min and AFM
imaging was performed in PBS at a scan rate of
0.5 frames/s. (A) Two-dimensional height image.
(B) Three-dimensional height image. (Inset) Height
analysis for the three adjacent oligomers (indicatedby the open dashed line). (C) Histogram showing
the height distribution for the (dark) lower and(bright) taller oligomers. (D) Enlarged band-passfiltered image of the hcp unit indicated by a hexagon
in panel A. (Arrows in panels A and D show theincomplete oligomers and the linkages between
oligomers, respectively.)oligomers suggests the membrane insertion of lysenin. Previously, the oligomer height in the prepore and pore states of
the cholesterol-binding cytolysin, perfringolysin O, wasdetermined using AFM (27). It is thus possible that the taller
oligomers are prepores and the lower ones are pores,
although we cannot infer whether these pores are functional
channels. Based on the crystal structure of lysenin, it is speculated that, in the prepore state, the N-terminus of lysenin
partially spans the membrane through its specific binding
to SM in the outer leaflet of the bilayer (9). This partial insertion might induce a conformational change in the core oflysenin, leading to pore formation. As an alternative, theC-terminus might bind the membrane first whereas the
N-terminus is tilted upwards in the hexameric prepore
state (see Fig. S3). Subsequent insertion of the N-terminus
might induce the conversion from prepore to pore state.
In addition to the ring-like oligomers, incomplete arcshaped oligomers were occasionally detected (Fig. 1 A,
indicated by arrows). The average height difference
between ring-like and arc-shaped oligomers was~0.25 nm, negligible compared to the height distribution
shown in Fig. 1 C. The presence of lower arc-shaped oligomers denotes that extended insertion into the membrane canoccur before completion of oligomerization (28). Nevertheless, the growth of oligomers seems to be faster thaninsertion, because the majority of lysenin oligomers arecomplete rings of similar diameter and height.
The band-pass filtered image revealed bridge-likestructures between neighboring oligomers (Fig. 1 D, indicated by arrows). Similar structural features have also
been observed in oligomers of other toxins, such as VacA,
sticholysin II, and cholera toxin (3,4,29). These bridgelike structures have not been characterized yet. It is specu-

lated that they may be formed by either the N-terminus or
the C-terminus of the toxin. De Colibus et al. (9) suggested
that the N-terminus of lysenin partially penetrates themembrane and is not subjected to reconfiguration in the prepore state, leaving the three two-stranded hairpins in the
C-terminus as potential candidates to form the bridgelike structures. As of this writing, the possibility that theobserved structures were caused by the AFM tip during
imaging cannot be completely ruled out.
Reorganization of lysenin during assembling
We then followed the time course of the assembling of lysenin on the SM/chol bilayer in situ at a scan rate of 3.3
frames/s over a scan area of 250  250 nm (see Movie S1and Fig. S4 in the Supporting Material). Initially, there
were only a few unfused vesicles (see Fig. S4 A, marked
by asterisk) on the membrane and lysenin monomers, moving too fast to be localized under our experimental conditions. The unfused vesicles were generally observed inaddition to the fused lipid bilayer in the acquired images.
After 2 min, oligomer-like assemblies of lysenin slowly
started to appear. A direct comparison of these assemblies
with the oligomers in Fig. 1 A is difficult due to their unclear
size and shape. Consequently, the term ‘‘cluster’’ will be
used instead of ‘‘oligomer’’ to describe and discuss thein-situ observations. In this context, clusters might be theintermediate assemblies of lysenin (dimer, trimer, tetramer,
open-ring, etc.) or rapidly associating/dissociating oligomers. With the increasing cluster number on the membrane,
an hcp structure was formed in <5 min.
Next, we tracked the assembling of lysenin on the SM/chol bilayer at a larger scan area of 400  300 nm and a
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lower scan rate of 0.5 frames/s (see Movie S2 and Fig. S5).
In selected images of this movie (Fig. 2 A) it is possible tosee the overall change in the packing of the lysenin clusters
and the change in the growth direction of the cluster arrays.
In addition to the round-shaped clusters, rapidly moving
irregular-shaped monomers/clusters can be observed.
During the course of the reaction, most of the round-shaped

Yilmaz et al.

clusters assembled into an hcp structure. Aside from lysenin
assembling on the SM/chol bilayer, lysenin also adsorbed
directly on mica, the lowest area in Fig. 2 A. The particle
radius of the membrane-bound lysenin after full coverage(36 s) was found to be very similar to hcp oligomers (seeFig. S6, A and C). In contrast, the size analysis for lyseninon mica yielded a much broader distribution, indicating

FIGURE 2 (A) Time-lapse AFM images of lysenin clusters forming the hcp structure on SM/chol(1:1) bilayer. Lysenin was injected into PBS andassembling of lysenin on SM/chol (1:1) bilayer
was followed at a scan rate of 0.5 frames/s. Time‘‘0 s’’ is the starting time of the observation
period. Asterisks indicate the unfused vesicles.
(B) Schematic illustration of the AFM images inpanel A showing the growth of domains of lyseninclusters in different regions. Each lysenin cluster
is depicted in green. Small dots represent irregularshaped clusters, perhaps the intermediates, andlarge dots show round shaped, presumably the final
oligomers. Black and blue frames indicate thecontinuously growing and unstable-disordereddomains, respectively. Red, orange and pink frames
correspond to the unstable-ordered domains. In
black-framed region, the initially formed clusters
in hcp structure are stable and new clusters bindto this domain, enlarging the stable hcp structure.
In blue-framed region, clusters show a dynamic
behaviour and an hcp structure does not form. In
red, orange and pink-framed regions, clusters might
associate/dissociate or diffuse laterally to arrange
into an hcp structure. (C) Change in the numberof randomly forming and close-packed clusters
within the observation time.

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Assembling of a Toxin on Lipid Membranethat lysenin exists on mica as a mixture of monomers andirregular-shaped aggregates (see Fig. S6, B and C). The
difference in particle size distribution between lyseninaggregates on mica and lysenin clusters on the membranestrongly suggests that lysenin cannot form oligomers on
membrane-free surface as described previously by YamajiHasegawa et al. (8). For a better understanding of the formation of the hcp structure, each round-shaped cluster isdepicted as a green dot in Fig. 2 B. The green dots wereplaced manually on the clusters that could be easily recognized by eye. The frames in different colors highlight thegrowth of the domains of lysenin clusters. Assembling oflysenin on the SM/chol bilayer exhibits the following
distinctive features:1401

with a stable hcp assembly of lysenin. The overlay of images
at 0, 2, 18, and 36 s at an opacity of 50% shows the change inthe position of clusters (Fig. 3 A). The lower contrast of theoverlay compared to the image at 36 s (Fig. 2 A) indicatesthe change in the position of clusters before the formationof the stable hcp assembly. The high-contrast domains in
the overlay imply the reduced dynamics of clusters duringassembling. The domains of lysenin clusters within the black,
red, and orange frames were analyzed to find the average

Random formation of clusters

In the initial stages of assembling (0 s in Fig. 2 B), either
individual or arrays of clusters formed randomly in different
regions of the membrane surface. The randomly forming
clusters were more dynamic than the close-packed clusters.
Most of the randomly forming clusters dissociated ordiffused along the membrane. The time-dependent change
in the number of randomly forming and close-packed clusters is shown in Fig. 2 C. As the number of close-packed
clusters increased by time, randomly forming clusters gradually diminished. This suggests that, at first, lysenin forms
individual clusters, which in turn diffuse along the membrane surface to arrange into an hcp structure.
Cooperative association/dissociation of clusters

Within a few seconds, while new clusters formed, bothfilling the gaps within the arrays and forming new arrays,
some of the existing clusters either dissociated or diffusedalong the membrane surface (e.g., the areas indicated by
the red arrow in Fig. 2 B at 0 and 2 s).
Change in the direction of cluster arrays

In some of the regions, the direction of the cluster arrays
changed. The clusters that formed within the top-left framein red at 2 s were aligned in a different direction than thosewhich formed at 0 s (Fig. 2 B). The directions of the cluster
arrays within the red frame at 0 s and 2 s are indicated by theblack arrows.
Similar packing density of clusters in different domains

A small domain of stable lysenin clusters formed within theblack-framed region at 0 s and continued growing (Fig. 2 B).
In the red, orange, and pink-framed regions, lysenin clusters
initially formed unstable domains in which the clusters either
dissociated/reassociated or changed their positions. We
named these domains ‘‘unstable-ordered’’ due to theirdynamic behavior during the formation of the hcp structure.
The clusters in the blue-framed region were both unstableand disordered. Ultimately, the domains in different regions
combined and the membrane surface was fully covered

FIGURE 3 Change in the packing density of lysenin clusters. (A)Overlay of images at 0, 2, 18, and 36 s at an opacity of 50%. (B) Averageneighboring distance as a function of time for the continuously growing
region 1, and the unstable-ordered regions 2 and 3. (Colors in panel B correspond to the regions in black, red, and orange frames in panel A.)
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neighboring distance between clusters. The change in the
average distance between two neighboring clusters calculated for three different regions on the membrane surface isshown together with the standard deviation as a function oftime in Fig. 3 B. The analyzed regions are indicated inFig. S5. The average neighboring distance could be calculated only for the domains of lysenin clusters showing anordered arrangement. The average neighboring distance forthese ordered domains did not greatly change in either spaceor time. There was a decrease in the standard deviation as theordered domains grew larger. The higher standard deviation
at the initial period of assembling is due to the randomly
forming clusters around those showing an ordered arrangement. Most of the lysenin clusters formed rapidly reversible/irreversible arrays or domains with the same averageneighboring distance instead of random distribution on themembrane even during the initial period of assembling.
The arrangement of lysenin clusters into an hcp structure
seems to be dependent on the liquid-ordered phase of the
SM/chol bilayer (30). The same process was also followedon pure SM bilayer; however, we did not observe the formation of an hcp structure (see Movie S3; scan rate: 1 frame/s,scan area: 500  500 nm, real scan time: 3 min 38 s). In
contrast to SM/chol, SM membrane exhibits a gel-likebehavior at room temperature (31). Previously, we reported
that the binding of lysenin to SM and SM/chol membranes
was similar; however, oligomerization was facilitated by thepresence of chol due to the change in the membrane fluidity(14). Therefore, in our study we suggest that the observed
difference in the assembling of lysenin on SM and SM/chol membranes is due to the difference in the membrane
fluidity, which affects oligomerization and hence formationof a well-ordered structure.
Subsecond association/dissociation of lyseninclusters
The association and dissociation of lysenin clusters from theedge of a domain of close-packed clusters was followed at ahigher magnification (85  40 nm) and a higher scan rate (6.7
frames/s) for a more careful examination of the dynamics ofclusters during assembling into an hcp structure (Fig. 4 A andsee Movie S4). The lysenin clusters at the domain edge are
depicted as green dots (Fig. 4 B). The cluster, indicated by
the white arrow at 0 s, disappeared at 0.15 s. This might bedue to the dissociation of the cluster into monomers, dimers,
etc., or the detachment of the cluster from the domain. Monomers, intermediates, and free clusters appear as spikes in theimages and cannot be clearly distinguished due to their rapid
diffusion along the membrane surface. The neighboring clusters (indicated by red arrows) also started to dissociate at
0.3 s. However, at 0.45 s simultaneous reassociation of threeneighboring clusters occurred. The cooperative association/
dissociation of clusters is marked with white dashed-line
rectangles. The dissociation and reassociation of the cluster
Biophysical Journal 105(6) 1397–1405

FIGURE 4 (A) AFM images showing the dynamics of lysenin clusters on
SM/chol (1:1) bilayer. Association/dissociation of clusters at the domainedge was followed at a scan rate of 6.7 frames/s. (B) AFM images in panel
A with the schematic outline of the association/dissociation of clusters at thedomain edge. (C) Histogram showing the number of observations fordifferent residence times before dissociation.

at the location indicated by the white arrow occurred at
0.15, 0.6, and 1.2 s, and 0.45, 0.75, and 1.35 s, respectively.
The residence time before dissociation of the clusters at theAssembling of a Toxin on Lipid Membrane

domain edge was analyzed (Fig. 4 C). In most cases, clusters
dissociated within 0.15 s after binding to the domain. The
average residence time was found to be ~0.45 s. This is comparable to the residence times of bacteriorhodopsin trimers
and aquaporin tetramers, previously studied by HS-AFM(16,32). In a report by Yamashita et al. (16), examinationof the effect of tip velocity on the lifetime of the trimers ofbacteriorhodopsin revealed that the dynamics of trimers
was not dependent on tip velocity. Hence, we think that theassociation/dissociation of lysenin oligomers was notaffected by the tip. In contrast to the clusters at the edge ofthe domain, those within the hexagonal structure were relatively stable. The instability of the clusters at the domainedge may result from the smaller number of interaction sites.
The clusters within the hexagonal structure, which seem tobe the oligomers, are linked to each other through six interaction sites, rendering them more stable than those at theedges. Another reason might be the conformational energy
barrier between lysenin molecules in adjacent clusters (33),
leading to the breakage of the intermolecular interaction
between the clusters.
Lysenin clusters did not always follow the same reorganization pathway (see Fig. S7). After the binding of new clusters, some of the clusters within the same domain
dissociated at random locations, leaving arrays of clusters
aligned in different directions. This implies that the organization of lysenin clusters was not unidirectional before theformation of the stable hcp structure.
Subsecond changes in cluster height
Besides monitoring the fast association/dissociation of lysenin clusters, the change in cluster height was also monitored. In Fig. 5, A–C, the height change of a cluster within
an hcp structure is shown for a scan area of 85  40 nm
and a scan rate of 6.7 frames/s. The height profiles along
the yellow line, marked at 0 s in Fig. 5 B, are shown in
Fig. 5 C. The cluster indicated by the arrow in Fig. 5 B dissociated at 0.15 s, leaving a depression of 1.4 nm, and reassociated at 0.3 s at the same height with the adjacent cluster
along the yellow line. After 0.3 s, the cluster dissociation/
reassociation took place with an increase in height. The
sequential pattern of changes was as follows: there was anincrease of 2.8 nm in the cluster height at 0.75 s, followed
by a diffusion of the cluster along the membrane surface,leaving a depression of 2.5 nm in the hcp structure at 0.9s, and finally a filling of the gap, along with a decrease in
its height at 1.5 s. The diffusion or dissociation of clusters
is an indication of their instability. The same cluster underwent similar changes after an additional dissociation at
4.65 s, followed by the appearance of a brighter oligomer
at 4.8 s and a decrease in its height within a time intervalof <0.45 s. The depressions observed at 0.15, 0.9, and4.65 s were of different heights. This might be due to theincomplete dissociation of the cluster when the image was1403

captured. Because lysenin is a water-soluble protein andforms oligomers only in the presence of SM, the bindingof lysenin to the SM-bound lysenin and its oligomerizationon the first layer seems to be unlikely. Thus, the observationof the (brighter) taller oligomers just before the dissociation
and after association supports the possibility of conformational changes in lysenin rather than multilayer formation.
Considering the size of lysenin, which is ~10 nm in length,a change in its height up to 3 nm seems to be resulting fromthe reversible partial insertion of clusters into the membrane
by vertical collapse. Therefore, the height change might be
an indication of the transition from prepore to pore. However,
we do not know the penetration depth of the clusters into themembrane before the formation of a stable hcp structure. The
transition of a complete pore to prepore seems unlikely.
Therefore, partial and weak insertion of clusters into themembrane is more plausible. After the completion of a stablehcp structure, some taller oligomers (see Fig. S8, indicated byarrows) showed a decrease in height whereas most oligomers
kept the same conformation. We did not observe any increasein the height of lower oligomers within the stable hcp
structure.
CONCLUSIONSThis work demonstrates the assembling of lysenin on an
SM-containing membrane. The high-speed AFM images
revealed the dynamics of the association of lysenin clusters into an hcp structure. Initially, both individual andsmall domains of lysenin clusters formed randomly at
different locations on the membrane. Although some ofthese domains grew continuously, most of them reorganized by subsecond dissociation/reassociation of the clusters. In addition to reorganization of the clusters, thechanges in the cluster height could also be followed.
The subsequent increases and decreases in the cluster
height can be an indication of the conformational changesof lysenin, resulting in its vertical collapse and membrane
insertion. After the membrane surface had been fully
covered with close-packed clusters, the hcp structure stabilized. The liquid-ordered behavior of the SM/chol membrane facilitates the diffusion of the clusters along themembrane and the formation of the hcp structure. The
similar size of the clusters and their occasionally observed
porous structure imply the formation of rapidly reversible/
irreversible oligomers of lysenin during its assembling on
SM/chol bilayer. Our results suggest that the assembling oflysenin on the membrane into an hcp structure is dependent on the rapid association/dissociation and diffusionof lysenin oligomers.
SM-rich lipid clusters do not randomly distribute in cell
membranes. Instead, they form domains whose radius is
60–80 nm on the outer leaflet of the plasma membrane(12). Lysenin-treated red blood cells produce globular protrusions rich in lysenin as revealed by immuno-electron
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FIGURE 5 (A) AFM images showing the height change for a particular lysenin cluster on SM/chol (1:1) bilayer. The height change and the association/
dissociation of the cluster were followed at a scan rate of 6.7 frames/s. (B) AFM images in panel A with the schematic outline of the association/dissociationof the cluster within the hexagonal structure and the time-dependent change in the cluster height. (Pale green) Taller oligomers. (C) Height profiles (along theyellow line) in panel B. (Dashed line) Baseline at 0 s.

microscopy (8). These protrusions contain aggregates ofround-shaped structures whose diameter is 10–15 nm (8).
Although the pores are not clearly observed, it is speculated
that these aggregates correspond to the hcp structure.
Formation of the hcp structure is advantageous for pore
formation because hcp stabilizes the pores.
SUPPORTING MATERIAL

This work was supported by the Lipid Dynamics Program of RIKEN and
Grants-in-Aid for Scientific Research Nos. 22390018 and 25293015(to T.K.) from the Ministry of Education, Culture, Sports, Science andTechnology of Japan.

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