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Current Medicinal Chemistry, 2013, 20, ????-????
1
The Sensing of Membrane Microdomains Based on Pore-Forming Toxins
M. Skoaj1, B. Bakra1, I. Kriaj2,3,4, P. Maek1, G. Anderluh1,5 and K. Sepi*,1
1
Department of Biology, Biotechnical Faculty, University of Ljubljana, Ljubljana, Slovenia; 2Department of Molecular and Biomedical Sciences, Joef Stefan Institute, Jamova 39, 1000 Ljubljana, Slovenia; 3Department of Chemistry and Biochemistry, Faculty of
Chemistry and Chemical Technology, University of Ljubljana, Slovenia; 4Centre of Excellence for Integrated Approaches in Chemistry and Biology of Proteins, Ljubljana, Slovenia; 5National Institute of Chemistry, Ljubljana, Slovenia
Abstract: Membrane rafts are transient and unstable membrane microdomains that are enriched in sphingolipids, cholesterol, and specific proteins. They are involved in intracellular trafficking, signal transduction, pathogen entry, and attachment of various ligands. Increasing experimental evidence on the crucial biological roles of membrane rafts under normal and pathological conditions require new
techniques for their structural and functional characterization. In particular, fluorescence-labeled cytolytic proteins that interact specifically with molecules enriched in rafts are of increasing interest. Cholera toxin subunit B interacts specifically with raft-residing ganglioside GM1, and it has long been the lipid probe of choice for membrane rafts. Recently, four new pore-forming toxins have been proposed
as selective raft markers: (i) equinatoxin II, a cytolysin from the sea anemone Actinia equina, which specifically recognizes free and
membrane-embedded sphingomyelin; (ii) a truncated non-toxic mutant of a cytolytic protein, lysenin, from the earthworm Eisenia
foetida, which specifically recognizes sphingomyelin-enriched membrane domains; (iii) a non-toxic derivative of the cholesteroldependent cytolysin perfringolysin O, from the bacterium Clostridium perfringens, which selectively binds to membrane domains enriched in cholesterol; and (iv) ostreolysin, from the mushroom Pleurotus ostreatus, which does not bind to a single raft-enriched lipid
component, but requires a specific combination of two of the most important raft-residing lipids: sphingomyelin and cholesterol. Nontoxic, raft-binding derivatives of cytolytic proteins have already been successfully used to explore both the structure and function of
membrane rafts, and of raft-associated molecules. Here, we review these four new derivatives of pore-forming toxins as new putative
markers of these membrane microdomains.
Keywords: Cholesterol, equinatoxin, fluorescent probe, liquid ordered phase, lysenin, membrane microdomains, membrane raft, ostreolysin,
perfringolysin O, sphingomyelin.
INTRODUCTION
Cell membranes are composed of a great variety of different
lipids and proteins [1]. According to the fluid mosaic model [2], it
has been long assumed that all of these molecules are relatively
randomly distributed in the homogeneous fluid lipid bilayer. The
current model of cell membrane structure, however, presumes that
the different lipids are de-mixed in the membrane plane according
to their various chemical and physical properties, to form distinctive membrane microdomains. In particular, sterols and sphigomyelin (SM) are responsible for the separation of the membrane lipid
domains into separate co-existing liquid-disordered (ld) and liquidordered (lo) domains [3]. The domains characterized by the lo phase,
in which the lipids are tightly packed but can still undergo fast lateral mobility, are the basis for the formation of membrane rafts, cell
membrane entities that are enriched in sphingolipids, cholesterol,
and specific proteins [4, 5].
Membrane rafts are believed to be transient, dynamic and unstable membrane microdomains that can be stabilized on binding
ligand molecules [6-8]. Their composition is also believed to
change on a sub-millisecond scale [8]. They have been shown to
exist as nanoscale clusters [6, 9] of various sizes [10, 11]. Experimental evidence of the crucial biological roles of membrane rafts
has dramatically increased in the last 15 years. Rafts have been
shown to be involved in several important biological functions,
including exocytosis and endocytosis, signaling, pathogen entry,
and the attachment of a variety of molecular ligands [1, 4, 12-15].
In particular, their involvement in the transduction of various signals has been shown to be important under a variety of disease conditions; e.g. Alzheimer’s disease, Parkinson’s disease, cardiovascular and prion diseases, systemic lupus erythematosus, and acquired
immunodeficiency syndrome [16]. All of these findings have made
membrane rafts interesting targets for further structural and functional studies of cell membranes and their constituents, as well as
*Address correspondence to this author at the Department of Biology, Biotechnical
Faculty, University of Ljubljana, Vena pot 111, 1000 Ljubljana, Slovenia; Fax: +3861-2573390; Tel: +386-1-3203419; E-mail: [email protected]
0929-8673/13 $58.00+.00
the for the cure and prevention of diseases by pharmacological
approaches. The development of new techniques and approaches
that would allow suitable visualization of these membrane microdomains will therefore be of great use.
Detergent insolubility has long been considered as the main basis for membrane raft isolation. In contrast to membrane domains in
the ld phase, lipids in the lo phase are resistant to solubilization by
many detergents [17], and therefore membrane rafts have often
been operationally called ‘detergent-resistant membranes’ (DRMs)
[4]. However, DRMs should not be assumed to describe biological
rafts in size, structure, composition or even existence, as detergents
themselves have been shown to be able to induce the formation of
ordered domains [17, 18].
Membrane rafts are not easily visualized, due to their temporal
instability and small size [11]. Several modern microscopic methods have been used recently for the visualization of membrane microdomains, including scanning probe techniques, such as nearfield scanning optical microscopy and atomic force microscopy,
and optical techniques, like fluorescence correlation microscopy,
fluorescence resonance energy transfer (FRET), stimulated emission and depletion microscopy, and stochastic reconstruction microscopy [19]. Other recent attempts to visualize particular membrane lipids and/or lipid domains have used fluorescent lipid analogs or lipid-binding probes, which have included proteins and
antibodies [20]. In particular, non-toxic recombinant fluorescencelabeled derivatives of natural toxins are gaining interest, which
include pore-forming toxins (PFTs), that interact specifically with
molecules enriched in membrane rafts.
Upon interaction with a membrane, the PFTs spontaneously
change their native conformation to enable the formation of transmembrane pores that consist of several protein monomers [21]. In
several cases, membrane rafts have been suggested to act as concentrating platforms that can promote the oligomerization of these
PFT proteins. Several PFTs have been found to interact with different molecules that are enriched in DRM domains that fulfill the
criteria of membrane rafts; e.g. SM, cholesterol, ceramides, gangliosides, or the glycan core of glycophosphatidylinositol (GPI)© 2013 Bentham Science Publishers
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Current Medicinal Chemistry, 2013, Vol. 20, No. 1
Skoaj et al.
the amino acids that participate in the binding of this headgroup
(labeled in green on Fig. 1) [44]. These amino acids are conserved
in the actinoporins and thus it appears that the mode of headgroup
binding is conserved across the actinoporins family [46].
anchored proteins [7, 20, 22-30]. Among the non-toxic fluorescent
derivatives of such natural toxins, the cholera toxin B-subunit (CTB) binds to the raft-enriched ganglioside GM1, and it has long been
the only such probe for membrane rafts [31]. Recently, however,
four new derivatives of PFTs that can bind to membrane SM and/or
cholesterol have been explored as new putative markers of membrane microdomains, and these are reviewed here (Table 1).
Formation of actinoporin pores proceeds through several steps,
which include membrane binding, transfer of the N-terminal region
to the membrane, oligomerization of several monomers, and the
pore formation itself, which brings the N-terminal regions of the
individual monomers across the membranes to form the final pore
[38-40, 45].
SPHINGOMYELIN-BINDING PORE-FORMING TOXINS
Sphingomyelin (N-acylsphingosine-1-phosphorylcholine, SM)
is a relatively abundant sphingolipid constituent of vertebrate cell
membranes that is mostly located in the outer leaflet of the plasmalemma [32]. In some cells, the SM concentration can account for
up to 60 mol% of their total phospholipid content; e.g. in erythrocytes, and ocular lens, peripheral nerve tissue and brain cells [33].
SM shares a phosphocholine moiety with phosphatidylcholine (PC),
whereas its ceramide backbone is specific for SM and glycosphingolipids. As well as its structural role in cell membranes, SM is
important in intracellular signaling pathways. Hydrolysis of membrane SM by activated sphingomyelinases leads to the formation of
ceramide, which serves as a secondary messenger in sphingomyelinase-dependent signal transduction [34], and can induce membrane endocytosis and blebbing [35]. Membrane SM has also been
shown to be involved in some disease conditions, e.g. in NiemannPick disease type A, an autosomal recessive disorder that arises due
to sphingomyelinase deficiency and that results in an overaccumulation of SM in cell membranes [36]. Membrane SM has
also been suggested to act as a non-receptor-based gate for the entry
of low-density lipoprotein, and thus to have a role in atherosclerosis
[37]. All of these observations make SM an interesting target for
specific probes that would allow further functional studies of cellular SM and its behavior in cell membranes under conditions of
health and disease. As SM is significantly enriched in membrane
rafts, specific probes that would sense higher local distributions of
SM would also be very important in structural and functional studies of membrane microdomains. In this regard, the PFTs from sea
anemones and earthworm coelomic fluid are the most promising
candidates for development of SM-specific protein-based probes.
The membrane-binding region of the actinoporins was defined
by functional studies [45, 47-51], which also provided insight into
the molecular recognition of SM by these PFTs. Surface plasmon
resonance and lipid dot-blot analyses have shown that EqtII binds
SM directly and specifically, and not any of the other related lipids,
like PC, dipropionylPC, or lysoPC [46]. The labeling of EqtII with
19
F on tryptophans indicated for the first time that the tryptophan at
position 112 in EqtII participates in the direct recognition of SM
[52]. Mutagenesis of this tryptophan, and of its neighboring Tyr113
residue, has shown that these two residues are the most important
ones for SM recognition [46]. Tyr113 and Trp112 of EqtII are located on a broad loop at the ‘bottom’ of the molecule (see Fig. 1),
in close vicinity to the phosphocholine binding site, and this structure might represent a generic three-dimensional structural motif
that can bind to a single SM molecule. Thus it appears that the discrimination between SM and PC occurs in the region directly below
the phosphorylcholine headgroup [46]. The preferential interaction
of EqtII with SM has also been inferred from 31P and 2H solid-state
nuclear magnetic resonance studies that used large unilamellar
vesicles composed of various mixtures of PC, SM and cholesterol
[53, 54].
To test the use of EqtII as a marker for cellular SM, a green
fluorescent protein fusion protein (EqtII-GFP) was produced and
expressed as a His6 protein in the Escherichia coli Rosetta (DE3)
pLysS strain. Addition of GFP to the C-terminus of EqtII did not
alter its SM-specific membrane activity. The EqtII-GFP fusion
protein was found to bind to the plasma membrane of MDCK II
cells when added extracellularly. Furthermore, the apical side of the
polarized MDCK cells was stained, but not the basolateral, in
agreement with the canonical cellular SM distribution [55]. When
EqtII-GFP was expressed intracellularly within the same cells, a
punctate perinuclear staining was revealed (Fig. 1). EqtII-GFP colocalized with markers for the Golgi apparatus, mainly at the cisGolgi, and did not co-localize with markers for the nucleus, endoplasmic reticulum, or mitochondria [55]. Recent investigations [56]
have shown that membrane binding of EqtII is rapidly accompanied
by extensive membrane reorganization into microscopic domains
that resemble coalesced membrane rafts, with the concomitant
blocking of endocytosis. Cholera toxin subunit B (CT-B) was also
seen to co-localize within these domains. In addition, sticholysin II
co-localized with raft-associated membrane fractions, and its lytic
activity was shown to be abolished by incubating CT-B with raftlike lipid vesicles (SM: dioleoylphosphatidylcholine [DOPC]: cholesterol; 1:1:1) [22]. Furthermore, with large unilamellar vesicles
The actinoporins are potent PFTs that have been purified from
sea anemones [38-40]. They are 20-kDa cysteine-less proteins that
have permeabilisation activities towards membranes that contain
SM. The most studied of the actinoporins are equinatoxin II (EqtII),
from the sea anemone Actinia equina, and sticholysins I and II,
from Stichodactyla helianthus [41]. The three-dimensional structure
of the actinoporins shows a tightly folded -sandwich that is
flanked on two sides by -helices. The N-terminal region of approximately 30 amino-acid residues contains one of the -helices,
and this is the only part of the molecule that can be dislocated from
the body of the molecule without disrupting its general folding [4244]. This N-terminal flexibility has been shown to be crucial for the
pore-forming activities of the actinoporins [45].
The structure of sticholysin II in a complex with phosphocholine, the headgroup of both SM and PC, allowed the definition of
Table 1.
Representative Overview of the Cytolytic Proteins and Their Lipid Binding Components that are Enriched in Membrane Rafts
Protein
Organism
MW
(kDa)
Binding Specificity
Reference
Equinatoxin II
Actinia equina
20
SM
Bakra et al., 2008.
Lysenin
Eisenia foetida
33.4
SM
Ishitsuka et al., 2004.
Perfringolysin O
Clostridium perfringens
54
Cholesterol
Ohno-Iwashita et al., 2004.
Ostreolysin
Pleurotus ostreatus
15
Cholesterol/SM
Sepi et al., 2004.
CT-B
Vibrio cholerae
11.5
GM1
Bacia et al., 2004.
Pore-Forming Proteins As Raft Markers
and lipid monolayers, the co-existence of the so+ld, and lo+ld phases
was found to favor the membrane insertion of EqtII, and epifluorescence measurements showed that the interfaces between co-existing
lipid phases can function as preferential binding sites for EqtII [57].
The binding of fluorescently labeled EqtII to the domain boundaries
was shown also in a giant unilamellar vesicle model system [58].
Current Medicinal Chemistry, 2013, Vol. 20, No. 1
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gomerization [62, 63]. Membrane-bound lysenin forms SDSresistant oligomers of about 250 kDa, and forms pores with a diameter of 3 nm [64]. Hemolytic activity occurs at concentrations of
a few nanograms of lysenin per milliliter [60]. Electron microscopy
of lysenin bound to equimolar SM: cholesterol liposomes has revealed the presence of non-uniformly distributed honeycomb-like
regular hexagonal structures on the membrane [64].
Lysenin and the homologous proteins LRP-1 and LRP-2 are secreted through the dorsal pores of the earthworm, and they are believed to act in the protection of E. foetida from predatory vertebrates [65]. Recombinant lysenin has also been shown to be active
against Bacillus megaterium (an E. foetida pathogen) in the nanomolar range, but not against other soil bacteria. The lysis of bacterial cells was shown to be independent of pore-formation [66]. Specific binding of lysenin to SM has been confirmed using several
approaches, including enzyme-linked immunosorbent assays, thinlayer chromatography immuno-staining, FRET, and SM-containing
lipid vesicles [60, 66]. Interactions of lysenin with related SM analogs (e.g. sphingosine, ceramide, sphingosinphosphorylcholine), or
with other phospholipids or glycosphingolipids, were not observed
[60, 67]. Experiments with planar lipid membranes also showed the
formation of voltage-dependent large conductance channels in a
SM-dependent manner [68, 69].
Fig. (1). Labeling of cellular SM by expressing EqtII-GFP in cells. EqtIIGFP and its mutants were expressed intracellularly in MDCK II cells. The
model of EqtII structure shows the position of the SM headgroup (sticks and
surface presentation) and the residues that participate in its binding (green).
The two residues responsible for SM specificity, Trp112 and Tyr113, are
shown in blue. Residues at position 8 and 69 are also shown. Change of
these two residues render EqtII hemolytically inactive upon disulfide formation. EqtII-GFP shows a spotted perinuclear distribution, as opposed to GFP
staining, which was used as the control. Mutation of Trp112 to alanine
changed the labeling pattern, which resembled that of the GFP control. The
plasma membrane was stained with rhodamine-conjugated wheat germ
agglutinin. Scale bar, 20 μm. Adapted from [55], with permission of the
American Society for Biochemistry and Molecular Biology.
It appears that EqtII, and the actinoporins in general, can be
used for probing for SM and might thus be helpful in other cellbiology studies. Moreover, as shown recently, treatment of human
immunodeficiency virus (HIV)-1 viral particles with the SMspecific toxins, EqtII and lysenin (see below), decreased the infectivity of the virus. The magnitude of this inhibition was dependent
on the SM content of the cells from which the HIV-1 was derived
[59].
Lysenin is a SM-specific cytolytic and hemolytic protein from
the coelomic fluid of the earthworm Eisenia foetida [60]. Lysenin is
a 33,440 Da protein composed of 279 amino-acid residues. When
analyzed with SDS-PAGE, lysenin behaves like a 41 kDa protein,
which is probably due to its glycosylation [61]. Recently determined 3D structure revealed that lysenin is a two-domain protein.
The C-terminal domain is composed of a -trefoil motif, while the
N-terminal domain is rich in -sheet and similar to proteins of
lysenin-like PFTs [62]. The structure revealed also binding site for
phosphocholine at the C-terminal domain and SM in the N-terminal
domain. The lysenin membrane binding thus begins with the interaction of its C-terminus with the membrane, followed by its Nterminus, which is necessary for its irreversible binding and oli-
The importance of membrane SM for lysenin activity is also reflected in its maximal activity towards sheep erythrocytes, which
contain a very high content of SM [64]. Several immunofluorescent
studies of cell-bound lysenin have been performed to determine its
potential as a specific SM marker. Lysenin binding to fibroblasts
from a Nieman-Pick disease type A patient yielded strong uniform
staining by lysenin, and this was impaired by pre-incubation of
lysenin with SM-containing lipid vesicles [60]. Furthermore, sphingomyelinase-pretreated CHO cells, and cells with defective sphingolipid synthesis, were resistant to the cytolytic action of lysenin,
which suggests that lysenin can be used as a tool for isolation of
SM-deficient cell mutants; e.g. cells deficient in sphingolipid synthesis or in ceramide transport from the endoplasmic reticulum to
the Golgi apparatus [70]. Lysenin has also been proposed as a tool
in de/remyelinization studies in the brain, due to its binding to the
surface of oligodendrocyte lineage cells that show increased membrane SM content during cell development [71].
However, it soon became clear that the presence of other lipids,
for example glycosphingolipids, can significantly influence lysenin
membrane activity. Epithelial cell lines (e.g. MDCK cells) that are
enriched in glycosphingolipids in their apical domains are highly
sensitive to lysenin when added from the basolateral side, although
they are resistant if lysenin is applied apically. Model membrane
systems have indicated that glycolipids alter the local density of
SM, and consequently decrease lysenin binding [72]. Another
molecule that can significantly affect lysenin binding is cholesterol.
The binding of lysenin to SM: cholesterol vesicles, and their permeabilization, were significantly enhanced when the vesicles contained at least 30 mole% cholesterol [60], which indicated the importance of a specific membrane phase for the activity of lysenin.
Ishitsuka and Kobayashi [73] further studied the effects of cholesterol in terms of the membrane activity of lysenin on SMcontaining membranes. The incubation of lysenin with SM-based
liposomes enriched in cholesterol inhibited the hemolytic activity of
lysenin. However, the presence of cholesterol in SM-rich membranes did not affect lysenin binding, and even promoted its oligomerization. The authors thus suggested that the effects of cholesterol might be linked to the fluidization of membrane SM, and consequently to a decreased number of membrane-bound lysenin
monomers, which would result in facilitated oligomerization. Similar observations were also made when lysenin binding to SM-rich
membranes was studied using a surface plasmon resonance technique. Addition of 50 mole% cholesterol to SM-membranes increased lysenin binding in comparison to pure SM membranes, but
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Current Medicinal Chemistry, 2013, Vol. 20, No. 1
did not affect the kinetic parameters (with KD values of 5.3 nM and
8.6 nM, respectively), which suggests that cholesterol can change
the distribution of SM in the membrane, and thus enhance its accessibility to lysenin [60]. Furthermore, surface plasmon resonance
analyses have indicated that only wild-type lysenin binds efficiently
to raft SM (e.g. that the binding is enhanced on SM/DOPC,
SM/DOPC/cholesterol and SM/dipalmitoylphosphatidylcholine [DP
PC]/cholesterol membranes, where phase separation occurs), while
the Trp20 mutant of lysenin loses this ability, and does not discriminate between free and clustered SM molecules [63].
The use of a fluorescent fused recombinant lysenin (His-Venuslysenin) further confirmed that lysenin not only specifically
‘senses’ SM molecules, but is particularly sensitive to the lateral
distribution of SM within the membrane. The binding of HisVenus-lysenin to SM plus C18:1-PC liposomes, where lipid immiscibility is observed and SM clusters are formed, is much higher
compared to its binding to SM plus C16:0-PC liposomes, where the
lipids are more or less homogeneously mixed. The fluorescence
was observed in aggregates, which suggests that lysenin binding
depends on the local concentration of SM (Fig. 2). Isothermal titration calorimetry revealed that one lysenin molecule can bind five
SM molecules [72]. This was also the first study in which lysenin
was proposed as a probe for the investigation of membrane rafts
and their heterogeneity. Studies that used a truncated lysenin mutant confirmed that this can also be used. This mutant lysenin,
which consists of the minimal amino-acid fragment necessary for
SM recognition, was tagged with GST and shown to be non-toxic
[74]. Double staining of Jurkat T cells with this GST-tagged truncated lysenin and CT-B showed no co-localization of these two
fluorescent probes, which suggested that SM-rich domains are spatially distinct from GM1-rich domains [74]. Indeed, in SMcontaining liposomes, the presence of GM1 abolished the binding of
Venus-lysenin, while the binding of CT-B was not affected by the
presence of SM [74], which implies that CT-B recognizes its target
(GM1) irrespective of the lipid distribution, while lysenin does not.
The absence of lysenin and CT-B co-localization was further confirmed by immunofluorescent studies. Similar results, indicating
different locations of cholesterol- and SM-rich microdomains on
the plasma membrane, were obtained using non-toxic derivatives of
lysenin and perfringolysin O as respective probes for membrane
SM and cholesterol. Both proteins were fused with a photoswitchable fluorescent protein, Dronpa, which allows the visualization of stained membrane microdomains by using the superresolution microscopy, PALM [75]. As well as shedding some light on
the structural heterogeneity of membrane rafts, the use of fluorescent lysenin has provided evidence that structurally different rafts
also show functional diversity. In Jurkat T cells, the cross-linking of
SM showed that SM-rich domains provide a functional signaling
cascade platform that is distinct from those characterized by T cell
receptors or by G M1 [74]. Observations performed by using the
superresolution microscopy and fluorescently labeled lysenin revealed the involvement of the SM-rich domains in the progression
of cytokinesis [76]. Very recently, the structural and functional
heterogeneity of SM membrane pools was inspected by comparatively staining COS-1 cells with lysenin and EqtII [77]. The use of
both SM probes revealed the existence of at least three different SM
pools in plasmalemma: the first stained only with lysenin, the second stained only with EqtII, and the third that could be labeled by
both probes. The same heterogeneity was found in intracellular
membranes, where lysenin could be found in late endosomes, while
EqtII stained both late and recycling endosomes.
CHOLESTEROL-BINDING PORE-FORMING TOXINS
Cholesterol is another abundant lipid in the membranes of vertebrate cells. Cholesterol can mediate membrane fluidity and can
increase the ordering of membranes through the formation of the lo
phase and the consequent formation of segregated lateral domains
Skoaj et al.
in the bilayer plane that have specific properties (i.e. membrane
rafts). Cholesterol is also involved in several important cell functions, as for example mediation of signal transduction, and exocytosis and endocytosis [78]. It is also a precursor in the synthesis of
steroid hormones, oxysterols, bile acids and neuroactive steroids
[79]. Any alterations in cholesterol metabolism can thus be reflected in a plethora of pathological conditions. This can cause
common diseases such as atherosclerotic cardiovascular syndrome,
or result in rare genetic malformation diseases, like Smith-LemliOpitz syndrome, lathosclerosis, desmosclerosis, chondroplasia
punctata, and congenital hemidysplasia [79]. Impaired cholesterol
trafficking can also lead to hyper-accumulation of cholesterol
within lysosomes and late endosomes, like in Niemann-Pick syndrome type C [80], and cholesterol also appears to be involved in
the pathogeneses of Alzheimer’s disease and Parkinson’s disease
[81]. Molecules that can be used to label membrane cholesterol are
thus extremely interesting from several points of view, as they
might allow studies of the distribution of cholesterol in the membrane plane, and to follow its intracellular trafficking and involvement in different cellular processes under normal and pathological
conditions.
Fig. (2). Influence of local sphingomyelin density on lysenin binding to
giant unilamellar vesicles. Confocal images of fluorescent His-Venuslysenin binding to GUVs composed of SM plus C18:1-PC, (molar ratio 3:7)
which show lipid immiscibility and the formation of SM clusters (A), and to
GUVs composed of SM plus C16:0-PC (molar ratio 3:7), in which the lipids
are more or less homogeneously mixed (B). Bar, 5 μm. Reproduced from
[127], with permission of Elsevier.
For a long time, the polyene antibiotic filipin was used as a
membrane marker for cellular cholesterol. Filipin can form complexes with free and membrane-embedded cholesterol, which can
be visualized upon excitation by UV light using fluorescent microscopy. The use of filipin as a membrane raft marker has several
drawbacks, however. First, filipin is only suitable for staining of
fixed cells, as otherwise it is cytotoxic. Secondly, filipin binds to
membrane cholesterol indiscriminately [82, 83 Fig. 3], which is
reflected also by the demonstration that after removing 30% cholesterol from membranes using methyl--cylcodextrin (MCD), filipin
binding is still significantly retained [82, Fig. 3].
Among other non-protein molecules that are suitable for direct
imaging of cholesterol and its trafficking, there is also a series of
fluorescent cholesterol analogs. Dehydrocholesterol is a naturally
occurring fluorescent cholesterol derivative that can undergo rapid
transbilayer movement, and it has been used for the study of trafficking of sterols [84, 85]. Currently, novel non-ionic amphipathic
cholesterol analogs, poly(ethylene glycol)cholesteryl esters (PEGcholesterols) are gaining much interest for the labeling of membrane cholesterol. These PEG-cholesterols show cholesterol-like
characteristics, but they are also water-soluble and only weakly
cytotoxic. Among these, fluorescein-tagged PEG-cholesterol
(fPEG-cholesterol) has been the most frequently used. fPEGcholesterol preferentially partitions into cholesterol-rich membranes, and co-localizes with the filipin fluorescence, as well as
with the membrane raft markers CD59 and GM1 [86]. fPEG-
Pore-Forming Proteins As Raft Markers
cholesterol was thus shown to be suitable for the study of the dynamics of cholesterol-rich membranes. In living cells, it is distributed into the outer membrane leaflet [86], and can be slowly internalized by a clathrin-independent pathway into endosomes, although only at low concentrations and at 37 °C.
Fig. (3). Fluorescence microscopy images of BC and filipin binding to
A431 cells. A431 cell monolayers were incubated with serum-free Dulbecco’s modified Eagle’s medium for 2 h, fixed, and then incubated with
either protease-nicked or biotinylated perfringolysin O (BC; (A)) or filipin
(B). BC-treated cells were then incubated with cy3-avidin. Left panels:
fluorescence staining of cells after incubation with perfringolysin O (A) or
filipin (B). Right panels: BC and filipin binding after treating the cells with
5 mM 2-hydroxypropyl--cyclodextrin. Depletion of cholesterol completely
abolished BC binding, whereas filipin binding was retained significantly.
Reproduced from [82], Copyright (2001) National Academy of Sciences,
USA.
Perfringolysin O (-toxin) is a cytolysin from the Grampositive bacterium Clostridium perfringens. Together with some
other known cytolysins and virulence factors, like streptolysin O
from Streptococcus pyogenes, pneumolysin from Streptococcus
pneumoniae, and listeriolysin O from Listeria monocytogenes, perfringolysin O belongs to a larger group of cholesterol-dependent
cytolysins [87-89]. All cholesterol-dependent cytolysins share a
high degree of amino-acid sequence similarity, and with the sole
exception of intermedialysin, they show specific binding to free and
membrane-embedded cholesterol [90]. It appears that cholesteroldependent cytolysins recognize the cholesterol hydroxyl group, as
shown by surface plasmon resonance for listeriolysin O [91], using
a motif of two conserved amino-acid residues (Thr490 and Leu491
according to perfringolysin O numbering) [92]. Representative
cholesterol-dependent cytolysins are well-characterized in terms of
their structure-function relationships. The perfringolysin O monomer consists of four domains (D1-D4). The C-terminus of the D4
domain has a tryptophan-rich motif that is responsible for the initial
recognition of membrane cholesterol. After the initial binding, 2050 toxin monomers oligomerize to form a pre-pore structure. This is
followed by the vertical collapse of the D2 domain, and the translocation of the D3 domain into the membrane bilayer, causing its
structural rearrangement and insertion of the perfringolysin O oligomer into the membrane, finally forming the transmembrane pore
[93].
Current Medicinal Chemistry, 2013, Vol. 20, No. 1
5
This specific interaction of perfringolysin O with membrane
cholesterol has stimulated a series of studies on the use of perfringolysin O as a marker for membrane cholesterol [94]. To date, several non-toxic perfringolysin O derivatives with retained cholesterol
binding have been produced. C and its methylated form, MC,
were obtained by limited proteolysis of perfringolysin O between
the 144-145 amino-acid residues, and they exist as a complex of 38
kDa and 15 kDa proteins. The binding to erythrocytes of both C
and MC is comparable to that of the native perfringolysin O; however, C and MC are not hemolytic, and do not oligomerize up to
20 °C and 37 °C, respectively [95, 96]. One of the most used of the
perfringolysin O derivatives is known as BC, which also has the
same binding properties as the native protein, but which is lytic
only above 37 °C. BC was obtained by proteolysis and biotinylation of perfringolysin O, and it is structurally comparable to biotinylated C. When coupled with FITC-conjugated avidin, BC
can be used for staining membrane cholesterol without the need for
antibodies [97]. Analyses of BC binding to erythrocytes and to V79 and HUVEC cells by flow cytometry and confocal microscopy
have showed that BC is a very suitable probe for visualization of
membrane cholesterol and for the evaluation of the topology and/or
distribution of cholesterol [97]. Further investigations of BC
membrane binding have revealed that BC is not only a suitable
probe for cholesterol, but also for the distribution of cholesterol in
cell membranes, as it interacts with membrane domains that have
higher local concentrations of cholesterol Fig. (3). Binding of BC
was shown to be enriched in low-density membrane fractions that
contain cholesterol, SM, and Src family kinases, thus fulfilling the
criteria for membrane rafts. This binding was retained even when
the raft-like domains were isolated without detergent extraction;
however, depletion of approximately 30 mole% cholesterol using
MCD abolished the binding of BC [82, Fig. 3]. Furthermore,
BC co-localizes with the NAP-22 protein, which is a major component of brain-derived rafts [98], and which decreases in aged
cells, suggesting that the density of cholesterol-rich membrane rafts
decreases during cell aging [99]. The labeling patterns of NAP-22
and caveolin do not coincide [100], which suggests that BC binds
to non-caveolar rafts, and thus reinforces the hypothesis of membrane raft heterogeneity [5, 6, 10, 101, 102].
This hypothesis is further supported by electron microscopy
analyses of membrane vesicles recovered in raft fractions of cells
previously incubated with BC, which has shown that not all of the
raft-fraction vesicles associate with BC [28]. BC coupled with
fluorescent avidin or colloidal gold-avidin has also been used as a
probe to analyze the distribution of membrane cholesterol by fluorescence and electron microscopy [97, 100, 105]. However, due to
the fluorescence of avidin, this was not suitable for real-time imaging of dynamic raft movements in living cells. Therefore, smaller
constructs of perfringolysin O have been prepared and tested for
their cholesterol binding and specificity.
The D4 domain is the -sheet-enriched domain of cholesteroldependent cytolysins that is located at the C-terminus, and it represents the minimal toxin fragment that binds to cholesterol with high
affinity. Surface plasmon resonance has revealed that the membrane
binding of D4 and perfringolysin O are similar. Thus, it has been
shown that the D4 domain retains its stable structure, selectively
binds cholesterol in membrane rafts, and is recovered in the lowdensity membrane fraction [106]. Finally, enhanced green fluorescent protein-tagged D4 (EGFP-D4) has been produced and used for
staining of MOLT-4 cells. Here, the EGFP-D4 membrane binding
was similar to that of Alexa-labeled D4, so the fusion with EGFP
does not influence the binding of the D4 domain. Depletion of cellular cholesterol with MCD abolishes this binding. Thus, EGFPD4 has been proposed for the monitoring of the distribution and
movement of cholesterol in cell membranes [106].
6
Current Medicinal Chemistry, 2013, Vol. 20, No. 1
Recently, new insights into the interactions of perfringolysin O
with membrane cholesterol have been published. Nelson et al. [93]
showed that the binding of perfringolysin O does not correlate with
the ability of certain sterols to promote the formation of rafts. Using
steady-state intrinsic tryptophan fluorescence with perfringolysin O
and artificial lipid vesicles with different compositions, Nelson et
al. [93] showed that tightly packed phospholipids and sterols in the
lo phase weaken the perfringolysin-O-membrane interactions. Similarly, phospholipids with smaller headgroups were shown to increase the perfringolysin O membrane activity. These two phenomena can be explained by the ‘umbrella model’ of lipid-raft structure;
i.e. the shielding of cholesterol by the phospholipid headgroups.
Overall, the insertion of perfringolysin O into the membrane was
shown to be triggered in a more loosely packed lipid environment.
Further investigations on membrane binding of perfringolysin
O to lipid vesicles have shown that the presence/ absence of membrane rafts does not correlate with perfringolysin O binding (e.g.
increased SM content in cholesterol-rich membranes decreases
perfringolysin O binding) [107]. Perfringolysin O binding is triggered only when the cholesterol concentration exceeds the association capacity with phospholipids, whereby the excess cholesterol at
the membrane surface is then free to associate with the perfringolysin O. Perfringolysin O thus appears to bind to cholesterol that
is membrane embedded, yet surface exposed. Similar trends, e.g.
the prerequisite of at least 30-40 mol% of membrane cholesterol for
effective binding of perfringolysin O, have also been observed with
listeriolysin O [91]. More recent investigations on the raft affinity
of perfringolysin O have suggested that perfringolysin O binds to
raft-associated domains, but is translocated to less-ordered membrane domains upon establishment of its transmembrane pore complex [108]. Very recently also, it was shown that mutations in the
amino-acid residues around the conserved Cys459, which is located
in the membrane-interacting domain D4, can alter the susceptibility
of perfringolysin O in terms of the cholesterol threshold that is necessary for perfringolysin O binding. Engineered mutants can bind to
artificial and biological membranes that have much lower cholesterol mole% than the native protein, reinforcing their future use as
molecular probes for studying the distribution and dynamics of
membrane cholesterol [109].
Ostreolysin is a 15-kDa acidic protein from the edible mushroom Pleurotus ostreatus, and it belongs to the aegerolysin protein
family that currently includes over 300 similar proteins from bacteria,
fungi and plants [110]; their exact biological roles are not known.
Mushroom aegerolysins are suggested to be involved in the
formation of mushroom fruit bodies [111, 112], while the
aegerolysins from bacteria and filamentous fungi might be involved
in host-pathogen interactions [110]. Some aegerolysins, including
pleurotolysin A [113] and its isoform ostreolysin (our unpublished
data), are membrane binding components of binary cytolysins,
along with another ca. 59 kDa protein with a membrane attack
complex/perforin (MACPF) domain. Our studies on the membrane
binding of ostreolysin [114, 115] have shown that ostreolysin specifically interacts with natural and artificial lipid membranes that
are highly enriched in cholesterol and SM or, alternatively, cholesterol and a fully saturated glycerophospholipid. In contrast to other
putative raft markers that bind the pure cholesterol, like perfringolysin [90] or SM, like the actinoporins [46] and lysenin, [60],
ostreolysin can recognize only mixtures of these two lipids, particularly with an excess of cholesterol, which suggests that it has a very
specific interaction with membrane rafts. This has been reinforced
by several studies that have shown specific interactions of ostreolysin with cholesterol. SM liposomes, ostreolysin binding to the
DRMs of both cells and equimolar cholesterol. SM liposomes
[115], and inhibition of ostreolysin binding after treatment of cell
membranes with MCD [101]. Immunofluorescence studies on the
binding of low concentrations of ostreolysin to the membranes of
chondrocytes [116] and of CHO cells [101, Fig. 4a] have shown
Skoaj et al.
binding and clustering of ostreolysin, which further suggests the
association of ostreolysin with distinct membrane microdomains.
As in the case of lysenin [74] and perfringolysin [100], double immunolabeling experiments have shown that there is no colocalization of ostreolysin and other raft markers, like caveolin and
CT-B [101], Fig. 4a which suggests that there are functionally and
structurally different raft-like regions of the plasma membrane.
These raft-like regions would comprise at least two lipid microdomain types: those containing (caveolae) or lacking caveolin [5].
Furthermore, by comparing the membrane activities of various
raft-binding cytolysins, it can be seen that ostreolysin needs a
higher local cholesterol concentration and a higher degree of membrane ordering for effective binding. In contrast to most other putative raft-binding cytolysins, which readily interact with vesicles
composed of equimolar mixtures of SM, cholesterol and palmitoyloleoyl-PC (POPC) or DOPC, or with membranes composed of
equimolar ratios of cholesterol and DOPC [22, 27], the presence of
even lower ratios of monosaturated or disaturated PC in equimolar
SM, cholesterol liposomes dramatically diminishes ostreolysin
membrane activity [115]. Using lipid vesicles composed of equimolar ratios of SM and various natural and synthetic sterols, it has
been further shown that the membrane binding of ostreolysin does
not coincide with the ability of specific sterols to induce the formation of the lo phase [117]. These combined data suggest that ostreolysin requires other structural/ physical membrane features to fully
exert its activity. To clarify the effects of membrane ordering on the
ostreolysin membrane activity, detailed analyses of a series of ostreolysin-susceptible and ostreolysin-resistant lipid vesicles were
carried out, using electron paramagnetic resonance and Fouriertransformed infrared spectroscopy [118]. Using GHOST analysis of
these electron paramagnetic resonance spectra, which provides a
condensation algorithm that filters and groups the solutions found
in optimization runs, this has revealed that regardless of their lipid
compositions, all ostreolysin-susceptible membranes have nearly
identical domain-distribution patterns. The most ordered domain
across all of these vesicle types, with an order parameter >0.72,
covered 80% or more of the membrane area. The distribution and
size of these domains, however, remain to be determined. It is also
interesting to note that the addition of higher concentrations of ostreolysin to susceptible lipid vesicles did not induce any detectable
changes in the electron paramagnetic resonance spectra [115], and
did not modify the membrane fluidity [114]. This suggests that
ostreolysin does not induce significant fluidity changes in the membrane upon binding.
As in the case of other raft-binding cytolysins, the membrane
lipid composition needed for optimal membrane ‘docking’ of ostreolysin remains to be clarified. One of the possible recognition sites
might be small cholesterol or cholesterol/SM clusters arranged in
the plane of the membrane. Such transient, nanoscale, lateral microheterogeneities of membranes where the stoichiometry varies
with the cholesterol concentration were recently suggested by molecular dynamics simulations of cholesterol. DPPC [119] and cholesterol. SM membranes [120]; these microheterogeneities were
found to be typical for cholesterol-and-SM-rich membranes and for
membrane rafts [1, 11].
Ostreolysin has recently been used to study the association of
the desmosomal cadherin desmocollin 2 (Dsc2) with cholesterolenriched membrane domains, and the influence of cholesterol on
desmosome assembly in MDCK epithelial cells. Biochemical
analysis showed the association of Dsc2 in cholesterol-enriched
fractions that contained the membrane raft markers caveolin-1,
flotillin-1 and ostreolysin, and immunofluorescence microscopy
confirmed this co-localization of Dsc2 and ostreolysin, which suggests that there is a pool of Dsc2 that is associated with membrane
rafts [121, Fig. 4b]. In the T24 (malignant) urothelial cancer cell
line, ostreolysin was also used to reveal the enrichment of cholesterol and SM membrane nanodomains along intercellular membrane
Pore-Forming Proteins As Raft Markers
Current Medicinal Chemistry, 2013, Vol. 20, No. 1
7
Fig. (4). Binding of ostreolysin to cholesterol-enriched raft-like domains of mammalian cell membranes. Confocal images of mouse somatotrophs (A)
and fluorescence microscopy of MDc-2 cells (B), double-labeled with antibodies against ostreolysin (Oly) (left panels; red), and cholera toxin B subunit (CTB) or YFP-desmocollin 2 (middle panels; green). Right hand panels show merged images of ostreolysin and CT-B, and ostreolysin and desmocollin 2. Scale
bars, 5 μm. Reproduced from [101, 121], with respective permissions of Elsevier and the American Society for Biochemistry and Molecular Biology.
nanotubes [122]. These highly curved tubular structures that connect neighboring cells are highly fragile and very difficult to isolate,
and therefore their exact molecular composition remains unknown.
GANGLIOSIDE-BINDING TOXINS
Cholera toxin subunit B (CT-B) is a part of cholera toxin from
Vibrio cholerae, and it has been used regularly as the probe of
choice for membrane rafts. CT-B assembles with the A subunit for
their ADP-ribosyltransferase and NAD +glycohydrolase activities,
which leads to the disruption of the intestinal epithelial barrier, and
to heavy diarrhea, as seen in the disease cholera [31]. CT-B is composed of five 11.5 kDa polypeptides that assemble into a stable
pentameric ring. By itself, CT-B is not toxic, and it binds with high
affinity (Kd <1 nM) to the ganglioside GM1, which is enriched in
membrane rafts. CT-B has been used for the detection of membrane
rafts in several studies. Kenworthy et al. [7] used FRET microscopy, which can measure molecular proximity on a scale <100 Å
between donor-labeled and acceptor-labeled antibodies. They compared the organization of three endogenous, raft-associated GPIanchored proteins labeled with antibodies (the folate receptor,
CD59, and 5'-nucleotidase) with GM1 labeled with CT-B. In most
cases, the FRET correlated with the surface density of CT-B.
Bacia et al. [123] used fluorescence correlation spectroscopy to
trace the mobility of CT-B (as a lo-phase marker) and dialkylcarbocyanine (as a ld-phase marker) in cells and in giant unilamellar vesicles (GUVs). This technique traces temporal fluorescence fluctuations from the diffusion of single fluorescent molecules, and it allows raft-associated and non–raft-associated molecules to be distinguished without the use of detergents. In GUVs composed of an
equimolar mixture of DOPC, SM: cholesterol, which allows phase
separation, CT-B and dialkylcarbocyanine showed different diffusional mobilities Fig. (6). On the other hand, in homogenous artificial GUVs composed of DOPC, the mobilities of CT-B and dialkylcarbocyanine were almost the same. When the cholesterol was depleted using MCD, this reduced the mobility of dialkylcarbocyanine, and increased the mobility of CT-B. In HEK 293 cells, the
same system saw no enhanced mobility of CT-B, which suggests
that cytoskeleton disruption is required for higher CT-B mobility.
Further studies on CT-B-induced phase separation in GUVs have
defined the dynamics of membrane phase separation, demonstrating
it to be a highly dynamic process, and showing that G M1 appears to
reside in the ld phase, and to be associated with the lo phase only
upon addition of CT-B [124].
CONCLUSIONS AND PERSPECTIVES
Membrane microdomains, including rafts, have been a subject
of several investigations over the last 15 years. Emerging evidence
of the implications of rafts in crucial biological functions and under
pathological conditions have required the development of new
techniques and markers that allow their visualization. However, in
addition to their heterogeneity, the spatial and temporal instabilities
of these membrane microdomains have hampered these efforts,
especially when studied in living cells. Non-lytic mutants or chemical derivatives of natural toxins, and particularly of the PFTs, either
fused with a fluorescent protein or dye, or labeled with a fluorescent antibody, represent tempting tools for membrane-raft visualization. These can specifically target and bind to a raft-residing
lipid, or lipid combination, or they can ‘sense’ a specific membrane
physical phase (e.g. the lo phase). We must, however, be careful in
our interpretation of data obtained with these external raft probes,
8
Current Medicinal Chemistry, 2013, Vol. 20, No. 1
as their binding can influence the membrane structure itself and can
induce local rearrangements of membrane microdomains. This has
been shown for the binding of the CT-B pentamer, which induces
clustering of five GM1 molecules [31], and for listeriolysin O, which
promotes the aggregation of several raft-associated molecules
[125]. The use of antibodies, with their characteristic larger size,
can lead to further clustering, as has been shown in the case of CTB [126]. Such adverse effects can lead to miss-interpretation of the
spatial and temporal architecture of these membrane microdomains.
In this regard, PFTs tagged with a fluorescent dye, fluorescently
fused PFTs, and engineered smaller binding domains of PFTs (as in
the case of perfringolysin O) might provide a more correct choice.
Furthermore, it is important to distinguish between direct interactions of a putative raft marker with a certain lipid, and its recognition of a certain physical state of the membrane (e.g. the lo phase)
that derives from the use of particular lipid mixtures. It has been
shown that the lipid phase can promote membrane insertion and
pore formation of EqtII even in membranes without SM [57]. However, EqtII binding was still low compared to that with membranes
with very low levels of SM.
Skoaj et al.
CONFLICT OF INTEREST
The author(s) confirm that this article content has no conflicts
of interest.
ACKNOWLEDGEMENTS
The authors gratefully acknowledge Dr. Chris Berrie for critical
reading of the manuscript, and the Slovenian Research Agency for
financial support (grants J1-4305 and P1-0207).
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