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Review
Pore formation by actinoporins, cytolysins from sea anemones☆
Nejc Rojko a, Mauro Dalla Serra b, Peter Maček c, Gregor Anderluh a,c,⁎a Laboratory for Molecular Biology and Nanobiotechnology, National Institute of Chemistry, Hajdrihova 19, 1000 Ljubljana, Sloveniab Istituto di Biofisica, Consiglio Nazionale delle Ricerche & Fondazione Bruno Kessler, via alla Cascata 56/C, 38123 Trento, Italyc Department of Biology, Biotechnical Faculty, University of Ljubljana, Jamnikarjeva 101, 1000 Ljubljana, Slovenia
Abbreviations: APs, actinoporins; CHOL, cholphosphatidylcholine; DPC, dodecylphosphocholine;fragaceatoxin C; IBS, interfacial binding site; Ld, liordered; PC, phosphatidylcholine; PFT, pore-formingSM, sphingomyelin; Stn, sticholysin.☆ This article is part of a Special Issue entitled: Pore-Fo
Dalla Serra and Franco Gambale.⁎ Corresponding author at: Laboratory for Molecular B
National Institute of Chemistry, Hajdrihova 19, 1000 LjublE-mail address: [email protected] (G. Anderluh).
Actinoporins (APs) from sea anemones are ~20 kDa pore forming toxins with a β-sandwich structure flanked bytwo α-helices. The molecular mechanism of APs pore formation is composed of several well-defined steps. APsbind tomembrane by interfacial binding site composed of several aromatic amino acid residues that allow bind-ing to phosphatidylcholine and specific recognition of sphingomyelin. Subsequently, theN-terminalα-helix fromthe β-sandwich has to be inserted into the lipid/water interphase in order to form a functional pore. Functionalstudies and single molecule imaging revealed that only several monomers, 3–4, oligomerise to form a functionalpore. In this model theα-helices and surrounding lipidmolecules build toroidal pore. In agreement, AP pores aretransient and electrically heterogeneous. On the contrary, crystallized oligomers of actinoporin fragaceatoxin Cwere found to be composed of eight monomers with no lipids present between the adjacent α-helices. Thisarticle is part of a Special Issue entitled: Pore-Forming Toxins edited by Mauro Dalla Serra and Franco Gambale.
Peptide and proteinaceous pore-forming proteins, operating astoxins (pore-forming toxins; PFTs), are very efficient in compromisingselective permeability of cellular membranes. They have evolved acrossall kingdoms of life from viruses and bacteria to eukaryotes to play animportant role in biologicalwarfare [1–9]. At themolecular level, pivotalto PFT's function are their sequential and coordinated conformationalchanges upon more or less selective interaction of monomers (or insome cases, of loosely associated oligomeric form of a PFT) with ac-ceptor and/or receptor molecule(s) on the membrane surface. A novel
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conformational state of the bound protein favours further reshaping ofits polypeptide chain. This results in oligomerisation of the monomersin themembrane plane, and coordinated protrusion of a certain amphi-pathic polypeptide region of each monomer across the lipid bilayer.Thus several α-helices (α-PFTs) or β-hairpins (β-PFTs) build a circular,transmembranewater-filled pore [5,6]. It is now accepted that architec-ture of a transmembrane pore is not restricted to protein-only com-plexes, which is typical for β-PFTs, but membrane lipids may beincluded as spacers between amphipathic α-helices lining the lumenof the pore of some but not all α-PFTs [10]. Such proteo-lipid poresare considerably less stable as compared to those of β-PFTs that are for-tified by a network of hydrogen bonding in a β-barrel structure. Evenhigher complexity of a transmembrane pore structure is sometimes ob-served, in example, with bi-component PFTs as reported for bacterialleukocidins [11] or mushroom pore forming proteins [12].
Cnidarians, including sea anemones (Actiniaria), produce a variety ofbioactive compounds such as for example neurotoxins, phospholipases,and cytolytic PFTs (extensively reviewed recently in [13]). On the basisof their pore forming activity, a group of 20 kDa cytolytic proteins isolatedfrom sea anemones were named actinoporins (APs) [14]. These proteinsare cysteinless and have a rather high isoelectric point (N8.8) except forone acidic cytolysin [15]. APs were found in about 40 different sea anem-one species [13,16,17]. However, APs and actinoporin-like proteins ap-pear distributed in other Cnidarians and even in evolutionary verydistant taxa [18]. To date, the best characterised APs are equinatoxin II(EqtII) from the sea anemone Actinia equina, sticholysin I and II (StnIand StnII) from Stichodactyla helianthus and fragaceatoxin C (FraC) fromActinia fragacea. The localization of APs in the body of sea anemonesother than stinging organelles, nematocysts, is not entirely clear.Caritoxins from Actinia cari and FraC for example were originally isolatedfrom the body fluid excreted from the living animal upon mechanicalstimulation [19,20]. In fact, only few APs have been proven to be locatedin nematocyst [16]. Equinatoxin was first isolated presumably from thenematocyst [21], however, several equinatoxin isoforms were also puri-fied from the body fluid of A. equina [22]. Furthermore, discharge of iso-lated A. equina nematocyst directly into erythrocyte suspension wasreported not to be haemolytic [23], even though purified native and re-combinant actinoporins lyse erythrocytes. Immunohistological stainingshowed that StnII is present in tentacles, mesenteric filaments andbasitrichous nematocyst of S. helianthus [24]. As APs from individual spe-cies belong to a multigene family [13,25], it is possible they evolved tofulfil the varied different feeding demands [25], defence purposes, intra-specific relationships such as spatial competition or even digestion [26].This wide range of putative biological roles might also explain why notonly nematocysts, but also other body parts may release APs [24]. Phar-macological effects of APs are discussed in detail elsewhere [27]. Briefly,cardiotoxic effect after intravenous equinatoxin delivery is considered tobe the key mechanism of the toxin's lethality in vertebrates, but theexact mechanism of APs' pathological activity in animals leading todeath remains unknown [28–30]. Induction of pulmonary edema wasalso shown on the isolated rat lungs [31]. In addition to widely observedhemolysis, APs also causeplatelet aggregation [32]. A veryweakphospho-lipase A2 activity of sticholysins, purified from the sea anemone extract,was reported [33]. However, enzymatic active site and in vivo relevanceof this observation has not been yet clarified.
APs’ mechanism of transmembrane pore formation proceedsthrough a succession of ordered steps. After binding to sphingomyelin(SM) containing natural or model lipid membranes, the N-terminalα-helix changes conformation and inserts into the lipid bilayer. Thisbinding processwas extensively characterised in the past [34–36], how-ever, recent research has focused mainly on the final oligomerisationand pore formation, which remained elusive for a long time. In this re-view, we focus on molecular details of membrane interactions andpore formation by APs. Enthusiastic reader is directed to other excellentreviews on APs, which cover some other aspects including their biolog-ical role [9,13,16,37–40].
2. Actinoporins structure
Early investigation of the actinoporin secondary structure in solutionwith circular dichroism and Fourier transform infrared spectroscopy re-vealed prevalence of β-sheet over α-helical structure [35,41–45]. Thiswas later confirmed by X-ray crystallography and NMR data for EqtII[46,47], sticholysins [48,49] and FraC [50]. APs are single-domain pro-teins. The protein core is a β-sandwich, composed of two β-sheetseach formed by five β-strands, and associated with an α-helix (Fig. 1A).Superposition of EqtII, StnII and FraC crystallographic models reveal ahigh degree of structural similarity with a RMS difference of less than0.9 Å despite the sequence identity is between 61 and 90% (Fig. 1B)[50]. First 30 residues, which include amphipathic N-terminal α-helix,lie in a hydrophobic groove on the face of the β-sheet (labelled blue inFig. 1A). Upon binding β-sandwich protein core is not disrupted [51],while the N-terminal α-helix is the only part of the molecule where sig-nificant conformational change takes place [34,52,53]. APs possess aprominent aromatic residues cluster on the surface of the protein,which was shown to be involved in lipid recognition and membranebinding (Trp112, Tyr113, Trp116, Tyr133, Tyr137 and Tyr138, EqtII andFraCnumbering) [34–36,53–56]. Interfacial binding site (IBS)wasdefinedby Bakrač et al. [36], which is composed of this cluster of aromatic aminoacid residues and some other residues that participate in lipid recognition(see below) (Fig. 1A and C). NMR structures of EqtII and StnI are in closeagreementwith the crystallographic data [47,49],moreover, itwas shown
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theβ-sheet core is rigid across awidepHand temperature range,whereasthe N-terminal part is more flexible [47].
3. Actinoporins membrane binding
3.1. Sphingomyelin recognition and binding
Binding ofwater soluble PFTmolecules to the targetmembrane is usu-ally driven by more or less tight interaction between a particular bindingsite located on protein and acceptor/receptor molecule(s) embedded inthe membrane. Due to the high pI of APs, electrostatic interactions be-tween positively charged protein and negatively charged membranescould be expected. However, high salt concentration lowered bindingby approximately 30% only [34] and modification of positively chargedresidues of EqtII did not significantly abrogate haemolytic activity [57,58]. Furthermore, surface plasmon resonance study of EqtII interactionwith lipid membranes revealed that neither negatively charged lipids inlipid membrane nor charge manipulation of IBS could significantly alterthe binding properties of proteins [36]. This suggests that the net chargeplays a minor role in actinoporin/membrane interactions, as furtherhighlighted also by the existence of actinoporinswith acidic pI [15].More-over, a closer look to the EqtII, StnII and acidic actinoporin from Sagartiarosea surface charge distribution reveals that charged residues are notconserved across the protein surface and absent from the region thatformsmembrane binding site (Fig. 1C). Instead, increased intrinsic trypto-phan fluorescence upon actinoporin interaction with lipid vesicles sug-gested involvement of Trp residues in binding [35,56]. Further mutatinghydrophobic residues Trp112 or Trp116 lowered binding by more than90%, implying that protein-membrane interaction is partially hydropho-bic at least [34,53,59]. 19F labelling of EqtII tryptophans enabled a detailedin vitro study of their importance for lipid binding. Accordingly, the mostaffected 19F resonance in the presence of phospholipidmicelles or bicelleswas that of Trp112, followed by Trp116 [55]. NMR studies of StnI con-firmed that aromatic amino acid residues from the IBS, Trp112, Tyr113,Trp116, Tyr133, Tyr137 and Tyr138 (EqtII and FraC numbering) areinserted into the dodecylphosphocholine (DPC) or dihexanoyl phosphati-dylcholine (DHPC)micelle, providing the first contact pointwith the lipidinterface [54,60].
Structural studies of APs in the presence of lipid analogues allowedfine mapping of IBS. Tanaka and colleagues have recently observedthat up to four DHPCmolecules can bind to FraCmonomer in the crystalstructures [53]. Two of these sites (L2 and L3, as denoted in Tanaka et al.[53]) were occupied in independent crystal structures more frequentlyand could represent the primary sites for protein attachment to lipidmembranes (Fig. 2A). Co-crystallization of StnII with phosphocholine(POC),which is a part of headgroup of both SMandphosphatidylcholine(PC), revealed important residues for binding which are also conservedin other APs (numbering according to EqtII and FraC): Ser54, Val87,Ser105, Pro107, Tyr113, Tyr133, Tyr137 and Tyr138 (labelled orangein Fig. 1C and Fig. 2B) [48]. Interestingly, POC bound in an orientationthat partially overlaps with L2 and L3 binding sites of FraC (Fig. 2Aand B). NMR study of StnI precisely mapped DHPC micelle binding siteresidues and most of them overlap with DHPC and POC binding sitesof FraC and StnII, respectively [54,60] (Fig. 2C). Very recently an NMRstudy enabled identifying residues involved in binding of EqtII to DPCmicelles [61], again mapping the IBS to the same region of themolecule(Fig. 2D). Altogether these results allowed identification of themost im-portant residues for lipid recognition (shown as sticks in Fig. 2F). Manyof these residues were previously identified in mutagenesis studies tobe important for APs permeabilising activity [34–36,53,59,62–66]. Therecent data, therefore, suggests that APs IBS allow interaction withseveral headgroups of membrane lipids [53]. The presence of multiplebinding sites for SM and PC headgroup could increase the affinity tolipid membranes during initial membrane interactions. Such multiva-lent interactions with membrane lipids clearly discriminate APs fromother PFTs [53].
It was evident very early that SM as a membrane acceptor plays akey role in actinoporin mechanism of action. Incubating sticholysinwith SMprior to erythrocyte addition reduces toxin haemolytic activity,the same effect was achievedwhen SMwas removed from erythrocytesby sphingomyelinase [67]. This was later shown also for EqtII [36]. Eventhough APs exhibit weak permeabilising activity on membranes com-posed of PC [45] or phosphatidylethanolamine alone [68], the additionof small SM fraction to planar lipid membrane increases the activityby several orders of magnitude [69,70]. Similarly, calcein leakage fromartificial lipid vesicles increases significantly when SM is part of thebilayer [36,45,70–73].
EqtII, however, can clearly discriminate between SM and otherlipids, most notably PC and cholesterol (CHOL), as shown in several in-dependent in vitro assays such as dot-blot assays [36], surface plasmonresonance [36] and lipid ELISA assay [74]. Site-directed mutagenesisrevealed residues Trp112 and Tyr113 to be responsible for specific SMrecognition. Because SM and PC share phosphorylcholine head group,other SM regions below phosphorylcholine head group must be in-volved in specific interaction with actinoporins [36]. Molecular dockingby using GOLD 5.0 suggested that Tyr135 in StnII (Tyr137 in EqtII) spe-cifically interacts with 2-NH and 3-OH groups of SM ceramide moietywhere Tyr111 and Tyr136 recognise phosphate oxygens (Tyr113 andTyr138 in EqtII) (Fig. 2) [72]. Similar approach backed-up with NMRdata suggested 2-NH and 3-OH groups are recognised by side chainsof Asp109 and Tyr113, and main chains of Pro81 and Trp112 [61]. Aloop with Tyr111 (Tyr113 in EqtII) could not be placed in the electrondensity map of StnII. It was proposed that Tyr111 induces high flexibil-ity of the loop, a feature important in protein adaptation to the mem-brane surface [48,65]. The importance of Tyr111 is underlined by thefact that this residue is conserved among all actinoporins. Trp112 alsocontributes to SM recognition and in addition stabilizes membrane-bound protein [36]. Notably, APs specificity for SM is biologicallyrelevant, as in sea anemones cell membrane SM is replaced by itsphosphono derivative with the phosphonocholine head group. In con-trast to SM, the derivative does not interact with APs, therefore makingsea anemones resistant to their own toxin [71].
Specific recognition of SM by EqtII was also shown in cellularmodels. Bakrač et al. demonstrated that fluorescently labelled EqtII spe-cifically binds cellular compartments containing SM [75], therefore,making it a suitable probe for cellular SM [76,77]. However, mecha-nisms underlying SM binding by proteins in cellular membranes arestill not well understood.When staining COS-1 cells with two SM specifictoxins, EqtII and lysenin, a SM-specific pore forming toxin from earth-worm, it was revealed that they tend to bind different plasmamembraneregions. This implies the importance of SM organization for binding avail-ability [74,76,78]. Both toxins are specific for SM in in vitro assays, howev-er, lysenin recognise clustered SM, while EqtII binds dispersed SM inmodel and cellular membranes [74]. There is also a possibility that otheror additional acceptors/receptors for APs are present in cellular mem-branes besides SM. This proposition is based on the observation thaterythrocytes and other mammalian cells were lysed at much lowertoxin concentration compared to artificial lipid vesicles containing SM[26,73]. Isolated cardiocytes were found to be strikingly sensitive andwere hyper-contracted even by ~pM EqtII [79]. It was suggested thanan RGD-motif close to the POC binding site could play a role in integrinrecognition at the cellular surface [80]. However, the proposed role ofthismotif seems unlikely as it is not conserved in APs [13], andmoreover,synthetic peptides with the RGD-motif in large excess were not efficientin suppression of the EqtII-induced haemolytic activity (P. Maček, unpub-lished observation).
3.2. Role of lipid domains
APs membrane interactions are, therefore, largely governed bySM binding. However, recent studies revealed that membrane bulkphysico-chemical properties, including coexistence of lipid domains,
Fig. 2.Molecular details of lipid recognition by actinoporins. Residues that participate in lipid binding are labelled on the surface of APs.Molecules are oriented in the sameway in all panels(A–D, F). A) Two DHPC molecules bound to FraC are presented by sticks. L2 binding site is labelled in red and L3 in blue [53]. B) Binding site of POC on StnII is labelled in orange [48].C) Residues that were affected upon DHPCmicelles addition to StnI are labelled in cyan [60]. D) Residues that were affected upon DPCmicelles addition to EqtII are highlighted in yellow[61]. E) Table that summarizes structural studies of APs interaction with lipid ligands (A–D). Black rectangle denotes participation of this particular residue in lipid binding. Residues thatare coloured in orange were shown previously bymutagenesis to be important for functional properties of APs (see text for details). Numbering is according to EqtII and FraC. F) The con-sensual residues for lipids recognition are denoted by red sticks (numbering according to EqtII and FraC).
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significantly modulate pore forming activity of APs. It was shown, forexample, that presence of membrane CHOL promotes APs pore forma-tion. Permeabilisation of COS-7 cells by StnII was decreased afterCHOL removal from the cellular membrane [81], whereas CHOL deple-tion from erythrocytes did not play a role in EqtII activity [36]. More-over, APs permeabilize PC:CHOL liposomes devoid of any SM [82,83],but the addition of CHOL, to different PC vesicles resulted only in slightlyenhanced binding as shown by SPR [36]. As APs do not directly bindCHOL [36], it is likely that CHOL modulates APs activity by ordering ef-fect on physiologically important fluid membrane state, promotinglipid segregation and formation of highly ordered membrane domains[84]. Lowering temperature has a similar ordering effect on the mem-brane, possibly explaining enhanced APs activity on PC:CHOL mem-branes at 4 °C [45,85].
EqtII was found to co-localize with lipid rafts, which are formed byspecific interaction of SM and CHOL [86]. Usingmodel lipid membranes
it was shown that supplementation of SM with CHOL in lipid vesiclesenhanced EqtII binding.When replacing CHOLwith ergosterol the effectof calcein leakage from lipid vesicleswas conserved,whereas substitutionwith cholestenone diminished it. CHOL and ergosterol, but notcholestenone, are promoters of liquid-ordered (Lo) domain formationand phase separation. This, together with observation of EqtII binding todomain borders, lead to suggestion that APs preferentially accumulateat lipid defect sites arising at the boundary between coexisting lipidphases, where lipids are more exposed [85]. Preferential domain bound-ary binding was later visualized also in giant unilamellar vesicles [87]and droplet interface bilayers [88] (Fig. 3A). These observationsmight ex-plain why EqtII and StnII both co-localize with raft membrane fractions[81,86], however, experimentswhich involve isolation of raftswith deter-gents are prone tomis-interpretations [89]. It is not clear if lipid defects atdomain boundaries expose SM head group for easier recognition by APsor expose hydrophobic acyl chains thus lowering the energy barrier for
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α-helix insertion [85,87,88,90]. Data suggests CHOL impact on APs activ-ity is mostly governed by its effect on SM head group tilt and dynamics,especially on the domain border, whereas bilayer fluidity plays only aminor role [91]. It is interesting to note that recent results do not supportthe importance of phase separation on StnI. Sterols induce its activity re-gardless of their ability to promote phase separation. Authors present amodel where sterols enhance APs pore formation not only bymodulatingmembrane fluidity and heterogeneity, but also by inducing negativemembrane curvature [92].
Counterintuitively, there is a growing evidence in support of APs in-sertion into disordered regions of the membrane, even though SM andCHOL are enriched in ordered domains [85,88,90,91]. A high contentof SM lowers the lateral diffusion, which reduced StnI binding andmembrane penetration [93], in agreement with previously observedlow binding to vesicles composed of SM alone [70]. Furthermore, func-tional pores are formed in liquid-disordered (Ld) domains as visualizedby ionicflux through individual pores inmodelmembrane, even thoughEqtII was concentrated at the domain boundary immediately aftermembrane binding [88] (Fig. 3B). It was suggested that decreasedlipid packing in disordered domains is a more suitable environmentfor actinoporin membrane insertion and pore formation [90,94], be-cause insertion into ordered phase is enthalpically unfavourable processdue to the breaking up of the tight lipid packing in the ordered domain[95]. An important factor governing protein distribution in the mem-brane is also the bilayer thickness. Due to the hydrophobicmismatch ef-fect, longer β-barrels, like that from perfringolysin O, tend to insert intogenerally thicker ordered domain, whereas β-barrels shortening lowersthis affinity. However, this is not the case with APs, because proteinswith single transmembrane segment can easily avoid the consequenceof mismatch by tilting the hydrophobic surface in order tominimize ex-posure to aqueous solution in thinner lipid domains [96]. In contrast tothe proposed role of themembrane physical state it was suggested thatPhe16 residue in FraC plays an active role in CHOL sensing. When onlyLo phase is available (i.e. high CHOL content), Phe16 is needed to facil-itate otherwise unfavourable FraC pore formation [97]. At high CHOLconcentration pore forming ability of N-terminal peptides from StnIand StnII was also reduced [98].
EqtII was also reported to promote expansion of Lo phase in modelmembranes [88], and to induce raft-like domains formation in livingcells [86]. In contrast, StnI and StnII in model membranes inducedlipid mixing after binding to domain boundary [90]. Similarly, EqtII
Fig. 3. Role of lipid domains for binding and pore formation by APs. Binding and pore formaseparatedmembranes EqtII bound preferentially to domain boundaries. Lipidmembraneswereing 0.1% 3,3′-dilinoleyloxacarbocyanine perchlorate as amarker of liquid disordered phase. EqtIin thefirst 300 s and then finally excluded from liquid ordereddomains. B) EqtII pore formationevident that protein covers liquiddisordered region completely and, hence, thenon-lytic formothe interior of the vesicle, where the calcium-sensitive dye Fluo-8H enabled imaging (green ddependent pore formation of equinatoxin II in droplet interface bilayers, 1630–1637, Copyrigh
ordered liquid disordered phases and disordered more ordered lipidphases in multilamellar vesicles as studied by NMR [99], but on theother hand phase behaviour in giant unilamellar vesicleswas not signif-icantly altered [74]. It is important to take into account the simplifi-cation of model lipid membranes, which uses binary or ternary lipidmixtures. Clearly, cellular membrane is much more complex, and Lo/Ldcoexistence in model membranes may not be adequate to explain lateralheterogeneity in cell membranes. It is now evident that transmembraneproteins are generally excluded from ordered phases in model mem-branes, even those thought to have affinity for raft domains in cell mem-branes [88,100].
4. Conformational change after membrane binding
In addition to the lipid-binding site, another part of the APs mole-cule, i.e. N-terminal α-helix, will be embedded in lipid membrane inlater stages of pore formation. Cysteine scanning mutagenesis of theN-terminal EqtII region, coupled with fluorescence labelling, has indi-cated that this part is translocated after binding step to the membraneinterphase [62,101]. Interestingly, the N-terminal region of APs is simi-lar to the honey bee venom component melittin, which has an amphi-pathic nature and inserts into lipid membranes [41]. Progressivedeletion of amino acids in the N-terminal does not significantly affecttoxin binding, but results in much weaker haemolytic activity [102].On the other hand peptides that correspond to the N-terminal se-quences of APs are several orders of magnitude less haemolytically ac-tive as the full length toxins [103,104]. Thus, while the bulk of theprotein is essential for efficient pore formation by facilitatingmembranebinding and stabilizing the oligomeric pore, theN-terminal region is thekey component of the pore walls. When the disulphide bridge is formedbetween theN-terminal region and theβ-sandwich, thus disabling con-formational change, the toxin is considerably less haemolytically activebut retainsmembrane-binding capability [34,51]. EqtII, therefore, in thefirst step recognises targetmembrane by the IBS as described above, andthen the second step is promoted by the N-terminalα-helix membraneinsertion [34]. Prior to pore formation, thisα-helix lies almostflat on themembrane-water interface [42,101,105,106]. Upon binding there is aslight increase in α-helical content in expense of the disordered struc-ture [44,107–109], consistent with the folding of unstructured regionfrom residue 10 to 15 as observed by electrophysiology of chemicallymodified mutants [101]. Similarly, the 32 residues-long N-terminal
tion of EqtII as visualized in droplet interface bilayer system [123,125,126]. A) In phase-composed of 1,2-diphytanoyl-sn-glycero-3-phospho-choline/egg SM/CHOL 4:1:1 contain-I was labelled with Cy3B dye andwas found initially to preferentially bind domain borderswas observed in liquid disorderedmembrane phase (stainedwith Cy3B-labelled EqtII). It isf the protein predominates. Poreswere visualizedbyfluxof calcium ions throughpores intoots). Reprinted from Biophysical Journal, Vol 106, Rojko N. et al., Imaging the lipid-phase-t (2014), with permission from Elsevier.
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fragment adopts α-helical structure from residue 6 to residue 28 whenbound to lipid micelles, making it nearly three times longer than in na-tive soluble protein [104]. Firstfive residues are not ofmajor importancefor pore formation [102], however they help to anchor the amphipathichelix on the trans side of themembrane and contribute to stabilizing thetransmembrane pore [52].
Very recently a crystal structure of oligomeric FraC embedded in de-tergent was determined (Fig. 4) [53]. This breakthrough was achievedby first incubating FraC with lipid vesicles, which allowed monomerbinding and pore formation. Vesicles were then solubilized with deter-gents, but oligomers remained stable and thus enabling pore isolationusing ion-exchange and size exclusion chromatography. The pore is afunnel-shaped oligomeric particle where N-terminal α-helices moveaway from the β-rich region, which remains essentially unchanged. Inagreement with other APs functional studies, upon detachment fromthe β-sandwich core the N-terminal α-helix is extended to encompassfinally residues 4–29. This is sufficient to punch a hole through themembrane thickness of about 5 nm [53,104]. The helices from the olig-omeric complex are placed approximately perpendicular, with hydro-phobic face oriented towards to the outside of the assembly andhydrophilic face towards the pore lumen [53]. Negatively charged resi-dues from this part of the helix modulate slight cation selectivity of thepore as revealed by electrophysiological measurements [52,70,73,101].
5. Oligomerisation and pore formation
5.1. Size and stoichiometry of the pore
Sufficiently high concentration of monomers on the membraneplane triggers pore formation. Kinetic experiments [70] and optical mi-croscopy [88] indicate that themajority of EqtII at the plane of themem-brane is in the monomeric form. A broad conductance distribution ofindividual pores implies a non-uniform size of the pore [73,101]. Fluo-rescent microscopy imaging recently confirmed that the ionic fluxthrough individual pores is indeed variable. Furthermore, individualpores appear to be unstable with quickly switching between multipleconductive states [88]. It is possible that subunits in the pore complex
Fig. 4. Conformational changes associated with formation of FraC octamer and structure of oligoctameric homo-oligomer (red). The only notable difference in3D structure between these twoβ-sandwich. B) FraC oligomeric structure. Eight protomers are labelled in grey. SMmolecules thSM binding by FraC protomers that clearly present fenestrations (openings) in the transmembsecondary structure elements. SM molecules are shown as orange spheres.
are exchanged with the monomeric membrane pool, similarly as ob-served in an anthrax protective antigen [110], but this is yet to be prov-en. An average pore radius was estimated to be 1 nm [70,73,82] anddoes not depend on toxin concentration [111]. The radius of the poreformed solely by the StnII N-terminal peptide was very similar to thatof the pore formed by the full-length StnII [103], indicating that thedimensions of the pore are inherent property of the helices arrange-ment in themembrane. In the crystal structure of FraC pore, inner diam-eter is reported to be 6 nm at the vestibule and 1.6 nm at its narrowestpoint at the cytoplasmic side, which is in agreement with functionalstudies [53].
Due to its instability the pore complex structure was elusive for along time and the stoichiometry was indirectly estimated from variousexperiments. Kinetic studies of EqtII, StnII and StnI induced pore forma-tion in lipid vesicles suggest predominantly three to fourmonomers perfunctional pore [70,73,82]. In the case of StnI, a single pore formationproceeds through 3 or 4 smaller conductance increases before reachingthe final pore configuration and this could be explained as a sequentialaddition of monomers to the final pore [112]. This functional data are inagreementwith tetrameric structures observedwith electronmicrosco-py of StnII bound to lipid interface [48]. Cross-linking experiments showmembrane bound oligomers mostly consisting of dimers and trimers,however, larger complexes were also observed [73,79,106]. Advancesin optical microscopy enabled direct observation of toxin distributionin lipid membranes. Single molecule tracking of fluorescently labelledEqtII in model membranes supported the earlier evidence of heteroge-neous oligomer distribution. The analysis shows a broad distributionof stoichiometries with a mean of 3.4 ± 2.3 EqtII protomers per oligo-mer, in agreement with a pore model built of 3–4 protomers. However,a small fraction of larger assemblies was also present. As expected fromearlier data [111], neither toxin concentration nor lipid composition haslarge effect on the distribution of means [113]. Similarly tomodelmem-branes, EqtII exists in the plasmamembrane as a mixture of monomers,dimers, tetramers, hexamers and also a small fraction of higher oligo-mers according to single molecule imaging [114]. In summary, experi-ments in model and cellular membranes with physiological proteinconcentrations indicate that only few protein monomers are enough
omeric FraC assembly. A) Overlay of FraC monomer in solution (blue) and taken from theforms is in theN-terminal region,whereα-helix elongates andmoves away from the stableat were found to be integral part of the pore are presented as orange spheres. C) Details ofrane pore structure. The transparent surface of FraC protomers is presented together with
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to form a functional pore (see Fig. 5 for a model) and that larger oligo-mers are not the predominate form of the protein at the membranesurface [70,88].
5.2. Lipids are involved in pore formation
Only three to four inserted α-helices are not enough to form a porewith radius of 1 nm and additional structural elements have to line thepore walls. The β-sandwich core of APs does not change conformationand is not exposed to the pore lumen [51,53]. All this led to the proposalthat APs form toroidal pores, where lipids are involved in the forming ofporewalls (Fig. 5). Thismodel was reinforced by the fact that lipidswithnegative curvature promote actinoporin activity [115] and that nega-tively charged lipids enhance cation selectivity [116]. Ion selectivity isgoverned by charged groups facing the conductive channel, thereforelipid headgroups must be present in the pore walls [116]. Moreover,APs induce “flip-flop” movement of lipids, in agreement with toroidalpore model. Lipid involvement as a pore structural element could alsoexplain pore instability and broad conductance distribution, a charac-teristic very different from the stable and well defined barrel-stavepores [9,112]. Actinoporins are not the only case where a toroidal porewas proposed, it seems this mechanism of pore formation is widelyspread feature among different α-PFT, like peptides, colicins and Baxapoptotic regulator, as well as β-PFT as recently reviewed in [10].
Recently, the toroidal pore model was challenged by crystallographicdata of FraC oligomers [50]. In this study, at a high protein concentration,FraC was crystallized in presence of detergent. For nonameric assembliesin the crystal lattice, it was suggested that thismay represent a biological-ly relevant oligomeric complex. Because the N-terminal α-helices werestill attached to the protein core, it was suggested that this crystal struc-ture could represent a non-lytic oligomer or a prepore. In this scenariooligomerisation precedes large conformational changes, a mechanismsimilar to many PFTs that generate β-barrel pores [117–121]. In contrast,kinetic dissection of APs pore formationmechanism does not imply a sta-ble oligomer formation prior to the N-terminal α-helix insertion in thelipid membrane. This was assessed by the use of a double cysteine EqtII
Fig. 5. Actinoporins pore models. Toroidal pore model, side (A) and top (B) views. Accordingstoichiometry is not fixed. Here we show fourmonomers to build a pore. Lipid head groups arebrane leaflet.Octameric poremodel, side (C) and top (D) views. Eightmonomers build a pore,meisolated points lipids acyl chains are exposed to pore lumen by fenestrations between monom
mutantwith restricted conformational freedombecause of the disulphidebond between the N-terminal α-helix and protein core. When this mu-tant first binds to the membrane and the disulphide is reduced after thebinding is completed, pore formation and conformational change aremuch slower compared to the wild type EqtII, most probably due tosterical reasons [106]. If a stable oligomer is formed before membranepenetration, one would expect addition of a reductant should speed upthe pore formation because the binding and oligomerisation are alreadyaccomplished, as shown in the case ofβ-PFTperfringolysinO [122].More-over, conformational change kinetics in EqtII is faster compared to poreassembly. It was suggested that themajor conformational change, the de-tachment of the N-terminal helix, occurs immediately after the bindingand must be accomplished before the oligomerisation to the final pore[106]. Accordingly, StnI monomers sequentially insert α-helices to thegrowing pores, what was observed as a subset of lower ion conductivepathway leading to the final tetrameric pore [112].
Very recently, Tanaka and colleagues crystallized FraC dimers withthe α-helices still attached to the protein core [53]. Even though largeconformational change was not observed, authors detected localizedminor conformational changes around Leu14 and Phe16, which couldtrigger a structure destabilization that leads to the helix detachmentand lying it down onto the membrane. The simultaneous insertion ofFraC twohelices fromeachdimer into the lipidmembrane could explainthe four sub-conductive states towards the formation of the finaloctameric pore, as reported for StnI [112]. However, microscopy studiesusing physiological concentrations of proteins revealed that EqtII in themembranes exist mostly as monomer, dimer, tetramer and hexamer,but only small fraction of higher oligomers, like octamer or nonamer,are present [113,114]. Dimers are also clearly visible after chemicalcrosslinking EqtII on themembrane [73,79,106]. Interestingly, StnI solu-ble dimers cross-linked with a disulphide bridge showed higher pore-forming activity in comparison to the monomeric protein [64]. Data al-together indicate that dimers could be intermediates in the pore forma-tion pathway of APs, as suggested for FraC and EqtII oligomer assembly[97,114]. This is another characteristics making difference between APspore forming mechanism and those of other PFTs. For example, no
to photobleaching and fluorescence correlation microscopy single molecule imaging thepart of the pore wall, resulting in a bent continuous monolayer from outer to inner mem-mbranemonolayers remain separated. Porewall ismostly built byN-terminalα-helices, aters. Membrane lipid SM is also part of the oligomeric assembly (see Fig. 3 for details).
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stable intermediates were observed in the pore formation process of anarchetypal β-PFT α-hemolysin [123].
In addition to the nonameric FraC assembly, octameric FraC pore-likestructures were also crystallized. Variable stoichiometry is a well-knownfeature among PFT. For FraC, octameric structure is probably moreabundant because protein–protein packing interface in octamer is morefavourable when compared to nonamer [53]. Mutational studies ofEqtII suggested that conserved Lys77 is the key residue involved inprotomer–protomer oligomerisation [44]. Surprisingly, small changes inRGD motif, which was initially hypothesised to represent the integrinbinding site, altered oligomerisation behaviour in solution but not thetoxin membrane binding [124]. However, Lys77 and RGD motif are notsignificantly involved in protomer-protomer interface as revealed by theoctameric FraC pore structure, but have a role in one of the SM bindingsites as deduced from the crystal structure and molecular simulation[53,61]. Strikingly, this specific SMmolecule in the crystal structure is lo-cated between two adjacent protein chains thus stabilizing oligomericstructure (Fig. 4B andC) [53]. In direct contactwith the adjacent protomeris Val60 as seen in FraC pore crystal structure (Fig. 1A).
Octameric or nonameric pores models do not predict a toroidal porestructure, where continuous lipid layer spans the cis and trans mem-brane side through the pore. However, space between the α-helices incrystal structure is occupied by the lipid acyl chains, a possible pathwayfor transbilayer movement of lipids [115] and the reason for the in-crease of electrical channel noise [9,112]. Indeed, negatively chargedlipids promote cation selectivity, which implies lipid head groups areexposed to the pore lumen [116].
5.3. Models of actinoporins pore formation
Twomost recent hypothetical pathways towards pore formation aredepicted in Fig. 6. The main difference regards the apparent lack ofprepore necessity, well reported in the traditional model based onEqtII and Stns (Fig. 6A). In this model the N-terminal α-helix startsearly its detachment from the protein and continues by inserting deeply
Fig. 6.Models of pore formation by actinoporins. A) Traditionalmodel of actinoporin pore formaα-helix is transferred to the lipid–water interface (M2), monomers subsequently oligomerizeα-helix inserts into the bilayer core prior to oligomerisation thus representing another monocrystallographic data [53]. This model involves dimerization (D1, D2) as an important intermedia(P′) where α-helices are still attached to the protein core. In this model it is not clear at whichQuestion marks denote steps which are not yet resolved by the data. α-helices are represented
in the lipid membrane during the oligomerisation step. The final, fullyactive state is reached by the structured aggregation of proteinmoietieswith α-helices already inserted [96,103]. Space in pore walls betweenα-helices is filled with lipid molecules, resulting in continuous lipidtorus. In the most recent model [53] the oligomerisation proceeds onthe lipid plane via aggregation of dimers, where the α-helices havestarted to partially unfold. This model supposes the formation of an in-active prepore where the α-helices are notably partially unfolded andslightly detached from the main protein. Only after the completion ofthe full ring, the helices are allowed to penetrate the lipid membrane.However, there may not be enough space to coordinately transfereight α-helices from the membrane surface to the other side of themembrane through octameric assembly. According to the structural di-mensions for the active pore as reported in [53], i.e. external diameter of11 nm and diameter of a single proteinmonomer of 2.8 nm, wemay es-timate the internal pore diameter, that should be 5.4 nm. This small di-mension is probably not big enough for permitting the simultaneousrearrangement of the eight α-helices, if considered each as rigid cylin-ders 3.5 nm long.Wemay suppose that the premature partial unfoldinginvolves the first half of the N-terminal region, shortening this way thelength of the rigid part which can therefore find enough space into thepore interior for tilting and penetration into themembrane. Aftermem-brane penetration the first andmore flexible N-terminal part can recon-stitute the helical structure. The twomodels are not so orthogonal, if wemay assume that the proteinmoiety able to trigger the helix insertion inthe traditional model, is actually composed by dimers, as reported inTanaka et al. [53].More structural information is necessary for unravellingthe nature of this basic structural element of the pore.
6. Conclusions
APs serve as an excellent model for α-PFT. The 20 kDa AP molecule,built from compact β-sheet core flanked by two α-helices, is capable ofrecognising SM, which acts as a receptor in the cell membrane. This fea-ture together with themolecule stability and ease of preparationmakes
tion. Solublemonomeric protein (S) binds SM (lipids in red) in the targetmembrane (M1).to form a pore (P), where α-helices traverse the membrane [101]. Recent data suggestsmer state on the membrane (M3) [106]. B) Alternative pore formation model based onte step, observed also on plasmamembranes of cells [114], and possibly prepore assemblystate helices transfer to the bilayer and across the membrane to form a functional pore.as the darker orange rectangles.
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actinoporins a handful molecular tool for studying membrane interac-tions and other steps involved in pore formation. Recent structuraland functional studies enabled updating of the model of APs pore for-mation by providing new details. It is clear now what is the structureof IBS and that conformational change after initial lipid-toxin associationis needed for functional pore. Some toxins need substantial rearrange-ments of water soluble structure to enable subsequent pore formation,however, APs undergo only minor structural transformations: theN-terminal α-helix elongates on behalf of disordered structure and sim-ply rotates away from stable β-sandwich core. The transition to a poreis a two-step process, where pore walls are formed after α-helices layflat on the membrane-water interface. It is also clear that different α-PFT may use different mechanisms for pore assembly. APs are clearlyunique as their pore forming efficiency evolved from combination of anamphipatic α-helix, resembling melittin, and the β-sandwich structurebearing several loops with an IBS providing selective lipid binding.
Acknowledgements
We dedicate this paper to the memory of our excellent colleague,friend and fine scientist Dr. GianfrancoMenestrina. Gianfranco has pro-vided many unique insights into actinoporin structural and functionalproperties and his work and human and professional attitude will beremembered with gratitude. The work in the laboratories of N. R., P.M. and G. A. is supported by the Slovenian Research Agency (Pro-gramme grants Molecular Interactions "P1-0391" and Toxins andBiomembranes "P1-0207"). M.D.S. acknowledges support from CNRand PAT (Autonomous Province of Trento) “Grandi Progetti 2012” Pro-ject “MaDEleNA”.
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