Structures of sea anemone toxins

Toxicon 54 (2009) 1075–1088

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Structures of sea anemone toxins Raymond S. Norton* Walter and Eliza Hall Institute of Medical Research, 1G Royal Parade, Parkville 3052, Victoria, Australia

a r t i c l e i n f o

a b s t r a c t

Article history: Available online 13 March 2009

Sea anemones produce a variety of toxic peptides and proteins, including many ion channel blockers and modulators, as well as potent cytolysins. This review describes the structures that have been determined to date for the major classes of peptide and protein toxins. In addition, established and emerging methods for structure determination are summarized and the prospects for modelling newly described toxins are evaluated. In common with most other classes of proteins, toxins display conformational flexibility which may play a role in receptor binding and function. The prospects for obtaining atomic resolution structures of toxins bound to their receptors are also discussed. Ó 2009 Elsevier Ltd. All rights reserved.

Keywords: Sea anemone Toxin Cytolysin Protease inhibitor Ion channel NMR Crystallography

1. Introduction Sea anemones possess numerous tentacles containing specialized stinging cells or cnidocytes, which are in turn equipped with organelles known as nematocysts that contain small threads which are everted forcefully following mechanical or chemical stimulation. Anemones use this venom apparatus mainly in the capture of prey (crustaceans, small fish) and for defence against predators, although possibly not for intraspecific aggression (Bartosz et al., 2008). In keeping with other venomous animals, sea anemones contain a variety of interesting biologically active compounds, including some very potent toxins (Be´ress, 1982). Peptides and proteins figure prominently amongst the sea anemone toxins characterized to date (Be´ress, 1982; Norton, 1991; Honma and Shiomi, 2006; Norton, 2006). Abbreviations: AP-A, anthopleurin-A; AP-B, anthopleurin-B; CD, circular dichroism; EqtII, equinatoxin II; HERG, human ether-a-go-go-related gene; Kv, voltage-gated Kþ channel; KCa, Ca2þ-activated Kþ channel; PC, phosphatidylcholine; ShK, Kþ channel toxin from Stichodactyla helianthus; SM, sphingomyelin; VGSC, voltage-gated sodium channels. * The Walter and Eliza Hall Institute of Medical Research, 1G Royal Parade, Parkville 3052, Australia. Tel.: þ61 3 9345 2306; fax: þ61 3 9345 2686. E-mail address: [email protected] 0041-0101/$ – see front matter Ó 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.toxicon.2009.02.035

The two most thoroughly characterized classes are the 5 kDa toxins that act by binding to the voltage-gated sodium channel and the 18–20 kDa cytolysins. This review focuses on the structures and structure–function relationships of these and other sea anemone toxins. Three-dimensional structures have been determined for other classes of sea anemone proteins, notably a range of fluorescent proteins that emit light in the far-red, red, yellow, cyan and blue regions of the visible spectrum (Chan et al., 2006), but these have no known toxic function and will not be discussed in this article. Protease inhibitors will be considered briefly because the Kunitztype fold adopted by some of these proteins has assumed ion channel inhibitory functions in sea anemones (Schweitz et al., 1995; Honma et al., 2008) and probably also in the closely related cnidarian Hydra magnipapillata (Sher et al., 2005). We begin with a brief description of biophysical methods available for studying peptide and protein structure and interactions. The structures of those toxins for which high-resolution structures have been determined are then described. Finally, we discuss the functional implications of these structures and the challenges remaining to defining the molecular basis for their various activities.

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2. Structural methods 2.1. Spectroscopy Various biophysical methods can provide low-resolution structural information more quickly and often utilizing much less protein than the higher resolution methods described below. Circular dichroism (CD) spectroscopy is a very powerful tool for characterizing protein secondary structure and detecting conformational changes over a range of conditions, which might include changes in pH, temperature and solvent or the presence of a binding partner (Kelly et al., 2005). While CD spectroscopic analysis of peptides and proteins in aqueous solution is relatively straightforward, studies in lipid vesicle systems are more challenging because of differential light scattering effects caused by the vesicles. Moreover, the high lipid:protein ratios usually required to mimic physiological conditions result in low protein concentrations and thus low intensity spectra. Synchrotron radiation CD (SRCD) spectroscopy is able to overcome these problems, and has the advantage that spectra can be recorded to lower wavelengths (around 175 nm instead of 190 nm), thereby providing more accurate secondary structure information (Miles and Wallace, 2006). As an example, SRCD has been used recently to examine membrane interactions of the pore-forming actinoporin equinatoxin II (EqtII) (Miles et al., 2008). Fourier transform infrared (FTIR) spectroscopy can also be used to monitor secondary structure, and is applicable to proteins bound to membranes (Barth, 2007). A recent study of the interaction of the actinoporin sticholysin II with membranes (Alegre-Cebollada et al., 2007) demonstrates that valuable information can be obtained by this approach, although overlap between protein and lipid signals can be problematic. Another drawback of infrared spectroscopy studies of proteins in aqueous solution is the strong absorbance of water in the mid-infrared spectral region (near 1645 cm1), which overlaps with several important protein backbone and side chain absorbance bands. This requires the use of short path lengths (around 5 mm) and high protein concentrations for aqueous samples. To overcome this problem, samples can be prepared in 2H2O since its absorbance band is shifted to w1210 cm1. Another form of vibrational spectroscopy which provides essentially the same information as FTIR but can be used in aqueous solution is Raman spectroscopy (Barth, 2007). This method has not been used extensively in studies of toxins but was applied in one of the early studies of sea anemone toxin conformation to assess secondary structure content and monitor side chain environments (Prescott et al., 1976). Raman optical activity, which is measured as a small circularly polarized component in Raman-scattered light from chiral molecules, is a promising extension of conventional Raman spectroscopy for studying proteins in aqueous solution (Barron, 2006) and may be useful in future studies of toxin structure and interactions. Fluorescence spectroscopy is a potentially valuable technique for probing the environment of residues that contribute to the observable signal, but suffers from the limitation that only a few residue types fluoresce and even these may be quenched in a native protein. The information

available from fluorescence studies of toxins bound to membranes has been reviewed recently using the cholesterol-dependent cytolysins from bacteria as one example (Johnson, 2005), and several applications to sea anemone cytolysins have been reported (e.g. Alvarez et al., 2001; Gutierrez-Aguirre et al., 2004). Fluorescence energy transfer is another powerful tool for studying molecular interactions and conformational changes, and single-molecule detection techniques promise to extend the reach of this and other fluorescence methods to unprecedented spatial resolution (Haustein and Schwille, 2004). 2.2. Atomic resolution structures The principal methods employed for the determination of atomic resolution structures of peptides and proteins are nuclear magnetic resonance spectroscopy and X-ray crystallography. Of these two methods, NMR spectroscopy has been used more widely for solving sea anemone toxin structures. NMR can be applied without isotopic labelling to small proteins up to 6–7 kDa in mass (Wuthrich,1986; Hinds and Norton, 1994) although the exact limit depends on the resolution of the 1H NMR spectrum and the magnetic field strength being used (generally speaking, the higher the better). Fortunately, a significant number of interesting toxins, not just from sea anemones but also other venomous creatures, fall within this mass range. For larger proteins, isotopic labelling is required (Yokoyama, 2003; Foster et al., 2007). Up to 10 kDa, 15N labelling coupled with a few straightforward three-dimensional NMR experiments should produce a good quality structure, but beyond this point double labelling with 15N and 13C is required. Isotopic labelling requires that the protein be expressed and refolded in a heterologous expression system that can grow on minimal media supplemented with 15N- and 13C-labelled sources of nitrogen and carbon, respectively. Bacterial systems are usually preferred, but the over-expressed toxin proteins often form inclusion bodies, in which case it is necessary to solubilize the expressed protein under denaturing and reducing conditions and then refold the protein in vitro (Gallagher and Blumenthal, 1992). Alternatively, folded toxins can be obtained by bacterial expression of soluble fusion proteins from which they are released by protease cleavage (Moran et al., 2006, 2007; Stehling et al., 2008). Yeast offers the advantage that the desired protein can be secreted with native disulfide bonds already formed, while insect cells are used much less often, but they can support isotopic labelling (Strauss et al., 2005). The enormous investments made in several countries in structural genomics/proteomics have benefitted all stages of the structure determination pipeline, whether by NMR or X-ray crystallography (Gileadi et al., 2007; Phillips et al., 2007). Methods for producing proteins, assessing whether they are suitable candidates for structure determination, collecting and analyzing the data, and evaluating the final structures obtained have all undergone significant improvements, which have in turn flowed back to the entire structural biology community, including those interested in toxins. It has been argued that structural genomics has less to offer toxinology because many potent toxins have similar folds but quite distinct biological

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activities (Norton, 2002), such that information about the fold per se is unlikely to provide clues to function. Nonetheless, structural studies of toxins have benefitted in other ways from structural genomics/proteomics. One significant advance in the field of protein NMR is the program CANDID (Herrmann et al., 2002), which allows high-resolution structures to be determined once NMR resonance assignments are complete. It is no longer necessary to manually assign the many hundreds of cross-peaks in a NOESY spectrum. In principle, CANDID is expected to be more reliable for larger proteins but we have found it to be valuable even for peptide and small protein structures such as analogues of the Kþ channel toxin ShK (Pennington et al., in press). When combined with isotopic labelling and 3D NMR methods it promises to be even more reliable. A recent report implies that high-resolution structures can be determined largely on the basis of chemical shift information alone (Shen et al., 2008). If this proves to be true for peptide and protein toxins it will accelerate the process of structure determination even further. X-ray crystallography, although a more mature method for determining high-resolution structures, continues to advance. Robotic crystallization trials have made it easier to identify suitable conditions for growing diffraction-quality crystals, while at the same time the minimum size of a crystal needed to provide a high-resolution structure continues to shrink as synchrotron radiation sources become more readily available (Sorensen et al., 2006; Manjasetty et al., 2008). Crystal structures of polypeptide toxins have traditionally been difficult to obtain because such toxins are often small and highly charged, so growing crystals has been a challenge. In fact, none of the sea anemone peptide toxin structures described in this article has been determined by X-ray crystallography, even though such toxins have been crystallized (Smith et al., 1984). By contrast, several crystal structures have been reported for peptide toxins from cone shells, both alone (Hu et al., 1996) and in complex with receptor homologues (Celie et al., 2005). Finally, we make note of another method for structure determination which is currently capable of producing only low-resolution structures, but is applicable to membranebound channels and receptors and may eventually be utilized for toxins bound to these molecules. Electron microscopy at cryogenic temperatures (electron cryomicroscopy or cryo-EM) and single-particle reconstruction have advanced to the point where there are now many examples of structures that have been solved to <10 Å resolution (Jiang and Ludtke, 2005). At this resolution, direct identification of ordered secondary structure elements is possible. Recently, a 4.5 Å resolution structure of a virus capsid was determined using this approach (Jiang et al., 2008). As discussed below, this method has been used to obtain low-resolution structures for the voltage-gated Naþ channel and a tetrameric pore-like structure of an actinoporin bound to a lipid monolayer. 3. Structures The peptide and protein toxins from sea anemones for which high-resolution structures have been determined are summarized in Table 1.

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3.1. Sodium channel toxins The first representatives of the Naþ-channel binding proteins were isolated in the early and mid-1970s by Laszlo Be´ress and co-workers at the University of Kiel and Ted Norton and co-workers at the University of Hawaii (Be´ress, 1982; Norton, 1991). It is now recognized that this group of toxins consists of at least three classes of peptides, two made up of molecules containing around 45–50 amino acid residues and one of shorter peptides containing 27–32 residues. ‘‘Long’’ peptides from the genera Anthopleura and Anemonia (family Actiniidae) have been classified as Type 1, and those from the Indopacific genera Heteractis (formerly Radianthus) and Stichodactyla (formerly Stoichactis), members of the family Stichodactylidae, as Type 2 (Kem, 1988; Kem et al., 1989). These two classes of peptides are similar with respect to the locations of the six half-cystines (which form three disulfide bonds), as well as several other residues thought to play a role in biological activity or maintenance of the tertiary structure. However, while there is extensive sequence homology (60%) within each class, there is only about 30% homology between the two classes (Norton, 1991). Interesting new representatives of these and other classes of anemone toxins continue to be isolated (Honma and Shiomi, 2006; Honma et al., 2008; Shiomi, 2009). The amino acid sequences of these and other sea anemone toxins described in this article are shown in figures in the Supplementary Material. These toxins bind to site 3 on voltage-gated sodium channels (VGSC), one of at least six toxin binding sites identified on vertebrate and insect channels (Ceste`le and Catterall, 2000; Catterall et al., 2007). The pore-forming asubunit of each VGSC consists of four homologous domains, each containing six putative transmembrane helices S1–S6, with the Naþ channel thought to be formed by the S5–S6 loops from all four domains (these S5–S6 linkers are further subdivided into the S5–P, P and P–S6 loops, with the P loop containing the SS1 and SS2 segments). Fujiyoshi and coworkers (Sato et al., 2001) have determined a low-resolution structure for the eel VGSC by means of cryo-electron microscopy; the channel has a bell-shaped outer surface, with several inner cavities connected to four small holes and eight orifices close to the extracellular and cytoplasmic membrane surfaces. Site 3, which the anemone toxins share with scorpion a-toxins, involves extracellular loops of domains I and IV of the VGSC (Thomsen and Catterall, 1989; Rogers et al., 1996), but the structure is not sufficiently well resolved to allow this (or other) toxin binding sites to be localized. The effect of site 3 toxin binding is to delay channel inactivation such that the channel remains open for a longer period and the action potential is prolonged (Wanke et al., 2009). It has been proposed that the bound toxin may interfere with conformational changes in the S3–S4 loop upon translocation of the S4 segment, which is an important part of the channel’s voltage sensor (Rogers et al., 1996). The structure of a Type 1 toxin, anthopleurin-A (AP-A) (Pallaghy et al., 1995), is shown in Fig. 1. It consists of a fourstranded, anti-parallel b-sheet linked by three loops, the first of which, spanning residues 8–16, is the largest and least well defined in solution, although it contains several residues essential for activity. The Type 2 toxin ShI (Fogh

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Table 1 Major classes of sea anemone toxins for which structures have been determined by NMR or X-ray crystallography. Class Naþ channel toxins Long Type 1

Long Type 2 Short (Type 3) Other Kþ channel toxins

Toxin

Species

Target

ATX I (As I) Anthopleurin-A (AP-A) Anthopleurin-B (AP-B) CgNa ShI ATX III (As III) APETx2

Anemonia sulcata Anthopleura xanthogrammica Anthopleura xanthogrammica Condylactis gigantea Stichodactyla helianthus Anemonia sulcata Anthopleura elegantissima

NaV1 NaV1 NaV1 NaV1 NaV1 (crustacean) NaV1 (crustacean, insect) ASIC3

ShK BgK BDS-I APETx1

Stichodactyla helianthus Bunodosoma granulifera Anemonia sulcata Anthopleura elegantissima

Kv1, IKCa Kv1, IKCa, molluscan Kþ channels Kv3.4 HERG

Equinatoxin II (EqtII) Sticholysin II (St II)

Actinia equina Stichodactyla helianthus

Membranes containing sphingomyelin Membranes containing sphingomyelin

ShPI AEI

Stichodactyla helianthus Anemonia sulcata

Proteases (Kunitz-type inhibitor) Proteases (‘non-classical’ Kazal-type elastase inhibitor)

Actinoporins

Protease inhibitors

The solution structure of a synthetic analogue of ShK, ShK-Dap22, has also been solved (Kalman et al., 1998), as has the crystal structure of an EqtII mutant containing a double cysteine mutant (Val8 to Cys, Lys69 to Cys) (Kristan et al., 2004). The amino acid sequence of EqtII is identical to that of tenebrosin-C from the anemone Actinia tenebrosa (Simpson et al., 1990). Protease inhibitors are included in this table even though they are not toxins; they may potentiate the actions of other toxins by inhibiting their proteolytic degradation and there are examples of ion channel inhibitors with the Kunitz-type fold in sea anemones (e.g. the kalicludines) and other species.

et al., 1990; Wilcox et al., 1993) has a very similar structure (Fig. 1). Both of these structures fall within the defensin family according to the structural classification database SCOP (Murzin et al., 1995). More recently the structure of CgNa (Sta¨ndker et al., 2006) has been solved (Salceda et al., 2007); this 47-residue toxin from the large Caribbean sea anemone Condylactis gigantea has greater sequence similarity to Type 1 toxins but displays structural homology with both Types 1 and 2 toxins. Calitoxin, from the anemone Calliactis parasitica, is a 46residue toxin with three disulfide bonds, but a sequence showing several significant differences from those of Types 1 and 2 toxins (Cariello et al., 1989) (Fig. S1). It nonetheless exerts effects on crustacean and insect giant axon sodium channels similar to those of the Types 1 and 2 toxins (Salgado and Kem, 1992). Gene sequencing has subsequently identified an analogue with a Glu6 / Lys substitution (Spagnuolo et al., 1994). Two shorter sea anemone toxins, ATX III (27 residues, three disulfide bonds) and PaTX (31 residues, four disulfides), also interact with the sodium channel, probably at site 3 (Norton, 1991). There is clear sequence similarity between them, but very little with the longer toxins (Fig. S2). More recently, three crab toxins of 30–32 residues were isolated from the anemones Dofleinia armata and Entacmaea ramsayi (Honma et al., 2003a), which were homologous to each other and also to PaTX, indicating that they belong to a distinct family of toxins. ATX III has a welldefined structure in solution (Manoleras and Norton, 1994), consisting of four reverse turns and two other chain reversals, but no regular a-helix or b-sheet structure (Fig. 1), and no close structural homologues. APETx2 is unrelated in sequence to the other Naþchannel toxins described here, but is closely related to the Kþ-channel toxins APETx1 and BDS described below

(Fig. S3). It inhibits the acid-sensing ion channel ASIC3, which is a proton-gated Naþ channel that has been implicated in pain transduction associated with acidosis in inflamed or ischemic tissues (Diochot et al., 2004). Despite the lack of sequence similarity, its structure (Chagot et al., 2005b) is similar to those of AP-A and ShI although the flexible loop found between the second and third Cys residues in AP-A and ShI (Fig. 1) is truncated in APETx2. 3.2. Potassium channel toxins A unique family of potassium channel blockers has been identified in anemones. The first representative to be isolated and characterized thoroughly was ShK, from Sticho˜ eda et al., 1995; Pennington et al., dactyla helianthus (Castan ˜ ada and Harvey, 2009) and since then 1995; Casten a number of others have been isolated (Norton et al., 2004; Hasegawa et al., 2006; Honma and Shiomi, 2006). The known sequences for these potassium channel blockers fall into two groups, with ShK, HmK and AETX K lacking the basic four-residue insert found in the N-terminal half of AeK, AsKS (also known as kaliseptine) and BgK but having two single-residue inserts in the C-terminal half (Fig. S4). The solution structure of ShK toxin (Tudor et al., 1996) represents a novel fold consisting of two short a-helices encompassing residues 14–19 and 21–24, and an Nterminus with an extended conformation up to residue 8 followed by a pair of interlocking turns that resembles a 310-helix (Fig. 1). Its structure contains no b-sheet and is thus quite distinct from the a/b fold found in scorpion Kþ channel blockers such as charybdotoxin (Bontems et al., 1992), but is similar to that determined subsequently for BgK toxin (Dauplais et al., 1997). Another family of peptides from anemones also acts on Kþ channels; the first members to be characterized were

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Fig. 1. Structures of various classes of sea anemone toxins. All of these structures were determined for the peptide in solution using NMR spectroscopic data. Category, toxin name, PDB accession code and references are as follows: Long Type 1 toxin, anthopleurin-A (AP-A), 1AHL (Pallaghy et al., 1995); Long Type 2 toxin, ShI, 1SHI (Wilcox et al., 1993); Short Naþ-channel toxin, ATX III, 1ANS (Manoleras and Norton, 1994); Kþ-channel blocking toxin ShK, 1ROO (Tudor et al., 1996); actinoporin equinatoxin II (EqtII), 1KD6 (Hinds et al., 2002) (a crystal structure has also been determined for EqtII, as described in RCSB ID code 1IAZ (Athanasiadis et al., 2001)). Structures are shown approximately to scale. Note the cluster of hydrophobic residues at the base of the ATX III structure. Based on Fig. 1 of Norton (2006).

BDS-I and -II, which were isolated some time ago before their target was defined, but have been shown recently to block the Kv3.4 Kþ channel (Diochot et al., 1998). The related toxin APETx1 specifically inhibits human ether-ago-go-related gene (HERG, Kv11.1) channels and shares 54% homology with BDS-I (Diochot et al., 2003), although they show no sequence homology with other Kþ channel toxins from sea anemones. Recently, Honma et al. (2005) described three new crab toxins from Antheopsis maculata, one of which, Am II, also belongs to this family. The precursor proteins of these three toxins consist of a signal peptide, a pro region concluding with a pair of basic residues (Lys-Arg), and the mature peptide; the Am I precursor protein contains up to six copies of Am I. The structures of BDS-I (Driscoll et al., 1989a,b), APETx1 and APETx2 (Chagot et al., 2005a,b) are similar to those of

the Naþ-channel toxins such as AP-A (Fig. 1) but quite different from the ShK/BgK family of Kþ-channel toxins. Clearly, anemones are capable of using a common structural scaffold to create blockers of distinct targets (AP-A, APETx1 and APETx2 act on VGSC, HERG and ASIC channels, respectively), while also using different scaffolds (all-b in APETx1 vs all-a in ShK) to block similar channels (HERG and Kv1, respectively). The targets of these toxins are Kþ channels, in particular the six-transmembrane channels, which include the voltage-gated Kþ channels (Kv) and the small-conductance and intermediate-conductance Ca2þ-activated Kþ channels (KCa) (Gutman et al., 2003; Catterall et al., 2007). In these channels the region between the fifth and sixth transmembrane segments (the pore loop) forms the ion conduction pathway, and four subunits come together to

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form a functional channel tetramer. The Kv channels are voltage-activated, opening in response to membrane depolarization. The surface of ShK involved in binding to Kv channels has been probed using alanine scanning and selected toxin analogues (Pennington et al., 1996; Rauer et al., 1999). Two residues, Lys22 and Tyr23, are crucial for activity, as also found subsequently for BgK toxin (Dauplais et al., 1997) but other residues also contribute to the Kþ channel binding surfaces. ShK toxin blocks Kþ channels by binding to a shallow vestibule at the outer entrance to the ion conduction pathway and occluding the entrance to the pore (Kalman et al., 1998; Lanigan et al., 2002). It appears that Lys22 and Tyr23 in the toxins represent a conserved dyad of residues that is essential for Kþ-channel blockade by a range of structurally unrelated peptide toxins from scorpions, snakes and cone snails (Dauplais et al., 1997; Norton et al., 2004). Me´nez and co-workers (Gasparini et al., 1998) proposed that this motif be more broadly defined as a lysine and a neighboring hydrophobic residue. ShK toxin blocks the voltage-gated channel Kv1.3 at very low (picomolar) concentrations. As Kv1.3 is crucial for the activation of terminally differentiated effector memory T cells, it is regarded as a promising target for the treatment of T-cell mediated autoimmune diseases such as multiple sclerosis and the prevention of chronic transplant rejection. ShK and its analogues are currently undergoing evaluation as leads in the development of new biopharmaceuticals for the treatment of multiple sclerosis and other T-cell mediated autoimmune disorders (Norton et al., 2004; Beeton et al., 2006). Understanding the structural basis for its activity is therefore of considerable benefit in designing more potent, selective and stable analogues (Pennington et al., in press). 3.3. Cytolysins The actinoporins are a family of sea anemone toxins that function by forming pores in cell membranes (Kem, 1988; Anderluh and Macˇek, 2002). These highly basic proteins, of mass 18–20 kDa (Fig. S5), display permeabilizing activity in model lipid and cell membranes that is markedly enhanced by the presence of sphingomyelin (SM). The actinoporins differ from bacterial pore-forming toxins in several respects: they are more potent, the pore they form does not have a stable structure and has not yet been visualized directly, and they are of smaller size and very stable toward proteolysis. In common with many pore-forming toxins, the actinoporins are highly water soluble, stable proteins, and yet their only known activity is the formation of oligomeric pores in membranes, consisting of three or, more likely, four monomers. The resulting pores have a radius of about 1 nm and are permeable to small molecules and solutes, with the resulting osmotic imbalance promoting cell lysis (Anderluh and Macˇek, 2002). SM plays a key role in the lytic activity of the actinoporins. The hemolytic activity of a cytolysin from S. helianthus was inhibited by pre-incubation with SM, and treatment of erythrocyte membranes with sphingomyelinase rendered them resistant to lysis (Bernheimer and Avigad, 1976). The hemolytic activity of equinatoxin II

(EqtII) was also inhibited by pre-incubation with SM. More recent studies with sticholysins I and II on model membranes confirmed that SM enhances lytic activity and suggested that cholesterol may have a minor role (de los Rios et al., 1998). The interaction of EqtII with large unilamellar vesicles containing phosphatidylcholine (PC) was reversible and did not involve major conformational changes (Caaveiro et al., 2001), but the presence of SM enabled irreversible membrane insertion and pore formation, which were associated with major conformational changes. Some EqtII-induced leakage was observed from large unilamellar vesicles containing only PC and cholesterol, again suggesting a minor contribution of this sterol lipid to actinoporin cytolytic activity. Fluorescence studies of EqtII binding to lipid vesicles showed that association was markedly enhanced by the presence of SM (Anderluh and Macˇek, 2002). Recent studies by Anderluh and co-workers have provided evidence for a direct interaction between EqtII and SM, with the two most important residues for SM recognition being the exposed aromatic residues Trp112 and Tyr113 (Bakracˇ et al., 2008). Their data also highlight the importance of phase boundaries for EqtII activity (Scho¨n et al., 2008), suggesting a dual role of SM as a specific receptor for the toxin and a promoter of the membrane organization necessary for EqtII action. Three-dimensional structures have been determined for EqtII by X-ray crystallography (Athanasiadis et al., 2001) and in solution by NMR (Hinds et al., 2002), and for ˜ o et al., sticholysin II by X-ray crystallography (Manchen 2003). EqtII consists of two short helices packed against opposite faces of a b-sandwich structure formed by two five-stranded b-sheets (Fig. 1). A number of studies have been carried out in an effort to elucidate the molecular mechanism of actinoporin pore formation. By site-directed mutagenesis, it was shown that at least two regions of EqtII become embedded in lipid membranes, the N-terminal region (residues 10–28) and the surface aromatic cluster including tryptophans 112 and 116 (Anderluh and Macˇek, 2002). The current model of pore formation proposes that EqtII binds to the membrane initially via the aromatic rich region (Fig. 2A), then the Nterminal region is transferred to the lipid membrane and across the bilayer to form a final functional pore (Hong et al., 2002; Anderluh et al., 2005). The structural details of the final oligomeric assembly are not known, but a hint as to how this might look is afforded by sticholysin II, which crystallized on lipid monolayers as tetrameric pore-shaped structures with fourfold symmetry. Three-dimensional reconstruction of electron microscopic images of these two-dimensional crystals produced a low-resolution model ˜ o et al., 2003). Such for the tetrameric pore (Manchen a structure would fit the concept of a toroidal pore in which the functional pore is lined with both lipid and protein (Valcarcel et al., 2001; Yang et al., 2001), as depicted in Fig. 2B. 3.4. Other anemone toxins Various protease inhibitors have been isolated from anemones, including several Kunitz inhibitors (Be´ress, 1982; Delfin et al., 1996; Honma and Shiomi, 2006) and,

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Fig. 2. (A) Possible models of the orientation of EqtII at the membrane surface, based on 19F NMR studies of EqtII with all five Trp residues replaced by 5-F-Trp. The Trp side chains coloured as follows: W112 red, W116 magenta, W117 gold, W149 green, W45 blue. This diagram is based on Fig. 7 of Anderluh et al. (2005). (B) Tentative model of the functional pore of St II. Left: top view of the putative functional St II pore, in which four St II monomers are shown in different colours. Right: front view of the same pore, with the yellow monomer omitted for clarity. The lipid headgroup regions are indicated as gray layers. Note that the walls of the pore would be lined by four helices and lipid molecules. Modified with permission from Fig. 7 of Manchen˜o et al. (2003). (For interpretation of colour in this figure, the reader is referred to the web version of this article.)

more recently, cysteine proteinase inhibitors such as equistatin (Lenarcˇicˇ et al., 1997). The solution structures of two of these protease inhibitors have been determined, a Kunitz-type inhibitor from S. helianthus (Antuch et al., 1993) and a ‘‘nonclassical’’ Kazal-type elastase inhibitor from A. sulcata (Hemmi et al., 2005). Protease inhibitors such as these are not toxic in their own right although they may potentiate the action of toxins in venom by blocking proteolysis of those toxins in prey or predator tissues and thus prolonging their effective lifetime. However, an intriguing finding has been made in some anemones, where proteins with this fold also display potassium channel blocking activity. This was first observed with the kalicludines (AsKC 1–3) from A. sulcata (Schweitz et al., 1995), which have both protease inhibitory and Kv1 blocking activities, although with less potency than bovine pancreatic trypsin inhibitor and toxins such as ShK, respectively. Recently, another example of a dual-activity

protein with a Kunitz fold has been found in SHTX III from Stichodactyla haddoni (Honma et al., 2008). As noted by Lazdunski and co-workers (Schweitz et al., 1995), ‘sea anemones appeared at least 800 million years ago and it may be that bifunctional molecules such as kalicludines are survivors of a remote past, and that evolution has gradually given rise, for efficiency, to molecules fully specialized either for trypsin inhibition or for blockade of the voltagesensitive Kþ channel’. It is possible that Kþ-channel blockers based on the Kunitz fold are an evolutionary vestige, with the function of Kþ-channel blockade in many anemone venoms being taken over by the ShK family of toxins. Several other classes of potentially toxic peptides and proteins have been identified in sea anemones, although structures have not yet been determined. One example is an epidermal growth factor-like toxin, gigantoxin I, from Stichodactyla gigantea (Honma et al., 2003b). Its precursor

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protein is composed of a signal peptide, a pro region and the mature peptide, similar to those of gigantoxins II and III. As these proteins are found in nematocysts, they may well function as toxins. 4. Relating structure to function It is beyond the scope of this article to discuss in detail the molecular basis for activity of the various classes of sea anemone toxin for which structures have been presented. In some cases, for example the Naþ-channel toxin ATX III, virtually nothing was known about which residues on the molecular surface are essential for activity (Manoleras and Norton, 1994) until a recent study of its interaction with insect channels (Moran et al., 2007) (note that in this study ATX III is referred to as Av3). In others, such as the long Naþ channel toxins, extensive studies by chemical modification and limited proteolysis have provided valuable information on structure–function relationships (Norton, 1991; Monks et al., 1994). Solid-phase peptide synthesis (Pennington et al., 1990a,b, 1994) offered a more systematic approach but was not pursued extensively for this class of toxins. For Type 1 toxins, site-directed mutagenesis of recombinant material proved to be the most informative approach, with numerous mutants of anthopleurin-B being described by Blumenthal and co-workers (e.g. Khera et al., 1995) and used to map interactions with site 3 on the voltage-gated Naþ channel (Benzinger et al., 1998; Smith and Blumenthal, 2007). In ShK toxin, the key residues for interaction with the Kþ-channel were well characterized using synthetic analogues (Pennington et al., 1996; Rauer et al., 1999), and models of how this and related toxins might bind to the target channel have been constructed based on extensive complementary mutational data (Kalman et al., 1998; Gilquin et al., 2002; Lanigan et al., 2002; Gilquin et al., 2005) (Fig. 3). Such models are not a substitute for experimental structures but in the absence of those they represent a valuable tool to guide further studies. Not all sea anemone toxins active against Kþ channels are pore blockers; a recent study indicates that APETx1 is a gating modifier toxin of the HERG channel (Zhang et al., 2007). A complete understanding of the molecular basis for activity of toxins, as with most other peptides and proteins, will come only from high-resolution structures of toxins in complex with their target receptors. This has proven to be a considerable challenge and indeed has not yet been achieved for any of the anemone toxins described in this article. However, it has been accomplished for a-conotoxins in complex with a receptor mimic, the acetylcholine binding protein from Aplysia californica (Celie et al., 2005; Hansen et al., 2005; Ulens et al., 2006), and the insights derived from these structures underscore their value in dissecting the molecular basis for toxin function. Solid-state NMR studies offer an alternative approach to understand toxin interactions with membrane-bound receptors. For example, Baldus and co-workers (Lange et al., 2006) have shown that high-affinity binding of the scorpion toxin kaliotoxin to a chimeric KcsA–Kv1.3 channel is associated with structural rearrangements in both the toxin and the channel and that conformational flexibility of the

interacting partners may therefore be important. Fig. 3B shows that significant chemical shift changes occurred for residues in both the pore helix and the selectivity filter upon kaliotoxin binding. The solid-state NMR data also showed that Asp64 in the vestibule of KcsA–Kv1.3 represents an important interaction site, a finding supported by the observation that mutation of the homologous residue in Kv1.3 channels reduced kaliotoxin affinity by more than three orders of magnitude (Aiyar et al., 1995). More recent studies described evidence of heterogeneity in the binding modes of kaliotoxin, which might enhance its affinity for KcsA–Kv1.3 via entropic stabilization (Zachariae et al., 2008). 5. Flexibility There is increasing evidence that protein dynamics in solution, particularly on microsecond to millisecond timescales, correlate closely with protein function. NMR relaxation measurements on a single-domain bacterial signalling protein demonstrated that there was a strong correlation between phosphorylation-driven activation and microsecond timescale backbone dynamics (Volkman et al., 2001). The unphosphorylated (inactive) and phosphorylated (active) states are in equilibrium, and phosphorylation shifts that equilibrium toward the active species. Thus, activation involves primarily a dynamic population shift between two preexisting conformations. A similar situation appears to apply for enzymes; in dihydrofolate reductase each intermediate in the catalytic cycle samples low-lying excited states whose conformations resemble the ground-state structures of preceding and following intermediates, with turnover rates being influenced by the dynamics of transitions between ground and excited states of the intermediates (Boehr et al., 2006). How does this apply to toxins? It has been known for some time that several of the long Naþ channel toxins contain two conformers in slow equilibrium with one another, manifest in the appearance of peak splitting in their NMR spectra (Gooley et al., 1984, 1988). Subsequently, it was shown in AP-A that this arises from cis–trans isomerism of the peptide bond preceding Pro41 (Scanlon and Norton, 1994) and that in the major form of the protein this bond adopts a cis conformation. Depending on the relative affinities of these two conformers for site 3 on the Naþ channel, this interconversion has the potential to affect both the kinetics and the thermodynamics of channel binding. Another type of conformational flexibility is also illustrated by the long Naþ channel toxins. These structures consist of a twisted, four-stranded, anti-parallel b-sheet linked by three loops (Fig. 1), the first and largest of which is less well defined by the NMR data than the bulk of the structure in all cases. Indeed, this flexible loop may be one reason why these toxin structures have not yielded to X-ray crystallography. This loop contains several residues known from chemical modification and site-directed mutagenesis studies, together with comparison of natural homologues, to be important for activity, at least in the case of the anthopleurins. Blumenthal and co-workers found that mutating Gly10 or Gly15 to Ala in AP-B diminished Naþ

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Fig. 3. (A) Side view of ShK docked into a model of the Kv1.3 channel. The channel subunit nearest the viewer has been removed. Toxin and channel side chains are coloured as follows: ShK Arg11 cyan, Lys22 and Arg24 blue, Tyr23 purple, Met21 yellow; Kv1.3 Asp386 red, Asp402 gold, His404 magenta and Ser379 orange. In this view the side chains of Tyr23 and Arg11 of ShK are partially obscured. The subunits of the channel model are coloured with different shades of green. This diagram is based on Fig. 2 of Lanigan et al. (2002). (B) Summary of solid-state NMR results for kaliotoxin binding to a KcsA–Kv1.3 chimera. Amino acid sequence of the KcsA–Kv1.3 pore region with secondary structures (boxes, s.f. stands for selectivity filter). Residues characterized by significant chemical shift changes upon complex formation are coloured red; those involved according to mutagenesis (Mut.) and molecular dynamics (MD) are marked with an asterisk. Blue, unperturbed residues; black, not determined. Perturbed (red) or unperturbed (blue) residues are indicated in surface plots of the channel (RCSB 1K4C). Models of two and four membrane-inserted KcsA subunits are also shown. Reproduced with permission from Lange et al. (2006). (For interpretation of colour in this figure, the reader is referred to the web version of this article.)

channel binding affinity, suggesting that the inherent flexibility of this loop may play an essential role in binding and isoform selectivity (Seibert et al., 2003). Assuming that the internal motions in this loop are largely frozen in the channel-bound state, the intrinsic flexibility in the ligand dictates that there will be a loss of conformational entropy upon binding which will reduce the binding affinity unless compensated for by increased flexibility elsewhere in the bound ligand or the channel. Examples of such compensatory changes have been reported (MacRaild et al., 2007). More subtle types of motion in toxins are revealed by NMR relaxation studies. This is illustrated using recent studies of the m-conotoxin m-SIIIA (Yao et al., 2008), although we expect the same considerations will apply to toxins generally. Internal motion of the backbone was detected over various timescales, with pyroGlu1, Asn2 and Ser9 showing larger magnitude motions on the ps–ns timescale and Cys4, Trp12 and Cys13 undergoing

conformational exchange on the ms to ms timescale. Even the presence of three disulfide bonds in a short polypeptide of just 20 residues was not sufficient to confer rigidity on the polypeptide backbone. Indeed, it may even be that conformational averaging associated with the disulfide bridges was the cause of the observed conformational exchange, as has been suggested for u-conotoxin MVIIA (Atkinson et al., 2000). An awareness of the presence of conformational exchange is valuable in several ways. Interpreting measured NOEs as structural restraints requires some caution if either or both of the interacting nuclei are from regions undergoing such exchange. Knowing which regions of a structure are potentially flexible is also important if modelling studies are to be undertaken based on that structure. And, most important of all, it must be taken into account when using structures determined in solution as a basis for understanding receptor binding or constructing mimetics (Baell et al.,

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2002). Whether conformational exchange serves any important biological function, for example in allowing the toxin to undergo required conformational rearrangements as it engages its cognate binding site, or whether it is simply an adventitious consequence of the presence of multiple disulfide bridges in certain environments, remains to be established. One way of addressing this question will be to replace some of the disulfide bridges with less flexible homologues, and means to achieve this are now available (Robinson et al., 2007). This approach may also stabilize toxins that are prone to disulfide exchange processes in serum (Armishaw et al., 2006).

6. Structure prediction Fashion models whose photos appear in magazines and newspapers usually bear little resemblance to real people. Is the same true of structural models that appear in the scientific literature? Are they any more credible? The various rounds of the CASP (Critical Assessment of Structure Prediction) project, where models are constructed for proteins for which the structure has been determined but not disclosed to the modellers, certainly show an encouraging improvement in the ability of modelling to predict the correct structure (Moult et al., 2007). Recently, we have had the opportunity to test the veracity of a toxin model constructed by homology modelling by comparing it with the actual experimental structure. The example is a m-conotoxin, but once again the conclusions are relevant to toxin modelling more generally. Models of m-conotoxins SIIIA and KIIIA were generated by comparative modelling methods using the solution structure of m-SmIIIA as a template, as described by Bulaj et al. (2005). Comparison with the solution structure of m-SIIIA determined subsequently (Yao et al., 2008) showed that the C-terminal halves of the two were very similar (backbone RMSD 1.1 Å for residues 11–20) and the helical region (residues 11–16) was conserved, but the N-terminal region was not so well determined, with a backbone RMSD of 2.6 Å over residues 1–10. The difference in the N-terminal region may be attributed to structural differences between m-SIIIA and m-SmIIIA, which was used as the starting point for the model. In fact, the simulated structure of m-SmIIIA showed some differences from the structure determined from NMR data in the N-terminal region (Keizer et al., 2003), and it may be that there is genuine flexibility in this part of the molecule. Thus, in this example at least, the homology modelled structure had some memory of its origins and its accuracy suffered as a result. It is likely that a model structure of m-SIIIA constructed from the recently determined structure of the more closely related m-conotoxin KIIIA (unpublished results) would match the actual structure more faithfully. This example is presented not with the intention of discouraging modelling of toxin structures but to emphasize the need to base the model on a structure as close as possible to the target structure and, preferably, to construct several models based on alternative starting structures. As the number of toxin structures in the Protein Data Bank grows, so do the prospects for generating accurate models of newly identified structures.

One final note of caution is appropriate, and that concerns disulfide bonding patterns. Recently, we reported the structure of the conotoxin i-RXIA (Buczek et al., 2007), the first representative of the I1 superfamiliy of conotoxins (Norton and Olivera, 2006). This toxin has four disulfide bridges, which were mapped both chemically and via structure calculations based on NMR data. The connectivity pattern proved to be I–IV, II–VI, III–VII and V–VIII, the same pattern seen in the Sydney funnel web spider toxin, robustoxin (Pallaghy et al., 1997). However, in a different spider toxin, J-atracotoxin, which shares an identical cysteine framework with I-conotoxins, NMR studies combined with a partial reduction/alkylation technique showed that the cysteine connectivities were I–VI, II–VII, III–IV, and V–VIII (Wang et al., 2000). Thus, the apparently conserved cysteine frameworks did not lead to the same disulfide connectivities. Indeed, i-RXIA is more similar to Jatracotoxin (in terms of spacing between cysteine residues) than to some other I-superfamily members characterized from Conus species and it remains to be seen which disulfide patterns are adopted by these other I-superfamily members. Another example of different disulfide connectivities occurring for toxins with similar amino acid sequences is provided by the m-conotoxins, where at least three different patterns of disulfide connectivity have been found in peptides with the same cysteine framework (Han et al., 2006; Du et al., 2007). The important point for this discussion, however, is that it is critical to ensure that the disulfide bridging pattern is known before embarking on model construction. 7. Perspectives The genome of one sea anemone, Nematostella vectensis, has already been determined (Putnam et al., 2007), and sequence information is available for another cnidarian, the hydra H. magnipapillata (Sher et al., 2005). As genome sequencing capacity expands and is applied in future to other cnidarians, the toxins community will be presented with an unprecedented array of targets for structural and functional studies. For those with recognizable sequence similarity to known toxins it should be straightforward to identify their major biological targets and to construct reasonably accurate structural models, with the caveat that the disulfide bonding patterns must be known. But there will also be many exciting new classes of toxins or toxin-like molecules to characterize. Here we will be able to take advantage of continually improving bioinformatic tools to assist in identifying likely targets and structures, and of the enhanced methodologies generated by structural genomics enterprises to solve toxin structures more efficiently. The ultimate goal of defining the structures of toxins bound to their cognate receptors remains a major challenge, but ongoing improvements in methods for producing biologically competent membrane-bound proteins and determining their structures, albeit at low resolution, suggest that even this will be achievable, although perhaps never routine, in the near future. We can look forward, therefore, to a greatly improved understanding of the molecular basis for activity of all classes of toxin. This

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information will also enhance efforts to harness the therapeutic potential of many of these toxins in treating a range of diseases. Acknowledgements I am very grateful to Ronelle Welton for assistance with sequence analyses, Jennifer Sabo for help with the figures and Charles Galea and Chris MacRaild for helpful comments on the manuscript. I am pleased to dedicate this article to my colleagues Laszlo Be´ress and Ted Norton (no relation to the author), formerly of the Universities of Kiel and Hawaii, respectively, whose enormous enthusiasm for sea anemone toxins helped to stimulate my interest in this field. Conflict of interest The author declares that there are no conflicts of interest. Appendix. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi: 10.1016/j.toxicon.2009. 02.035

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