Nematotoxicity of Marasmius oreades Agglutinin (MOA) Depends on Glycolipid Binding and Cysteine Protease Activity

Fruiting body lectins have been proposed to act as effector proteins in the defense of fungi against parasites and predators. The Marasmius oreades agglutinin (MOA) is a Galα1,3Gal/GalNAc-specific lectin from the fairy ring mushroom that consists of an N-terminal ricin B-type lectin domain and a C-terminal dimerization domain. The latter domain shows structural similarity to catalytically active proteins, suggesting that, in addition to its carbohydrate-binding activity, MOA has an enzymatic function. Here, we demonstrate toxicity of MOA toward the model nematode Caenorhabditis elegans. This toxicity depends on binding of MOA to glycosphingolipids of the worm via its lectin domain. We show further that MOA has cysteine protease activity and demonstrate a critical role of this catalytic function in MOA-mediated nematotoxicity. The proteolytic activity of MOA was dependent on high Ca(2+) concentrations and favored by slightly alkaline pH, suggesting that these conditions trigger activation of the toxin at the target location. Our results suggest that MOA is a fungal toxin with intriguing similarities to bacterial binary toxins and has a protective function against fungivorous soil nematodes.

Fruiting body lectins have been proposed to act as effector proteins in the defense of fungi against parasites and predators. The Marasmius oreades agglutinin (MOA) is a Galα1,3Gal/GalNAc-specific lectin from the fairy ring mushroom that consists of an Nterminal, ricin B-type lectin domain and a Cterminal dimerization domain. Latter domain shows structural similarity to catalytically active proteins, suggesting that, in addition to its carbohydrate-binding activity, MOA has an enzymatic function. Here, we demonstrate toxicity of MOA towards the model nematode Caenorhabditis elegans. This toxicity depends on binding of MOA to glycosphingolipids of the worm via its lectin domain. We further show that MOA has cysteine protease activity and demonstrate a critical role of this catalytic function for MOA-mediated nematotoxicity. The proteolytic activity of MOA was dependent on high Ca 2+ concentrations and favoured by slightly alkaline pH, suggesting that these conditions trigger activation of the toxin at the target location. Our results suggest that MOA is a fungal toxin with intriguing similarities to bacterial binary toxins and has a protective function against fungivorous soil nematodes.
A wide variety of lectins with different structures and specificities has been isolated from fungi (1). Recently, cytoplasmic fungal lectins, also referred to as fruiting body lectins, were proposed to be part of a fungal defense system against parasites and predators. This hypothesis is based on the carbohydrate-binding dependent toxicity of several fungal lectins towards nematodes, insect larvae and amoebae (2). The exclusive and abundant expression of these lectins in fruiting bodies or sclerotia of the host fungus may result in a special protection of these reproductive organs. It is proposed that predators such as fungivorous nematodes ingest the cytoplasmic lectin when feeding on the content of the fungal cell and that binding of the lectin to specific glycans in the intestine of the worm induces toxicity by an unknown mechanism (2-5).
The Marasmius oreades agglutinin (MOA) is a type B erythrocyte agglutinating lectin that was isolated from fruiting bodies of the fairy ring mushroom. The dimeric lectin consists of 293residue protomers and has been shown to specifically recognize Galα1,3-containing structures, exhibiting highest affinity for the branched blood group B trisaccharide Galα1,3(Fucα1,2)Gal (6). Crystal structures of MOA in complex with the xenotransplantation epitope and the blood group B trisaccharide revealed an N-terminal ricin B-type lectin domain with three canonical carbohydrate binding sites and a C-terminal domain that is involved in dimerization. The C-terminal domain showed structural homology to catalytically active proteins such as peptide-N-glycanases, Nacetyltransferases, transglutaminases and cysteine proteases. This structural similarity was limited to a putative catalytic triade consisting of Cys215, His257 and Glu274 (7,8). However, no catalytic activity of MOA has been reported so far. Moreover, the biological target glycan and the function of the lectin remained unclear. Like other fruiting body lectins, MOA lacks a classical secretion signal and is thus predicted to be localized in the cytoplasm. An endogenous function of the lectin is therefore unlikely as glycosylated structures are generally secreted or localized in the extracellular space (9). Accordingly, no glycoconjugate recognized by MOA has thus far been identified in fungi.
In this study, we investigated the possible role of MOA as a fungal defense molecule against nematodes, using C. elegans as a genetically tractable model system. First, we demonstrate carbohydrate-binding dependent nematotoxicity of MOA and identify glycosphingolipids as the ligands of MOA in C. elegans. Second, we show that MOA-mediated toxicity is dependent on the previously suggested catalytic function that we identified as Ca 2+ dependent cysteine protease activity. Our results suggest that MOA is a fungal toxin that protects the fruiting body from predators such as fungivorous nematodes.

EXPERIMENTAL PROCEDURES
Strains, cultivation conditions, cloning and site-directed mutagenesis. Strains, primer sequences and experimental procedures can be found in SI Materials and Methods.
C. elegans toxicity assays and statistical analysis. Biotoxicity assays with C. elegans were performed as described (10). Results were analyzed using a t-test for pairwise comparisons.
Protein expression, purification and biotinylation. MOA and MOA(C215A) were expressed recombinantly in E. coli and purified as described (6,11). Detailed information can be found in SI Materials and Methods. t-Cry5B was prepared as previously described (12). For TLC overlay analysis, purified MOA and t-Cry5B were labeled with EZ-Link Sulfo-NHS-Biotin (Pierce) using a 20-fold molar excess of labeling reagent. Labeling proceeded for two hours on ice followed by desalting on a PD-10 column.
Screen for MOA-resistant C. elegans mutants using Mos1 insertional mutagenesis and light microscopy of C. elegans. Mos1 insertional mutagenesis, stereomicroscopy and differential interference contrast (DIC) microscopy of C. elegans was performed as described (3).
Preparation and analysis of glycolipids from C. elegans. Glycolipid extraction from C. elegans N2, bre-2 and bre-4 strains and TLC overlay staining was performed as described by Barrows et al (13). For overlay analysis biotinylated MOA and t-Cry5B were used at a concentration of 150nM.
In vitro protease assay. Briefly, 20 μg of denatured RNase A was incubated with 0.1 ng to 1 μg of purified recombinant MOA or MOA(C215A) in assay buffer (40 mM Tris-HCl pH 8.0, 1 mM CaCl 2 , 1% NP-40) in a final reaction volume of 30 μl for 1 h at 37 °C. Samples were analyzed by SDS-PAGE and Coomassie blue staining. Detailed information is provided in SI Materials and Methods.
Cleavage site specificity. 40 µg of bovine asialofetuin, bovine casein and human hemoglobin were digested with 100 ng of MOA in 40mM Tris-HCl pH8.0, 1mM CaCl 2 for 1h at 37°C. The reaction was stopped by addition of 25 mM EDTA. Samples were desalted using C18 ZipTips (Millipore) and analyzed on an LTQ-Orbitrap XL mass spectrometer (Thermo Fischer Scientific, Bremen, Germany) coupled to an Eksigent-Nano-HPLC system (Eksigent Technologies, Dublin (CA), USA). Detailed information on MS analysis and data evaluation is provided in SI Materials and Methods.

MOA inhibits C. elegans development and amoebal growth dependent on its carbohydratebinding ability.
Based on the results with other fruiting body lectins (2), we tested MOA for toxicity towards the nematode Caenorhabditis elegans and the amoeba Acanthamoeba castellanii. These biotoxicity assays involved feeding of the organisms with MOA-expressing E. coli as described previously (10) and revealed a clear toxicity of MOA towards C. elegans and A. castellanii ( Fig. 1; Fig. S1). Both C. elegans and A. castellanii were fed with E. coli cells expressing either authentic MOA, the C-terminal deletion mutant MOA(C) or the carbohydratebinding deficient variant MOA(Q46A,W138A) (7). In a control experiment, test organisms were grown on vector-containing E. coli transformants. In case of C. elegans, severe inhibition of larval development was observed for N2 wildtype and pmk-1(km25) mutant animals feeding on MOA-expressing bacteria compared to the vector control (p<0.05) (Fig.  1A). The pmk-1(km25) mutation in the p38 mitogen-activated protein kinase (MAPK) pathway was previously shown to cause hypersensitivity to different kinds of stress including exposure to fungal lectins (3). In contrast, MOA(Q46A,W138A) had no significant effect on the development of the worms (p>0.05). Similar results were obtained with A. castellanii (p<0.05) (Fig. S1). Previous studies showed that residues Q46 and W138 are involved in glycan coordination (7,8) and that the binding activity of the double mutant to thyroglobulin is reduced compared to wild-type MOA (14). The lacking toxicity of MOA(Q46A,W138A) suggests that nematotoxicity of MOA is dependent on the carbohydrate-binding activity of the protein. This is also supported by the decreased nematotoxicity of MOA(C) (p<0.05), a mutant that showed reduced carbohydrate-binding activity possibly due to loss of dimerization (14) (Fig. 1A).
Physiological effects of intoxication were studied by comparative light-and differential interference contrast (DIC) microscopy of C. elegans L4 larvae fed with MOA-expressing or vector-containing E. coli cells. MOA-intoxicated animals showed an expanded lumen of the anterior intestine compared to control animals ( Fig. 1B). A similar phenotype was observed with other nematotoxic fungal lectins (3) and bacterial toxins (15).
The ligand of MOA in C. elegans is a glycosphingolipid. In order to identify the target glycoconjugate of MOA in C. elegans, a forward genetic screen for mutations conferring MOAresistance was performed. Strain pmk-1(km25) was used to increase the sensitivity of the screen. Four MOA-resistant mutants were isolated in the screen, all of which carried mutations in exons of the bre-3 gene. Bre-3(op504) and bre-3(op505) mutants had a Mos1 insertion in the seventh exon of bre-3. Bre-3(op506) mutants carried a Mos1 insertion in the fifth exon of the same gene. In an independent screen, one mutant (op508) with a Mos1 insertion site identical to the one of bre-3(op504) was identified ( Fig. 2A). Bre-3 encodes for a glycosyltransferase that acts in a biosynthetic pathway building up an invertebrate-specific glycosphingolipid that is recognized by the Bacillus thuringiensis crystal toxin Cry5B (16). The oligosaccharide core of the ceramide-linked glycan is synthesized by the sequential action of BRE-3 (β1,4mannosyltransferase), BRE-5 (β1,3-GlcNActransferase), BRE-4 (β1,4-GalNAc-transferase) and BRE-2 (putative β1,3-galactosyltransferase). Bre-1 encodes for an enzyme involved in the conversion of GDP-mannose into GDP-fucose (17). Given the resistance of bre-3 mutant animals we tested worms with mutations in the other bre-genes for sensitivity towards MOA. We observed that also mutations in bre-4(ye27) and bre-5(ye17) genes conferred resistance to MOA (p<0.05) whereas bre-1(ye4) and bre-2(ye31) mutant worms were sensitive to intoxication with MOA (p>0.05) (Fig. 2B). These results suggest that fungal MOA, like bacterial Cry5B, targets a glycosphingolipid but that the species is different from the one recognized by Cry5B.
In order to confirm the genetic data, direct binding of MOA to glycolipids isolated from N2, bre-2(ye31), and bre-4(ye27) mutant worms was assessed in a thin layer chromatography (TLC) overlay. For this purpose lipids were extracted from C. elegans, partitioned into lower phase lipids and upper phase glycolipids and subsequently purified as described by Barrows et al. (13). Purified glycolipids were separated by TLC using chloroform:methanol:water (4:4:1) as a solvent and visualized using Orcinol sulfate (Fig. 2C, lane 2-4). The main glycolipid bands were observed in positions similar to those reported previously (12,16). TLC overlay staining of upper phase glycolipids with MOA revealed binding to N2 and bre-2(ye31) but not to bre-4(ye27) glycolipids (Fig. 2C, lane 5-7). This is consistent with MOA-susceptibility of N2 and bre-2(ye31) worms and resistance of bre-4(ye27) animals. To identify the glycolipid species that is recognized by MOA, comparative overlay staining with truncated Cry5B (t-Cry5B) as described by Ideo et al. (12) was performed. The pattern of the Orcinol staining and the t-Cry5B overlay of N2-derived glycolipids (Fig.  2C, lane 1) indicated that the main glycolipid species recognized by MOA corresponds to component D according to the classification of Griffitts et al. that has been previously characterized (16,18). Binding to the glycan part of this glycosphingolipid of the arthroseries (Galα1,3GalNAcβ1,4GlcNAcβ1,3Manβ1,4Glc, Fig. 2D) (16,18) is in agreement with the Galα1,3Gal specificity of MOA. Finally, MS and MS/MS analysis of glycolipids extracted from the main TLC band recognized by MOA further supported the presence of component D including the ceramide moiety composed of a saturated hydroxy fatty acid with 22 and 24 carbon atoms, respectively (Fig. S2). Besides component D, MOA is able to bind to a number of unidentified glycolipid species from both N2 and bre-2(ye31) animals (Fig. 2C, lanes 5, 6). These glycolipid species are absent from the bre-4(ye27) animals.
MOA shows in vitro protease activity. X-ray crystallographic studies of MOA in complex with the xenotransplantation epitope and the blood group B trisaccharide revealed that the C- terminal part of MOA adopts a fold that is observed in catalytically active proteins such as peptide N-glycanases, N-acetyltransferases, transglutaminases and cysteine proteases (7,8). As the highest homology was observed to a peptide N-glycanase (PNGase) we tested purified recombinant MOA in an in vitro PNGase assay described by Suzuki (19). However, instead of deglycosylation of the glycoprotein RNase B, degradation of the substrate was observed, indicating that MOA may have protease-rather than peptide Nglycanase activity. To confirm this hypothesis, we performed an in vitro protease assay with non-glycosylated RNase A as substrate (Fig. 3). MOA efficiently degraded denatured RNase A within 1 h at 37 °C. In contrast, MOA(C215A), in which the cysteine residue of the putative catalytic triad was replaced by an alanine, had no effect on RNase A, confirming that the observed protease activity was MOA-dependent. In accordance with these results, degradation of the substrate protein by MOA was abolished in the presence of 50 μM E-64, an irreversible, highly selective cysteine protease inhibitor, but not in the presence of 2 mM PMSF, an irreversible serine protease inhibitor or 2 mM AgNO 3 , a potent trypsin and chymotrypsin inhibitor (Fig.  3A). Based on these results, we conclude that MOA has, in addition to its glycan-bindingproperties, cysteine protease activity.
Structural homology to catalytically active proteins was first observed in a crystal structure of MOA in complex with the xenotransplantation epitope (7). A second crystal structure of MOA complexed with the blood group B trisaccharide revealed that binding of two Ca 2+ ions close to the hypothetical active site in the C-terminal domain causes a conformational change opening up a large cleft containing the potential active site (8). To investigate the influence of divalent cations on MOA protease activity, reactions in assay buffer supplemented with 1mM CaCl 2 , MgCl 2 , MnCl 2 , and ZnCl 2 , respectively, were performed. In the presence of Ca 2+ ions proteolytic degradation of RNase A by MOA was observed. In contrast, MOA showed hardly any activity in buffers supplemented with MgCl 2 , MnCl 2 or ZnCl 2 or in the absence of divalent cations (Fig. 3B). This observation is in accordance with the structural data and suggests that Ca 2+ binding activates the catalytic activity of MOA. We found that a minimal concentration of 50 μM was required for MOA to exhibit protease activity and that this activity further increased upon raising the Ca 2+ concentration up to 5 mM (Fig. S3).
To test the influence of the pH on the reaction, protease assays in buffers of different pH-values were performed. Complete degradation of RNase A by MOA was observed at pH 8.0. At higher pH, MOA only partially degraded the substrate whereas under acidic conditions the enzyme was not active (Fig. 3C). Therefore MOA protease activity is favoured by a slightly alkaline pH. Presence or absence of the carbohydrate ligand Galα1,3Galβ1,4GlcNAc did not affect the proteolytic activity of MOA (Fig.  S4).
In order to characterize the substrate specificity of MOA, we tested its activity towards different denatured and native protein substrates in the in vitro assay. Bovine serum albumin (BSA), fluorescein isothiocyanate (FITC) labeled casein, asialofetuin, and hemoglobin were degraded when denatured before incubation with MOA (Fig. S5A). In the native state, only FITC-casein, asialofetuin and hemoglobin were susceptible to MOA protease activity whereas BSA, RNase A and RNase B were not degraded (Fig. S5B). Proteolytic degradation of FITC-casein by MOA could also be detected in a fluorometric assay, where it showed a 3.6 fold higher activity/mole at pH 8.0 than the cysteine protease papain at pH 7.5 (Fig.  S6). As MOA did not act on every tested native protein substrate we conclude that the protease has a certain substrate specificity.
To investigate the cleavage site specificity of MOA, native asialofetuin, casein or hemoglobin were digested with purified MOA and the resulting peptides were analyzed by LC-ESI-MS/MS. This procedure yielded 83 cleavage sites that were used to create an iceLogo (20) of the preferred sequence specificity, where the cleavage occurs between position 4 and 5 (Fig.  3D). The results of this analysis suggest that MOA prefers proline and proline or valine at position 0 and 2, respectively. A digest of C. elegans proteome with MOA yielded similar results (data not shown), confirming the specificity observed for the model substrates.
Protease dimers like wild-type MOA as confirmed by analytical gel filtration (Fig. S7). The toxicity of MOA(C215A) was significantly reduced in N2 animals and pmk-1(km25) mutants compared to wild-type MOA (p<0.05) (Fig. 4A). Similar results were obtained for A. castellanii (p<0.05) (Fig. S1). Based on these results, we conclude that, in addition to carbohydrate-binding, the cysteine protease activity of MOA is necessary for full toxicity. In C. elegans, the contribution of the catalytic domain to the toxicity of MOA is also apparent by the fact that pmk-1(km25) mutant and N2 wild type worms are equally susceptible towards wild-type MOA (Fig 1A, Fig  4A). This is in contrast to nematotoxic fungal lectins lacking a catalytic domain which display a higher toxicity to pmk-1(km25) mutant worms (2,3). Accordingly, abolishment of the catalytic activity, as in the case of MOA(C215A) and MOA(C), results in higher susceptibility of pmk-1(km25) worms compared to N2 worms (( Fig 1A, Fig 4A). The higher toxicity of MOA(C215A) in comparison to MOA(C) can be explained by the lack of dimerization in latter case. These results suggest that the toxicity mechanism of MOA differs from the one of nematotoxic lectins without a catalytic domain.
As the protease activity of MOA could be inhibited by E-64 in vitro, pmk-1(km25) animals were grown on MOA expressing bacteria in the presence of 10 to 500 μM of this highly selective cysteine protease inhibitor. The cell-permeable inhibitor significantly reduced MOA-mediated toxicity in a dose-dependent manner. On the other hand, E-64 did not impair development of worms feeding on vector-containing E. coli (Fig.  4B). These results confirm the role of MOA cysteine protease activity in nematotoxicity. As observed for the catalytic site mutant in the case of pmk-1(km25) worms, toxicity was not completely abolished by inhibiting protease activity even at the highest concentration of E-64.
Remaining toxicity at inhibitor concentrations higher than 50 μM can be referred to the toxicity mediated by the lectin domains of the dimer, which are not affected by E-64.

DISCUSSION
Fruiting body lectins have been proposed to be part of the fungal defense system against predators and parasites (2). In agreement with such a function, we demonstrate that the fruiting body lectin MOA is toxic towards the nematode C. elegans and the amoeba A. castellanii. As MOA mutants deficient in carbohydrate-binding exhibited no or significantly reduced toxicity for both organisms, we conclude that the lectinactivity of the protein is essential for its toxicity. Morphological changes of MOA-intoxicated worms include an enlargement of the intestinal lumen, an effect that has previously been reported for the fungal lectin CGL2 from C. cinerea (3) and the B. thuringiensis crystal toxin Cry5B (15). These results suggest that binding of the lectin to a target glycan in the worm intestinal epithelium is essential for nematotoxicity. We hypothesize that the lectin is stored in the cytoplasm of the fungal cell and is released and transferred to the intestine of fungivorous nematodes upon predation.
Using C. elegans genetics, we could identify the target glycoconjugate of MOA in this organism that is responsible for its toxicity. Similar to the nematotoxic Cry5B toxin from Bacillus thuringiensis, MOA binds to different species of the invertebrate-specific arthroseries of glycosphingolipids in C. elegans. Based on TLC overlays and MS analysis, the main species recognized by MOA is component D according to the annotation by Griffitts et al. (16). This finding is consistent with the genetic results as bre-3, bre-5 and bre-4 (but not bre-2) encode glycosyltransferases involved in the biosynthesis of this specific glycosphingolipid. In addition to this previously identified glycosphingolipid species, MOA bound to several unknown glycolipids present in N2 and bre-2(ye31) but not in the bre-4(ye27) mutant worms. These results indicate the presence of multiple additional glycosphingolipid species carrying terminal Galα1,3Gal in C. elegans. The full sensitivity of bre-1(ye4) worms suggests that in contrast to glycosphingolipid-binding Cry5B, fucosylation of the lipid is not required for MOA binding.
The putative catalytic activity of MOA was investigated in in vitro assays with purified recombinant protein. In these assays, MOA showed proteolytic activity towards several native and denatured model substrates. The lack of activity towards some native protein substrates suggests a certain substrate specificity. The proteolytic activity of MOA was dependent on a cysteine residue in the predicted catalytic site, on the presence of Ca 2+ and had a pH optimum of 8.0. The C-terminal domain of MOA does not show homology to any existing proteinase family currently classified in the MEROPS database. Interestingly, the peptidase domain of MOA is the smallest in terms of residues that has ever been described and is even smaller than the HIV1 retropepsin. The crystal structure suggests that the active site cysteine is very close to the calcium binding sites, and the glutamine that would be an ideal candidate for forming the oxyanion hole is also a calcium ligand. Thus, the chemistry around the active site of this protease domain appears to be very 'busy' (N. D. Rawlings, personal communication).
Intriguingly, we could show that, in addition to carbohydrate-binding, the proteolytic activity of MOA is required for full toxicity towards C. elegans as toxicity of a catalytic site mutant is decreased and toxic effects of MOA are reduced in the presence of the highly specific cysteine protease inhibitor E-64. The equal susceptibility of N2 and pmk-1(km25) worms towards wild type MOA, as opposed to catalytically inactive MOA variants, suggests that the catalytic activity adds an additional twist to this lectin with regard to its mechanism of toxicity.
As determined by in vitro digests of different model substrates, MOA preferentially cleaves sequences with proline or valine at position P2. Interestingly, a similar sequence specificity has been described for calpains, a family of calciumdependent, non-lysosomal cysteine proteases that are expressed ubiquitously in mammals and many other organisms (21) and are involved in regulation of signal transduction pathways, cell motility, and apoptosis (22,23). In contrast to calpain that proteolyzes its substrates in a limited manner exhibiting modulatory functions by cutting interdomain regions, our analysis using model substrates suggests that MOA digests protein substrates to small peptides. The specificity for sequences containing proline may allow MOA to act on a wide range of proteins, as proline residues are generally located in turns at the protein surface. However, further studies with more physiological substrate proteins are required to strengthen this hypothesis.
Chimeric lectins homologous to MOA are present in the basidiomycetes Polyporus squamosus (PSL) (24) and Schizophyllum commune (SCA) (25). In PSL, the catalytic triad and the four Asp residues taking part in Ca 2+binding are conserved. The lectin domain of PSL, however, has been shown to be specific for Neu5Acα2,6Galβ1,4Glc/GlcNAc groups of N-glycans (24,26,27). In contrast, SCA has a similar carbohydrate-binding specificity as MOA (will be published elsewhere) but the catalytic residues Cys, His and Glu of MOA are replaced by Ser, Ser and Asp, respectively, and only one of the four Asp residues involved in Ca 2+binding is conserved suggesting that the cysteine protease activity is not conserved in this homologue.
As one possible toxicity mechanism, binding of MOA to glycosphingolipids in the nematode intestinal epithelium may result in digestion of surrounding proteins and ultimately lead to disintegration of the intestinal epithelium. A similar mechanism has been proposed for the maize cysteine protease Mir1-CP, which has been shown to impair the growth of fall armyworm (Spodoptera frugiperda) larvae by disrupting the insect's peritrophic matrix in the intestine (28)(29)(30). Such proteases are used by plants, besides lectins and protease inhibitors, as defense molecules against herbivores (31) in line with our hypothesis of MOA being involved in fungal defense. Interestingly, Mir1-CP was recently shown to synergize the insecticidal activity of Bacillus thuringiensis toxins (32).
On the other hand, the carbohydrate-binding deficient but protease-proficient variant MOA(Q46A,W138A) was absolutely non-toxic even at the high lectin concentrations in the biotoxicity assay with recombinant bacteria (Fig.  1) and the lumen of the C. elegans intestine is thought to be slightly acidic based on the pH optima of digestive hydrolases (33), arguing against such a mechanism. In addition, the nature of its ligand (receptor) as well as the structural and functional organization of MOA are strikingly similar to bacterial AB 5 toxins such as the Cholera and the Shiga toxin. These proteins consist of a non-toxic B-subunit which binds to specific glycolipids and an enzymatically active A-subunit that induces toxicity by blocking the GTPase-activity of G α and activating adenylate cyclase (Cholera toxin) or by inactivating ribosomal protein biosynthesis (Shiga toxin). The intoxication process by these proteins is initiated by attachment of the toxin via the B-subunit followed by internalization of the AB 5 -receptor complex by receptor mediated endocytosis and retrograde transport to the endoplasmic reticulum (ER) (34). Finally, the Asubunit of these toxins is translocated to the cytoplasm by the sec translocon. This final step is omitted in the case of the subtilase cytotoxin SubAB that induces cell death by exhibiting highly specific protease activity towards the ERresident chaperone BiP (35). It is remarkable that SubAB produces pathological features in mice that strongly resemble the hemolytic uraemic syndrome (HUS) in humans (35), a characteristic that has also been observed for MOA (36). These results suggest that SubAB and MOA may have a similar toxicity mechanism. In accordance with such a hypothesis, the combination of neutral pH (37) and a Ca 2+ concentration of more than 100 μM in the endoplasmic reticulum (38) meet the requirements of MOA to exhibit protease activity. On the other hand, MOA is not expected to be catalytically active at the neutral pH but low Ca 2+ concentration (~ 100 nM) of the cytosol (37,39) which allows storage of this toxic protein in an inactive form in the cytoplasm of the fungal cell. Similarly, the slightly acidic pH in the intestinal lumen of the nematode should inhibit the proteolytic activity of MOA despite of the presumably high Ca 2+ concentrations in this compartment (33). Such a regulation of its proteolytic activity via pH and Ca 2+ concentration could explain the apparent lack of autoproteolytic activation or of pronounced substrate specificity of MOA. We therefore hypothesize that, upon binding to the glycosphingolipid receptor present on intestinal epithelial cells in the nematode, MOA is internalized to the early endosomal compartment by endocytosis and transported via the Golgi to the ER, where its catalytic domain is activated and may induce toxicity by degrading proteins that are in the folding process (Fig. S8). Further experiments are required to corroborate this hypothetical toxicity mechanism.
expressing MOA mixed with E-64 diluted in M9 buffer at the indicated concentrations. Development of the animals from L1 to L4 was analyzed as outlined above. As a control the experiment was performed with E. coli BL21(DE3) containing empty vector. Data shown represent the average of 9 biological replicates. Error bars indicate standard deviations.