Afp18 gene harbours a putative glycosyltransferase domain

Y. ruckeri Afp18 is a component of the prophage tail-like injection machinery (Afp) and exhibits similarities with Serratia Afp18 in terms of size and amino-terminal architecture, but differs in the carboxyl-terminal toxic domain (Fig. 1a,b, scheme). The toxic domain of Yersinia Afp18 comprises a putative glycosyltransferase domain (Fig. 1b, green coloured region), which exhibits significant sequence similarity to glycosyltransferase toxins from Legionella pneumophila (Lgt1–3), Photorhabdus asymbiotica (PaTox) and clostridial glycosylating toxins, including toxin A and B from Clostridium difficile (Fig. 1b, sequence alignment)29. All these glycosyltransferases contain a conserved DxD-(aspartic acid- × -aspartic acid) motif, which is essential for sugar donor substrate binding and, thus, crucial for enzymatic activity. Mutations of this motif result in catalytic defective enzymes7,30,31.

Afp18G severely affects early zebrafish embryo development

To assess the cellular effects and toxicity of Afp18, we isolated the DNA and cloned the variable carboxyl-terminal fragment (Afp18G; amino acids 1,771–2,123) comprising the putative glycosyltransferase domain. In our studies, we used Y. ruckeri isolated from an infected rainbow trout in Idaho, USA. This strain is identical to Y. ruckeri recently isolated from a wound infection of a 16-year-old male patient in Belgium32. We purified the Afp18G protein in E. coli and microinjected the recombinant protein into zebrafish zygotes at one-cell stage. In addition, we constructed an Afp18G mutant with an exchange of the DxD motif against an enzymatically non-functional NxN, which we injected as control. Afp18G NxN injected embryos developed normally and were indistinguishable from non-injected or buffer injected control embryos (Fig. 1c). Afp18G-injected embryos performed the first three to four cell divisions with morphologically visible cleavage planes between the dividing blastomeres (Fig. 1c, 16-cell stage, arrows), albeit progress of development was delayed compared with controls. At the 256-cell stage, 2.5 h post fertilization (h.p.f.), control embryos showed normal development of the blastoderm positioned on top of the large vegetal yolk cell. In contrast, Afp18G-injected embryos failed to establish the typical multilayered organization of the blastoderm and large sections of the blastoderm were devoid of morphologically discernible cell boundaries (Fig. 1c, 256-cell stage, arrow). About 1 h later, at the onset of gastrulation, control embryos initiated epiboly, a coordinated cell movement, in which the static blastomeres became motile and spread vegetalward to cover the yolk cell. In contrast, Afp18G-injected embryo did not initiate epiboly, the blastoderm disrupted and in most severe cases the yolk cell and (or) blastomeres lysed and embryos completely disintegrated (Fig. 1c, 30% epiboly, arrowhead and asterisk, respectively). These data reveal that the glycosyltransferase domain of Afp18 is crucial for its severe toxic effect on early zebrafish development.

Dose-dependent effects of Afp18G on embryo development

To rule out contribution of effects by contaminants from the E. coli-derived protein preparation, we microinjected mRNA encoding the Afp18G protein. Dilution series of Afp18G encoding mRNA facilitated to determine effects of a wider range of toxin dosage. We injected 0.1–100 pg in vitro transcribed Afp18G or Afp18G NxN mRNA per embryo, or as control GFP mRNA. Morphological phenotypes were documented at the 1,000-cell stage (3.3 h.p.f.; Fig. 2a,b quantitative analysis; Supplementary Fig. 1). Embryos injected with 0.1 pg Afp18G mRNA did not develop significantly different from control embryos. About 30% of the embryos injected with 0.5 pg Afp18G mRNA developed a disintegrated blastoderm, with irregular cell shape, local loss of blastomere boundaries and abnormal cell sizes (arrowhead in Fig. 2c, 0.5 pg). Injection of 1 pg Afp18G mRNA per embryo affected cell morphology in all analysed embryos. Frequently, larger cells detached from the blastoderm and abnormal vesicles were formed (arrowhead in Fig. 2c). When 10 or 100 pg Afp18G mRNA were injected per embryo, cell morphology was severely affected, tissue integrity progressively lost during gastrulation and blastoderms frequently disintegrated (Fig. 2c and Supplementary Fig. 1a). In contrast, microinjection of mRNA encoding the glycosyltransferase-deficient mutant Afp18G NxN had no effect on embryonic development, revealing that the effects were indeed caused by the glycosyltransferase activity. Given that the effects of Afp18G on cellular functions cannot be analysed when embryos disintegrate early, we chose to inject 0.5 pg Afp18G mRNA for further analyses.

Figure 2: Afp18G disturbs zebrafish early development. (a) Live images of non-injected, GFP, Afp18G NxN mRNA (each 100 pg per embryo), or different amounts (0.1–100 pg per embryo) of Afp18G mRNA-injected embryos at 1,000-cell stage (3 h.p.f.). Embryos oriented animal to the top. Scale bar, 500 μm. (b) Quantification of the blastoderm phenotype of control and Afp18G-injected embryos shown in a at 1,000-cell (non-injected, n=41; GFP mRNA, n=37; Afp18G NxN mRNA, n=52; 0.1 pg Afp18G, n=35; 0.5 pg Afp18G, n=45; 1 pg Afp18G, n=60; 10 pg Afp18G, n=62; 100 pg Afp18G, n=60). Only values for 100 pg per embryo injected GFP and Afp18G NxN mRNA are shown. Developing live embryos were classified into categories ‘normal’ (WT like), ‘disrupted blastoderm’ (cellular structure of blastoderm abnormal) or ‘disintegrated embryos’ (blastoderm and (or) yolk cell lysed). The distribution of phenotypes was analysed for significant differences using Fisher exact probability test, revealing significant differences (*P values <0.0001) between Afp18G NxN control and Afp18G samples. (c) Optical image planes of live blastoderm regions at 1,000-cell (3 h.p.f.) of non-injected, GFP, Afp18G NxN or Afp18G mRNA-injected embryos (pg per embryo) oriented animal to the top. Control embryos show normal development of blastomeres, while the blastoderm progressively loses cellular integrity with increasing amounts of Afp18G mRNA injected. Arrowheads mark the disrupted regions without visible blastomere boundaries, abnormal sized and irregular shaped cells. Asterisks mark detached blastomeres and disintegrated blastoderm. Scale bar, 100 μm. (d) Live images of control, full-length Afp18 and full-length Afp18 NxN mRNA (each 50 pg per embryo) injected zebrafish embryos. Afp18-injected embryos disintegrate at 1,000-cell stage (3 h.p.f., upper row, embryos are oriented animal to the top) and degrade early in gastrulation (lower row 40% epiboly, 5 h.p.f.). Scale bar, 500 μm. For five different concentrations each of Afp18 (non-injected n=80; 2.5 pg n=34; 5 pg n=50; 12.5 pg n=65; 25 pg n=85; 50 pg n=38) and Afp18 NxN (2.5 pg n=43; 5 pg n=58; 12.5 pg n=50; 25 pg n=76; 50 pg n=23) mRNA injections, the bar graph shows percentage of embryos, which at 5 h.p.f. develop like WT controls or degrade. Full size image

The G domain is the major pathogenicity determinant in Afp18

Next we wanted to clarify whether parts of Afp18 other than the glycosyltransferase (G) domain affect zebrafish embryo development. Therefore, we injected mRNA encoding the full-length Afp18 protein, or the enzymatically non-functional Afp18 NxN mutant, and analysed embryos and early larvae to identify potential morphological alterations in embryogenesis. Full-length Afp18 caused severely degraded embryos during early gastrulation (40% epiboly; Fig. 2d), while Afp18 NxN injected embryos developed indistinguishably from non-injected wild-type (WT) control embryos (Fig. 2d). These data revealed that the glycosyltransferase domain is the Afp18 protein domain mediating toxicity in this assay.

Afp18G disrupts the actin cytoskeleton

To analyse the in vivo cellular components affected by the Afp18G toxin, we co-injected mRNAs encoding Lifeact-GFP (green fluorescent protein) to fluorescently label the actin cytoskeleton and histone H2B-dsRed to label nuclei in living embryos. On Afp18G mRNA injection, blastomeres were severely enlarged (Fig. 3a, asterisk). In addition, blastomeres frequently contained two or more nuclei. We assumed that these Afp18G-induced morphological alterations were caused by deregulation of F-actin cytoskeleton organization. The Lifeact-GFP signal suggested a reduced amount of polymerized actin in Afp18G-expressing embryonic cells (Fig. 3a, Supplementary Movie 1). To analyse the effects of Afp18G on the actin cytoskeleton in more detail, we introduced recombinant 6xHis-tagged glycosyltransferase domain protein into zebrafish ZF4 cells (embryo-derived fibroblast-like cell line)33, using protective antigen (PA; the binding and translocation component of anthrax toxin) as a delivery system34, and stained filamentous actin with TRITC-phalloidin. Cells to which the glycosyltransferase-deficient NxN-mutant was delivered (Fig. 3b, middle panel) had an actin cytoskeleton indistinguishable from untreated control cells. In contrast, cells to which Afp18G was delivered showed a massive loss of actin fibres (Fig. 3b, bottom right panel) and finally rounded up (Fig. 3b top right panel). Live imaging of ZF4 cells transfected with GFP-actin revealed a complete disassembly of the actin cytoskeleton after delivery of Afp18G (Fig. 3c, time series bottom row; top row Afp18G NxN-treated control cell). The actin-depolymerizing effect was remarkably rapid. During 60 min of incubation, actin fibres disappeared and ZF4 cells collapsed. Comparable effects were observed using human HeLa cells (Supplementary Fig. 3a). Thus, the glycosyltransferase activity of Afp18 appears to severely affect the regulation of cellular actin.

Figure 3: Afp18G affects the actin cytoskeleton. (a) Live images of Afp18G NxN or Afp18G mRNA (each 0.5 pg per embryo) injected embryos at 30% epiboly. Embryos are co-injected with mRNA encoding Lifeact-GFP (green) and H2B-dsRed (magenta; 100 pg per embryo each) labelling the F-actin cytoskeleton and the nuclei. Afp18G NxN embryos developed indistinguishable from WT embryos. Differential interference contrast (DIC) images of blastomeres are shown in the upper row, the corresponding confocal epifluorescence images below. Asterisk marks abnormally large blastomere; the arrow marks a blastomere with two nuclei. Single-plane image, scale bar, 20 μm. (b) Fluorescent micrographs of ZF4 cells treated with 6xHis-tagged Afp18G (right panel) or glycosyltransferase-deficient mutant Afp18G NxN (middle panel) proteins in combination with anthrax protective antigen (PA) as translocation system for His-tagged proteins. Top row shows phase-contrast images. Bottom row shows TRITC-phalloidin staining (red) of the actin cytoskeleton and a DAPI nuclei staining (magenta) of ZF4 cells fixed after 2 h. Arrows indicate regular stress fibres of F-actin in Afp18G NxN cells compared with disrupted F-actin fibres in Afp18G treated cells. Scale bar, 50 μm (top panel), 10 μm (bottom panel). (c) Time-lapse microscopic images of GFP-actin expressing HeLa cells intoxicated with Afp18G NxN (top row) and Afp18G (bottom row) as described in b. Scale bar, 10 μm. DAPI, 4′,6-diamidino-2-phenylindole. Full size image

Afp18G affects cytokinesis in early development

We examined whether the completion step of cell division, involving coordinated actin rearrangement, was affected by Afp18. We found that cytokinesis proceeded normally in Afp18G NxN expressing control embryos (Fig. 4a, upper row, arrow; Supplementary Movie 1). In contrast, cytokinesis including the assembly of the actomyosin ring and formation of the cleavage furrow was severely impaired in blastomeres of Afp18G mRNA-injected embryos, resulting in cells frequently containing more than one nucleus (Fig. 4a, lower row, arrow; Supplementary Movie 1). We evaluated cortical and cytoplasmic deposition of F-actin via measurement of the integrated density of Lifeact-GFP fusion protein fluorescence (Fig. 4b). The ratio of cytoplasmic versus cortical F-actin localization at three developmental time points (sphere, 30% epiboly, 50% epiboly) was almost constant in blastomeres of Afp18G NxN control injected embryos, when measured in 30 min time windows. In contrast, cortical actin was significantly enriched in blastomeres of Afp18G mRNA-injected embryos, indicating a deregulation of the dynamic organization of the actin cytoskeleton. However, the cell cycle phases involving proper assembly and organization of microtubule-based mitotic spindle progressed normally. Time series of Lifeact-GFP and H2B-dsRed labelled embryos revealed the accurate composition of the metaphase, with the sister chromatids moved to the opposite spindle poles for both, Afp18G NxN (Fig. 4a, upper row) and Afp18G (Fig. 4a lower row) mRNA-injected embryos. In summary, Afp18G seemed to selectively affect actin-dependent filament organization and dynamics during cytokinesis, while microtubule dynamics and karyokinesis appeared to progress normally.

Figure 4: Afp18G blocks cytokinesis leading to multinucleated cells. (a) Cytoskeleton and cytokinesis analysed by time series of Afp18G NxN (upper row) or Afp18G (lower row) mRNA (0.5 pg per embryo each) injected embryos at dome stage (4.3 h.p.f.). Embryos were co-injected with mRNA encoding Lifeact-GFP (green) and H2B-dsRed (magenta; 100 pg per embryo each) labelling the F-actin cytoskeleton and the nuclei. Asterisk marks dividing blastomeres. Afp18G NxN and Afp18G mRNA-injected blastomeres complete microtubule mediated mitotic phases including chromosome segregation. Arrows indicate the actin ring contraction during cytokinesis, which is strongly affected in Afp18G-dividing blastomeres. Confocal z-stack projection of 10-μm depth; scale bar, 10 μm. (b) Quantification of cytoplasmic versus cortical Lifeact-GFP integrated epifluorescence signal density at indicated developmental stages. Confocal image of Afp18G NxN (left side) and Afp18G (right side) injected embryo showing an example blastomere with manually defined areas of cortical (region depicted in between red and yellow selection) and cytoplasmic (region outlined by yellow selection) F-actin and analysed using Fiji-ImageJ software measure function (sphere, n=12 each; 30% epiboly n=12 each; 50% epiboly n=12 each). Scale bar, 10 μm. Values are average±s.e.m. Statistical significance was evaluated using Mann–Whitney test. Full size image

Bleb formation is impaired by Afp18G

The organization of the cortical actin network controls cell membrane protrusions, and, thus, has a strong influence on cell motility35,36. It was shown that spherical protrusions, called blebs, are formed dynamically at the membrane of migrating cells. We scored bleb formation of blastomeres of Afp18G and Afp18G NxN mRNA-injected embryos at the onset of epiboly. Fig. 5a and Supplementary Movie 2 show a time series of a forming bleb documented by differential interference contrast transmitted light (upper row, black arrowhead) and fluorescence microscopy of Lifeact-GFP (lower row, white arrowhead) of a control injected embryo (upper two rows). The three phases of a bleb life cycle—initiation (after 10 s), expansion (up to 30 s) and retraction (40 to 70 s)—were clearly visible. Initially, blebs form and grow devoid of actin, while during retraction fluorescently labelled Lifeact-GFP-actin signal appeared slightly enhanced. Blebbing was severely reduced in numbers but also in bleb size in Afp18G mRNA-injected embryos (Fig. 5a lower two rows), as quantified by bleb counts (Fig. 5a graph). In addition, we analysed other actin-driven protrusion behaviours of blastomeres, excluding blebbing. Time series of both Afp18G and Afp18G NxN mRNA-injected embryos showed blastomeres, which form lamellipodia driven by actin remodelling (marked by black and white arrows in differential interference contrast and fluorescence images, respectively; Fig. 5b and Supplementary Movie 2). Quantification revealed that blastomeres did not significantly differ in lamellipodia number (membrane protrusions with actin remodelling—see graph in Fig. 5b) when Afp18G and Afp18G NxN expressing embryos were compared.

Figure 5: Afp18G blocks cell blebbing without affecting lamellipodia and filopodia formation. (a) Bleb formation analysed at 30% epiboly (4.7 h.p.f.) by time series of embryos injected with 0.5 pg per embryo of Afp18G NxN mRNA (upper two rows) or Afp18G mRNA (lower two rows). Embryos were co-injected with mRNA encoding Lifeact-GFP (100 pg per embryo) labelling the F-actin cytoskeleton. DIC images are shown in the upper row and corresponding epifluorescence images in the lower row. Black arrowheads indicate spherical membrane protrusions of a blastomere, known as bleb formation, which are initially devoid of actin (white arrowheads). In contrast, blastomeres of Afp18G-injected embryos show significantly less blebbing and blebs appear smaller. Graph shows the quantification of bleb formation of blastomeres in Afp18G mRNA compared with Afp18G NxN control injected embryos (0.5 pg per embryo) during a time series (frames: 10-s intervals for 300 s; Afp18G NxN n=75; Afp18G n=75 cells analysed of three different embryos, 25 blastomeres each). Values are average±s.e.m.; statistical significance was analysed using Mann–Whitney test. Scale bar, 10 μm. (b) Protrusive activity analysed in time series of Afp18G NxN (upper two rows) or Afp18G (lower two rows) mRNA (0.5 pg per embryo each) injected embryos at 30% (4.7 h.p.f.). Embryos were co-injected with mRNA encoding Lifeact-GFP (100 pg per embryo) labelling the F-actin cytoskeleton. DIC images are shown in rows 1 and 3 and corresponding fluorescent image in the rows 2 and 4. Both Afp18G NxN and Afp18G mRNA-injected embryos show blastomeres forming sheet-like membrane protrusions resembling lamellipodia, filled with actin bundles and actin branches (arrows). Scale bar, 10 μm. Quantification shows the percentage of analysed blastomeres forming blebs without actin, blastomeres forming sheet-like membrane protrusions with actin or blastomeres without membrane protrusion activity during the captured time series (Afp18G NxN n=41; Afp18G n=37 cells analysed from three different embryos). The statistical significance (P=0.0091) of differences between the proportion of Afp18G NxN and Afp18G blastomeres showing these specific behaviours was computed using the Kruskal–Wallis test. The analysis revealed that the distribution of cell behaviour types indeed differs significantly, caused by changes in blebbing activity but not in lamellipodia formation. DIC, differential interference contrast. Full size image

Blebs have been reported to be primarily controlled by RhoA and its effector ROCK-I37,38. In contrast, Rac and Cdc42, which increase actin-dependent lamellipodia formation, inhibit bleb formation and amoeboid migration39,40. Therefore, our in vivo data suggested that Afp18G predominantly targeted RhoA, whereas Rac1 and Cdc42 might be less affected. To determine whether Afp18 may co-localize with RhoA, we used DNA vector injection to generate mosaic embryos in which individual cells were expressing enzyme-deficient EGFP-tagged Afp18G NxN and RHOA. Anti-EGFP and anti-RHOA immunofluorescence revealed that EGFP-Afp18G NxN appeared to target the cell membrane, and evaluation of line scans of fluorescent profiles shows co-localization with RHOA (Supplementary Fig. 2).

Afp18G utilize UDP-N-acetylglucosamine to modify Rho GTPases

To identify the cellular targets of Afp18 in zebrafish, we elucidated the sugar donor for this reaction by enzyme-catalysed UDP-[14C]sugar hydrolysis and found that Afp18G efficiently hydrolysed UDP-[14C]GlcNAc (Fig. 6a). Using UDP-[14C]GlcNAc in a glycosylation reaction with Afp18G and cell lysate, we identified proteins with an electrophoretic mobility corresponding to 23 kDa, which were labelled with 14C-GlcNAc and migrated similarly to the GTPase RhoA (Fig. 6b). As Rho GTPases are known regulators of the actin cytoskeleton and putative substrate candidates, we applied RhoA, Rac1 and Cdc42 to an in vitro glycosylation reaction with 1 nM Afp18G and obtained a strong modification of RhoA. Signals of Rac1 and Cdc42 were hardly visible under these conditions (Fig. 6c). When we applied higher amounts of Afp18G (100 nM), we could also observe the glycosylation of RhoB, RhoC, Rac2 and Rac3, and Cdc42 (Supplementary Fig. 3b). Other subfamily members of the Rho family were not modified and also Ras proteins did not serve as substrates. Using mass spectrometric analysis of Afp18G glycosylated RhoA, Rac1 and Cdc42, we could confirm that these GTPases were modified by a single covalently attached N-acetylhexosamine (HexNAc) moiety, which resulted in a mass increase of 203 Da in the corresponding switch I peptides (Fig. 6d). Again, RhoA was modified most efficiently among these GTPases. Thus, RhoA may be the primary target of Afp18G and might explain the previous results obtained in RhoA-, Rac1- and Cdc42-mediated actin dynamic analysis in zebrafish embryos and ZF4 cells. To prove that the NxN mutant of Afp18 is inactive and not able to glycosylate RhoA, we used UDP-GlcNAz as donor in a click chemistry reaction with biotin alkyne and radiolabelled UDP-[14C]GlcNAc in a glycosylation reaction and show that the mutant indeed is deficient in glycosyltransferase activity (Fig. 6e). Furthermore, we analysed the nucleotide status of RhoA which is glycosylated by Afp18 and found that Afp18G preferentially modified GTP (GTPγS)-bound RhoA in comparison to GDP-bound or nucleotide-free RhoA (Supplementary Fig. 4a).

Figure 6: Afp18G selectively modifies RhoA using UDP-GlcNAc. (a) Donor substrate specificity of Afp18G determined by UDP-glycosidase activity. Percentage of hydrolysed UDP-[14C]sugars was determined by PEI thin-layer chromatography and autoradiography after incubation for 10 min at 30 °C. Data are representative of three independent experiments. (b) Autoradiography of the SDS–PAGE from cell lysate incubated with Afp18G and indicated radiolabeled UDP-sugars. Western blot of RhoA (right panel) showed similar electrophoretic mobility as radiolabelled proteins. (c) Time course of in vitro GlcNAcylation of RhoA, Rac1 and Cdc42 by Afp18G (1 nM). Inserts show representative autoradiograms (upper panel) and Coomassie-stained SDS–PAGE (bottom panel). Error bars indicate s.e.m.’s of three technical replicates. (d) Extracted ion chromatograms of thermolysin-digested GST-RhoA, GST-Rac1 and Cdc42 treated with Afp18G (lower chromatogram) or untreated control (upper chromatogram). The molecular mass [M+2H]2+ of the switch I peptides (RhoA: 26 SKDQFPEVYVPT 37 ; Rac1: 24 TTNAFPGEYIPT 35 ; Cdc42: 24 TTNKFPSEYVPT 35 ) are indicated. Afp18G-modified GTPases (lower chromatogram) show a mass shift of 203 Da revealing a modification with a single N-acetylglucosamine (shifted blue curves). In comparison with Rac1 and Cdc42, RhoA was modified most efficiently. (e) The DxD motif of Afp18 is crucial for GlcNAcylation of RhoA. Recombinant RhoA was incubated in the presence UDP-GlcNAz or UDP-[14C]GlcNAc with Afp18G, Afp18G NxN or without toxin. Modified proteins were analysed by click chemistry using biotin alkyne and western blotting and autoradiography, respectively. Anti-RhoA served as input control. Full size image

GlcNAcylated RhoA is observed in an inhibited conformation

To identify the site of modification and gain insights into the structural alteration caused by GlcNAcylation of RhoA, we crystallized Afp18-modified RhoA in the presence of magnesium and GDP and solved the X-ray structure at 2.0 Å resolution (Fig. 7a, Table 1). In the RhoA structure, we could clearly assign additional electron density at the hydroxyl group of tyrosine-34 to an attached N-acetylglucosamine moiety (Fig. 7b, Supplementary Fig. 3c). Interestingly, despite the presence of magnesium in the crystallization conditions, the structure revealed an unusual GDP-bound but magnesium-free conformation. The overall fold of the crystal structure of GlcNAcylated RhoA is very similar to known structures of Rho GTPases. However, the switch regions, especially the switch I region, adopt a conformation with tyrosine-34 positioned away from the nucleotide binding pocket, resulting in a structure distinct from the structures of RhoA bound to GDP41 or GTP42, which is generally not compatible with effector and regulator interaction (Fig. 7c).

Figure 7: Structural consequences of RhoA GlcNAcylation at tyrosine-34. (a) Crystal structure of glycosylated RhoA at tyrosine-34. Switch I and switch II regions are highlighted in blue. GDP is shown as sticks and balls in black. The GlcNAc moiety attached to tyrosine-34 is shown as sticks and balls in yellow. (b) Electron density map of a section of 2F o −F c protein (grey), contoured at a level of 1σ, showing GlcNAc moiety attached to tyrosine-34 of RhoA in the alpha configuration of the glycosidic bond (marked). Additional residues of the switch I region were omitted for clarity. (c) Superposition of the effector loops of GlcNAcylated RhoA with structures of non-glycosylated active GTPγS-bound RhoA (pdb code 1A2B)42 and inactive GDP-bound RhoA (pdb code 1FTN)41. The switch I and II regions of GlcNAc-modified RhoA adopt distinct open conformations. (d) Autoradiograms and Coomassie stainings of Afp18G-catalysed in vitro14C-GlcNAcylation of WT GST-RhoA and the indicated mutants. (e) Cytoplasmic OGA is unable to revert GlcNAcylation of Rho. RhoA was radioactively preglycosylated by Afp18G and applied to a deglycosylation reaction with GST-OGA. TAB1, preglycosylated by OGT, served as positive control. Autoradiographs (top panels) of 14C-GlcNAcylated RhoA and GlcNAcylated TAB1 are shown. Coomassie staining of RhoA, TAB1 and GST-OGA are shown as input controls. Full size image

Table 1 Data collection, phasing and refinement statistics for RhoAY34(GlcNAc). Full size table

Tyrosine-34 is located within the effector loop region conserved in all Rho family GTPases. In in vitro glycosylation experiments with Afp18, followed by tandem mass spectrometric (LC–MS–MS) analyses, we confirmed that also Cdc42 was GlcNAcylated at tyrosine-32 (Supplementary Fig. 3d). Furthermore, site-directed mutagenesis and in vitro glycosylation experiments with Y32(34)F mutants of RhoA, Rac1 and Cdc42 confirmed switch I tyrosine-32(34) as the acceptor site of modification (Fig. 7d, Supplementary Fig. 3e). No other hydroxyl-containing amino acid (threonine or serine) or recently discovered glycosyl acceptor residues as tryptophan43 or arginine44,45 were able to substitute tyrosine in RhoA as an acceptor residue for glycosylation (Fig. 7d).

Afp18 forms deglycosylation-resistant α-glycosidic bonds

The defined electron density of the sugar attached to the hydroxyl group of tyrosine-34 revealed the α-anomeric configuration of the glycosidic bond (Fig. 7b). This finding implies that the glycosylation mechanism proceeds under retention of the stereochemistry of D-α-GlcNAc. Thus, Afp18 can be grouped into the family of retaining glycosyltransferases. The stereochemistry of the glycosidic bond has most likely no influence on the functional consequences of Afp18-mediated glycosylation, but might have an influence on the stability of the glycoside inside the host cytoplasm. Only one enzyme, namely O-GlcNAcase (OGA) exists in eukaryotic cytoplasm, which is able to remove mono-O-GlcNAc moieties from proteins. OGA predominantly cleaves sugars attached in the β-configuration. To clarify whether the glycosidic bond on RhoA was resistant to hydrolysis of OGA, we tested 14C-GlcNAcylated RhoA in a deglycosylation reaction. As a control protein, we used TGF-beta-activated kinase 1 (TAB1) preglycosylated by nucleocytoplasmatic O-GlcNAc transferase (OGT), which was efficiently deglycosylated by OGA (Fig. 7e). Notably, OGA did not deglycosylate Afp18-modified RhoA. We confirmed these results by using CpNagJ, another highly efficient mono-O-GlcNAcase from Clostridium perfringens (Supplementary Fig. 3f,g). Taken together, we found that Afp18 is a mono-O-glycosyltransferase, which selectively uses UDP-GlcNAc as sugar donor substrate to modify Rho GTPases site-specifically at a switch I tyrosine residue.

RhoA GlcNAcylation blocks regulator and effector interaction

To elucidate the molecular consequences of tyrosine-34 GlcNAcylation of RhoA, we analysed the nucleotide binding and nucleotide exchange of RhoA by mant-GDP fluorescence spectroscopy. Whereas the Afp18-catalysed glycosylation of RhoA did not affect nucleotide binding kinetics of mant-GDP or mant-GppNHp, a non-hydrolysable GTP analogue, (Fig. 8a), RhoA modification blocked leukaemia-associated RhoGEF (LARG)-catalysed nucleotide exchange (Fig. 8b). This effect was not caused by competition of Afp18G with GEF (Supplementary Fig. 4b), because GEF-stimulated nucleotide exchange was only blocked by the addition of UDP-GlcNAc. Thus, RhoA GlcNAcylation by Afp18 prevented the activation step of the GTPase. In addition, the interaction of RhoA to its GTPase-activating protein p50RhoGAP was impaired by Afp18G-mediated glycosylation of tyrosine-34 (Supplementary Fig. 4c).

Figure 8: Functional consequences of tyrosine-34 glyosylation of RhoA. (a) GlcNAcylation of RhoA by Afp18G does not alter nucleotide binding. Fluorimetric analysis of mant-GDP (mGDP) or mant-GppNHp (mGppNHp) binding to WT RhoA or glycosylated RhoA (RhoA-GlcNAc) bound to GDP. Nucleotide exchange was monitored by the increase in fluorescence on mant-nucleotide binding to RhoA. Data are representative of two independent experiments. (b) Nucleotide exchange was measured by mant-GppNHp exchange with WT RhoA, Afp18G-GlcNAcylated RhoA in the presence or absence of LARG. Data are represented as means±s.d. of three technical replicates. (c) Western blot analysis of RhoA, Rac1, Cdc42 pull-down experiments with zebrafish (ZF4) cells treated with Afp18G (plus PA for delivery), Afp18G NxN (plus PA for delivery) and C. difficile toxin B (TcdB). After Rho GTPase activation with CNF1, active RhoA was pulled down with ROCK II-coupled beads and Rac1 and Cdc42 with PAK-coupled beads. Bound GTPases were detected by western blotting using anti-RhoA, anti-Rac1, and anti-Cdc42 antibodies, respectively. Immunoblot of total RhoA is the loading control. (d) Live images at 30% epiboly (4.7 h.p.f.) of control non-injected, GFP mRNA (50 pg per embryo), RHOA mRNA (50 pg per embryo), RHOA Y34F (50 pg per embryo)+Afp18G mRNA (0.5 pg per embryo), Afp18G mRNA (0.5 pg per embryo)+RHOA mRNA (50 pg per embryo) or Afp18G (0.5 pg per embryo)+RHOA Y34F mRNA (50 pg per embryo) injected embryos. GFP mRNA (50 pg per embryo) was also included in each injection mix, and GFP fluorescence used to examine for homogenous expression: panels show transmitted light images in left columns and corresponding GFP fluorescence images in right columns. GFP, RHOA and RHOA Y34F injected embryos developed indistinguishable from non-injected control embryos. Scale bar, 500 μm. (e) Quantification of percentage embryos that develop normally or were severely affected regarding the organization of the blastoderm in experiment shown in d. Co-injection of RHOA or RHOA Y34F mRNA together with Afp18G mRNA significantly reduces the fraction of disrupted and degraded blastoderm phenotypes, and increases the fraction of embryos developing normally at 30% epiboly. Values are average mean±s.e.m. of four biological replicates. Significance (P values <0.001) of changes in phenotype distribution was valuated using the Kruskal–Wallis test. Full size image

Next, we analysed the interaction of Rho, Rac1 and Cdc42 with its downstream effectors Rho kinase α (ROCKII)- and p21-associated kinase (PAK) and used zebrafish (ZF4) cells in an effector pull-down assay using ROCKII- and PAK-coupled beads (Fig. 8c). To activate Rho GTPases independently of intracellular GEFs, we pretreated cells with cytotoxic necrotizing factor 1 (CNF1), a toxin that activates Rho GTPases constitutively by deamidation46. Subsequent cell intoxication with Afp18G totally blocked RhoA interaction with ROCKII, whereas RhoA could efficiently be precipitated from cells treated with the glycosyltransferase-deficient mutant Afp18G NxN. Toxin B from Clostridium difficile (TcdB), which glycosylates threonine-37 in RhoA and thereby inhibits Rho effector interaction, served as a control. Impaired effector interaction catalysed by Afp18G was also observed for Rhotekin, an effector of RhoA, in human HeLa cells (Supplementary Fig. 4d). Intriguingly, the interaction of Rac1 or Cdc42 with their effector PAK could not be blocked by Afp18-mediated GlcNAcylation, whereas TcdB treatment of ZF4 cells efficiently prevented PAK interaction (Fig. 8c). This is consistent with our findings in time-lapse microscopy of Afp18-intoxicated ZF4 cells, which showed a rapid degradation of filamentous stress fibres but a persistence of membrane dynamics like membrane ruffling and filopodia formation, which are regulated by Rac1 and Cdc42, respectively (Fig. 3c, Supplementary Movie 3). It seems that Afp18 specifically inactivates RhoA signalling, but less signalling of Rac or Cdc42.

Afp18G-phenotype suppression by RhoA overexpression

To determine whether Afp18G predominantly acts through the glycosylation of RhoA in vivo, we tried to rescue the toxin phenotype by co-injection of mRNA encoding human WT RHOA or the non-glycosylatable mutant version RHOA Y34F. Zebrafish RhoA as well as other Rho GTPases share over 95% sequence conservation with their human homologues16. Embryos injected with human RHOA or RHOA Y34F mRNA (up to 50 pg per embryo) alone developed normal, indistinguishable from non-injected or GFP mRNA-injected control embryos (Supplementary Fig. 5a). We co-injected RHOA or RHOA Y34F mRNA with 0.5 pg Afp18G mRNA and found the disrupted blastoderm phenotype rescued to a large extent, with more embryos developing normally compared with Afp18G mRNA-injected embryos (Fig. 8d and quantification Fig. 8e). Taken together, both human RHOA and RHOA Y34F overexpression were able to rescue development of early embryos from Afp18G toxicity.