A gene horizontally transferred from bacteria protects arthropods from host plant cyanide poisoning

1Plants have developed a remarkable diversity of chemical defenses to deter herbivores from feeding. Cyanogenesis is one of the most ancient and widespread of these defenses, and more than 2500 plant species are known to synthesize cyanogenic glucosides and cyanolipids as phytoanticipins (constitutive defense compounds present in the plant prior to herbivore attack). Upon tissue disruption by herbivore feeding, cyanogenic glucosides are degraded by the plant β-glucosidases and α-hydroxynitrile lyases, which results in the release of toxic hydrogen cyanide (HCN) and other toxic products such as their aglycones (;;;; Figure 1). The released cyanide is a potent inhibitor of the mitochondrial respiratory chain and has a disruptive effect on various metabolic pathways, thus providing a broad defense against generalist herbivores.

2In their arms race with plants, arthropods have evolved several mechanisms to overcome plant cyanogenesis. A well-documented case of co-evolution is found in insect lepidopteran specialists that sequester the ingested cyanogenic glucosides in their own defense against predators. Remarkably, cyanogenic compounds have become so crucial for some species such as burnet moths (Zygaenidae) that they not only sequester, but also synthesize these compounds de novo by convergent evolution of the biosynthetic pathways. Next to sequestration, other mechanisms have evolved to cope with the toxic effects of HCN, such as avoiding the ingestion of cyanogenic compounds or detoxifying HCN upon plant release. In animals, HCN is thought to be detoxified by two main pathways. The enzyme rhodanese converts cyanide into thiocyanate, but this biochemical reaction is not very common and thought to be inefficient. Alternatively, the conversion of HCN and cysteine into β-cyanoalanine and sulfide has been suggested as the main detoxification pathway in arthropods (Figure 1). This is supported by several biochemical surveys showing a correlation between β-cyanoalanine synthesis and HCN exposure in lepidopteran species tolerant to HCN. However, the enzyme that catalyzes this crucial reaction in arthropods has not been identified to date. The conversion of HCN into β-cyanoalanine by an enzyme called β-cyanoalanine synthase (CAS) has been best studied in bacteria and plants that need to protect themselves from HCN during the synthesis of cyanogenic glucosides or ethylene. The enzymes responsible for CAS activity also have cysteine synthase activity (CYS) and are referred to as CYS or CAS depending on their substrate specificity. CYS catalyzes the conversion of O-acetylserine into cysteine, an essential final step in the cysteine biosynthesis pathway unique for plants and bacteria. Animals synthesize cysteine by a different pathway and use related enzymes such as cystathionine-β-synthase (CBS) and cystathionine-γ-lyase (CGL), which form together with CAS and CYS a group of pyridoxal-5′-phosphate dependent enzymes (; Figure 1). Here we identify the enzyme responsible for detoxification of cyanide to β-cyanoalanine in a spider mite and show that this enzyme is horizontally acquired from bacteria and is also widely distributed in Lepidoptera.

3In order to gain a better insight into arthropod defenses against plant cyanogenesis, we used the two-spotted spider mite T. urticae as a generalist herbivore model in a host plant adaptation experiment. This species is one of the most polyphagous arthropod pests known to date, and feeds on more than 1100 plant species from more than 140 plant families, including many cyanogenic plants. We transferred a spider mite strain reared on acyanogenic bean plants (Phaseolus vulgaris) to a cultivar of Phaseolus lunatus containing high levels of well characterized cyanogenic glucosides such as linamarin and lotaustralin and allowed the strain to adapt to this host plant for more than 30 generations. Gene expression differences between mites feeding on P. vulgaris and on P. lunatus were then determined using genome-wide microarrays.

4In contrast to the broad response previously detected after host plant changes from Fabaceae to Solanaceae and Brassicaceae, only a limited set of 28 genes (absolute fold change ≥2, Benjamini-Hochberg corrected p-value <0.05) was found differentially expressed between the parental and adapted lines (Supplementary file 1). Within this small set of genes, 18 had an increased expression in the adapted line, while 10 exhibited a lower expression. The most differentially expressed genes after transfer to cyanogenic lima bean encode closely related, small cytoplasmic proteins of unknown function. They were also seen in the transcriptional response after transfer to tomato suggesting that these genes could be part of a broad general stress response of T. urticae. Among the genes with an increased expression were three cytochrome P450 genes, known to respond readily to host plant changes. Moreover, we identified a gene (tetur10g01570, Tu-CAS) encoding a predicted cytosolic protein with high similarity to bacterial cysteine synthases (Conserved Domain Database (CDD): COG0031). The reported microarray data have been deposited at the Gene Expression Omnibus.

5Within chelicerates, we only detected close homologues of Tu-CAS in two closely related tetranychid mites. By sequencing PCR amplicons (see below, this section) and a tBLASTn-search in a published transcriptome, CAS genes were identified in Tetranychus evansi and Panonychus citri, respectively. A tBLASTn-search in published genomes of mesostigmatid mites and of ticks (Metastigmata) did not reveal homologues (Supplementary file 2). Broadening the search to arthropods by tBLASTn-searches in NCBI databases and additional arthropod genome portals (Supplementary file 2), we only detected close homologues of Tu-CAS in lepidopteran genomes (Bombyx mori, Danaus plexippus, Heliconius melpomene, Manduca sexta and Plutella xylostella) (Figure 2). On average, Tu-CAS showed 75% similarity with lepidopteran protein sequences, and genes encoding these proteins were intronless in both mites and insects. By searching additional NCBI EST-databases and published lepidopteran transcriptomes, a total of 20 (complete and partial) homologous sequences were identified in arthropods.

6A phylogenetic reconstruction of these arthropod proteins with CYS, CAS and CBS enzymes of plants, fungi, oomycetes, bacteria and Metazoa indicated that these homologous arthropod sequences might be monophyletic. They were embedded with high node support within bacterial cysteine synthase sequences, indicative of a horizontal gene transfer (Figure 2). The most closely related sequences are from bacteria that belong to the α- and β-Proteobacteria, two of which are Methylobacterium species, free-living epiphytic bacteria known to establish endophytic colonies. These species are reported to be transferred from the plant to phytophagous insects and to survive inside arthropod hosts, facilitating a potential horizontal gene transfer. Alternatively, Proteobacteria are known endosymbionts of arthropods and often reside in the reproductive organs for vertical transmission to following generations. Because of this intimate relationship, successful horizontal gene transfer is more likely to occur. The mite and insect sequences formed a branch in bacterial cysteine synthase enzymes, some of which have documented dual CYS and CAS activities (Figure 2). The arthropod protein sequences and their closest bacterial homologues shared a unique 9 amino acid insertion not present in cysteine synthases of other organisms, but the residues known to be crucial for substrate and cofactor binding in plants and bacteria showed conservation. The lysine residue (Lys95, Glycine max CAS numbering, ) that forms a Schiff base linkage to the cofactor pyridoxal-5′-phosphate was also conserved in arthropods (Figure 2—figure supplement 1).

7To exclude the possibility that the Tu-CAS sequence was derived from contaminating bacterial DNA, we examined its position in the T. urticae genome into more detail. Tu-CAS is located on a 3 Mb large scaffold (scaffold 10) and is flanked by typical eukaryotic genes, tetur10g01550 and tetur10g01580. Both genes contain introns with splice sites confirmed by EST or RNA-seq data (, GenBank: LIBEST_025606, Figure 3). Tetur10g01580 encodes a nudix hydrolase (CDD: cl00447) highly similar to other arthropod nudix hydrolases (BLASTp hits with E-value <1e−45). To rule out that the linkage of Tu-CAS to these spliced genes is a genome assembly artefact, we took several independent genomic approaches. First, we remapped the T. urticae Sanger reads used for the genome assembly of the London strain of T. urticae and examined the Illumina read coverage of the Tu-CAS region in two re-sequenced strains (EtoxR and Montpellier) that are geographically distinct from the London strain. Neither Sanger nor Illumina-reads revealed any inconsistencies in the Tu-CAS region in any of these strains (Figure 3). Second, using a PCR approach, we amplified a 6 kb genomic region bracketing Tu-CAS with the surrounding spliced genes (tetur10g01550 and tetur10g01580, Figure 4, Figure 4—figure supplement 1). Using a similar strategy we also amplified a 6 kb region in the closely related spider mite T. evansi (Figure 4, Figure 4—figure supplement 1) and a nucleotide dot plot between the amplified region of these two species showed clear synteny and the absence of discontinuity around the CAS gene (Figure 4). This would not be expected by bacterial contamination of either genome. Last, gene compositions of T. urticae and bacteria were analyzed by determining both the GC-content at the synonymous third codon position (GC3) and the overall GC-content (GC) of the genes to look if amelioration occurred. Amelioration is the process by which the DNA composition of the newly acquired gene becomes homogenized to match the composition of the recipient genome (all genes in the recipient genome are subject to the same mutational processes). Indeed, the GC/GC3 content of Tu-CAS was most similar to the GC/GC3 content of genes from the T. urticae genome, and quite distinct from the GC/GC3 content of genes (including the CYS/CAS genes) from the three annotated bacterial genomes in the closest sister clade to the apparent monophyletic arthropod clade (Figure 5). Taken together, these data provide strong evidence that Tu-CAS is a sequence integrated in the genome of T. urticae and does not represent bacterial contamination. Sequence data is available at Genbank (accession numbers: KF981736 and KF981737).

8In order to functionally test whether arthropod enzymes are active and still able to catalyze both reactions after horizontal gene transfer, we recombinantly expressed Tu-CAS in Escherichia coli and obtained mg quantities after affinity purification. Subsequent biochemical assays confirmed that Tu-CAS catalyzes both reactions (Figure 6). In order to determine which of the two reactions (cysteine synthesis, CYS and β-cyanoalanine synthesis, CAS) was favored by the Tu-CAS enzyme, we calculated the ratio of the specificity constants for each reaction. The specificity constant kcat/Km defines, at any concentration, the specificity of an enzyme for a particular substrate. The CAS/CYS ratio of the specificity constants was 33.7, showing that CAS activity was strongly favored over CYS activity (Table 1). (The ratio was calculated from the respective values of Vmax/Km, as kcat = Vmax/[E] and as the reactions were measured with the same enzyme preparation.) This preferred CAS activity is typical of known plant CAS enzymes and clearly different from CYS enzymes (Table 1;;;; ). Enzymatic activity was dependent on pyridoxal-5′-phosphate as a cofactor, and the substrate-dependent formation of β-cyanoalanine was further confirmed by thin layer chromatography (TLC) and LC-MS (Figure 7).

9The very high sequence similarity between Tu-CAS and the lepidopteran proteins strongly suggests that the lepidopteran proteins can catalyze the same two reactions (cyanide detoxification and cysteine synthesis) and that these proteins are responsible for the known wide occurrence of CAS activity in lepidopteran species.

10In arthropods, the ability to detoxify HCN plays a crucial ecological role and is thought to have allowed the exploitation of cyanogenic plants by circumventing the toxic effects of HCN. We have shown here that the two-spotted spider mite increases transcript levels of a horizontally transferred β-cyanoalanine synthase upon adaptation on cyanogenic bean. Phylogenetic evidence alone does not constitute strong evidence for horizontal gene transfer (HGT), because in the absence of introns in the sequence, a contamination of the mite genome sequence with a sequence from a bacterial symbiont or commensal cannot be excluded. However, the results of mite genomic analysis, codon amelioration, synteny and genomic PCR, combined with the phylogenetic evidence, unambiguously prove that T. urticae CAS has a bacterial origin and was laterally transferred and incorporated into the mite genome prior to the divergence in the Tetranychidae. A homologous lateral gene transfer has also occurred in Lepidoptera (Figure 2) raising the question of the time and number of HGT events needed to explain the phylogenetic pattern of distribution of the CYS/CAS genes. Several hypotheses can be discussed. (a) A single HGT to a common ancestor of mites and insects, followed by selective losses resulting in the present phylogenetic distribution. The broad sampling of arthropod species that revealed the absence of any CYS/CAS-like sequence (even as a distant recognizable trace) argues against the origin of the gene in the common ancestor of tetranychid mites and Lepidoptera, followed by multiple independent losses. These losses would have to be numerous: seven in the hexapods (Trichoptera, Diptera, Hymenoptera, Coleoptera/Strepsiptera, Hemiptera/Heteroptera/Thysanoptera/Phthiraptera, Orthoptera, Odonata/Ephemeroptera) and six further in other arthropods (Copepoda/Branchiopoda, Myriapoda, Aranea, Scorpiones, Metastigmata and Mesostigmata). This figure of 13 losses is an absolute minimum that implies that the loss occurred each time at the origin of the lineage, that is before any speciation beyond the point of coalescence. A loss later in the history of each lineage would rapidly increase the total number of losses. We strongly believe that this hypothesis is not parsimonious, and therefore that there was no CYS/CAS gene in the common ancestor of mites and Lepidoptera. (b) Alternatively, two HGT events might have occurred (one to a mite ancestor, one to a lepidopteran ancestor) or (c) a single HGT from a donor bacterial species, followed by a transfer between a mite and a lepidopteran. These two hypotheses are impossible to distinguish based on the topology of the tree (Figure 2). The sampling of bacterial species phylogenetically close to the presumed donor species is too shallow at present. The apparent monophyly of the arthropod CAS gene may be due to the fact that both the mite and the insect gene came from very closely related bacteria, but these bacteria are not represented in the tree. A future survey of Proteobacteria likely to be associated with arthropods (directly or through a plant host) may resolve this question by finding several potential bacterial donor species that would invalidate the apparent monophyly (i.e., split the tree at the point of arthropod coalescence). The transfer from a lepidopteran to mite is improbable, because the mite sequence would branch with the closest relative to the lepidopteran donor, rather than being basal. Conversely, we cannot exclude the transfer from a mite to a lepidopteran, as sampling within the Prostigmata is extremely limited. We therefore favor hypothesis (b), with a very old transfer to an ancestral lepidopteran, and a second transfer to an ancestral mite. Current sampling of mite species is insufficient to give a good estimate of time, but we note the absence of a CAS enzyme in Mesostigmata (typically parasitic and predatory mites) and Metastigmata (ticks), and transfer is likely to have occurred after the split of Prostigmata in the Lower Devonian about 400 MYA. The hypothesis of multiple transfers is also compatible with the presence of a CYS-like sequence in the mealybug Planococcus citri, apparently derived from its endosymbiont. In that case, the phylogenetic tree clearly distinguishes the HGT event from that under study here.

11In Lepidoptera, the CAS gene was subsequently duplicated in H. melpomene and D. plexippus that have 3 and 2 copies of the gene, respectively. The importance of the laterally acquired gene for HCN detoxification in those species was not previously apparent. These species of Papilionoidea, like burnet moths, not only thrive on cyanogenic plants, but have themselves evolved the ability to synthesize cyanogenic compounds de novo that now have crucial functions in their life history. Cyanogenesis serves in predator defense by releasing HCN, but also stores reduced nitrogen that can be mobilized for chitin synthesis, and plays a role in mate choice by determining the attractiveness of nuptial gifts from male to female partners. Our results support the early idea that CAS activity is needed for the exploitation of cyanogenic glucosides in insects whether they are sequestered from the plant, or synthesized de novo. Higher β-cyanoalanine synthase activity in Spodoptera eridania than in Trichoplusia ni is related to higher cyanide tolerance, and this enzyme activity is widespread in Lepidoptera.

12Moreover, it was recently shown that specialist pierid butterflies that feed on Brassicales, release equimolar concentrations of HCN upon metabolism of benzylglucosinolates, turning the ‘mustard oil bomb’ into a ‘cyanide bomb’. When Pieris rapae feeds on a cyanogenic (dhurrin-containing) plant that this species does not normally consume, an increased production of β-cyanoalanine and thiocyanate is observed, thus implicating both a CAS and a rhodanese activity. It was therefore proposed that the ability of P. rapae to metabolize HCN allowed the primary host transfer from Fabales to Brassicales. The gene for either CAS or rhodanese has not been identified in arthropods before, and their respective role in detoxification of HCN is not formally demonstrated. There is no close homologue of the known rhodanese (thiosulfate sulfurtransferase) gene in T. urticae or in Lepidoptera. However, we identified Tu-CAS and functionally demonstrated that the enzyme it encodes converts HCN to β-cyanoalanine in vitro. Such evidence is difficult to obtain in vivo with mites, as it would be difficult to exclude the possibility that a plant or a bacterial enzyme rather than the mite enzyme catalyzes the reaction in vivo. We argue that the presence of the same gene in lepidopteran species that display this activity in vivo is strong evidence for the function of the laterally transferred CAS gene. It will be of great interest to confirm that the homologous CAS genes of Lepidoptera that we identified indeed encode a β-cyanoalanine synthase, and to provide evidence of its protective role against HCN poisoning.

13Next to the detoxification function, the CYS activity acquired after horizontal gene transfer may also have enhanced the sulfur amino acid economy of mites and lepidopterans (Figure 1). To date, nematodes were the only animal species thought to synthesize cysteine independently from methionine by CYS activity. However, nematode CYS sequences grouped with plants and oomycetes, clearly outside the arthropod-bacterial clade, suggestive of a different origin of CYS between metazoan subgroups (Figure 2). Duplications of cys genes were observed in nematodes, and indeed genetic evidence suggests that in Caenorhabditis elegans cyanide resistance is conferred by the cysl-2 gene, probably encoding an enzyme with CAS activity while cysl-1 is a classical cysteine synthase gene. The acquisition of an alternative cysteine biosynthesis route fits into previously documented horizontal gene transfers in T. urticae that include a cobalamin-independent methionine synthase gene, genes for carotenoid biosynthesis, as well as laterally acquired genes that likewise respond to host plant change such as intradiol ring-cleavage dioxygenases and a cyanase gene. The latter encodes an enzyme that decomposes cyanate (CNO−), a bacterial or photochemical decomposition product of cyanide, to carbon dioxide and ammonia. This enzyme may serve as a second line of spider mite defense against cyanogenic plants, or alternatively may have a regulatory function in the amino acid and pyrimidine metabolism as previously suggested. For a polyphagous herbivore, horizontal gene transfer might play an important role in gaining independence from the varying plant nutrients and defense compounds. It remains unclear which reaction (CAS/CYS) of Tu-CAS provides the strongest adaptive advantage, but the CYS activity might be one of the reasons why these horizontally transferred genes have been retained in organisms that are at present not living on cyanogenic plants.

14In conclusion, a horizontal gene transfer from a bacterial ancestor underlies the exploitation of cyanogenic host plants in some arthropod lineages and made the subsequent evolution of a convergent pathway for synthesis of cyanogenic glucosides possible, as shown in burnet moths.

15The London strain of T. urticae was maintained on acyanogenic P. vulgaris L. cv ‘Prelude’. The strain London-CYANO originated from this population and was transferred to cyanogenic P. lunatus as previously described. Before the start of experiments the cyanogenic potential of both plant species was determined, confirming the acyanogenic nature of P. vulgaris and revealing high levels of cyanogenic precursors produced in the P. lunatus cultivar. For this study, young adult female mites were collected for gene-expression analysis 35 generations after the initial host shift. The T. evansi strain was maintained in the laboratory on Solanum lycopersicum L. cv ‘Moneymaker’ as previously described. All strains were maintained in climatically controlled rooms at 26°C, 60% RH and 16:8 hr light:dark photoperiod.

16Total RNA samples were isolated with the RNeasy minikit (Qiagen, Belgium) and were subsequently treated with DNase (Turbo DNA-free kit, Ambion, Belgium). RNA was extracted from 100–120 young adult female mites in four replicates. Cy5- or Cy3-labeled cRNA was produced using the Low Input Quick Amp Labeling Kit (Agilent Technologies, Belgium) as previously described. Microarray hybridization and scanning procedures were performed as previously described, using the GE2_107_Sep09 protocol. The data was transferred to GeneSpring GX 11.0 software (Agilent Technologies) for statistical analyis. Probes were flag filtered (only probes that had flag-value ‘present’ in 50% of all replicates were retained) and linked to T. urticae genes using the ‘Create New Gene-Level Experiment’ option. Differentially expressed genes were identified by a Student's t test with the cutoff for Fold Change (FC) and corrected p-value (Benjamini-Hochberg correction) set at 2 and 0.05, respectively. The array design is accessible under the GEO-platform format GPL16890.

17A full-length Tu-CAS (tetur10g01570) homologue was retrieved in T. evansi by sequencing PCR products bracketing the CAS gene with neighboring genes (Supplementary file 4). Additional Tu-CAS homologues were identified by conducting BLASTp and/or tBLASTn searches in NCBI, UniProt, P. citri transcriptome and diverse arthropod genome portals (Supplementary file 2) using Tu-CAS as query. As several best BLAST-hits (E-value ≤1e−90) included members of the order Lepidoptera, transcriptome databases from Lepidoptera not included in the NCBI database were also mined for Tu-CAS homologues (Supplementary file 3). This approach yielded 35 arthropod and bacterial protein sequences, which according to the Conserved Domain Database all contained a motif typical for cysteine synthases (COG0031). This dataset was further complemented with cysteine synthase M (cysM) protein sequences from bacteria and cysteine synthase protein sequences from fungi, Chromalveolata, plants, nematodes, and Planococcus citri and its three best BLASTp hits, harboring a cysteine synthase CDD motif (COG0031 or PLN2565). Finally, a diverse set of cystathionine-β-synthase protein sequences, related to cysteine synthases and also belonging to the group of pyrodixal-5′-phosphate dependent β-substituted alanine synthases were added as an outgoup.

18The final dataset contained 90 protein sequences. Accession numbers of protein sequences, their trivial name, CDD classification and cellular localization are listed in Supplementary file 3. Protein sequences were aligned with MUSCLE using default settings. Model selection was done with ProtTest 2.4 and according to the Akaike information criterion the model LG+I+G was optimal for phylogenetic analysis. A maximum-likelihood analysis was performed using Treefinder with edge-support calculated by 1000 pseudoreplicates (LR-ELW). Resulting trees were midpoint rooted prior to further analysis. Phylogenetic trees were visualized and edited using MEGA5 and CorelDraw X6 (Corel inc., UK), respectively.

19Paired-end T. urticae Sanger reads (available in the Trace Archive at the NCBI website, http://www.ncbi.nlm.nih.gov/Traces/home/) were remapped to the T. urticae genome using Bowtie 2.1.0 and the preset parameter option ‘–very-sensitive’. Resulting SAM files were converted into BAM files using SAMtools. Illumina-reads from genomic DNA sequencing of the London, Montpellier and EtoxR strains of T. urticae and Illumina RNA-seq reads from adult T. urticae polyA selected RNA were mapped as previously described. Read alignments and coverage were visualized with IGV 2.3 using the most recent T. urticae genome annotation (‘Tetur_gff3_20130708’, accessible at http://bioinformatics.psb.ugent.be/orcae/-overview/Tetur); for display, Sanger reads were arranged in Adobe Illustrator CS5 while maintaining alignment coordinates.

20Genomic DNA was collected from T. evansi and T. urticae by phenol-chloroform extraction. Primer pairs were designed to amplify a genomic region of Tu-CAS and adjacent genes on either the 5′ or the 3′ end on the 10th scaffold of the T. urticae genome (Supplementary file 4). The Expand Long Range PCR kit (Roche, Belgium) was used to conduct PCR, and fragments were sequenced with primers listed in Supplementary file 4. Some primer pairs designed on the T. urticae genome sequence also successfully amplified genomic fragments of T. evansi (Supplementary file 4). The resulting fragments were sequenced by primer walking (Supplementary file 4). A nucleotide dot-blot between the two spider mite species was constructed using the MEGALIGN program of DNASTAR software, allowing 5% mismatch in a 21 bp window. We analyzed overall GC contents and at the third codon position (GC3) of whole coding nucleotide sequences using UGENE of all coding sequences of T. urticae and Achromobacter xylosoxidans, Methylobacterium radiotolerans and Methylobacterium sp. GXF4. These three bacterial genomes were selected based on our phylogenetic analysis as the closest fully annotated bacterial genomes to the arthropod clade (Figure 2).

21Recombinant Tu-CAS was produced by the GenScript Corporation (Piscataway, NJ, USA). After codon optimization of the Tu-CAS coding sequence (Supplementary file 5), an E3 expression vector was used to transform E. coli cells. The transformed cells were cultured in 3 l LB. Using a Ni2+-column, the N-His-tagged Tu-CAS protein was purified from the supernatant. After sample sterilization via a 0.22 µm filter, the recombinant protein was stored in a buffer containing: 50 mM Tris, 150 mM NaCl, 2 mM DTT, 10% glycerol at a pH of 8.0 and finally kept at −80°C. The concentration and purity of the recombinant protein sample was determined respectively by a Bradford protein assay and a densitometric analysis of a Coomassie Blue-stained SDS-PAGE gel.

22Chemicals for the activity assays were purchased from Sigma–Aldrich (Belgium), except β-cyanoalanine, which was acquired from VWR, Cayman Chemical. All reactions were carried out in gastight 7 ml vials with a screw cap having a PTFE/rubber septum (Supelco–Sigma–Aldrich, Belgium). Reagents were added to the reaction volumes using gastight syringes (Hamilton, series 1700, Gastight, 1750RN, VWR Belgium). Prior to measuring enzyme activity, recombinant Tu-CAS was incubated at 30°C for 10 min in the appropriate reaction buffer containing 500 µM pyridoxal-5′-phosphate. For measuring CAS activity, reactions were executed in a 0.2 M Tris buffer at pH 8.5. The CAS activity assay was a modification of the method of Hendrickson. The standard reaction was started with 0.5 ml of 0.01 M cysteine, 0.5 ml of 0.01 M sodium cyanide and 1.5 µg recombinant Tu-CAS. All CAS assays were performed at 37°C on a mechanical shaker. CAS activity was quantified by spectrophotometrically measuring the H2S formed at 650 nm (PowerWavex340, BioTek Instruments Inc., Winooski, VT, USA) by the method of Siegel. The CYS activity assay was based on using 0.5 µg recombinant Tu-CAS per reaction. Standard substrate concentrations were 10 mM and 2 mM for O-acetylserine and sodium sulfide, respectively. The reaction product cysteine was quantified by measuring the absorbance at 560 nm by the method of Gaitonde. For both activity assays, the spontaneous formation of the measured reaction product and its potential presence in protein preparations was corrected by respectively non-enzyme controls and zero time point controls. Each experimental assay condition was analyzed using three independent and three technical replicates. Kinetic data was fitted to the Hill equation, from which the Km and Vmax values for O-acetylserine and cysteine were calculated for respectively the CYS and CAS reaction.

23Recombinant Tu-CAS was incubated at 30°C for 10 min in a 1 mM phosphate buffer containing 500 µM pyridoxal-5′-phosphate. The standard reaction was executed in a 320 µl reaction volume at 37°C on a mechanical shaker in 1 mM phosphate buffer pH 8.5, with 80 µl of 7.5 mM cysteine and 80 µl of 15 mM sodium cyanide. Each reaction was started by adding 3.75 µg of recombinant protein and was terminated by snap freezing at different time points. No-substrate and no-enzyme controls were included in the analysis. Reaction mixtures were defrosted at 4°C and a 20 µl aliquot was spotted on a thin-layer chromatograph (HPTLC Silica gel 60 F254, Merck, Darmstadt Germany) and run with a mobile phase of (ethanol/28% ammonium hydroxide/water) with a (18/1/4) ratio. Five µg of β-cyanoalanine and cysteine were spotted as standards. After drying, the TLC plate was treated with a ninhydrin solution (20 g ninhydrin in 600 ml ethanol) for amino-acid visualization. In this validated TLC set-up, cysteine and β-cyanoalanine can be identified both by color and relative mobility (Rf-value). β-cyanoalanine consistently displayed a blue-green color with a Rf value of around 0.8. In contrast, cysteine colored red and exhibited a consistent different Rf value.

24The identification of β-cyanoalanine as a blue spot at 0.8 Rf was further confirmed by LC-MS analysis. After TLC separation, the zone around 0.8 Rf was scrapped from the TLC plate. After scrapping, the plate was colored as described above to confirm that the correct blue/green zone was collected. The collected silica was mixed with 200 µl ddH2O, vortexed and centrifuged at 21000×g for 5 min. The supernatant was collected and directly analyzed using an Agilent 1100 Series LC-MSD with a diode array detector operating at 220 nm. The column oven was programmed at 35°C using a Phenomenex Luna 5u column (particle size) C18(2), 100A (pore size) with a column size of 250 × 3.0 mm. A gradient elution program driven by a quarternary pump was used at a flow rate of 0.5 ml/min (injection volume: 20 µl). The acetonitrile/water gradient used was 0–2 min (5% acetonitrile); 2–17 min (5–100% acetonitrile); 17–22 min (100% acetonitrile); 22–24.5 min (100–5% acetonitrile); 24.5–27 min (5% acetonitrile). The mass spectrometer was operated using the SCAN mode in the electrospray ionization mode. The analyzing sector contained a quadrupole analyzer and an electron multiplier detector.