ParST is a widespread toxin–antitoxin module thattargets nucleotide metabolismFrank J. Piscottaa, Philip D. Jeffreyb, and A. James Linka,b,c,1

aDepartment of Chemical and Biological Engineering, Princeton University, Princeton, NJ 08544; bDepartment of Molecular Biology, Princeton University,Princeton, NJ 08544; and cDepartment of Chemistry, Princeton University, Princeton, NJ 08544

Edited by Marlene Belfort, University at Albany, Albany, NY, and approved December 4, 2018 (received for review August 27, 2018)

Toxin–antitoxin (TA) systems interfere with essential cellular pro-cesses and are implicated in bacterial lifestyle adaptations such aspersistence and the biofilm formation. Here, we present structural,biochemical, and functional data on an uncharacterized TA system,the COG5654–COG5642 pair. Bioinformatic analysis showed thatthis TA pair is found in 2,942 of the 16,286 distinct bacterial speciesin the RefSeq database. We solved a structure of the toxin boundto a fragment of the antitoxin to 1.50 Å. This structure suggestedthat the toxin is a mono-ADP-ribosyltransferase (mART). The toxinspecifically modifies phosphoribosyl pyrophosphate synthetase(Prs), an essential enzyme in nucleotide biosynthesis conservedin all organisms. We propose renaming the toxin ParT for PrsADP-ribosylating toxin and ParS for the cognate antitoxin. ParTis a unique example of an intracellular protein mART in bacteriaand is the smallest known mART. This work demonstrates that TAsystems can induce bacteriostasis through interference with nucle-otide biosynthesis.

toxin–antitoxin system | ADP-ribosylation | posttranslational modification

Our understanding of toxin–antitoxin (TA) systems has pro-gressed significantly since their identification nearly 35 y

ago. In the pioneering work, ccdAB was shown to enhance thestability of a mini-F plasmid in Escherichia coli by killingdaughter cells lacking the plasmid and also established that theccdA genetic element could provide an antidote for the post-segregational killing (PSK) of ccdB (1). An investigation into theE. coli R1 plasmid revealed a similar system hok/sok that alsoimproved plasmid stability, where again one element, sok, couldneutralize the PSK activity of the other, hok (2). Results such asthese led to the hypothesis that the role of such genetic regionswas to ensure the inheritance of plasmids during cell division. Itwas later discovered, however, that these systems resided notonly in plasmids but also on bacterial chromosomes and that,while not conferring replicon stability, they may play an impor-tant role in regulating cell growth (3–5). Further investigationsinto bacterial TA systems have also implicated them in biofilmgrowth and persister formation. The toxin gene mqsR was firstshown to be up-regulated in E. coli biofilms and later confirmedto promote biofilm formation through regulation of quorumsensing (6, 7). One of the most ubiquitous TA systems, hipBA,was the first to be implicated in persister formation; the well-characterized mutant hipA7 was shown to increase E. coli per-sistence up to 1,000-fold, due to a weakened interaction with itsantitoxin (8). Various other toxins have also been implicated inbiofilm and persister formation, including RelE, YafQ, andMazF (9). As the importance of biofilms and persisters in clinicalsettings grows, the study of TA systems provides an avenue forbetter understanding the mechanisms driving these processes, aswell as a possible path to intervention.Although most downstream effects of these toxins are gener-

ally similar (bacteriostasis or cell death), the methods by whichthese systems work are remarkably diverse. TA systems arebroadly grouped into six classes, type I through VI, based on thenature of the TA interaction, although type I and II comprisemost known TA systems and have been extensively studied. Type

I toxins are typified by an RNA-level TA interaction, where anantisense RNA binds toxin mRNA to inhibit its translation. TypeII TA systems function via protein–protein interactions, wherebinding of the antitoxin to the toxin inhibits its activity (10). Ofthe two earliest known TA systems mentioned above, hok/sok isan example of a type I system, while ccdAB is type II (11, 12).Upon translation, type I toxins are small, hydrophobic peptidesthat lead to cell lysis through disruption of the plasma mem-brane. Type II toxins, however, function through a variety ofmechanisms. A large group of these are endoribonucleases thatcleave RNA in either a ribosome-dependent or independentmanner. The well-studied HigB, RelE, and MazF toxins, amongmany others, all fall into this group (13–15). Beyond these,however, there is much variation. HipA phosphorylates glutamyl-tRNA synthetase (GltX), inactivating it, while CcdB engages ingyrase-mediated double-stranded DNA cleavage (16, 17). An-other toxin, FicT, also interferes with DNA gyrase, but rather byinactivation through adenylation (18). More recently, DarT wasfound to ADP-ribosylate single-stranded DNA (19). The con-tinued discovery of new toxins like DarT emphasizes the role ofTA systems in cell biology.Type II TA systems are quite prevalent throughout bacterial

genomes. In a comprehensive analysis by Makarova et al. (20) in2009, of the 750 reference genomes searched, putative type IITA systems were found in 631, with on average 10 loci in each hit(6,797 total). The number of occurrences in a single genome canbe much larger, however; the deadly pathogen Mycobacteriumtuberculosis has over 88 TA loci in its genome, while the soilbacterium and plant symbiont Sinorhizobium meliloti containsan astounding 211 predicted type II loci (21, 22). Makarovaet al. proposed the existence of 19 new putative type II TA pairs,

Significance

Toxin–antitoxin (TA) systems are pairs of genes found throughoutbacteria that function in DNA maintenance and bacterial sur-vival. The toxins in these systems function by inactivatingcritical cell processes like protein synthesis. This work describesa TA system found in 17% of all sequenced bacteria. A high-resolution crystal structure and an array of biochemical testsrevealed that this toxin targets an essential enzyme in nucle-otide biosynthesis, a unique way in which TA systems can haltbacterial growth.

Author contributions: F.J.P. and A.J.L. designed research; F.J.P., P.D.J., and A.J.L. per-formed research; F.J.P., P.D.J., and A.J.L. analyzed data; and F.J.P. and A.J.L. wrotethe paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.

Data deposition: The atomic coordinates and structure factors have been deposited in theProtein Data Bank, www.wwpdb.org (PDB ID codes 6D0I and 6D0H).1To whom correspondence should be addressed. Email: ajlink@princeton.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1814633116/-/DCSupplemental.

Published online December 31, 2018.

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including 12 new toxins and antitoxins, again highlighting thevariety that exists within the type II class. To organize the rapidlyincreasing amount of data available, the TA database was cre-ated in 2010 and is an invaluable resource in parsing what weknow about these systems (23).Of the 19 predicted TA systems, one stood out to us in par-

ticular. The COG5654 (RES domain)-COG5642 TA family isone of five putative TA systems from this paper where bothcomponents were uncharacterized. We had encountered thispair in genome mining for natural products studied by our lab-oratory and were intrigued by how poorly understood it wasconsidering how widespread it appeared to be; it was present in250 of the 750 genomes analyzed by Makarova et al. Followingits identification in 2009, we found only one study that examinedthe putative COG5654–COG5642 TA pair, a genetic study onS. meliloti. This organism has 211 putative TA loci as mentionedabove, with roughly 100 of these on the organism’s two mega-plasmids. The authors demonstrated through a series of geneticdeletions that only four of these were actively functioning as TAsystems, including a COG5654–COG5642 locus, which con-firmed the hypothesis put forth in Makarova’s work (22).Here, we report a detailed physiological, structural, and bio-

chemical study of the COG5654–COG5642 TA system, startingwith a computational approach that demonstrates its prevalencein bacterial genomes. We confirm that it functions as a TA sys-tem in E. coli, an organism with no native COG5654–COG5642pairs, suggesting a target well-conserved in bacteria. We then report ahigh-resolution crystal structure of the protein complex, which in-

cludes the structure of an RES domain protein. Solving this structureallowed us to determine that the toxin is a mono-ADP-ribosyl-transferase (mART). We further show, both in vivo and in vitro thatthe toxin ADP-ribosylates phosphoribosyl pyrophosphate synthetase(Prs), an essential enzyme involved in nucleotide biosynthesis. Thisreport expands the set of bacteriostatic mechanisms employed byTA systems to include protein-modifying ADP-ribosylation.

ResultsSphingobium sp. YBL2 Bears Multiple COG5654–COG5642 TA Systems.Our investigation into Sphingobium sp. YBL2 began when agenome-mining algorithm developed in-house for discoveringnew lasso peptide producers identified this organism as havingtwo lasso peptide gene clusters of interest (24). It was immedi-ately evident that one of these gene clusters differed from theexpected architecture, containing within it genes encoding twohypothetical proteins, yblI and yblJ (Fig. 1A). A BLAST of thesesequences revealed them to be a COG5654–COG5642 putativeTA pair, first identified in a computational study in 2009 andlater confirmed to function as a TA pair in S. meliloti (20, 22). Toour knowledge, this is the last published work on this TA family.COG5654 toxins contain a roughly 150-aa RES domain,

named for a highly conserved Arg-Glu-Ser motif. In addition, thedomain also features two strongly conserved Tyr and His resi-dues. An alignment of the Sphingobium sp. YBL2 toxin withhomologs from various pathogens illustrates that outside of theseand neighboring residues, there is little shared sequence similarity(Fig. 1B). The function of the RES domain is unknown, and a

Fig. 1. Bioinformatic analysis of the COG5654–COG5642 TA family. (A) Lasso peptide gene cluster containing the TA pair studied here (yblIJ). (B) Sequencesimilarity of the COG5654 toxin from Sphingobium sp. YBL2 and several bacterial pathogens. Residues with >70% similarity are shaded. Similarity is clusteredaround the strictly conserved Arg-31, Tyr-41, and Glu-52 residues. (C) Distribution of this TA family among bacterial phyla is shown at Left; Proteobacteriahave by far the largest number of examples of this TA system. A further breakdown within Proteobacteria is shown at Right.

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BLAST of these toxins offers no homologous proteins withknown activities. COG5642 antitoxins meanwhile have a roughly60-aa C-terminal domain of unknown function, DUF2384, whichfollows an N-terminal DNA binding domain. Antitoxins fromtype II TA modules generally have C-terminal toxin bindingdomains, making this a likely function of DUF2384 (25).As Fig. 1B shows, COG5654 toxins appear in the genomes of

many pathogenic bacteria. To better understand the prevalence ofthe COG5654–COG5642 system, we devised code (Dataset S1)that searched all Refseq genomes for genes encoding an RESdomain protein downstream of a DUF2384 protein. The results ofthis search are summarized in Fig. 1C. Of the 108,787 bacterialRefseq genomes searched, there are 15,354 organisms that con-tain at least one copy of this TA family (14.1%) and 16,964 totalinstances. Some organisms contained many copies, with Spirosomaspitsbergense and Spirosoma luteum having 12 and 10, re-spectively. Sphingobium sp. YBL2 itself has five loci, three onits chromosome and two on plasmids. An alignment of the fivetoxins reveals high identity (45%) and similarity (53%) betweentwo of the chromosomal toxins, but little between the remainingthree, including the one we originally identified (SI Appendix,Table S1).To avoid overrepresentation of bacteria with many strains

characterized (e.g., M. tuberculosis or Bordetella pertussis), we con-solidated each into a single species entry. After doing so, the TAfamily was found in 2,942 unique bacterial species of 16,826(17.5%). Within these, we found the COG5654–COG5642 pair in15 bacterial phyla, although it was by far the most abundant andoverrepresented in Proteobacteria, occurring in 2,527 of 7,483 spe-cies (33.8%) (Table 1). The phyla also included Verrucomicrobia,Deinnococcus-Thermus, Spirochaetes, Planctomycetes, andFirmicutes, all of which were not observed earlier in Makarovaet al. Within Bacteroidetes, we identified 254 species and noticeda remarkably high overrepresentation in the Chitinophagia,Cytophagia, and Sphingobacteriia classes, occurring in 27/33(81.8%), 90/115 (78.3%), and 45/65 (69.2%) organisms, respec-tively. There were 86 Actinobacteria identified, including 55Mycobacterial species. Curiously, of the 3,406 Firmicutes ge-nomes, only a single instance (0.03%) of this TA family wasfound in Exiguobacterium sp. S17, which may warrant furtherinvestigation. Most Proteobacterial species were Alpha- (1,007/2,244, 44.9%), Beta- (542/1,167, 46.4%), and Gammaproteo-bacteria (949/3,617, 26.2%), with many from orders of knownpathogens, such as Burkholderiales, Legionellales, and Pseudo-monadales. A selected and full breakdown of the results can be

found in SI Appendix, Table S2 and Dataset S2, respectively. TheTA family was not observed in any archaeal genomes, in agree-ment with Makarova’s findings in 2009. The results of our searchconfirm that the COG5654–COG5642 system is widespread inbacteria, even more so than originally thought.

The COG5654–COG5642 Pair Functions as a TA System in E. coli. Tomore easily study the COG5654–COG5642 TA pair, we movedthe component genes from Sphingobium sp. YBL2 into E. coli.Although laboratory strains of E. coli harbor dozens of TA loci,the COG5654–COG5642 pair is not one of them, and thus, ourfirst aim was to confirm that this system can still function in thisnonnative organism (10, 25). The toxin was expressed from alow-copy pBAD33 plasmid (26), and upon induction, exerted abacteriostatic effect as measured by OD600 (Fig. 2A). We alsoexamined cell viability by colony forming unit (cfu) count. Onehour after toxin induction, the cfu count had decreased dra-matically, although it began to increase at 3 h (SI Appendix, Fig.S1). This is similar to what was observed with RelE, a well-knownbacteriostatic toxin (27). Coexpression of the COG5642 antitoxinrestored the normal growth phenotype, confirming that thisnonnative TA system functions normally in E. coli (Fig. 2A).We then performed an alanine scan on the highly conserved

residues of the RES domain described in the previous section.Substitution of alanine for Arg-31, Glu-52, and His-56 resulted inthe elimination of the toxic phenotype (Fig. 2B). Conversely,substitution of alanine for Tyr-41 and Ser-122 had no effect ontoxicity (Fig. 2B). While this is surprising in both cases, it is lessso for Ser-122, where the change is more conservative. Indeed,the alignment from Fig. 1B reveals that the corresponding aminoacid in M. tuberculosis is itself alanine. We subsequently exam-ined the mildly conserved Ser-21, 44, and 45 to exhaust candi-dates for the namesake serine in the “RES” domain, but thesesubstitutions also had no effect on toxicity (SI Appendix, Fig. S2).To confirm that the loss of toxicity of the R31A, E52A, and

H56A toxins was not driven by low expression, we cloned thesevariants into high-copy pQE80 vectors. The R31A toxin expressedat a high level as judged by SDS/PAGE and no growth defect wasobserved, while the E52A toxin recovered the toxic phenotype,likely due to the increased copy number of pQE80 relative topBAD33 (SI Appendix, Fig. S3) (28). Neither the wild-type nor theH56A toxin could be cloned into pQE80. This suggests that thereis still residual toxicity of the H56A variant due to leaky expres-sion. Taken together, these results indicate that while Glu-52 andHis-56 play a role in toxicity, Arg-31 is of paramount importance.

Crystallography Reveals Structural Similarity to mARTs. With noreasonable starting point for elucidating the function of theCOG5654 toxin, we instead set our sights on determining itsstructure, hoping to later infer its activity from this result.Because the wild-type toxin can only be made at a high levelwhen expressed alongside the cognate antitoxin, we set outto crystallize the protein complex first. For easily controlledcoexpression, we cloned toxin and antitoxin into a pRSFduetvector. The complex was expressed for 3 h at 37 °C before nativepurification, with typical yields of >5 mg/L. In our experimentalsetup, although only the toxin bears an affinity tag, the untaggedantitoxin copurifies due to strong protein–protein interactionscharacteristic of TA systems (Fig. 2C) (29, 30). Furthermore, theproteins elute as a single peak during size exclusion chromatog-raphy, with a retention time indicating a 2:2 stoichiometry (a TAdimer) (SI Appendix, Fig. S4).This purified TA complex had a solubility limit of 1 mg/mL

and was unsuitable for crystallization. The general understandingof TA systems is that the antitoxin is inherently less stable thanthe toxin, demonstrated previously in vitro by the selective totaldegradation by trypsin of the VapB antitoxin from a foldedVapBC complex (10, 31). We adapted this procedure for our

Table 1. Breakdown of bacterial Phyla containing theCOG5654–COG5642 TA family

TA-bearingspecies

Uniquespecies searched Percentage, % Phylum

14 27 51.85 Acidobacteria86 3,469 2.48 Actinobacteria254 1,334 19.04 Bacteroidetes1 8 12.50 Balneolaeota6 17 35.29 Chlorobi25 302 8.28 Cyanobacteria11 62 17.74 Deinococcus-Thermus1 3,406 0.03 Firmicutes2 3 66.67 Gemmatimonadetes1 1 100 Tectomicrobia3 41 7.32 Planctomycetes2,527 7,483 33.77 Proteobacteria1 5 20 Rhodothermaeota2 128 1.56 Spirochaetes6 40 15 Verrucomicrobia

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system and observed by LC/MS that even after overnight trypsintreatment, a 72-aa C-terminal antitoxin fragment was intact de-spite the existence of multiple possible cut sites (SI Appendix,Fig. S5). This, we hypothesized, was due to extensive TA contactsmaking the remaining sites inaccessible. We constructed a seriesof antitoxin truncations that left the C-terminal portion intactand found that even the shortest antitoxin fragment we tested(amino acids 99–159) could bind and neutralize toxin whencoexpressed (SI Appendix, Fig. S6). Upon trypsin digestion, thetoxin lost only its two most C-terminal amino acids. This trypsin-treated complex exhibited solubility greater than 10 mg/mL andshowed promise in crystal screening.The trypsin-digested toxin contains only a single methionine

(Met) residue. To increase Met content for selenomethionine(SeMet) substitution and single-wavelength anomalous disper-sion (SAD), we used a L48M toxin variant, which we confirmedretained toxicity (SI Appendix, Fig. S7). We successfully crystal-lized the trypsin-treated SeMet-substituted L48M TA complexand determined its structure to 1.55 Å using SAD (PDB ID code:6D0I). The complex crystallized in the P1 space group with unitcell dimensions of 41.98 Å, 51.32 Å, and 57.94 Å. From this, wesubsequently determined the structure of the trypsin-treatedwild-type complex to 1.50 Å (PDB ID code: 6D0H; Fig. 3A).A full description of data collection and refinement parameterscan be found in SI Appendix, Table S3. The complex crystallizesas a dimer, where the C terminus of each antitoxin is buriedin a putative toxin active site cleft housing all five highly con-served residues discussed above (Fig. 3B).We used PDBePISA to analyze the interactions between TA

complex subunits (32). The surface area of the TA interface is1,369.6 Å2 with an average ΔiG (free energy gain on interfaceformation, excluding hydrogen bonds and salt bridges) of−13.7 kcal/mol, indicating a strong affinity between the twoproteins. Extensive hydrogen bonding occurs between 13 anti-toxin and 14 toxin residues. Interestingly, salt bridging occursbetween the C-terminal carboxyl of the antitoxin and both theHis-56 and critical Arg-31 of the toxin (Fig. 3C). This interaction

places Arg-31 in a conformer directed away from the active site,also engaging it in hydrogen bonding with the toxin Ala-94 backbone oxygen. The strength of the TA interface is illus-trated experimentally by urea-induced dissociation, where puri-fied TA complex required incubation in 6 M urea to separate thecomponent proteins (SI Appendix, Fig. S8).A significant interface also exists between the two toxin sub-

units, with an interfacial surface area of 762.2 Å2 and ΔiG of−7.1 kcal/mol. The network of hydrogen bonding is less extensive,but notably there is an interaction between the highly conservedHis-56 of one toxin subunit and Tyr-153 of the other (Fig. 3D).Salt bridging between the toxin subunits occurs at Glu-127 andGlu-128 of one subunit and Arg-46 and Arg-149 of the other (Fig.3D). To determine if these salt bridges are necessary for toxicity,we constructed a series of mutations at the two glutamate residues.Single mutations to alanine had little effect on toxicity, but adouble mutation to even sterically similar glutamines rendered theprotein nontoxic (SI Appendix, Fig. S9). Thus, although a singlepair of salt bridges can be disrupted, the presence of at least one iscritical for interfacial integrity and proper function. This stronglysupports the idea of the toxin working as a dimer in vivo. Lastly,there is no discernable interface between the two antitoxin frag-ment subunits, suggesting these two exclusively interact with thetoxins they bind.Following the elucidation of the complex structure, we employed

the DALI structural similarity server to determine if any simi-larity to known proteins existed (33). The top result from thissearch was the catalytic domain of diphtheria toxin fromCorynebacterium diphtheriae, which had a Z-score of 7.0 andrmsd of 3.0 Å, indicating some structural similarity despite only11% sequence identity (SI Appendix, Fig. S10). Diphtheria toxinis a secreted mART capable of cleaving NAD+ into nicotinamideand ADP-ribose. It transfers the latter to a posttranslationallymodified histidine, diphthamide, in eukaryotic elongation factor-2 (EF-2), which inhibits protein synthesis (34). Other mARTssuch as cholix toxin from Vibrio cholerae and exotoxin A fromPseudomonas aeruginosa also exhibited strong Z-scores (SI Appendix,

Fig. 2. Heterologous expression in E. coli of the Sphingobium sp. YBL2 COG5654–COG5642 TA system. (A) Growth curves showing the effect of toxin ex-pression with and without antitoxin induction 30 min prior. The toxin exerts a bacteriostatic effect that is prevented by antitoxin coexpression. (B) Growthcurves of toxin variants with alanine substitutions for five highly conserved amino acids. R31A, E52A, and H56A variants are all nontoxic. Asterisks in A and Bindicate the timepoint of induction of toxin. (C) SDS/PAGE gel from a native purification of N-terminally His-tagged toxin coexpressed with untagged an-titoxin. Its interaction with the toxin results in copurification despite the lack of a His tag.

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Table S4) (35, 36), and we hypothesized based on these results thatthe COG5654 toxin is also a mART. Of note, the results alsocontain several poly-ADP-ribosyltransferases (pARTs) includingtankyrase-1 and -2, although these generally have lower Z-scoresthan the mARTs (Dataset S3).

Phosphoribosyl Pyrophosphate Synthetase Is a Putative Toxin Target.To test for mART activity, we required an isolated toxin sample.As described in the previous section, this could not be achievedby a trypsin digestion due to the strong interactions betweentoxin and antitoxin. A typical procedure for toxin isolation formany TA families involves complex denaturation, toxin repur-ification, and refolding by dialysis (16, 29, 30). We adapted thismethod for our system, noting that the toxin would only suc-cessfully refold when arginine was present in the dialysis buffer(37). When dialysis was complete, arginine could then be removedwithout causing toxin aggregation.As a first test of mART activity, we incubated 1 μM toxin with

1 mM NAD+. mARTs such as diphtheria toxin and iota toxinfrom Clostridium perfringens have been shown to exhibit weakNADase activity in the absence of a protein target (38–40). Weobserved not only a decrease in NAD+ and increase in freeADP-ribose, but also the appearance of an ADP-ribosylatedtoxin (SI Appendix, Fig. S11). A tryptic digest revealed that thismodification occurred on a single peptide that contains the im-portant Glu-52 and His-56 residues (SI Appendix, Fig. S11C).Although we are unsure of its significance for this system, auto–ADP-ribosylation is a known regulatory mechanism for severalmARTs and may serve a similar function here, regulating toxinactivity once disassociated from antitoxin in vivo (41, 42). Werepeated this reaction using the inactive R31A toxin variant and

observed that after 3 h, toxin ADP-ribosylation had greatlydecreased relative to the wild type (SI Appendix, Fig. S12),supporting the idea that the loss of toxicity of the R31A toxin isa result of a loss of transferase activity. The reactions were alsocarried out with wild-type toxin using an NADP+ substrate, and weobserved similar levels of glycohydrolase activity and automodifi-cation as for NAD+ (SI Appendix, Fig. S13A). This result was sur-prising as mARTs generally show high specificity toward NAD+.Encouraged by these results, we sought to identify a cellular

substrate for the toxin. We hypothesized based on toxin self–ADP-ribosylation that the target was a protein and not anucleic acid as was recently observed with DarT (19). Here, weemployed a pulldown where cell lysates from toxin-expressingcultures were incubated with macro domain protein AF1521from Archaeoglobus fulgidus and subject to His-tag affinity pu-rification. Macro domain proteins such as AF1521 have beenused previously to isolate ADP-ribosylated proteins from com-plex mixtures such as cell lysates (43, 44). Several proteins werepulled down via this technique, and all were identified after anin-gel tryptic digest, followed by LC/MS and Mascot peptidemass fingerprinting (SI Appendix, Fig. S14). These proteins,YfbG, GlmS, and EF-Tu, are all common contaminants associ-ated with His-tag affinity purifications (45, 46). We were in-trigued, however, by the last protein we identified, phosphoribosepyrophosphate synthetase (Prs), which is not a known affinitypurification contaminant. Prs is a homohexameric protein enco-ded by the essential prs gene, which is found in both bacteria andeukaryotes and strictly conserved in all nonparasitic organisms(47). Prs catalyzes the conversion of ribose 5-phosphate to phos-phoribose pyrophosphate, a precursor in the synthesis of nucleotides,histidine and tryptophan, NAD+, and NADP+ (48). Its critical position

Fig. 3. Crystal structure of the trypsin-treated TA complex. (A) Overall structure of the TA dimer (toxin, orange; antitoxin, blue) determined to 1.50-Åresolution. (B) Close-up of the putative active site. The five highly conserved residues examined in the alanine scan are highlighted. (C) Bonding network ofthe critical toxin His-56 and Arg-31. The arginine sidechain forms salt bridges with the C-terminal carboxyl of the antitoxin, which establishes a rotamer thatalso results in hydrogen bonding with the toxin’s own Ala-94. (D) Structure of the toxin-toxin interface. Shown are the intersubunit salt bridges between Arg-46 and Glu-128′ and Arg-149 and Glu-127′, as well as hydrogen bonding between Tyr-153 and the functionally important His-56′.

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in metabolism and strong conservation across a range of or-ganisms make it a logical target for cellular toxins.

The COG5654 Toxin ADP-ribosylates Prs in Vitro. We next sought toreconstitute Prs ADP-ribosylation in vitro. Reactions of 10 μMPrs, 5 μM toxin, and 10 mM NAD+ were set up at 37 °C, and theproducts were analyzed by LC/MS at 3 h and 20 h. After 3 h,about one-third of the Prs was ADP-ribosylated (Fig. 4A). Al-though most of the modified Prs was singly ADP-ribosylated(∼25% of the total), a small amount of Prs had acquired a sec-ond ADP-ribose. When the reaction was run for 20 h, the amountof ADP-ribosylated Prs had increased, with roughly equalamounts of unmodified, singly modified, and doubly modified Prspresent (Fig. 4A).Holding the concentration of Prs constant, we also performed

in vitro reactions at lower toxin (1 μM) and/or NAD+ (1 mM)concentrations. This concentration of NAD+ is below the phys-iological level in E. coli (49). We were interested in both how thiswould affect the rate of reaction, as well as if these conditionswould still drive the formation of doubly ADP-ribosylated Prs.After 3 h, about 10–15% of Prs had been ADP-ribosylated (downfrom ∼33%) and little to no doubly modified Prs was present.Additionally, the effect was independent of whether toxin orNAD+ (or both) concentration had been lowered (SI Appendix,Fig. S15). By increasing the reaction time to 20 h, singly modifiedpeaks doubled in size, now accounting for ∼25% of total Prs. Aswith high toxin and NAD+ concentration, doubly modified spe-cies showed the most increase from 3 h to 20 h, now present atroughly the same amount as singly modified Prs. This longer timecourse suggests that the additional modification is not merely afunction of high reactant concentration. Also of note, we ob-served that the extent of toxin auto–ADP-ribosylation decreasedin the presence of Prs, supporting the idea that self–ADP-ribosylation may be a form of regulation in the absence of target(SI Appendix, Fig. S16). Collectively, this data shows that thetoxin can ADP-ribosylate Prs twice, a surprising result giventhat mARTs generally ADP-ribosylate only a single amino acidon their target.

To confirm that these Prs modifications are not occurringnonspecifically, we used higher energy collision dissociation(HCD) LC/MS2 to analyze tryptic digests of the in vitro reactions(50). Analysis with Scaffold proteomics software revealed thatthe majority of ADP-ribosylation occurred at either Lys-182 orSer-202 of the E. coli Prs, showing that these modifications arespecific (Fig. 4B and SI Appendix, Fig. S17). Lys-182 is conservedamong many bacterial Prs and has been shown to participate inATP binding, placing it at the Prs active site (51). The Prs fromSphingobium sp. YBL2 has an aspartic acid (also a mART target)at this position, which is also present in M. tuberculosis and otherbacteria, as well as in all three human Prs isozymes (51). Amutation of this residue to histidine in human isozyme one hasbeen implicated in dysregulation in patients with Prs superac-tivity, suggesting that an ADP-ribose modification could alsocritically alter Prs in bacteria (52). Ser-202 shows some conser-vation but, to our knowledge, is not critical for catalytic activityor regulation. It is, however, part of a flexible loop involved incatalysis (53), and its modification could prevent this region fromentering a productive conformation. Although the correspondingvaline in Sphingobium sp. YBL2 Prs cannot be ADP-ribosylated,a targetable threonine is adjacent. Further investigation intothese modifications is necessary, both to determine if one is apreferred substrate and what effect the modification has on Prsactivity. Although we wished to repeat these reactions with theSphingobium sp. YBL2 Prs, the protein expressed poorly in E.coli and coeluted with large quantities of GroEL (SI Appendix,Fig. S18). Based on the toxin’s activity, we propose to rename theRES domain family of toxins to ParT (Prs ADP-ribosylatingtoxin) and the corresponding antitoxin family to ParS.Noting ParT’s ability to cleave NADP+, we also attempted to

modify Prs with this substrate, but no pADP-ribosylation couldbe detected (SI Appendix, Fig. S13B). This agrees with what istypically observed for mARTs and demonstrates that here, al-though there may be some relaxed specificity for glycohydrolysis,transferase activity still requires the traditional NAD+ substrate.

Fig. 4. In vitro ADP-ribosylation of E. coli Prs. (A) ESI mass spectrometry of the reaction of Prs (10 μM) with toxin (5 μM) and NAD+ (10 mM). After 3 h,conversion to a singly ADP-ribosylated Prs is ∼25%, with a doubly ADP-ribosylated peak beginning to appear. At 20 h, ∼65% of the Prs is ADP-ribosylated,with ∼35% singly modified and ∼30% doubly modified. (B) Crystal structure of the E. coli Prs hexamer (PDB ID code 4S2U). The ADP-ribosylated residuesidentified by LC/MS2, Lys-182 and Ser-202, are shown in gray. The Lys-182 sidechain is directed toward the Prs active site, while Ser-202 is located on acatalytic flexible loop.

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An Inactive Prs Variant Attenuates the ParT-Induced Growth Defect.To confirm that ParT targets Prs in vivo, we planned to sup-plement Prs on a plasmid. An analogous experiment with HipAdemonstrated that overexpression of its target, GltX, allowed forrecovery of normal cell growth (16). It is known, however, thatPrs hyperactivity is toxic to human cells (54), and indeed, weobserved a growth defect in E. coli when overexpressing the wild-type protein, making direct supplementation of Prs impossible(SI Appendix, Fig. S19A). To overcome this issue, we insteadsupplied an inactive H131A variant of E. coli Prs, which alters ahistidine involved in binding the γ-phosphate of the ATP sub-strate (55, 56). As expected, overexpression of this H131A Prscaused no negative effects on cell growth (SI Appendix, Fig.S19B). Supplying an inactive Prs is not ideal, but we reasonedthat the overexpressed protein could at act as a decoy, with ParTtargeting the abundant inactive Prs rather than the productivenative enzyme. Although eventually the native Prs would also getmodified, we believed this should at least attenuate ParT toxicity.We expressed H131A Prs for 1 h before ParT induction. Rather

than the immediate growth arrest that is seen in cultures expressingonly ParT, cultures also expressing H131A Prs continued to growfollowing toxin induction (Fig. 5A). Although growth is slower thanin ParT-uninduced cultures, final OD600 is considerably higher inH131A Prs coexpressing cultures than in those expressing the toxinalone (Fig. 5A). This result agrees with our hypothesis that althoughH131A Prs can mitigate toxicity, it cannot completely overcome it.A similar effect was observed in cultures expressing H131A Prs foronly 30 min before ParT induction, but to a lesser degree.To rule out the other proteins identified in the macro domain

pulldown as purely contaminants and not possible ParT targets,we repeated this coexpression with EF-Tu in place of Prs. Wechose EF-Tu because of its association with multiple other post-translational modifications, including phosphorylation, methylation,and acetylation (57–59). Unlike in the case of Prs, expression of EF-Tu for 1 h before ParT induction led to a phenotype that was in-distinguishable from cultures expressing ParT only (SI Appendix, Fig.S20). This result strongly suggests that EF-Tu is not a target of ParT.

To confirm that the change in OD600 observed in culturescoexpressing toxin and H131A Prs was not due to cell morphologychanges or effects unrelated to ParT, we also examined cell via-bility. Cfu counts were taken immediately before ParT induction,as well as 15 min and 1 h after induction. Within 15 min, the vi-ability of E. coli expressing only ParT had fallen drastically by fourorders of magnitude (Fig. 5 B and C). Cultures that had beenexpressing H131A Prs before ParT induction showed a singleorder of magnitude decrease in cell viability, markedly better thanParT-only cells. This result strongly agrees with the growth dataand confirms that the increased cell density is due to improvedviability caused by the presence of H131A Prs.

DiscussionHere, we provide biochemical and structural insights into the na-ture of the COG5654–COG5642 TA system, identifying the toxinas a mART that targets the essential metabolic enzyme Prs. Wehave thus renamed the toxin parT and the cognate antitoxin parS.We determined a high-resolution crystal structure of the ParSTcomplex, which revealed that the complex crystalizes as a dimer(Fig. 3A). Disruption of the dimeric interface between the toxinsleads to a loss of the toxic phenotype (SI Appendix, Fig. S9), illus-trating that ParT is a mART that functions as an obligate dimer.The C terminus of each ParS antitoxin is buried in the active sitecleft of the cognate toxin, forming a salt bridge with both His-56 and Arg-31 of ParT. Among mARTs, this is an example witha high-affinity inhibitor that directly binds the mART active site.Although a BLAST of ParT reveals sequence similarity only to

other RES domain proteins, a DALI search revealed multiplemARTs with similarities in secondary structure, including exo-toxins such as diphtheria, cholera, and pertussis toxins (DatasetS3). However, key differences emerge between ParT and theseknown bacterial mARTs, making it a unique addition to thisfamily of enzymes. First, the exotoxins are all secreted and haveeukaryote-specific targets. ParT, however, acts in the cytoplasmof its producing cells and its target, Prs, is conserved amongall free-living organisms. The secretion of mART exotoxins is

Fig. 5. Coexpression of ParT with an inactive Prs variant suppresses ParT toxicity. (A) Growth curves of cells coexpressing H131A Prs and ParT. H131A Prs wasinduced at 4 h (**) and toxin induced 1 h later (*). Cultures that accumulated H131A Prs showed a marked increase in OD600 following toxin induction. (B) Cellviability as measured by cfu count. Within 15 min, cfu count drops dramatically (∼10,000-fold) when cultures express only ParT. In contrast, cultures allowed toaccumulate H131A Prs show only mildly decreased viability (∼10-fold). (C) Cfu count 15 min after toxin induction. Cultures were diluted 10,000-fold before100 μL was plated on LB supplemented with the appropriate antibiotics and grown overnight at 37 °C.

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accomplished with an N-terminal signaling sequence, while up-take into eukaryotic cells is mediated in most cases through adistinct recognition domain. This is not the case for ParT,however, which is comprised solely of the catalytic RES domain.In this sense, ParT is most like C3 bacterial mARTs, which arealso single-domain proteins, although they retain an N-terminalsignal sequence (60). Furthermore, despite the similar domainarchitecture, C3 toxins (∼25 kDa) are substantially larger thanParT (∼18 kDa) and a structural alignment reveals little similarity.To our knowledge, ParT is the smallest example of a mART and,thus, may represent a mART that appeared early in evolution.The catalytic motif of ParT bears a resemblance to both diph-

theria toxin-like (DT) and cholera toxin-like (CT) mART motifsbut is distinct from both. The defining DT HYE motif featurestwo highly conserved tyrosines, a histidine, and glutamate, allcritical for toxicity (60). ParT and other RES domain proteinsinclude each of these residues in their active site, with the Glu andHis residues important for toxin function (Fig. 2B). ParT alsocontains one conserved Tyr residue, but it is not critical for toxicity(Fig. 2B). The HYE motif of DT lacks an arginine residue com-parable to the critical Arg-31 residue in ParT (Figs. 2B and 3B),highlighting the differences between these two mART active sites.The CT RSE motif is defined by a critical arginine, an STS se-quence, and an (E/N)XE sequence (60). Despite being almostidentical in name to the RES domain in ParT, there are minimalsimilarities between these active sites besides the critical arginineresidue. Current structural and biochemical data on the mecha-nism of mART exotoxins suggests that mART active sites stabilizethe cation formed upon glycohydrolysis of NAD+ (36, 61, 62).Given that the ParT active site differs from both of the canonicalmART active sites, it is an attractive target for further mechanisticanalysis of the ADP-ribosyltransferase reaction.Recently, the DarT toxin was shown to ADP-ribosylate thy-

midines on single-stranded DNA, establishing nucleotidyl ADP-ribosylation as yet another bacteriostatic mechanism of the di-verse type II toxins (19). In contrast, our work demonstrates thattype II toxins can also be protein mARTs. Intracellular proteinmARTs in bacteria are exceptionally rare, with the only priorexample a nitrogenase mART from Rhodospirillum rubrum (63).ParT exerts its bacteriostatic effect via modification of a cellulartarget, Prs. This represents an essential cellular function, nucleotidebiosynthesis, targeted by TA systems. Further distinguishing theParST system from DarTG is that ParS exhibits toxin neutraliza-tion via a protein–protein interaction, rather than the modification-reversing activity of ADP-ribosyl glycohydrolase DarG. While thestructure of DarG was determined, there is no structure of DarT tocompare with the ParT structure. Furthermore, DarT and ParTshare little sequence similarity and DarT is 230 aa compared withthe much smaller ParT (161 aa). A recent review article postulatedthat additional TA systems could harbor mARTs that modify pro-teins (64). Our work here bears out that prediction and shows thatthis type of TA system is exceptionally widespread across bacteria.

MethodsPlasmid Construction. All cloning was performed in E. coli XL1 Blue cells.Genes encoding toxin and antitoxin were amplified from the genome ofSphingobium sp. YBL2. The organism was a gift from Xin Yan’s laboratory atNanjing Agriculture University, Nanjing, China. Antitoxin was cloned into highcopy pQE80-L, and toxin was cloned into low copy pBAD33. For coex-pression, the proteins were cloned into pRSF-duet. Macro domainAF1521 as well as E. coli and Sphingobium sp. YBL2 Prs were cloned intopQE80-L. Mutations were introduced using PCR site-directed mutagene-sis. A full list of primers and plasmids used in this study can be found in SIAppendix, Tables S5 and S6.

COG5654–COG5642 Genome Search. The code used to search for COG5654–COG5642 TA pairs can be found in Dataset S1 in SI Appendix.

Protein Expression and Purification. All His-tagged protein expression wasinduced in midlog phase BL21(DE3) E. coli (ΔslyD) and carried out for 3 h at37 °C. Proteins were purified via Ni-NTA native affinity purification accord-ing to the manufacturer’s protocol (Qiagen). For toxin isolation, TA complexwas first purified as described above. The complex was dissociated in 6 Murea, and denatured toxin was repurified by Ni-NTA affinity purification.Arginine was then added to a final concentration of 200 mM, and thesample was dialyzed against TBS supplemented with 200 mM arginine usinga Slide-alyzer cassette (GE) followed by buffer exchange into TBS.

Growth Curves. Growth curves were measured using a Synergy 4 plate reader(Biotek). In isolated toxin expression, expression was induced at the begin-ning of exponential phase by the addition of arabinose. In coexpressions withother proteins, the nontoxic protein was induced 1 h before the toxin. Ex-pressions were carried out for at least 4 h.

Trypsin Digestion of Native TA Complex. Trypsin digestion was carried out withsequencing grade modified trypsin according to the manufacturer’s protocolwithout a denaturation step (Promega). The digested complex was analyzedby SDS/PAGE and LC/MS to determine the extent of degradation (LC: Agilent1260 Infinity with Zorbax 300SB-C18 column, MS: Agilent 3560 Accurate-Mass qTOF).

SeMet Protein Expression and Purification. Overnight cultures were used toinoculate M9 minimal media supplemented with 50 mg/L selenomethionineand 40mg/L of all amino acids except for methionine. Cultures were grown toearly log phase, at which point a solution of Ile, Leu, Val, Lys, Phe, and Thr(10 mg/mL each) was added to suppress Met biosynthesis. After 20 min,protein expression was induced. Expression, purification, and trypsin di-gestion of the complex were all carried out as described above.

Protein Crystallization. A 10 mg/mL solution of trypsin-digested TA complexwas subject to screening using a Phoenix crystallization robot (Art Robbins).On scaleup, crystals were grown at 20 °C using the sitting-drop vapor dif-fusion method. Drops were 1.5 μL:1 μL protein solution:MPD precipitantsolution (0.1 M sodium acetate trihydrate, 10 vol% MPD, pH 5.0). Crystalstypically reached maximum size in 2 wk, were harvested, and dipped incryoprotectant (MPD solution and 10% glycerol) before vitrification. SeMet-substituted L48M TA complex was crystallized using identical methods. Datafor SeMet L48M TA complex was collected at the Brookhaven NationalLaboratory National Synchrotron Light Source II, and phasing was de-termined using SAD. The SeMet L48M TA structure was used to solve thewild-type TA complex, with data collected in the Princeton macromolecularcrystallography core facility with a Rigaku Micromax-007HF X-ray generatorand Dectris Pilatus 3R 300K area detector.

Macro Domain Pulldown and Prs Identification. E. coli expressing toxin for 1 hwere harvested and lysed by sonication. Separately, His-tagged macrodomain protein was incubated with Ni-NTA slurry to saturate the resin andreduce nonspecific binding. The macro domain-slurry mixture was thenincubated with the toxin-induced lysate for 2 h at room temperature be-fore purification. The elutions were analyzed by SDS/PAGE and subject toan in-gel trypsin digest according to the manufacturer’s protocol (ThermoFisher Scientific). Digests were analyzed by LC/MS against the MASCOTprotein database.

In Vitro ADP–Ribosylation. Reactions were performed in TBS at 37 °C. Prsconcentration was held constant at 10 μM, while NAD(P)+ concentration wasvaried between 1 mM and 10 mM and toxin between 1 μM and 5 μM.Samples were taken after 3 h, and the reactions continued overnight (20 htotal). For intact protein analysis, samples were injected directly onto theAgilent LC/MS system described above. Trypsin-digested samples wereused to determine the modification sites using a coupled Easy 1200 UPLCand Fusion LUMOS mass spectrometer at the Princeton University corefacility.

Cell Viability (Cfu Count). Cells transformedwith toxin, toxin and blank pQE80-L,or toxin and H131A Prs were grown to the beginning of log phase, at whichpoint nontoxic proteins were allowed to express for 1 h before toxin ex-pression was then induced. Samples of culture were taken immediatelybefore toxin induction, as well as 15 min and 1 h after induction. These wereserial diluted in LB and plated on LB with the appropriate antibiotics. Plateswere incubated overnight at 37 °C and colonies were counted manually thenext day.

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ACKNOWLEDGMENTS. We thank Dr. Tharan Srikumar of the Princeton Univer-sity Mass Spectrometry Facility for assistance with HCD LC/MS2 experiments, thestaff of beamline AMX at the National Synchrotron Light Source II for assistance

with data collection, and Jordan Ash of Ryan Adams’ laboratory at PrincetonUniversity for assistance with writing and implementing code for the ParST ge-nome search. Funding was provided in part by NIH Grant GM107036 (to A.J.L.).

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