crossm High-Resolution X-Ray Structures of Two Functionally Distinct Members of the Cyclic Amide Hydrolase Family of Toblerone Fold Enzymes Thomas S. Peat,a Sahil Balotra,b,c Matthew Wilding,c Carol J. Hartley,c Janet Newman,a Colin Scottc CSIRO Biomedical Manufacturing, Parkville, Victoria, Australiaa; Research School of Chemistry, Australian National University, Canberra, Australian Capital Territory, Australiab; CSIRO Land and Water, Black Mountain, Canberra, Australian Capital Territory, Australiac

ABSTRACT The Toblerone fold was discovered recently when the first structure of the cyclic amide hydrolase, AtzD (a cyanuric acid hydrolase), was elucidated. We surveyed the cyclic amide hydrolase family, finding a strong correlation between phylogenetic distribution and specificity for either cyanuric acid or barbituric acid. One of six classes (IV) could not be tested due to a lack of expression of the proteins from it, and another class (V) had neither cyanuric acid nor barbituric acid hydrolase activity. High-resolution X-ray structures were obtained for a class VI barbituric acid hydrolase (1.7 Å) from a Rhodococcus species and a class V cyclic amide hydrolase (2.4 Å) from a Frankia species for which we were unable to identify a substrate. Both structures were homologous with the tetrameric Toblerone fold enzyme AtzD, demonstrating a high degree of structural conservation within the cyclic amide hydrolase family. The barbituric acid hydrolase structure did not contain zinc, in contrast with early reports of zinc-dependent activity for this enzyme. Instead, each barbituric acid hydrolase monomer contained either Na⫹ or Mg2⫹, analogous to the structural metal found in cyanuric acid hydrolase. The Frankia cyclic amide hydrolase contained no metal but instead formed unusual, reversible, intermolecular vicinal disulfide bonds that contributed to the thermal stability of the protein. The active sites were largely conserved between the three enzymes, differing at six positions, which likely determine substrate specificity.

Received 17 December 2016 Accepted 15 February 2017 Accepted manuscript posted online 24 February 2017 Citation Peat TS, Balotra S, Wilding M, Hartley CJ, Newman J, Scott C. 2017. High-resolution X-ray structures of two functionally distinct members of the cyclic amide hydrolase family of Toblerone fold enzymes. Appl Environ Microbiol 83:e03365-16. https://doi.org/ 10.1128/AEM.03365-16. Editor Rebecca E. Parales, University of California—Davis © Crown copyright 2017. The government of Australia, Canada, or the UK (“the Crown”) owns the copyright interests of authors who are government employees. The Crown Copyright is not transferable. Address correspondence to Colin Scott, [email protected]

IMPORTANCE The Toblerone fold enzymes catalyze an unusual ring-opening hydro-

lysis with cyclic amide substrates. A survey of these enzymes shows that there is a good correlation between physiological function and phylogenetic distribution within this family of enzymes and provide insights into the evolutionary relationships between the cyanuric acid and barbituric acid hydrolases. This family of enzymes is structurally and mechanistically distinct from other enzyme families; however, to date the structure of just two, physiologically identical, enzymes from this family has been described. We present two new structures: a barbituric acid hydrolase and an enzyme of unknown function. These structures confirm that members of the CyAH family have the unusual Toblerone fold, albeit with some significant differences. KEYWORDS atrazine, evolution, hydrolase, phylogenetic analysis, structure-activity relationships, triazine


n 2013, we described a unique fold for AtzD, the cyanuric acid hydrolase (CAH) from Pseudomonas sp. strain ADP, with the first X-ray structure of this enzyme (1). CAH belongs to the recently identified cyclic amide hydrolase (CyAH) family, members of May 2017 Volume 83 Issue 9 e03365-16

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FIG 1 Reaction schemes for cyanuric acid hydrolase (CAH) and barbituric acid hydrolase (BAH). CAH and BAH catalyze analogous ring-opening hydrolyzes with cyanuric acid (CA) and barbituric acid (BA), respectively. The product of CAH-mediated hydrolysis (1-carboxybiuret) is unstable in water and decomposes to form biuret. Spon., spontaneous.

which perform novel ring-opening reactions with triazines and pyrimidines (Fig. 1) (1–8). To date, the only reactions characterized for members of this family of enzymes are the hydrolysis of cyanuric acid (CA) or barbituric acid (BA). Interestingly, no enzyme has been described that will hydrolyze both CA and BA, despite the high degree of similarity in the two substrates (1, 4, 6–8). The physiological role of BAH is in the catabolism of pyrimidines. BAH catalyzes the opening of the BA ring to yield ureidomalonic acid, which is then further hydrolyzed by an ureidomalonic acid hydrolase (8). BA is formed in the oxidative catabolic pathway for pyrimidines (8); this is one of three known pathways for pyrimidine catabolism, the others being the more common reductive and Rut (pyrimidine utilization) pathways (9–13). The physiological role of CAH is in the decomposition of CA, often as part of a catabolic pathway for atrazine and related herbicidal s-triazines (1). CAH hydrolyzes CA, forming the unstable metabolite 1-carboxybiuret, which decomposes to biuret spontaneously in an aqueous environment (Fig. 1). Biuret is mineralized by the action of two further enzymes (biuret hydrolase and allophanate hydrolase) (1, 4, 14–16). Although CA is occasionally formed during the oxidative damage of DNA, its abundance in the environment has increased considerably since the late 1950s as a result of the intensive use of s-triazine-based compounds in herbicides (e.g., atrazine) and polymers (e.g., melamine) (1). Based on their biochemical similarities, high sequence identity (⬎50%), and physiological roles, it has been proposed that CAH evolved from BAH (1, 4, 8). CAH is a homotetramer, the monomer of which possesses a novel fold that we dubbed the Toblerone fold (1). Although not obvious from the protein sequence, each CAH monomer possesses 3-fold rotational symmetry, for which the repeated structural motif is structurally similar to that of the YjgF superfamily of proteins (two ␣ helices and a four-strand antiparallel ␤-sheet), despite having low sequence similarity (⬍15%). This suggests that the Toblerone fold evolved from concatenated YjgF family proteins (1). Each CAH monomer coordinates a single noncatalytic cation (most likely Mg2⫹ based on a combination of biochemical and crystallographic evidence) via a unique metalbinding site (1). The structure of a second CAH, from Azorhizobium caulinodans, has also been described and is essentially identical to that of CAH from Pseudomonas sp. strain ADP (2). Interestingly, an enzyme with low sequence identity with the CyAH family, chorismatase (FkbO), has been shown to share a degree of architectural similarity to the CAH structures (17), which suggests that the Toblerone structural family is larger than the CyAH family. The 3-fold symmetry of each CAH monomer extends to the active site, which contains three Arg-Lys-Ser triads, which bind the substrate via an extensive hydrogen bonding network (1). Biochemical analysis revealed that serine and lysine are essential May 2017 Volume 83 Issue 9 e03365-16

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for catalytic competence, and it is likely that one or more of the active-site Ser-Lys dyads are catalytically active, with the serine acting as a nucleophile that opens the ring, forming a covalent intermediate that is subsequently hydrolyzed to regenerate the active site. Crystallographic and proteomic evidence suggested that Ser85 and Lys42 were the catalytic dyad; however, genetic and phylogenetic evidence presented in a later study suggest that Ser226 and Lys156 comprise the catalytic center (1, 2). To date, no BAH structure has been available to confirm that CAH and BAH share the same fold; however, it seems highly likely based on their level of sequence conservation (1, 4). Reports from Soong et al. have suggested that, unlike CAH, BAH is a zincdependent metalloprotease (8). Although protein folds can accommodate a diversity of catalytic centers and reaction mechanisms (e.g., the TIM-barrel fold [18–21]), the sequence conservation of the proposed CAH catalytic residues, homology modeling of BAH based on the CAH structure, and similarities in the substrates and products of the two enzymes suggest that there is a shared catalytic mechanism between BAH and CAH (1, 4). It should be noted that the noncatalytic metal in CAH was not suspected until the crystal structure was obtained (1), and it is possible that noncatalytic Zn2⫹ in BAH is responsible for the loss of activity observed in BAH when treated with chelating agents. Homology modeling of BAH based on the CAH structure had suggested that the difference in the two enzymes’ substrate specificities was determined by amino acid residues at positions 194, 320, 324, 344, and 345 (per CAH numbering) (1). The distribution of the predicted activities correlated with the phylogenetic distribution of the CyAH family (Fig. 2): CAH activity is predicted in three of the six classes (i.e., classes I, II, and III), and BAH activities were predicted in a fourth class (i.e., class VI) (1). Classes IV and V, however, appear to have different amino acid compositions for their specificity-determining motif, suggesting that there are additional physiological functions for members of this protein family, although the substrate specificities of enzymes in all of the classes have not been assessed empirically (1). Here, we have confirmed that enzymes from classes predicted to contain CAH and BAH have the predicted enzymatic activities, with one exception: a BAH that has promiscuous CAH activity. We have also solved the X-ray structures of a BAH and a class V CyAH of unknown function from Frankia sp. strain Eul1, allowing further analysis of structure/function relationships within this unique family of enzymes. RESULTS AND DISCUSSION Phylogenetic relationships correlate with physiological function in the CyAH family. The relationship between the phylogenetic distribution of cyclic amide hydrolases and their biochemical functions was investigated in previously uncharacterized CyAH enzymes, which were tested against a range of substrates (Table 1). The observed enzymatic activities were largely consistent with those which we had predicted previously: enzymes from classes I to III were found to be CAHs, with second-order rate constants ranging from 2.8 ⫻ 104 to 2.7 ⫻ 106 M⫺1 · s⫺1 (Table 1), and members of class VI possessed BAH activity, with second-order rate constants in the range of 3.6 ⫻ 107 to 4.6 ⫻ 108 M⫺1 · s⫺1 (Table 1). The Nocardioides sp. strain JS614 genome carries two paralogous class VI genes (GenBank accession numbers ABL81019.1 and ABL83767.1; 71% identical), the products of which were both tested for activity. Interestingly, one of the two encoded enzymes (YP925454) had low-level CAH activity (1.1 ⫻ 103 M⫺1 · s⫺1), in addition to its predicted BAH activity (3.6 ⫻ 107 M⫺1 · s⫺1), while its paralog possessed no detectable CAH activity (BAH activity of 8.9 ⫻ 107 M⫺1 · s⫺1) (Table 1). This is the first enzyme from the CyAH family reported to possess a promiscuous activity, a property that has been shown to be important in the evolution of new enzyme function (22–26) and is consistent with previous suggestions that CAH have evolved from an ancestral BAH (1, 4, 6). Classes IV and V were predicted to contain enzymes that did not possess either CAH or BAH activities (1), and evidence from a class IV enzyme from Azorhizobium caulinodans ORS 571 was consistent with this prediction, with no detectable activity with May 2017 Volume 83 Issue 9 e03365-16

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FIG 2 Phylogeny of the Toblerone fold cyclic amide hydrolase (CyAH) family. The CyAHs are distributed into six classes. Classes I to III are cyanuric acid hydrolases, class VI includes barbituric acid hydrolases, and the functions of classes IV and V are not yet known. Enzymes for which there are solved structures are marked with asterisks. Enzymes with potential disulfides are marked with a “d,” and those with potential vicinal disulfides are marked with a “v.” Enzymes that do not possess a conserved metal-binding motif are shown with an “m.” The substrates for enzymes that have been biochemically characterized are also shown: C, cyanuric acid; B, barbituric acid; and N, neither cyanuric acid nor barbituric acid.

either BA or CA substrates. Despite concerted efforts, we were unable to express soluble enzymes from the class IV CyAH-encoding genes from Acidithiobacillus ferrooxidans ATCC 53993 and Sulfobacillus acidophilus DSM 10332 or from the class V CyAHencoding gene from Bacillus cellulolyticus DSM 2552 and therefore were unable to test their substrate specificities. However, we were able to express the class V CyAH from Frankia sp. strain Eul1. Neither CAH nor BAH activities were found for the Frankia sp. strain CyAH, so the enzyme was tested for activity against a range of pyrimidines and triazines, including BA, CA, alloxan, thymine, uracil, methyl uracil, dihydroorotate, and orotic acid, but no detectable activity was found with any of the compounds tested. Structural comparison of CAH, BAH, and the Frankia CyAH. To date, the only crystal structures from the CyAH family have been the CAHs from Pseudomonas sp. strain ADP and A. caulinodans (1, 2), which have a high degree of sequence and structural similarity to each other. In the work presented here, we have empirically determined the structures of a BAH and the CyAH from Frankia (Fig. 3 and Table 2), extending our knowledge of this structural family. Comparison of the three structures shows that the Toblerone fold is conserved, with the main differences restricted to the loop regions between the main secondary May 2017 Volume 83 Issue 9 e03365-16

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TABLE 1 Kinetic properties of the members of the CyAH family characterized to date Class I II II II II III III III III III V V VI

Genome Pseudomonas dioxanivorans CB1190 Pseudomonas sp. strain NRRLB-12227 (TrzD) Moorella thermoacetica ATCC 39073 Pseudomonas sp. strain ADP Bradyrhizobium japonicum USDA110 Pseudomonas pseudoalcaligenes CECT 5344 Azorhizobium caulinodans ORS 571b (BAF89890) Bradyrhizobium sp. strain ORS 375 Rhizobium leguminosarum bv. trifolii WSM1325 Rhizobium leguminosarum bv. viciae 3841 Azorhizobium caulinodans ORS 571 (BAF89201) Frankia sp. strain Eul1 Nocardioides sp. strain JS614 (ABL81019.1)


Nocardioides sp. strain JS614 (ABL83767.1) Rhodococcus erythropolis JCM3132



kcat/Km (Mⴚ1 · sⴚ1) 3.2 ⫻ 105 5.9 ⫻ 105 3.3 ⫻ 104 4.0 2.8 1.3 4.0 2.7 3.8

⫻ ⫻ ⫻ ⫻ ⫻ ⫻

104 104 105 104 106 104

3.6 1.1 8.9 4.6

⫻ ⫻ ⫻ ⫻

107 103 107 108

Km (␮M) 176 159 140 79 370



Reference and/or source This work This work, 41 This work, 5 1 4 This work 4 This work This work 4 4 This work This work This work This work, 7

cyanuric acid; BA, barbituric acid; ND, not determined.

structure features. The quaternary structure is also conserved, as all three of the enzymes form a D2 tetramer despite the space group promiscuity: the Pseudomonas CAH was observed in R32, BAH crystallized in three space groups (I222, P41212, and P212121), and the Frankia CyAH crystallized in the P1, I2, and P212121 space groups. The monomeric structures were aligned using the SSM algorithm in Coot. The CAH and BAH structures share 44% sequence identity and a 1.5-Å root mean square deviation (RMSD) in C␣ positions across 346 aligned residues (9 gaps), the Frankia enzyme has 36% sequence identity with CAH and a 1.9-Å RMSD in C␣ positions across 343 aligned residues (11 gaps), and the Frankia CyAH structure has 41% sequence identity and 1.3-Å RMSD in C␣ positions across 336 aligned residues (12 gaps) to BAH. The active sites were well conserved and often occupied by buffer molecules in the absence of substrate or inhibitor [CAH has phosphate and BAH has an N-(2acetamido)iminodiacetic acid (ADA) molecule]. The composition of the active sites are the following (given in the order CAH/BAH/Frankia CyAH; amino acid identities are given for CAH and where they differ between the three enzymes) (Fig. 4): Lys40/41/54, Arg54/53/66, Ser83/83/97, Gly84/84/98, Lys162/162/177, Arg194/Asn194/Asn213, Ser232/ 230/251, Ala233/Ser231/Ser252, Lys295/299/319, Thr320/His324/His342, Arg324/Lys328/ Lys346, Ser343/347/365, Gly344/Val348/Gly366, and Gly345/Ser349/Gly367. The Frankia CyAH active site is somewhat larger than those of CAH and BAH, and it incorporates an additional amino acid reside (Glu368), although the significance of these differences is unclear in the absence of a known substrate.

FIG 3 Structures for the monomers of the three functionally distinct CyAH family enzymes determined to date. ␣-Helices are shown in red, ␤-strands are in yellow, and loops are in green. The structural metals of cyanuric acid hydrolase and barbituric acid hydrolase are shown as gray spheres. May 2017 Volume 83 Issue 9 e03365-16

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70.3, 82.6, 114.3 90, 90, 90

41.3–1.71 (1.74–1.71) 0.103 (0.799) 0.041 (0.330) 0.998 (0.729) 13.2 (2.3) 99.7 (93.8) 7.2 (6.4)

41.5–1.71 34,725 15.1/19.4

3,128 2,815 13 295

19.6 19.2 19.2 26.6

0.019 1.912

Cell dimensions a, b, c (Å) ␣, ␤, ␥ (°)

Resolution (Å) Rmerge Rpim CC1/2 Mean I/␴I Completeness (%) Redundancy

Refinement statistics Resolution (Å) No. of unique reflections Rwork/Rfree (%)

No. of atoms Total Protein Inhibitor/buffer Water

B-factor (Å2) Protein Inhibitor/buffer Water

RMSD Bond lengths (Å) Bond angles (°) 0.010 1.196

28.7 28.8 28.1 30.8

2,994 2,755 9 226

41.2–2.01 21,081 18.6/22.3

41.2–2.01 (2.06–2.01) 0.117 (0.816) 0.032 (0.226) 0.999 (0.869) 24.6 (3.5) 99.6 (95.2) 14.4 (13.5)

69.4, 82.4, 114.6 90, 90, 90

5HY1 BAH with CA I222 1

0.017 1.729

25.2 25.0 28.1 30.2

6,132 5,470 26 617

41.5–1.83 62,328 21.1/25.2

45.7–1.83 (1.87–1.83) 0.130 (0.907) 0.026 (0.200) 0.999 (0.832) 22.2 (3.5) 98.1 (81.0) 26.6 (20.5)

83.4, 83.4, 216.2 90, 90, 90

5HXU BAH P41212 2

0.004 0.713

31.4 32.3 33.1 22.7

11,036 10,843 26 163

45.3–2.36 61,605 21.5/24.8

45.3–2.36 (2.42–2.36) 0.184 (0.746) 0.072 (0.295) 0.991 (0.741) 8.0 (2.6) 99.9 (98.6) 7.4 (7.3)

81.6, 83.7, 215.1 90, 90, 90

5HXZ BAH P212121 4

0.015 1.682

26.7 27.6 NA 17.1

10,739 10,377 NA 350

41.5–2.40 61,420 19.4/23.6

46.3–2.40 (2.46–2.40) 0.144 (0.934) 0.085 (0.557) 0.989 (0.603) 24.0 (3.8) 97.5 (83.1) 3.9 (3.8)

73.6, 85.7, 87.2 96.5, 114.9, 111.8

5HY0 CyAH P1 4

0.010 1.259

54.4 55.5 NA 33.6

4,783 4,752 NA 31

41.5–2.60 26,209 20.2/24.1

46.6–2.60 (2.72–2.60) 0.126 (1.023) 0.049 (0.404) 0.997 (0.766) 14.0 (2.1) 99.9 (99.1) 7.5 (7.4)

93.3, 72.8, 133.1 90, 92.2, 90

5HY2 CyAH I2 2

0.013 1.522

53.9 55.6 NA 32.0

19,841 19,585 NA 256

46.0–2.56 90,462 22.9/26.3

46.7–2.56 (2.60–2.56) 0.124 (1.181) 0.048 (0.457) 0.998 (0.631) 12.5 (1.8) 99.5 (98.9) 7.5 (7.6)

92.8, 105.8, 299.1 90, 90, 90

5HY4 CyAH P212121 8


not applicable. Values in parentheses are statistics for the high-resolution bin. asymmetric unit; CC1/2, correlation coefficient when one-half of the data are compared to the other half; I/␴I, signal-to-noise ratio; Rmerge and Rpim, quality factors for data; Rwork and Rfree, quality factors for the model/structure.


5HWE BAH I222 2

Value(s) for PDB entrya:

Parameterb Protein Space group No. in AS

TABLE 2 X-ray statistics Peat et al. Applied and Environmental Microbiology

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FIG 4 Active sites of CAH (left), BAH (middle), and the Frankia CyAH (right). Amino acid residues conserved between the three enzymes are shown in green, and those not conserved are shown in blue. The main chain has been hidden for clarity for all amino acids except glycine.

The 3-fold symmetric Ser-Lys pairs are conserved in each active site, with the first pair, Ser83-Lys40 (CAH numbering), very well aligned (C␣ to C␣ distances of 0.6 to 0.9 Å) and maintaining the same rotamers/orientation among the three enzymes. The second most similar pair is Ser343-Lys295 (CAH), which has the same basic orientation in all three cases and differs in C␣ positions by 0.7 to 1.4 Å. The Ser232-Lys162 pair varies the most in rotamer orientation and varies in distance between C␣ atoms by 0.7 to 1.3 Å. Of the three Arg residues found sharing 3-fold symmetry in the CAH structure, only one (Arg52) is conserved across the other two enzymes (Arg53 in BAH and Arg66 in the Frankia CyAH). Arg66 in the Frankia enzyme is found in two orientations, while only one rotamer is found in the other two enzymes for this residue. Arg324 in CAH is found as Lys328/Lys346 (BAH/Frankia CyAH), and Arg194 is found as Asn194/Asn213 (BAH/Frankia CyAH). In all three enzymes, the specificity determinants form a single surface in the active site. In CAH, this surface is identical to the two other surfaces of the active site, engendering 3-fold rotational symmetry that matches that of its substrate. In BAH and the Frankia CyAH, this surface is nonidentical to the other two surfaces, and the active sites of these enzymes consequently have 2-fold mirror symmetry (Fig. 4). The additional active-site residue in the Frankia CyAH enzyme (Glu368) is also found on this face of the active site, which suggests that it also has a role in determining substrate specificity. Although analysis by Soong et al. (8) suggested that BAH was dependent upon Zn2⫹, there was no indication of zinc in the structure: synchrotron X-ray fluorescence scans failed to detect a peak corresponding to zinc, and there was no indication of an atom with sufficient electron density in the resulting high-resolution structure. BAH does contain a metal in a position equivalent to the cation in CAH, but the electron density identifies this cation as either Mg2⫹ or Na⫹ (as it is in CAH) and not zinc. The structural metal found in both CAH and BAH is absent from the Frankia CyAH. The electron density seen for residues 371 to 376 in the Frankia CyAH, the turn where the metal is found in both CAH and BAH, is weak or missing (in one protomer), and we hypothesize that the metal provides stability to this part of the structure for the CAH and BAH enzymes. Indeed, the average B-factor for the whole Frankia CyAH structure is about 27 Å2, whereas it is ⬃80 Å2 for this set of residues. Conversely, the B-factors of residues involved in this turn for CAH and BAH are lower than the average for those structures. There is a conserved glycine-proline pair (Gly350 Pro351 in CAH, Gly354 Pro355 in BAH, and Gly374 Pro375 in the CyAH from Frankia) between the three enzymes that makes a tight turn structure which chelates the metal in the cases of CAH and BAH. In the Frankia CyAH, we find a second proline, 376, which distorts the loop from the geometry seen in the other enzymes, and the one chelating residue outside the turn in May 2017 Volume 83 Issue 9 e03365-16

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FIG 5 Superposition of the CAH, BAH, and Frankia CyAH metal-binding loops. CAH is in green, BAH in cyan, and the Frankia CyAH in magenta. The bound metal is shown as a gray or gold sphere.

CAH and BAH (Glu297/301) is not present in the CyAH from Frankia (Val321) (Fig. 5). It is noteworthy that this Glu297/301 is just two residues removed from the conserved lysine in the third Ser-Lys pair of the active site, and we previously suggested that the structural metal of CAH was important in stabilizing this component of the active site (1). The conserved metal binding residues are found in both the Pseudomonas CAH as well as in the 4NQ3 structure of CAH from A. caulinodans (2). The Frankia CyAH also lacks the metal-coordinating residues Ala347 and Gln349. Although the metal-binding residues are conserved throughout most of the CyAH family, several residues are not conserved in a small subset of CyAHs (Fig. 2): Micromonas sp. strain RC299, Bacillus alcalophilus ATC 27647, Bacillus cellulolyticus, Sulfobacillus acidophilus DSM 10332, A. caulinodans ORS571 (class V), Rhodococcus sp. strain Mel, and Gordonia sp. strain KTR8. The consequences of the amino acid substitutions in these sequences with respect to metal binding are currently unknown; however, it is interesting that two of the four class I and all five of the class V CyAHs lack the canonical metal-binding motif. An unusual vicinal cysteine motif in the Frankia CyAH. One of the notable differences in the structure of CyAH from Frankia compared with the structures of CAH and BAH is the presence of a pair of vicinal cysteines (303 and 304) (Fig. 6) in a loop. This loop is close to the same loop in a neighboring monomer, and the vicinal cysteines of one monomer each make a disulfide bond with the corresponding residue in the neighboring monomer. The corresponding loop in the CAH enzyme is shorter and does not make contact in this dimer interface, whereas the BAH enzyme has a similar loop

FIG 6 Vicinal disulfide in the Frankia CyAH. The disulfide forms between two monomers in the Frankia CyAH tetramer, such that each tetramer contains two sets of vicinal disulfides. The two monomers are shown in green and yellow, and the disulfides are shown as sticks. May 2017 Volume 83 Issue 9 e03365-16

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FIG 7 SDS-PAGE of the reduction of oxidized Frankia CyAH with dithiothreitol (DTT). The Frankia CyAH was oxidized with 2 mM hydrogen peroxide (ox) before buffer exchange to remove the peroxide and reduction with 1 to 100 mM DTT. The Frankia CyAH monomer is ⬃40 kDa, and the dimer is ⬃80 kDa.

structure and does make contact at this interface, although the contribution of this loop to the stability of the BAH dimer interface is less than that of the covalent bonds formed by the double disulfide bond found in the Frankia CyAH interface. The disulfides could be reversibly reduced (Fig. 7) with dithiothreitol and reoxidized with hydrogen peroxide. Differential scanning fluorimetry (DSF) analysis showed a significant difference (⬃10°C) in the stability of oxidized Frankia CyAH compared with the reduced form, suggesting that vicinal disulfide bonds stabilize the enzyme. Reversible disulfide bonds are found in other proteins and have been best studied in redox-sensitive transcriptional regulators (27–30) and redox-dependent regulation of enzyme activity (31, 32). Vicinal cysteines are also present in the same loop in the Micromonas sp. strain RC299 CyAH and TrzD from Pseudomonas sp. strain NRRLB-12227. Moreover, the CyAHs of Bradyrhizobium WSM1253, Gluconacetobacter sp. strain SXCC-1, Sulfobacillus acidophilus DSM 10332, Acidithiobacillus caldus SM1, and Nocardioides sp. strain JS614 (Fig. 2) possess a single cysteine in the same loop, which suggests that a single disulfide also can be used to stabilize the structure (and potentially regulate enzyme function). However, there appears to be no obvious phylogenetic conservation of this disulfidecontaining loop, suggesting that it has evolved several times or that it is an ancient motif that has been lost in the majority of CyAH sequences. Without an identified catalytic activity, it was not possible to ascertain the effect of the disulfide on the activity of the Frankia CyAH. Conclusions. The biochemical function of the CyAH family appears to correlate closely with their phylogenetic distribution, as previously predicted. Surprisingly, the CyAH enzymes surveyed in this study revealed the first BAH that has been shown to have low levels of promiscuous CAH activity, supporting the hypothesis that CAH activity evolved in an ancestral enzyme that may have had a physiological role as a BAH. Interestingly, the presence or absence of a canonical metal-binding motif also appears to be correlated with the phylogeny of the CyAHs, with half of class I and all five class V enzymes partially lacking this motif. We have also solved the structures of two CyAHs that are phylogenetically and biochemically distinct from the CAH structures previously obtained. The new structures reveal that the Toblerone fold is conserved within this family of proteins, as is the quaternary structure (i.e., a tetramer). Although there is considerable conservation of the active sites of the three enzymes, one of the three active-site surfaces varies in composition in a manner that corresponds to the phylogenetic distribution of the CyAHs and which appears to correspond to substrate specificity of the enzyme. Two major structural differences within the family appear to be the presence of a structural metal, found in CAH and BAH but not the Frankia CyAH, and the presence of unusual vicinal disulfides in the Frankia CyAH, which are absent from CAH and BAH. The functions of the metal and disulfides have yet to be fully elucidated; however, both contribute to the stabilities of their respective proteins. May 2017 Volume 83 Issue 9 e03365-16

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Unfortunately, we were unable to identify the substrate of the Frankia CyAH, although we have shown that it does not possess CAH or BAH activities. This is the second CyAH for which no activity could be identified, the other being another class V CyAH from A. caulinodans ORS571 (4). This suggests that the CyAH family of enzymes is more functionally diverse than reported to date (i.e., just CAH and BAH activities). MATERIALS AND METHODS DNA methods. Synthetic genes encoding proteins from the six classes from the phylogeny predicted in Peat et al. (1) were purchased from GeneArt (Life Technologies, Regensburg, Germany). The accession numbers of proteins encoded by those genes, along with the source organism and predicted functionality, are the following: ACH83170 (Acidithiobacillus ferrooxidans ATCC 53993, no predicted functionality), WP_013487839 (Bacillus cellulosilyticus DSM 2522, no predicted functionality), CCD93500.1 (Bradyrhizobium sp. strain ORS 375, CAH), WP_041260875.1 (Frankia sp. strain EuI1c, no predicted functionality), YP_430955 (Moorella thermoacetica ATCC 39073, CAH), ABL81019.1 (Nocardioides sp. strain JS614, BAH), CDM41754.1 (Pseudomonas pseudoalcaligenes CECT 5344, CAH), P0A3V4 (Pseudomonas sp. strain NRRLB-12227, CAH), AEA25384.1 (Pseudonocardia dioxanivorans CB1190, CAH), ACS61202.1 (Rhizobium leguminosarum bv. trifolii WSM1325, CAH), AEW05774.1 (Sulfobacillus acidophilus DSM 10332, no predicted functionality), CAC86669 (Rhodococcus erythropolis, BAH), and ABL83767.1 (Nocardioides sp. strain JS614, BAH). These genes were provided as NdeI/BamHI inserts in the pMA-RQ vector of GeneArt with ampicillin resistance. The genes were cloned into pETcc2 expression vectors (described elsewhere [1]) using T4 DNA ligase (New England BioLabs, Ipswich, MA). An in-frame N-terminal extension that included a hexa-his tag and thrombin cleavage site (MGSSHHHHHHSSGLVPRGSH) was added to each enzyme sequence as a result of the subcloning into this vector. Protein expression and purification. Chemically competent Escherichia coli BL21(DE3) (Invitrogen) was used for protein expression. These cells were transformed with plasmids (pETcc2) harboring the desired gene and grown in Miller’s LB broth (33) or on Miller’s LB broth with agar (15 mg · ml⫺1 agar; Merck), and samples were then incubated at 37°C overnight. A single colony from these plates was used to inoculate the desired volume of LB medium and grown overnight at 37°C while being shaken at 200 rpm. All of the growth media used for growing cells was supplemented with 100 ␮g · ml⫺1 ampicillin (Sigma-Aldrich, MO, USA). The soluble expression of all of the CAH/BAH homologues (exceptions noted below) was achieved at 30°C using the following protocol. A starter culture was prepared by inoculating LB medium with a single colony of transformed BL21(DE3) cells and shaking (200 rpm) at 30°C overnight. A 1:20 dilution of the overnight starter culture was used to inoculate 1 liter of Miller’s LB broth in 2-liter shaker flasks, followed by incubation at 30°C with shaking at 200 rpm until an optical density at 600 nm of 0.6 to 0.8 was obtained. Protein expression was induced by the addition of 200 ␮M isopropyl-beta-Dthiogalactopyranoside (IPTG; Astral, NSW, Australia), followed by overnight incubation at 30°C with shaking at 200 rpm. The homologues from Pseudomonas pseudoalcaligenes CECT 5344 and Moorella thermoacetica were expressed as soluble proteins by inducing at 16°C and 28°C, respectively. The following homologues failed to express soluble proteins: Bacillus cellulolyticus DSM 2552, Sulfobacillus acidophilus DSM 10332, and Acidithiobacillus ferrooxidans ATCC 53993. Cells from the induced cultures were harvested by centrifugation at 4,000 ⫻ g for 10 min using an Avanti J-E centrifuge (Beckman Coulter, Indianapolis, IN). The cells expressing barbiturase from Rhodococcus erythropolis were resuspended in 15 ml of lysis buffer (20 mM potassium phosphate, pH 7.5, 1 mM dithiothreitol [DTT], 10% ethylene glycol) per liter of harvested culture. The cells expressing all other homologues were resuspended in 50 mM HEPES, 100 mM NaCl buffer at pH 7.5. The cells were lysed by three passages through a Microfluidics M-110P homogenizer (Microfluidics, MA, USA) at 137,900 kPa. The cellular debris was removed by centrifugation at 21,000 ⫻ g for 15 min using an Avanti J-E centrifuge. Enzyme purification was carried out by using metal affinity chromatography. Soluble cell extract was loaded onto a 5-ml nickel-nitrilotriacetic acid (Ni-NTA) column (Qiagen, Venlo, Netherlands), and the column was washed with five column volumes of 50 mM imidazole in the lysis buffer followed by another six-column-volume wash with 65 mM imidazole in the same buffer. The bound enzyme was finally eluted with six column volumes of 250 mM imidazole in lysis buffer. The purified enzymes were concentrated in an Amicon Ultra-15 centrifugal filter unit with an Ultracel-10 membrane (Millipore, Carrigtwohill, Ireland) and snap-frozen in liquid nitrogen in 50-␮l aliquots. The final purity of all of these homologues and mutants was estimated to be 98% from a Coomassie-stained SDS-PAGE gel. Protein crystallization and structure solution. (i) BAH (Rhodococcus erythropolis). The BAH protein was concentrated to 15.5 mg · ml⫺1 in 20 mM potassium phosphate, pH 7.5, 1 mM DTT, 10% ethylene glycol and snap-frozen in 50-␮l aliquots. This protein was used to run a thermal melt analysis to identify a formulation more appropriate for crystallization trials. A standard buffer/salt analysis identified the formulation 50 mM N-(2-acetamido)iminodiacetic acid (ADA), pH 6.5, 50 mM NaCl as a possible alternative, as it showed a melting point (Tm) increase from 42.6°C (phosphate-ethylene glycol formulation) to 51.2°C (ADA-NaCl formulation). Protein was dialyzed into the ADA-NaCl formulation overnight and set up in two 96-condition crystallization screens, JCSG⫹_C3 and PS_gradient, and incubated at 20°C (a detailed description of these screens is available at http://c6.csiro.au). Droplets consisted of 150 nl protein at 15 mg/ml and a 150-nl reservoir, equilibrated in SD2 crystallization plates (Molecular Dimensions, United Kingdom) against 50 ␮l of reservoir solution. The second subwell on the plate was set up with protein treated in situ with thrombin, where 100 ␮l protein was added to 10 ␮g of lyophilized thrombin before setting up in crystallization trials. Crystals started appearing overnight and May 2017 Volume 83 Issue 9 e03365-16

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grew to full size within days, particularly under salt (ammonium sulfate or sodium malonate)-containing conditions. Crystals of the malonate-grown crystals, while visually perfect, diffracted poorly (to about 5 Å) at the Australian Synchrotron (AS) MX-1 beamline. Ammonium sulfate-grown crystals were used to produce microseeds, and a matrix microseeding experiment was performed (JCSG⫹_C3, c3_2, and c3_6 screens), using both the tagged and thrombin-treated protein in ADA-NaCl. A crystal grown from 2.45 M ammonium sulfate, 10% glycerol was cryoprotected by the addition of a further 10% glycerol and flash-cooled in liquid nitrogen. Data were collected at the MX-2 microfocus beamline of the Australian Synchrotron, yielding a 99.8% complete data set to 1.71 Å. The crystal displayed I222 symmetry, with 1 protomer in the asymmetric unit. A second crystal form, P41212, grown in 2.5 M ammonium sulfate with 10% (vol/vol) MMT buffer at pH 9 and cryoprotected with AP/E core 150 base stock (Mobil, Australia), diffracted to 1.83 Å at the MX-1 beamline and had two protomers in the asymmetric unit. A third crystal form was found in 2.5 M ammonium sulfate with 0.1 M bis-Tris chloride buffer at pH 6.5, and the data were indexed in the P212121 space group and diffraction extended to 2.36 Å. A crystal grown in 2.5 M ammonium sulfate, 0.1 M bis-Tris chloride, pH 6.5, was soaked with cyanuric acid, cryoprotected with AP/E core 150 base stock (Mobil, Australia) and used to collect a data set at the MX-1 beamline which extended to 2.01 Å and had one protomer in the asymmetric unit in space group I222. Structures and structure factors were deposited in the RCSB Protein Data Bank under codes 5HWE (high resolution, I222), 5HY1 (cyanuric acid, I222), 5HXZ (P212121), and 5HXU (P41212). (ii) Frankia CyAH. The CyAH protein was initially concentrated to 4.2 mg/ml in 50 mM HEPES, pH 7.5, 100 mM NaCl, and the standard thermal melt analysis did not identify any buffer system which was more stabilizing (Tm in HEPES-NaCl of 48.8°C). The protein was set up in 4 96-well crystallization screens (shotgun at 8°C and 20°C, PS_gradient and PACT_C3 at 20°C), both as provided, and with an in situ thrombin treatment. Droplets were 150 nl protein with a 150-nl reservoir, set up in SD2 plates against a 50-␮l reservoir. Disk-shaped crystals started appearing within hours under many polyethylene glycol (PEG)-based (with either magnesium or calcium) conditions with the in situ thrombin-treated protein. The protein was diluted down to 3 mg · ml⫺1 and was set up with microseeding in optimization experiments, including fine screening and additive screening. Various potential binders were tested in cocrystallization experiments: dCTP, dTTP, atrazine, melamine, barbituric acid, and cyanuric acid. Data at a crystal diffraction of 2.40 Å were collected from a P1 crystal grown at 8°C from 2.1 mg · ml⫺1 thrombin-treated protein and the Silver Bullet Bio (SBB; Hampton Research, USA) additive screen (200 nl protein, 100 nl SBB screen condition C1 [2=-deoxyguanosine 5=-monophosphate sodium salt hydrate, ethanolamine, theophylline, isopropyl-1-thio-beta-D-galactopyranoside, oxalacetic acid, HEPES sodium, pH 6.8], 100 nl of base condition, 20%, wt/vol, PEG 3350, 0.2 M magnesium acetate). The crystal was cryoprotected by the addition of extra PEG 3350 to bring the concentration to 25%, and 720 frames of 0.5° oscillations were collected at the MX-2 (microfocus) beamline of the Australian Synchrotron. A second space group, I2, was found to grow in 22% PEG 3350, 86 mM sodium acetate when the protein had been cleaved with thrombin to remove the His tag and treated with atrazine prior to crystallization. This crystal diffracted to 2.6 Å, and 360 frames of 0.5° oscillation (180°) were collected at the MX-2 beamline of the AS. A third space group, P212121, was also found for crystals of Frankia CyAH, which diffracted to 2.56 Å, and 180° of data were collected at the MX-2 beamline of the AS. This crystal was from protein at 4.2 mg · ml⫺1 with a reservoir of 20% PEG 3350 and 0.2 M magnesium acetate. For the two initial structures of BAH and Frankia CyAH, XDS (34) was used to index the data, Aimless (35) was used to determine the space group and scale the data, Phaser (36) was used for molecular replacement (the CAH structure [PDB code 4BVQ] was used as the MR model), Bucaneer (37) was used to automatically build the initial structures, Coot (38) was used to manually rebuild the models, and Refmac (39) was used for refinement. For the subsequent structures, the initial (“native”) structure of either BAH or Frankia CyAH was used for molecular replacement in the new space group. Structures and structure factors for the Frankia CyAH were deposited with RCSB under codes 5HY0 (P1), 5HY2 (I2), and 5HY4 (P212121). Enzyme assays. The enzyme assays in this study were conducted at 28°C and were initiated by the addition of an enzyme, and the reaction rate in linear range was used for obtaining the kinetic constants. CAH activity assays were conducted in 1 mM TAPS [N-[Tris(hydroxymethyl)methyl]3-aminopropanesulfonic acid] (Sigma-Aldrich) buffer at pH 8.5. The substrate was dissolved in 1 mM TAPS buffer and the pH was adjusted to 8.5 prior to its addition to the reaction mixture. CAH activity was measured by monitoring the decrease in absorbance at 214 nm associated with the loss of CA by hydrolysis (1, 4) using a SpectraMax M2 spectrophotometer (Molecular Devices, CA, USA). BAH activity assays were conducted in 20 mM Tris-HCl [2-amino-2-(hydroxymethyl)-1,3-propanediolhydrochloride; Sigma-Aldrich] buffer at pH 8.0. BA stock was prepared in 20 mM Tris-HCl buffer and was adjusted to pH 8.0. The reaction was monitored via the loss of absorbance at 256 nM, resulting from the hydrolysis of BA (4, 8). The enzyme expressed from Frankia sp. strain EuI1c (WP_041260875.1) from class V was predicted not to have BA or CA activities, so it was tested against the following compounds as its potential substrates: BA, CA, uracil, 1-methyluracil, thymine, alloxan, L-dihydroorotic acid, and orotic acid. The enzyme assays with orotic acid as a substrate were conducted at 248 nM under the same conditions as those mentioned for the barbituric acid assay. Oxidation and reduction of the Frankia CyAH. Oxidation of the CyAH from Frankia was by incubation with hydrogen peroxide (2.5 ␮M to 20 mM) for 5 min. The final concentration of the enzyme under these conditions was 1.05 ␮M. Peroxide-oxidized Frankia CyAH was reduced by the addition of dithiothreitol (1 to 100 mM) after buffer exchange into 50 mM HEPES, 100 mM NaCl, pH 7.5. Samples were analyzed by SDS-PAGE (nuPAGE; Life Technologies, USA) under nonreducing conditions. May 2017 Volume 83 Issue 9 e03365-16

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The Frankia CyAH was analyzed for stability under reducing and oxidizing conditions separately by thermal melt analysis. Melt curves for the protein in the presence of either reducing agent (5 mM dithiothreitol) or oxidizing agent (5 mM, hydrogen peroxide) were generated using differential scanning fluorimetry. The samples was tested in a suite of different buffers and pHs in triplicate (40). The assay was performed in a CFX96 reverse transcription-PCR (RT-PCR) machine (Bio-Rad) with 19.6 ␮l of each screening condition, 300 nl protein at 1 mg · ml⫺1, and 300 nl of a 1:10 (aqueous) dilution of SYPRO orange dye (Sigma). Accession number(s). Structures and structure factors were deposited in the RCSB Protein Data Bank under codes 5HWE, 5HY1, 5HXZ, 5HXU, 5HY0, 5HY2, and 5HY4.

ACKNOWLEDGMENTS We thank Robyn Russell and Andrew Warden (CSIRO Land & Water) for their constructive comments during the preparation of the manuscript. We also thank the beamline scientists at the Australian Synchrotron for their help in data collection. Crystals were grown in the CSIRO Collaborative Crystallisation Centre (http://www.csiro.au/C3) at CSIRO Manufacturing.

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High-Resolution X-Ray Structures of Two Functionally Distinct Members of the Cyclic Amide Hydrolase Family of Toblerone Fold Enzymes.

The Toblerone fold was discovered recently when the first structure of the cyclic amide hydrolase, AtzD (a cyanuric acid hydrolase), was elucidated. W...
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