Structural Analysis of the Glycine Oxidase Homologue CmiS2 Reveals

Communication Cite This: Biochemistry 2019, 58, 2706−2709

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Structural Analysis of the Glycine Oxidase Homologue CmiS2 Reveals a Unique Substrate Recognition Mechanism for Formation of a β‑Amino Acid Starter Unit in Cremimycin Biosynthesis Daisuke Kawasaki, Taichi Chisuga, Akimasa Miyanaga,* Fumitaka Kudo, and Tadashi Eguchi* Department of Chemistry, Tokyo Institute of Technology, 2-12-1 O-okayama, Meguro-ku, Tokyo 152-8551, Japan

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polyketide backbone (Figure S1).5−7 Therefore, it has been suggested that these β-amino fatty acids are synthesized via the same mechanism. Mechanistic insights into the catalytic reaction and substrate recognition of CmiS1-type enzymes have been elucidated through structural analysis of the CmiS1 homologue SAV606.4,8 However, the substrate recognition mechanism of CmiS2 remains elusive. The sequence of CmiS2 is similar with that of FADdependent glycine oxidases, including ThiO from Bacillus subtilis.9 ThiO is well studied from a mechanistic and structural point of view9−15 and has been suggested to participate in the early stage of thiamine biosynthesis.16 In the proposed reaction mechanism of ThiO, the pro-S hydrogen of the glycine Cα atom is transferred to FAD as a hydride to form a glyoxyl imine intermediate, which is thought to be used for formation of the thiazole ring prior to hydrolysis.16 It is expected that CmiS2 utilizes a similar reaction mechanism; that is, CmiS2 catalyzes the oxidation of the N-carboxymethyl moiety of CMANA via hydride transfer to FAD, followed by spontaneous hydrolysis to produce ANA and glyoxylic acid (Figure S2). However, it is likely that CmiS2 employs a different substrate recognition mechanism to glycine oxidase, given that the chemical structure of CMANA is considerably different from that of glycine. In this study, we carried out biochemical and structural analyses of CmiS2 to elucidate the substrate recognition mechanism. The stereochemistry at the β-position of the substrate (CMANA) and product (ANA) in the CmiS2 reaction was unclear, because we used the racemic compound as a substrate for CmiS2 in a previous study.3 Therefore, we first determined the stereochemistry of the product by carrying out a large-scale reaction with CmiS1, CmiS2, glycine, and non-2-enoyl Nacetylcysteamine. After methyl esterification of the enzymatic reaction product, the resulting methyl ester was treated with (S)α-methoxy-α-(trifluoromethyl)phenylacetyl chloride [(S)MTPA-Cl] and then analyzed by 1H nuclear magnetic resonance according to Mosher’s method.17 Analysis of the MTPA derivative of racemic ANA revealed distinguishable signals of the (R)- or (S)-ANA-derived diastereomers (Figures S3−S6). The MTPA derivative of (R)-ANA showed signals at 3.69 ppm (attributable to OMe in the methyl nonanoate moiety) and 3.45 ppm (attributable to OMe in MTPA), indicating that the product was an R enantiomer (Figure S6). This suggests that

ABSTRACT: The flavin adenine dinucleotide-dependent oxidase CmiS2 catalyzes the oxidation of N-carboxymethyl-3-aminononanoic acid to produce a 3-aminononanoic acid starter unit for the biosynthesis of cremimycin, a macrolactam polyketide. Although the sequence of CmiS2 is similar with that of the well-characterized glycine oxidase ThiO, the chemical structure of the substrate of CmiS2 is different from that of ThiO substrate glycine. Here, we present the biochemical and structural characterization of CmiS2. Kinetic analysis revealed that CmiS2 has a strong preference for N-carboxymethyl-3-aminononanoic acid over other substrates such as N-carboxymethyl3-aminobutanoic acid and glycine, suggesting that CmiS2 recognizes the nonanoic acid moiety of the substrate as well as the glycine moiety. We determined the crystal structure of CmiS2 in complex with a substrate analogue, namely, S-carboxymethyl-3-thiononanoic acid, which enabled the identification of key amino acid residues involved in substrate recognition. We discovered that Asn49, Arg243, and Arg334 interact with the carboxyl group of the nonanoic acid moiety, while Pro46, Leu52, and Ile335 recognize the alkyl chain of the nonanoic acid moiety via hydrophobic interaction. These residues are highly conserved in CmiS2 homologues involved in the biosynthesis of related macrolactam polyketides but are not conserved in glycine oxidases such as ThiO. These results suggest that CmiS2-type enzymes employ a distinct mechanism of substrate recognition for the synthesis of βamino acids.

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remimycin is a 19-member macrolactam antibiotic that is produced by Streptomyces sp. MJ635-86F5 and contains a 3-aminononanoic acid (ANA) moiety in its polyketide skeleton.1 ANA is synthesized from non-2-enoyl-acyl carrier protein (ACP) thioester that is constructed via the polyketide pathway (Figure 1).2−4 The thioesterase CmiS1 catalyzes the Michael addition of glycine to the β-position of non-2-enoylACP and subsequent hydrolysis of the thioester to Ncarboxymethyl-3-aminononanoic acid (CMANA). The flavin adenine dinucleotide (FAD)-dependent oxidase CmiS2 then catalyzes the oxidation of CMANA to produce ANA and glyoxylic acid. Homologues of cmiS1 and cmiS2 are present in the biosynthetic gene clusters for BE-14106 (becU and becI), ML-499 (mlaU and mlaI), and heronamide (herU and herI), all of which contain a long-chain β-amino fatty acid within their © 2019 American Chemical Society

Received: May 20, 2019 Revised: May 31, 2019 Published: June 2, 2019 2706

DOI: 10.1021/acs.biochem.9b00444 Biochemistry 2019, 58, 2706−2709

Communication

Biochemistry

Figure 1. Illustration of the biosynthetic pathway of 3-aminononanoic acid (ANA) in cremimycin biosynthesis. The ANA unit is colored red. CmiP4, CmiP3, and CmiP2 represent polyketide synthase (ACP, acyl carrier protein; AT, acyltransferase; DH, dehydratase; ER, enoylreductase; KR, ketoreductase; KS, ketosynthase; KSQ, ketosynthase-like decarboxylase).

Figure 2. (A) Overall structures of ligand-free CmiS2 (left) and CmiS2 complexed with S-carboxymethyl-3-thiononanoic acid (CMTNA) (right). The FAD-binding and substrate-binding domains are colored magenta and orange, respectively. Yellow and cyan sticks indicate FAD and CMTNA, respectively. (B) Substrate-binding pocket of the complex. Substrate recognition residues are shown as sticks. An Fo − Fc electron density map contoured at 2.5σ was constructed before incorporation of the CMTNA molecule.

FAD-binding domain (residues 1−85, 148−222, and 309−368) and a substrate-binding domain (residues 86−147 and 223− 308). The CmiS2 molecule contains one noncovalently bound FAD molecule. The adenosine diphosphate-ribosyl moiety is sequestered in the FAD-binding domain, and the reactive isoalloxazine ring is located at the interface between the two domains. The overall structure of CmiS2 is similar to those of ThiO [PDB entry 1NG3; root-mean-square deviation (RMSD), 1.9 Å; sequence identity, 25%] and glycine oxidase18 from Geobacillus kaustophilus (PDB entry 4YSH; RMSD, 1.9 Å; sequence identity, 32%). CmiS2 is also structurally similar to sarcosine oxidase19 from Corynebacterium sp. U-96 (PDB entry 3AD7; RMSD, 2.5 Å; sequence identity, 21%), D-amino acid oxidase (DAAO)20 from Rhodosporidium toruloides (PDB entry 1C0L; RMSD, 2.9 Å; sequence identity, 23%), and the glycine oxidase-like domain (MnmC1) of MnmC21 from Escherichia coli (PDB entry 3PS9; RMSD, 2.5 Å; sequence identity, 19%), all of which belong to the FAD-dependent D-amino acid oxidase family.22 To understand the substrate recognition mechanism, we soaked a CmiS2 crystal in a solution containing CMANA. However, we were unable to observe clear electron density corresponding to CMANA. Therefore, we synthesized a racemic substrate mimic, S-carboxymethyl-3-thiononanoic acid

CmiS1 produces (R)-CMANA via Michael addition of glycine and subsequent hydrolysis, and CmiS2 uses this compound as a substrate. This is consistent with a previous report that stated that SAV606, a CmiS1 homologue, produces (R)-N-carboxymethyl-3-aminobutanoic acid [(R)-CMABA].8 To investigate the substrate specificity of CmiS2, we carried out kinetic analysis using a horseradish peroxidase-coupled assay (Figure S7).10,14 The Km and kcat values for racemic CMANA were 0.169 ± 0.033 mM and 1.40 ± 0.13 s−1, respectively (Table S1). These values are comparable with those of ThiO for glycine (Km = 0.99 mM, and kcat = 1.3 s−1) and sarcosine (Km = 0.22 mM, and kcat = 1.6 s−1).9 We found CmiS2 to have a 130-fold larger Km value (21.7 ± 1.3 mM) and an 8-fold lower kcat value (0.165 ± 0.004 s−1) for racemic CMABA compared with those for CMANA, suggesting that CmiS2 can distinguish alkyl chain length. Furthermore, CmiS2 showed a 14-fold higher Km value (302 ± 29 mM) for sarcosine compared with that for CMABA and showed no detectable activity against glycine, indicating that the substrate specificity of CmiS2 is different from that of ThiO. These results suggest that CmiS2 recognizes the nonanoic acid moiety of the substrate. We determined the crystal structure of CmiS2 [Protein Data Bank (PDB) entry 6J38] at 2.30 Å resolution (Figure 2A and Table S2). The CmiS2 molecule consists of two domains, a 2707

DOI: 10.1021/acs.biochem.9b00444 Biochemistry 2019, 58, 2706−2709

Communication

Biochemistry (CMTNA) (see Figures S8 and S9), which was found to be a competitive inhibitor (Ki = 6.0 ± 0.1 μM) (Figure S10). A CmiS2 crystal was soaked in a solution supplemented with CMTNA, which enabled successful determination of the crystal structure of CmiS2 in complex with CMTNA (PDB entry 6J39) at 2.45 Å resolution (Figure 2A and Table S2). Electron density corresponding to (R)-CMTNA was clearly observable in the vicinity of the isoalloxazine ring of FAD (Figure 2B). Binding of CMTNA led to an ∼2.5 Å movement of the η-helix 1 (η1; Asn49−Leu52) backbone toward the substrate-binding pocket (Figure S11), although no significant conformational change in the polypeptide backbone was observed in other regions (RMSD of 0.34 Å for the Cα atom of chain A). In the ligandfree structure, the side chain of Glu50 is oriented away from the substrate-binding pocket and forms a salt bridge with Arg243; in the CMTNA-bound structure, the side chain of Glu50 rotates approximately 90° toward the substrate-binding pocket and forms a salt bridge with Arg334. The η1 region might function as an active-site lid, switching between open and closed conformations to control substrate binding and product release in a manner similar to that of other FAD-dependent enzymes such as D-amino acid oxidase.23 The substrate-binding pocket is comprised mainly of nine residues: Pro46, Asn49, Leu52, Arg243, Tyr252, Tyr267, Arg308, Arg334, and Ile335 (Figure 2B). The methylene carbon of the S-carboxymethyl moiety is positioned 3.2 Å from the N5 atom of FAD (Figure S12). The carboxyl group of the Scarboxymethyl moiety of CMTNA forms hydrogen bonds with Tyr252 (2.5 Å) and Tyr267 (3.2 Å) and a bidentate salt bridge with Arg308 (2.4 and 2.6 Å). The carboxyl group of the nonanoic acid moiety interacts with Arg243 (3.2 Å), Arg334 (2.5 Å), and Asn49 (2.9 Å). In addition, the alkyl chain of the nonanoic acid moiety is positioned in the hydrophobic pocket that is formed by Pro46, Leu52, and Ile335. The presence of this nonanoic acid moiety seems to be important for the correct placement of the glycine moiety of the substrate, which could explain the lower activity of CmiS2 toward CMABA and sarcosine (Table S1). Structural comparison with the ThiO−N-acetylglycine complex (PDB entry 1NG3) revealed that the S-carboxymethyl moiety of CMTNA occupies almost the same position in the CmiS2 structure as the glycine moiety of N-acetylglycine in the ThiO structure (Figure 3). Tyr252, Arg308, and Arg334 of CmiS2 are conserved in ThiO (Tyr246, Arg302, and Arg329, respectively). Furthermore, Tyr246 and Arg302 of ThiO are located close to the carboxyl group of the glycine moiety of Nacetylglycine. However, the side chain of Arg329 of ThiO interacts with the carboxyl group of the glycine moiety in ThiO, whereas Arg334 of CmiS2 forms a salt bridge with the carboxyl group of the nonanoic acid moiety. This difference might be due to the presence of Tyr267 in CmiS2, which is equivalent to Met261 in ThiO. In CmiS2, Tyr267 interacts with the Scarboxymethyl moiety of CMTNA that prevents Arg334 from accessing the S-carboxymethyl moiety (Figure 3). Residues Pro46, Asn49, Leu52, Arg243, and Ile335 of CmiS2 are not conserved in ThiO (Figure S13) and are responsible for selective recognition of the nonanoic acid moiety by CmiS2. The η1 position of CmiS2 shows an ∼5 Å displacement compared with the corresponding η1 of ThiO, which contributes to the relatively expanded substrate-binding pocket of CmiS2 (Figure 3). Movement of η1 enables the alkyl chain of the nonanoic acid moiety to be accommodated in the substrate-binding pocket of CmiS2. Structural comparison with the DAAO−D-alanine

Figure 3. Structural comparison of the CmiS2 complex (magenta and orange) and the ThiO−N-acetylglycine complex (dark gray). FAD molecules in the CmiS2 and ThiO structures are shown as yellow and light gray sticks, respectively. Ligand molecules in the CmiS2 and ThiO structures are shown as cyan and black sticks, respectively.

complex (PDB entry 1C0P) showed that the orientation of the S-carboxymethyl moiety of CMTNA is also similar to that of D-alanine in the DAAO structure (Figure S14). Tyr252, Tyr267, and Arg308 of CmiS2 are conserved in DAAO (Tyr223, Tyr238, and Arg285, respectively), although Pro46, Asn49, Leu52, Arg243, Arg334, and Ile335 of CmiS2 are not conserved. Tyr223, Tyr238, and Arg285 similarly interact with the carboxyl group of D-alanine in the DAAO structure.20 However, the position of Tyr223 of DAAO shows an ∼2 Å displacement compared with the corresponding Tyr252 of CmiS2. This displacement appears to provide sufficient space to accommodate the methyl group of D-alanine in the substrate-binding pocket of DAAO. Comparison of the sequence of CmiS2 with those of the CmiS2 homologues BecI, MlaI, and HerI (levels of sequence identity of 57%, 54%, and 57%, respectively) as well as with those of glycine oxidases (Figure S13) revealed that Tyr252 and Arg308 of CmiS2 are completely conserved in all enzymes. These residues are important for recognition of the carboxyl group of glycine. Residues involved in recognition of the nonanoic acid moiety (Pro46, Asn49, Leu52, Arg243, and Ile335) are almost conserved among CmiS2 homologues but not conserved in glycine oxidases. This implies that all CmiS2type enzymes recognize β-substituted long-chain fatty acids in a similar manner. In conclusion, we have elucidated the structural basis of substrate specificity of the novel FAD-dependent oxidase CmiS2 in cremimycin biosynthesis. Although CmiS2 shows sequence and structural similarities to glycine oxidases such as ThiO, the substrate specificity of CmiS2 is different from that of glycine oxidases. Furthermore, structural analysis showed that key residues involved in the recognition of fatty acid moieties of βamino acid substrates are conserved only among this class of macrolactam biosynthetic enzymes. 2708

DOI: 10.1021/acs.biochem.9b00444 Biochemistry 2019, 58, 2706−2709

Communication

Biochemistry

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Biosynthesis of macrolactam BE-14106 involves two distinct PKS systems and amino acid processing enzymes for generation of the aminoacyl starter unit. Chem. Biol. 16, 1109−1121. (6) Jørgensen, H., Degnes, K. F., Dikiy, A., Fjærvik, E., Klinkenberg, G., and Zotchev, S. B. (2010) Insights into the evolution of macrolactam biosynthesis through cloning and comparative analysis of the biosynthetic gene cluster for a novel macrocyclic lactam, ML-449. Appl. Environ. Microbiol. 76, 283−293. (7) Zhu, Y., Zhang, W., Chen, Y., Yuan, C., Zhang, H., Zhang, G., Ma, L., Zhang, Q., Tian, X., Zhang, S., and Zhang, C. (2015) Characterization of heronamide biosynthesis reveals a tailoring hydroxylase and indicates migrated double bonds. ChemBioChem 16, 2086−2093. (8) Chisuga, T., Miyanaga, A., Kudo, F., and Eguchi, T. (2017) Structural analysis of the dual-function thioesterase SAV606 unravels the mechanism of Michael addition of glycine to an α,β-unsaturated thioester. J. Biol. Chem. 292, 10926−10937. (9) Nishiya, Y., and Imanaka, T. (1998) Purification and characterization of a novel glycine oxidase from Bacillus subtilis. FEBS Lett. 438, 263−266. (10) Job, V., Marcone, G. L., Pilone, M. S., and Pollegioni, L. (2002) Glycine oxidase from Bacillus subtilis. J. Biol. Chem. 277, 6985−6993. (11) Molla, G., Motteran, L., Job, V., Pilone, M. S., and Pollegioni, L. (2003) Kinetic mechanisms of glycine oxidase from Bacillus subtilis. Eur. J. Biochem. 270, 1474−1482. (12) Pedotti, M., Ghisla, S., Motteran, L., Molla, G., and Pollegioni, L. (2009) Catalytic and redox properties of glycine oxidase from Bacillus subtilis. Biochimie 91, 604−612. (13) Jamil, F., Afza Gardner, Q., Bashir, Q., Rashid, N., and Akhtar, M. (2010) Mechanistic and stereochemical studies of glycine oxidase from Bacillus subtilis strain R5. Biochemistry 49, 7377−7383. (14) Settembre, E. C., Dorrestein, P. C., Park, J.-H., Augustine, A. M., Begley, T. P., and Ealick, S. E. (2003) Structural and mechanistic studies on ThiO, a glycine oxidase essential for thiamin biosynthesis in Bacillus subtilis. Biochemistry 42, 2971−2981. (15) Mörtl, M., Diederichs, K., Welte, W., Molla, G., Motteran, L., Andriolo, G., Pilone, M. S., and Pollegioni, L. (2004) Structure-function correlation in glycine oxidase from Bacillus subtilis. J. Biol. Chem. 279, 29718−29727. (16) Jurgenson, C. T., Begley, T. P., and Ealick, S. E. (2009) The structural and biochemical foundations of thiamin biosynthesis. Annu. Rev. Biochem. 78, 569−603. (17) Dale, J. A., and Mosher, H. S. (1973) Nuclear magnetic resonance enantiomer reagents. Configurational correlations via nuclear magnetic resonance chemical shifts of diastereomeric mandelate, O-methylmandelate, and m-Methoxy—trifluoromethylphenylacetate (MTPA) Esters. J. Am. Chem. Soc. 95, 512−519. (18) Martínez-Martínez, I., Navarro-Fernández, J., García-Carmona, F., Takami, H., and Sánchez-Ferrer, Á . (2008) Characterization and structural modeling of a novel thermostable glycine oxidase from Geobacillus kaustophilus HTA426. Proteins: Struct., Funct., Genet. 70, 1429−1441. (19) Moriguchi, T., Ida, K., Hikima, T., Ueno, G., Yamamoto, M., and Suzuki, H. (2010) Channeling and conformational changes in the heterotetrameric sarcosine oxidase from Corynebacterium sp. U-96. J. Biochem. 148, 491−505. (20) Umhau, S., Pollegioni, L., Molla, G., Diederichs, K., Welte, W., Pilone, M. S., and Ghisla, S. (2000) The x-ray structure of D-amino acid oxidase at very high resolution identifies the chemical mechanism of flavin-dependent substrate dehydrogenation. Proc. Natl. Acad. Sci. U. S. A. 97, 12463−12468. (21) Kim, J., and Almo, S. C. (2013) Structural basis hypermodification of the wobble uridine in tRNA by bifunctional enzyme MnmC. BMC Struct. Biol. 13, 5. (22) Fitzpatrick, P. F. (2010) Oxidation of amines by flavoproteins. Arch. Biochem. Biophys. 493, 13−25. (23) Ball, J., Gannavaram, S., and Gadda, G. (2018) Structural determinants for substrate specificity of flavoenzymes oxidizing Damino acids. Arch. Biochem. Biophys. 660, 87−96.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.9b00444. Experimental details, Figures S1−S14, and Tables S1 and S2 (PDF) Accession Codes

Protein Data Bank entries 6J38 and 6J39.

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Akimasa Miyanaga: 0000-0003-2219-6051 Fumitaka Kudo: 0000-0002-4788-0063 Tadashi Eguchi: 0000-0002-7830-7104 Author Contributions

A.M., F.K., and T.E. designed the research. D.K., T.C., and A.M. performed the experiments. D.K., T.C., A.M., F.K., and T.E. analyzed the data. D.K., A.M., F.K., and T.E. wrote the manuscript. Funding

This work was supported in part by a Ministry of Education, Culture, Sports, Science and Technology Grant-in-Aid for Scientific Research in Innovative Areas (16H06451 to T.E.) and the Japan Society for the Promotion of Science A3 Foresight Program. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was performed with the approval of the Photon Factory Program Advisory Committee (Proposal 2016G624). ABBREVIATIONS ACP, acyl carrier protein; ANA, 3-aminononanoic acid; CMABA, N-carboxymethyl-3-aminobutanoic acid; CMANA, N-carboxymethyl-3-aminononanoic acid; CMTNA, S-carboxymethyl-3-thiononanoic acid; DAAO, D-amino acid oxidase; FAD, flavin adenine dinucleotide; MTPA, α-methoxy-α(trifluoromethyl)phenylacetate; RMSD, root-mean-square deviation.

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REFERENCES

(1) Igarashi, M., Tsuchida, T., Kinoshita, N., Kamijima, M., Sawa, R., Sawa, T., Naganawa, H., Hamada, M., Takeuchi, T., Yamazaki, K., and Ishizuka, M. (1998) Cremimycin, a novel 19-membered macrocyclic lactam antibiotic, from Streptomyces sp. J. Antibiot. 51, 123−129. (2) Amagai, K., Kudo, F., and Eguchi, T. (2011) Biosynthetic pathway of macrolactam polyketide antibiotic cremimycin. Tetrahedron 67, 8559−8563. (3) Amagai, K., Takaku, R., Kudo, F., and Eguchi, T. (2013) A unique amino transfer mechanism for constructing the o-amino fatty acid starter unit in the biosynthesis of the macrolactam antibiotic cremimycin. ChemBioChem 14, 1998−2006. (4) Miyanaga, A. (2019) Michael additions in polyketide biosynthesis. Nat. Prod. Rep. 36, 531−547. (5) Jørgensen, H., Degnes, K. F., Sletta, H., Fjærvik, E., Dikiy, A., Herfindal, L., Bruheim, P., Klinkenberg, G., Bredholt, H., Nygård, G., Døskeland, S. O., Ellingsen, T. E., and Zotchev, S. B. (2009) 2709

DOI: 10.1021/acs.biochem.9b00444 Biochemistry 2019, 58, 2706−2709