Aminoacyl-tRNA-Utilizing Enzymes in Natural Product Biosynthesis

Review pubs.acs.org/CR

Aminoacyl-tRNA-Utilizing Enzymes in Natural Product Biosynthesis Mireille Moutiez,† Pascal Belin,† and Muriel Gondry*

Chem. Rev. 2017.117:5578-5618. Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 11/22/18. For personal use only.

Institute for Integrative Biology of the Cell (I2BC), CEA, CNRS, Univ. Paris-Sud, Université Paris-Saclay, 91198, Gif-sur-Yvette Cedex, France ABSTRACT: Aminoacyl-tRNAs were long thought to be involved solely in ribosomedependent protein synthesis and essential primary metabolism processes, such as targeted protein degradation and peptidoglycan synthesis. About 10 years ago, an aminoacyl-tRNA-dependent enzyme involved in the biosynthesis of the antibiotic valanimycin was discovered in a Streptomyces strain. Far from being an isolated case, this discovery has been followed by the description of an increasing number of aminoacyltRNA-dependent enzymes involved in secondary metabolism. This review describes the three groups of aminoacyl-tRNA-dependent enzymes involved in the synthesis of natural products. Each group is characterized by a particular chemical reaction, and its members are predicted to share a specific fold. The three groups are cyclodipeptide synthases involved in diketopiperazine synthesis, LanB-like dehydratases involved in the posttranslational modification of ribosomal peptides, and transferases from various biosynthesis pathways.

CONTENTS 1. Introduction 1.1. The aa-tRNA Scaffold: An Adaptor for Ribosome-Dependent Protein Synthesis 1.2. aa-tRNA: An Unusual Substrate for Diverse Enzymes 2. aa-tRNA-Dependent Synthases: The Cyclodipeptide Synthases (CDPSs) 2.1. CDPSs and Their Cyclodipeptide Products 2.2. 3D-Structure and Catalytic Mechanism 2.2.1. A Common Architecture with a Rossmann-Fold Domain 2.2.2. A Common Sequential Ping-Pong Mechanism 2.3. Molecular Basis of the Interaction between CDPSs and Their Substrates 2.3.1. Specificity Determinants of the Two aatRNA Substrates 2.3.2. The Binding Sites of the Two Substrates 2.3.3. Toward the Prediction of CDPS Specificity 2.4. CDPS-Dependent Pathways 3. aa-tRNA-Dependent Dehydratases in RiPP Synthesis 3.1. Full-Length LanB Dehydratases in Lanthipeptides Biosynthesis 3.1.1. Mechanism and Structure of LanB 3.1.2. LanBs Are Partners of a Multiprotein Complex 3.1.3. To Be or Not To Be Substrate 3.2. Split LanBs in Thiopeptide and Goadsporin Biosynthesis 3.2.1. Characteristics of TpdBs and TpdCs in the Clusters Described © 2017 American Chemical Society

3.2.2. What We Know about the Mechanism of Split LanBs: TpdB and TpdC 3.2.3. What We Know about Specificity 3.3. “Small LanB” Proteins in Biosynthesis Clusters 4. aa-tRNA-Dependent Transferases 4.1. Biosynthesis of the Azoxy Compound Valanimycin: The First Involvement of aa-tRNAs in Secondary Metabolite Biosynthesis To Be Described 4.2. The tRNA-Dependent Biosynthesis of Phosphonopeptides 4.2.1. The Biosynthetic Pathway of Dehydrophos 4.2.2. Putative tRNA-Dependent Biosynthetic Pathways for Other Phosphonopeptides 4.3. Biosynthesis of Uridyl Peptide Antibiotics (UPAs) 4.3.1. The Transferase PacB in the Biosynthesis of PACs 4.3.2. PacB Homologues Encoded in the Biosynthetic Gene Clusters of UPAs 4.4. Biosynthesis of the Streptothricin Analogue BD-12 4.5. Similarities and Differences between aatRNA-Dependent Transferases 5. The Availability of aa-tRNAs for Functions Outside of Translation 6. Conclusion Author Information Corresponding Author

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Special Issue: Unusual Enzymology in Natural Products Synthesis

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Received: August 8, 2016 Published: January 6, 2017 5578

DOI: 10.1021/acs.chemrev.6b00523 Chem. Rev. 2017, 117, 5578−5618

Chemical Reviews ORCID Author Contributions Notes Biographies Acknowledgments References

Review

tRNA part of the molecule has a characteristic cloverleaf secondary structure that folds into an L-shaped tertiary structure6−8 (Figure 1a,b). One end of the L-shaped molecule carries the trinucleotide anticodon that specifically interacts with mRNA codons by base pairing, whereas the other end bears the attachment site for the cognate amino acid. Amino acid attachment is catalyzed by specific aminoacyl-tRNA synthetases (aaRSs) in a two-step reaction.9 The amino acid is first activated with adenosine triphosphate (ATP) by the direct linkage of its carboxyl group to the AMP moiety. The resulting aminoacyladenylate is then transferred to the 2′- or 3′-hydroxyl group of the adenosine at the 3′-end of the tRNA moiety, leading to the formation of an activated aminoacyl−ester bond (Figure 1c). Once aminoacylation is completed, the aa-tRNA is generally captured by elongation factor EF-1A (or EF-Tu), which carries the aa-tRNA to the dedicated ribosome site.10 However, the aatRNA can evade the translation machinery and serve as a substrate for other cellular processes.

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1. INTRODUCTION The amazing chemical diversity of natural products (NPs) is reflected in the huge array of catalytic enzymes involved in their biosynthesis. Some of these enzymes catalyze unusual reactions or have unusual properties (see this thematic issue). In this context, the number of enzyme families shown to use aminoacyltransfer ribonucleic acids (aa-tRNAs) as substrates for the catalysis of diverse chemical reactions via different mechanisms is steadily increasing. This review will begin with a brief presentation of aa-tRNA substrates and the enzymes that use them for diverse cellular functions, and will then describe the aatRNA-dependent enzymes involved in NP biosynthesis.

1.2. aa-tRNA: An Unusual Substrate for Diverse Enzymes

aa-tRNA-dependent enzymes are involved in various cellular processes in primary and secondary metabolism (for reviews, see refs 5 and 12−18) (Figure 2). Several families of aminoacyl transferases catalyze the transfer of the aminoacyl moieties of aatRNAs to diverse acceptors for essential cellular functions. These enzymes include the Fem transferases, which catalyze the formation of amide and peptide bonds during peptidoglycan biosynthesis,19−21 the aminoacyl-phosphatidylglycerol synthases (aaPGSs), which catalyze the addition of a single amino acid, Ala or Lys, to the polar headgroup of phosphatidylglycerol in the bacterial inner membrane during adaptation to changing environmental conditions,21−24 and the aminoacyl protein transferases, which transfer a phenylalanyl, leucyl, or arginyl

1.1. The aa-tRNA Scaffold: An Adaptor for Ribosome-Dependent Protein Synthesis

aa-tRNAs are ubiquitous molecules originally identified as the compounds responsible for delivering amino acids for the mRNA-guided synthesis of proteins at the ribosome. They function as adaptors between the mRNA codons and the growing polypeptide chain.1−3 They are composed of a tRNA of about 80 nucleotides in length attached to an aminoacyl moiety consisting of a single amino acid. Several bases constituting the tRNA undergo species-specific posttranscriptional modifications that are important for folding, stability, translational efficiency, and fidelity, and for diverse regulatory processes4,5 (Figure 1a). The

Figure 1. Structures of the tRNAPhe from Escherichia coli and the phenylalanyl-tRNAPhe linkage. (a) Cloverleaf secondary structure of the tRNAPhe. The main parts of the tRNA are highlighted in different colors: the acceptor arm is shown in blue, the D arm in orange, the anticodon arm in dark red, the variable arm in magenta, and the TΨC arm in green. Mature tRNAPhe is 76 nucleotides long (nucleotides 1, 72, and 76 are indicated), and it contains 10 modified nucleosides: 4-thiouridine (s4U), dihydrouridine (D), pseudouridine (Ψ), 2-methylthio-N6-isopentenyladenosine (mA), 7-methylguanosine (mG), 3-(3-amino-3-carboxypropyl)uridine (acpU), and 5-methyluridine (T). The 5′- and 3′-ends of the molecule are indicated, and the 3′-end bears the attachment site for the cognate phenylalanine. (b) Cartoon representation of the L-shaped tertiary structure of the tRNAPhe (PDB ID, 3L0U). The tRNAPhe molecule is shown as a semitransparent molecular surface, with its main parts shown with the same colors than those used in panel a. The structure was solved with unmodified tRNAPhe, but the overall fold of the tRNA is essentially similar to that of mature tRNA.11 The nucleotides 74−76 were omitted from the model due to lack of electron density. (c) The attachment of the phenylalanyl moiety to the tRNA. The carboxyl end of the amino acid forms an ester bond to the 2′- or 3′-hydroxyl group of the adenosine 76. 5579


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Figure 2. Diversity of aa-tRNA-utilizing enzymes. The aa-tRNA pool is primarily used for ribosomal protein synthesis. Some aa-tRNAs are hijacked for use as substrates by diverse enzymes (schematically represented by external circles) involved in various cellular processes (indicated in boxes). Blue circles indicate enzymes belonging to the GNAT protein superfamily.41,42 tRNA-dependent modifying enzymes are detailed in ref 15.

residue to the N-terminus of a protein for targeted degradation.25−28 The enzymes of the glutamyl-tRNA reductase family reduce the aminoacyl moiety of Glu-tRNAGlu to form glutamate 1-semialdehyde, the first precursor in the biosynthesis of tetrapyrroles such as hemes and chlorophylls.29−31 Enzymes from other families catalyze modification of the aminoacyl moiety of aa-tRNAs (i.e., while this moiety is still attached to tRNAs) to generate aa-tRNAs loaded with asparagine, glutamine, formylmethionine, cysteine, or selenocysteine.15,32 The past decade has seen the identification of new aa-tRNA-dependent enzyme families, all of which are involved in the biosynthesis of microbial secondary metabolites,33−40 referred to hereafter as NPs (Figure 2). The enzyme families involved in NP biosynthesis can be classified into three main groups according to their catalytic activities and their structurally conserved fold (Figure 3). The first includes the cyclodipeptide synthase (CDPS) family, which

uses two aa-tRNAs for the formation of various cyclodipeptide products,34 which serve as the precursors of many bioactive compounds.43 CDPSs have similar architectures, with a Rossmann-fold domain,44 closely similar to the catalytic domain of class-Ic aaRSs.45−47 The second group was defined on the basis of recent biochemical and structural studies on lantibiotic dehydratases (LanBs)39,48 and LanB-like enzymes,49,50 which are involved in the biosynthesis of several classes of ribosomally synthesized and posttranslationally modified peptides (RiPPs).51,52 These enzymes dehydrate specific serine or threonine residues of a precursor peptide in a Glu-tRNAGludependent manner during the production of class I lanthipeptides,39,48 thiopeptides,49 or the atypical goadsporin50 (Figure 3). The third group consists of aminoacyl transferases belonging to the structural GCN5-related N-acetyltransferase (GNAT) protein superfamily,41,42 along with Fem transferases, aaPGSs, and aminoacyl protein transferases (Figure 2). These enzymes

Figure 3. aa-tRNA-utilizing enzymes involved in NP biosynthesis. These enzymes form three groups: synthase enzymes with a Rossmann-fold domain (purple circle), dehydratase enzymes with a LanB-like fold (green circles), and transferase enzymes with a GNAT fold (blue circles). They are involved in the synthesis of several classes of NPs (indicated in boxes). 5580


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Figure 4. CDPS-dependent biosynthesis of diketopiperazines. (a) Schematic representation of the albonoursin (2) biosynthetic pathway. (b) Schematic representation of a generic CDPS-dependent pathway with examples of DKPs produced: nocazine XR334 3, pulcherriminic acid 4, mycocyclosin 5, doubly-N-methylated cyclo(L-Trp-L-Trp) 6, and nocardioazine B 7. aa1 and aa2 are the first and the second amino acid, respectively, and R1 and R2 are the corresponding side chains. The DKP ring is in blue.

in length, displaying only moderate amino acid sequence identity, with most displaying less than 30% sequence identity.55 They are distributed on two main branches of the CDPS phylogenetic tree (Figure 5), reflecting the existence of two subfamilies. These subfamilies differ in the composition of a specific couple of catalytic residues and are thus referred to as the NYH and XYP subfamilies (see section 2.2.2). The active members of the CDPS family produce a large collection of about 55 different cyclodipeptides with 17 proteinogenic aminoacyl residues55,65,67(Figure 6). The cyclodipeptides produced mostly contain hydrophobic amino acids, and the acidic Asp and basic Arg and Lys residues have never been detected in these molecules (Figure 6). Almost half of the known CDPSs display high substrate specificity, producing only one cyclodipeptide, as is often the case for enzymes synthesizing cyclo(L-Trp-L-Trp)55,63,65 or cyclo(L-Cys-L-Cys).55 Other members of this group display a certain degree of substrate promiscuity and generally produce one major cyclodipeptide and several minor cyclodipeptides of the general formula cyclo(AA1-X), in which AA1 is the preferred amino acid and X indicates the variability of the incorporated amino acids.34,55

transfer the aminoacyl moieties of selected aa-tRNAs during the biosynthesis of various antibiotics such as valanimycin, which is also an antitumoral compound,33 pacidamycin,36 dehydrophos,37 and their respective homologues (Figure 3).

2. aa-tRNA-DEPENDENT SYNTHASES: THE CYCLODIPEPTIDE SYNTHASES (CDPSS) The group of synthase enzymes with a structurally conserved Rossmann-fold domain currently contains only one family of enzymes, the CDPSs (Figure 3). In 2002, the first member of this family was identified during characterization of the albonoursin biosynthetic pathway in Streptomyces noursei.53 This CDPS, AlbC, uses Phe-tRNAPhe and Leu-tRNALeu to form the cyclodipeptide cyclo(L-Phe-L-Leu) 1,34 which is subsequently modified by cyclic dipeptide oxidase (CDO) to generate albonoursin 2 (cyclo(α,β-dehydroPhe-α,β-dehydroLeu) or cyclo(ΔPhe-ΔLeu))54 (Figure 4a). Many other CDPSs have since been identified,55 and most are associated with one or several cyclodipeptide-tailoring enzymes in biosynthetic pathways producing a diverse range of 2,5-diketopiperazines (DKPs)56−60 (Figure 4b). The physiological roles of the DKPs produced remain unclear, but these molecules have attracted considerable interest because of their diverse pharmacological activities.43

2.2. 3D-Structure and Catalytic Mechanism

2.2.1. A Common Architecture with a Rossmann-Fold Domain. The crystallographic structures of three CDPSs belonging to the same NYH subfamily (see section 2.1 and Figure 5; see also section 2.2.2 and Figure 10)AlbC f rom S. noursei (PDB ID, 3OQV), Rv2275 from Mycobacterium tuberculosis (PDB ID, 2X9Q), and YvmC from Bacillus licheniformis (PDB ID, 3OQH, 3OQI, 3OQJ, and 3S7Y)have been solved at resolutions of 1.9 Å, 2.0 Å, and 1.7−2.8 Å, respectively.45−47 These structures were obtained with the enzymes alone45 or in complex with buffer components.46,47 The crystal structure of AlbC in complex with a reaction intermediate analogue (PDB ID, 4Q24) was recently determined at a resolution of 2.9 Å.68 Despite displaying only about 27% sequence identity, these three CDPSs have a common architecture, consisting of a monomer with a compact α/β fold comprising a central β sheet with five parallel β strands bound on either side by nine α helices and two β strands (Figure 7a). This monomer contains a Rossmann-fold domain,44 formed by strands β3−β5 and helices α2 and α4, that can be superimposed in the three structures (Figure 7a). A striking feature of the CDPS structures obtained to date is the

2.1. CDPSs and Their Cyclodipeptide Products

CDPSs use aa-tRNAs as substrates for the formation of the two peptide bonds of cyclodipeptide scaffolds. This reaction has been demonstrated for CDPSs from various microorganisms, through heterologous approaches in Escherichia coli, both in vivo (recombinant CDPSs use E. coli aa-tRNAs to produce cyclodipeptides that are then released into the culture medium) and in vitro (recombinant purified CDPSs are incubated in the presence of tRNAs and aaRSs from E. coli (or Saccharomyces cerevisiae) to generate aa-tRNAs in situ).34 The CDPS family currently contains 64 functionally characterized members34,55,61−65 (Figure 5), and about 450 putative members can be retrieved by searching the National Center for Biotechnology Information (NCBI) protein database. The characterized CDPSs essentially originate from three bacterial phyla (Actinobacteria, Firmicutes, and Proteobacteria). Only one of these enzymes originates from a eukaryotic phylum,61 and none have been found in Archaea (Figure 5). CDPSs are small enzymes of about 200−300 residues 5581


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Figure 5. Phylogenetic tree of the characterized CDPSs. The tree was generated using the PhyML program (v 3.1)66 based on the maximum likelihood method. CDPS names consist of the corresponding accession numbers in the NCBI database and the host organisms in which they were found. The eight CDPSs found to be inactive in standard experimental conditions are indicated by stars. The two main branches corresponding to the XYP and NYH subfamilies are shown in blue and gray, respectively. CDPS names are shown in color, according to the taxonomic group from which they originate: actinobacteria (dark blue), proteobacteria (purple), firmicutes (green), chlamydiae (cyan), cyanobacteria (brown), bacteroidetes (pink), and eukaryotes, including 1 fungus and 1 metazoan (black). The putative CDPS identified by a metagenomic approach is shown in orange.

presence of a deep surface-accessible pocket, P1 (see section 2.3.2), bordered by the catalytic residues (defined in section 2.2.2).

The CDPSs display strong structural similarity to the catalytic domains of class Ic TyrRSs and TrpRSs,45−47 despite there being less than 15% sequence identity between the enzymes of these 5582


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outside of tRNA aminoacylation are known71−74 and have been reviewed.75,76 2.2.2. A Common Sequential Ping-Pong Mechanism. The catalytic mechanism of CDPSs has been extensively investigated for the three enzymes for which crystal structures have been determined (see section 2.2.1), and the overall catalytic cycle has been elucidated for AlbC.46,68 AlbC hijacks Phe-tRNAPhe and Leu-tRNALeu (or a second Phe-tRNAPhe)34 and uses them as substrates for a ping-pong mechanism involving the formation of two successive acyl-enzyme intermediates68 (Figure 8). The first aa-tRNA binds to the enzyme such that its aminoacyl moiety is accommodated in the P1 pocket.46 This moiety is then transferred onto a conserved serine residue (Ser37) to form an aminoacyl-AlbC intermediate. The formation of this intermediate has been demonstrated by trypsin digestion and peptide map fingerprinting (PMF) on AlbC incubated with Phe-tRNAPhe.46 The second aa-tRNA binds to the enzyme such that its aminoacyl moiety is positioned in a wide pocket, P2, located close to P168 (see section 2.3.2). The phenylalanyl-AlbC then reacts with the second aa-tRNA to generate a dipeptidylAlbC intermediate, from which the final cyclodipeptide is obtained by intramolecular cyclization (Figure 8). This second intermediate has been trapped by modifying a conserved tyrosine (Tyr202) involved in the cyclization process, and detected by PMF analyses carried out with the Tyr202Phe variant.68 In addition to the two conserved residues, Ser37 and Tyr202, four other residues from the P1 pocket have been identified as catalytic residues: Tyr178, Glu182, Asn40, and His203. The roles of these residues, proposed on the basis of site-directed mutagenesis and chemical biology studies combined with analysis of the crystal structure of a dipeptidyl-AlbC analogue,46,68 are detailed in ref 68 and shown in Figure 9. Briefly, Tyr178 and Glu182 are involved, through polar contacts, in accommodating and stabilizing the aminoacyl moiety of the first substrate throughout the catalytic cycle (steps 1−5). Glu182 also acts as an essential catalytic base, by deprotonating the ammonium group of the aminoacyl-enzyme (step 3), enabling it to attack the carbonyl group of the second substrate to form the dipeptidyl-enzyme (step 4). Asn40 and His203 participate in a hydrogen bond network responsible for the accurate positioning of catalytic residues (steps 1−5). In particular, Tyr202 is located

Figure 6. Diversity of the cyclodipeptides produced by characterized CDPSs. The 20 amino acids are indicated in the top row and the righthand column. The cyclodipeptides produced by CDPSs are indicated by orange squares at the intersection of a row and a column.

two families. The Rossmann-fold domains present in the two families can be superimposed with ease, as exemplified by a comparison of the structures of AlbC and TyrRS from Methanococcus jannaschii (PDB ID, 1J1U69) (Figure 7b). In particular, the surface-accessible P1 pocket in CDPSs is positioned similarly to the aminoacyl binding pockets in the two aaRSs.45−47 However, there are several key differences between CDPSs and class Ic aaRSs.45−47 The ATP-binding motifs present in aaRSs are not present in CDPSs, consistent with the use, by CDPSs, of amino acids that have already been activated. TyrRSs and TrpRSs are homodimers, in which the two active sites are interdigitated at the dimer interface,70 whereas CDPSs are active as monomers,46,47,61 as is Rv2275 (personal data). It therefore appears likely that CDPSs evolved from class I aaRSs, gaining features enabling them to use the products of aaRSs as their substrates for the catalysis of amide bond formation.56 A few other aaRS paralogues with biological roles

Figure 7. Comparison of the crystal structures of CDPSs. (a) Superimposition of the cartoon structures of AlbC (PDB ID, 3OQV; green), Rv2275 (PDB ID, 2X9Q; gray), and YvmC (PDB ID, 3OQH; magenta). The secondary structure elements are labeled; strand β1 is masked in this view. (b) Superimposition of the cartoon crystal structures of AlbC and TyrRS of M. jannaschii in complex with its tyrosine substrate (PDB ID, 1J1U). AlbC is in green; the catalytic and tRNA-binding domains of TyrRS are in blue and light blue, respectively, and the tyrosine substrate is shown in stick format, in orange. Reprinted with permission from ref 46. Copyright 2011 Oxford University Press. 5583


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Figure 8. Overall catalytic cycle for the production of cyclodipeptides by the CDPS AlbC. Reprinted with permission from ref 57. Copyright 2012 Royal Society of Chemistry. AlbC (in green) uses two aa-tRNAs produced by aaRSs (in red) and is involved principally in protein synthesis at the ribosome (in blue). AlbC proceeds through a sequential mechanism involving the formation of aminoacyl and dipeptidyl enzymes.

Figure 9. Proposed roles of the catalytic residues of AlbC. Reprinted with permission from ref 68. Copyright 2014 Macmillan Publishers, Ltd. The first and the second aa-tRNAs are represented in blue and orange, respectively. Possible interactions are shown by dashed lines.

at an ideal position to deprotonate the ammonium group of the dipeptidyl-enzyme (step 5), enabling it to attack the enzyme ester bond to form the cyclodipeptide (Figure 9). One intriguing issue relating to the Ser37 residue remains to be resolved. As

mentioned above, a role for this residue has been clearly established in the formation of the aminoacyl-enzyme, but no relevant residues for its activation (as its hydroxyl group must be deprotonated to act as a nucleophile) have been clearly 5584


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substrates. The interaction of AlbC with its first substrate can be observed specifically, by studying the first step in the catalytic reaction, the formation of the acyl-enzyme intermediate. This was achieved with a set of aa-tRNAs either carrying wild-type E. coli sequences or harboring mutations in the acceptor stem. The interaction with the second substrate was studied through a combination of diverse biochemical assays with different isoacceptors,78 Leu-tRNALeu and mutated tRNALeu or tRNAPhe. Posttranscriptional tRNA modification was not required for AlbC activity, as tRNAs extracted from E. coli (i.e., maturated) and tRNAs produced by in vitro transcription from E. coli genes as templates (i.e., nonmaturated) were used similarly by the CDPS. These experiments demonstrated that AlbC first interacts specifically with Phe-tRNAPhe to form a phenyalanyl-enzyme. The specificity of the CDPS for its first substrate is mediated by the aminoacyl moiety. However, the tRNA moiety is essential in other ways: for activation of the carboxylate function of the amino acid, facilitating the formation of the acyl-enzyme, and to enhance the binding of the substrate to the CDPS via nonspecific electrostatic interactions.78 This mode of recognition is associated with a very high specificity for the first substrate. Most of the CDPSs seem to interact first with a single preferred aminoacyl loaded onto tRNA, which is preferentially incorporated into the different cyclodipeptides synthesized by the enzyme when several products are detected.57 Consistent with this finding, AlbC essentially synthesizes cyclo(L-Phe-L-X) compounds. tRNAs loaded with D-Phe are not transformed by AlbC, suggesting that only L-configured amino acids can be used for cyclodipeptide biosynthesis.79 The specificity of AlbC for its second substrate involves both the aminoacyl moiety and some of the bases from the acceptor stem in the tRNA moiety. The interaction of the second substrate with AlbC depends on the presence of a pair G1-C72 in the tRNA sequence.78 This pair of bases is found in many tRNA sequences from prokaryotes but is not a necessary and sufficient condition for an efficient and productive interaction. Specificity for this second substrate is rather relaxed, but AlbC does not incorporate charged or polar amino acid into cyclodipeptides. Instead, it preferentially recognizes large hydrophobic amino acids. However, few characterized CDPSs produce only one product, suggesting that specificity for the second aa-tRNA substrate can also be highly specific.55,63−65 These enzymes include cyclo(LTrp-L-Trp)-synthesizing CDPSs. This specificity may reflect a unique feature of most bacterial tRNATrp molecules, which combine an A1-U72 base pair and a G73 base.78 Some CDPSs also exclusively synthesize a cyclodipeptide containing a charged or a polar amino acid (cyclo(L-Ala-L-Glu) for example).55 The particular physical and/or chemical characteristics of the second amino acid used by these CDPSs probably results in the presence of residues able to take part in specific interactions, excluding the binding of other residues. Most of the CDPSs characterized were studied in E. coli or with E. coli tRNAs. However, the sequences of E. coli tRNAs may differ significantly from those of the tRNAs of the species from which the CDPS originates. It follows that some CDPSs may be inactive in E. coli or produce only minor cyclodipeptides distantly related to their major product, due to a lack of appropriate substrates. This may account for the very low activity observed for the only eukaryotic CDPS characterized. This CDPS originating from Nematostella vectensis synthesizes cyclo(L-Trp- L-X) cyclodipeptides in very small amounts when expressed in E. coli.61 tRNATrp is one of the few tRNAs for which there are significant differences in the acceptor stem sequence between eukaryotes

identified. Its activation probably results from substrate-assisted catalysis involving the free hydroxyl group of the ribose moiety from the first aa-tRNA (during step 1). The free hydroxyl group of ribose from the aa-tRNA was proposed to participate in catalysis through a proton shuttle mechanism for FemX from Weissella viridescens (FemXWv)20 and ribosomal peptidyl transferase.77 The catalytic cycle with two successive acyl-enzyme intermediates elucidated for AlbC is likely used by all CDPSs

Figure 10. CDPS functional sequence signatures. Reprinted with permission from ref 55. Copyright 2015 Macmillan Publishers, Ltd. (a) Functional signature for all characterized CDPSs. The signature is built with sequence logos corresponding to the frequency plot of the catalytic residues at positions 37, 40, 178, 182, 202, and 203 (AlbC numbering). (b) Functional subsequence signatures for the NYH and XYP subfamilies. A potential third subfamily, with one member that has a Gln residue at position 203, was previously reported,55 but this protein is now considered to belong to the XYP subfamily (see WP 005863245.1 in Figure 5).

(Figure 8). Indeed, the catalytic residues Ser37 and Glu182, involved in the formation of the aminoacyl and dipeptidyl enzymes, respectively, are strictly conserved in almost all active CDPSs (the exceptions being two CDPSs in which the role of Glu182 could be fulfilled by an Asp)55 (Figure 10a). The formation of the aminoacyl-enzyme has also been demonstrated for the CDPSs Rv2275, YvmC,45,47 and pSHaeC06 from Staphylococcus hemolyticus.78 The Tyr178 and Tyr202 residues, responsible for anchoring the aminoacyl moiety of the first substrate and cyclization of the final product, respectively, are also well conserved (Figure 10a). By contrast, the Asn40 and His203 residues involved in the positioning of the two loops bearing the catalytic residues68 are not conserved: about half of the characterized CDPSs have the couple of residues “Asn40, His203” whereas the other half have the couple “X40, Pro203″, with X being a nonconserved residue55 (Figure 10a). This couple of residues is different in each of the two phylogenetically distinct subfamilies of CDPSs (see section 2.1 and Figure 5) and underline the naming of the subfamilies as NYH or XYP according to the “X40, Tyr202, X203” sequence55 (Figure 10b). Although the NYH and XYP CDPSs probably share the same catalytic mechanism, they could adopt two different solutions for the accurate positioning of the catalytic residues. The testing of this hypothesis will require characterization of the structures of XYP subfamily members, because all three CDPSs for which structural determinations have been carried out to date are NYH subfamily members.45−47 2.3. Molecular Basis of the Interaction between CDPSs and Their Substrates

2.3.1. Specificity Determinants of the Two aa-tRNA Substrates. A study carried out on AlbC, which synthesizes the hetero cyclodipeptide cyclo(L-Phe-L-Leu) 1 as a major product, showed that CDPSs discriminate between their first and second 5585


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Figure 11. Binding sites of AlbC for the two substrates. (a) Molecular surface of the AlbC−ZPK complex (PDB ID, 4Q24).68 The loops bearing the catalytic residues, CL1 (residues 36−43) and CL2 (residues 202−207), are in yellow. The basic residues of helix α4 are in blue, and those shown to participate in tRNA binding during the first step of the reaction are shown in dark blue and labeled. Residues shown to be involved in tRNA binding during the second step are shown in pink. (b) Close-up view of the binding site of the first amino acid; the residues delineating P1 are shown as sticks. (c) Close-up view of the binding site of the second amino acid with P2 residues shown as sticks. Reprinted with permission from ref 68. Copyright 2014 Macmillan Publishers, Ltd.

through electrostatic interactions mediated by basic residues is common among RNA-binding proteins.83 However, such binding may not be the rule for all CDPSs. Multiple sequence alignments for CDPSs from the XYP subfamily revealed a much smaller number of basic residues in the region corresponding to the α4-helix.55 XYP enzymes have a large number of serine, threonine, glutamine, asparagine, aspartate, and/or glutamate residues instead. All these residues are also known to interact specifically with nucleotide bases84 and may participate in the specific recognition of some substrates by interacting with only a restricted number of tRNA sequences. However, this hypothesis remains to be tested. The aminoacyl moiety of the second substrate lies in a wide, open cavity, P2, delineated by the two loops bearing the catalytic residues, CL1 and CL2, and parts of helices α6 and α7 (Figure 11a,c). This cavity is not present in the structures of unliganded CDPSs45 or CDPSs harboring solvent molecules in P1,46,47 and probably results from a local reorganization of CL1 and CL2 during formation of the acyl-enzyme intermediate. Its large size is consistent with the relaxed specificity for the second aminoacyl moiety observed for many CDPSs.55,57 The tRNA moiety of the second substrate has been shown to interact with residues from the loops α6−α7 and β6−α878 (Figure 11a). The equivalent loops are also involved in binding the acceptor stem of the tRNA substrate in class Ic TyrRSs and TrpRSs and in specific recognition of the N1-N72 base pair.70,80,85 This suggests that CDPSs may have retained some properties of the class Ic aaRSs in terms of their interactions with the tRNA moiety of the second substrate. Further studies are required to validate this hypothesis. 2.3.3. Toward the Prediction of CDPS Specificity. The growing number of available sequences encoding putative CDPSs (about 450 in June 2016) raises questions about how to predict substrate specificities and products. Any prediction model will require precise determination of the pockets accommodating the substrates and the biochemical characterization of a large enough number of CDPSs. Such an approach was successfully used in the development of the first prediction tools for the substrate specificity of the adenylation domains of nonribosomal peptide synthetases (NRPSs).86,87 The availability of about 60 biochemically characterized CDPSs and the first

and prokaryotes. In particular, differences at positions 1, 72, and 73 are largely responsible for the very low cross-reactivity between eukaryotic and bacterial TrpRS-tRNATrp pairs (i.e., a bacterial TrpRS cannot load a eukaryotic tRNATrp and vice versa).80 The use of a more appropriate tRNA set for the study of the eukaryotic CDPS would probably provide a different view of its activity. Finally, experiments carried out on AlbC have shown that apparent discrepancies can be observed between the amino acids preferentially recognized by the CDPS in vitro and the major cyclodipeptide produced in vivo. AlbC interacts preferentially with Phe-tRNAPhe as both a first and a second substrate,78 producing cyclo(L-Phe-L-Phe) 8 as the major cyclodipeptide in vitro, but it synthesizes predominantly cyclo(L-Phe-L-Leu) 1 in both E. coli34 and S. noursei,81 although cyclo(L-Phe-L-Phe) 8 is also observed. One likely reason for this observation is the relative abundances of tRNAs in E. coli (or S. noursei). Whatever the growth rate, the tRNALeu used as a substrate by AlbC is about seven times more abundant than tRNAPhe in E. coli cells.82 The reconstitution of this ratio in in vitro experiments with AlbC led to the predominant synthesis of cyclo(L-Phe-L-Leu) 1 over cyclo(L-Phe-L-Phe) 8.78 2.3.2. The Binding Sites of the Two Substrates. Recent studies have shown that the two substrates of CDPSs are accommodated at different binding sites.68,78 The crystal structure of AlbC complexed with a reaction intermediate analogueN-carbobenzyloxy-L-Phe-methyl ketone (ZPK), which mimics the Phe-Phe dipeptidyl intermediaterevealed the presence of two different cavities at the surface of the protein that accommodate the aminoacyl moieties originating from the first and second aa-tRNAs (Figure 11a). The first aminoacyl moiety was buried deep in the hydrophobic pocket P1 (Figure 11b) that corresponds to the aminoacyl-binding pocket of class I aaRSs and bordered by the catalytic residues. This location is consistent with the findings of mutagenesis experiments showing that the substitution of a single residue situated at the bottom of this pocket affects the nature of the amino acid recognized by a CDPS.46 Another study on AlbC demonstrated the involvement of basic residues from helix α4 (Figure 11a) forming a basic patch at the surface of the protein in the binding of the tRNA moiety of the first substrate to the CDPS.78 The binding of RNA molecules 5586


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Figure 12. Organization of the characterized gene clusters for the CDPS-dependent biosynthesis of DKPs. (a) Albonoursin 2 biosynthetic gene cluster in S. noursei.53,54 (b) Nocazine biosynthetic gene cluster in N. dassonvillei.62 A similar cluster has been identified in Nocardiopsis alba.62 (c) Pulcherriminic acid 4 biosynthetic gene cluster in B. subtilis.34,93,97 (d) Mycocyclosin 5 biosynthetic gene cluster in M. tuberculosis.34,94−96 (e) Biosynthetic gene cluster of the simply and doubly N-methylated cyclo(L-Trp-L-Trp) 6 in A. mirum.63 Genes are colored according to the function of the encoded proteins. Genes annotated as regulator, transporter, or involved in resistance have not been shown as belonging to the biosynthetic pathways.

prediction efficiency of the current tool. The second improvement of this tool will result from determination of the precise binding sites for the tRNA moieties. The tRNA does not seem to contribute to recognition specificity for the first substrate, but, together with the aminoacyl moiety, it plays an important role in the specific recognition of the second substrate. This role is thought to be essential in enzymes synthesizing only cyclo(LTrp-L-Trp),55,63−65 and it remains to be explored.

molecular description of the two substrate-binding sites paved the way for the development of the first prediction model.55 This model is based on determination of the residues constituting the P1 and P2 pockets and does not take into account interactions of the tRNA moieties of the two substrates. These residues are identified from multiple alignments with structurally characterized CDPSs, manually adjusted with HHpred predictions. HHpred is a free protein structure prediction server for the detection of remote protein homology.88,89 Starting from multiple sequence alignments with the characterized CDPSs, this work led to the construction of sequence logos indicating the relative frequencies of amino acids in the various positions, for both the P1 and P2 aminoacyl binding pockets, and the association of these logos with the production of a cyclodipeptide, taking only the major cyclodipeptide synthesized by a given CDPS into account. A minimum of five different CDPSs from the same subfamily (NYH or XYP) and synthesizing the same major cyclodipeptide were used to determine the association of a logo with an activity. Six different groups were identified, synthesizing cyclo(L-Trp-L-Trp), cyclo(L-Leu-L-Leu), and cyclo(L-Cys-L-Cys) in the NYH subfamily, and cyclo(L-AlaL-Glu), cyclo(L-X-L-Glu), and cyclo(L-Ala-L-X) in the XYP subfamily.55 The validity of this approach was demonstrated by determining the activity of another 10 CPDSs predicted to synthesize one of these cyclodipeptides. Since this study, the number of putative CDPSs has increased by more than 50%. By determining the putative residues constituting P1 and P2 in these new enzymes, it is possible to group them on the basis of their P1 and P2 pocket compositions. Determination of the activity associated with these new groups and the availability of new three-dimensional structures of CDPSs will improve the

2.4. CDPS-Dependent Pathways

CDPS-dependent pathways are dedicated to the biosynthesis of cyclodipeptides and their modified derivatives, DKPs (Figure 4b). Cyclodipeptides are produced by the CDPSs and modified by tailoring enzymes encoded by genes generally clustered with the CDPS genes.56,57,59 Only three CDPS-dependent pathways synthesizing DKPs produced by the corresponding host organisms have been fully characterized to date: those directing the biosynthesis of albonoursin 2 in S. noursei,81 the nocazine family members nocazine XR334 3 and E (a 3 derivative with only one α,βdehydrogenation) in Nocardiopsis dassonvillei and Nocardiopsis alba,90,91 and pulcherriminic acid 4, the precursor of the red pigment pulcherrimin formed in the presence of FeIII in Bacillus subtilis92 (see Figure 4). The albonoursin 2 gene cluster contains three genes, encoding the CDPS AlbC and the two subunits of the tailoring CDO (Figure 12a). CDO is a flavin-dependent α,βdehydrogenase with a relaxed substrate specificity that forms the two double bonds of albonoursin 2 (Figure 4a).53,54 The nocazine XR334 3 and E clusters contain four genes encoding the CDPS, the two subunits of a CDO homologue and an Omethyltransferase (Figure 12b). The CDPS produces mostly 5587


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the final RiPP. This precursor usually consists of an N-terminal leader peptide fused to a C-terminal core peptide containing the various posttranslational modification sites. Many posttranslational processing enzymes specifically recognize the leader peptide while being permissive with respect to the sequence of the core peptide. The modified core is finally released by one or more peptidases, and cyclized if appropriate, to yield the mature active compound.51,52 A role for an acylated tRNA in the biosynthesis of different classes of RiPPs51 was recently established and shown to involve exclusively glutamyl-tRNAGlu.39,48−50 The acylated tRNA is mainly involved in the processing steps of two different

cyclo(L-Phe-L-Tyr), which undergoes one or two α,β-dehydrogenations, followed by methylation of the hydroxyl group of the tyrosine residue.62 Another gene annotated as O-methyltransferase is found upstream from the CDO gene (Figure 12b) and is thought to be responsible for the O- and N-methylations of the DKP ring observed in other nocazine family members.62 The gene cluster for pulcherriminic acid 4 is composed of two genes encoding a CDPS and a cytochrome P450 enzyme (Figure 12c). The CDPS synthesizes cyclo(L-Leu-L-Leu),34 and the cytochrome Cyp134A1 catalyzes oxidative reactions leading to double N-oxide formation with aromatization of the DKP ring.93 Two other CDPS-dependent pathways are well documented, but the final DKPs produced in the host organisms remain to be determined. The CDPS gene rv2275 clusters with a gene encoding the P450 CYP121 enzyme in M. tuberculosis94,95 (Figure 12d). Rv2275 synthesizes mostly cyclo(L-Tyr-L-Tyr),34 and CYP121 has been shown to catalyze the formation in vitro of a carbon−carbon single bond between the two meta positions of the tyrosyl side chains of the cyclodipeptide,96 generating mycocyclosin 5. The second pathway has been identified in Actinosynnema mirum, the genome of which contains a gene cluster composed of a CDPS gene and two genes encoding an Nmethyltransferase and a fatty acyl-CoA ligase63 (Figure 12e). The CDPS produces cyclo(L-Trp-L-Trp), which is modified by the methyltransferase to generate a cyclodipeptide with one or two methylations 6 of the DKP-ring nitrogens. No function could be assigned to the ligase.63 It has recently been suggested that two CDPS homologues from Nocardiopsis sp. CMB-M0232 are involved in the biosynthesis of nocardioazines A and B 7.64,65 Both CDPSs synthesize cyclo(L-Trp-L-Trp), which could be further tailored by prenyltransferase, methyltransferase, and cytochrome P450 enzymes. By contrast to the biosynthetic pathways described above, the genes required for nocardioazine biosynthesis are not grouped into a single gene cluster.64 Finally, a large number of putative CDPS-dependent pathways remain to be characterized. Bioinformatic analysis of the genetic surroundings of each known and putative CDPS gene often reveals the presence of one or several genes encoding diverse putative tailoring enzymes capable of introducing a large set of modifications in cyclodipeptides.57,59 This strongly suggests that CDPS-dependent pathways are responsible for the biosynthesis of a wide variety of DKPs. The only other biosynthetic route for the biosynthesis of DKPs involves NRPSs, multimodular megaenzymes responsible for generating the vast majority of nonribosomal peptides (for a recent review, see ref 98). To date, 12 NRPS-dependent pathways have been identified for DKP biosynthesis,99−108 including two pathways producing DKPs as side products during the biosynthesis of longer peptides.109,110 These pathways were described and compared with CDPS-dependent pathways in ref 57.

Figure 13. (a) Glu-tRNAGlu-dependent dehydration of serine and threonine residues by LanB enzymes. Adapted with permission from ref 39. Copyright 2015 Macmillan Publishers, Ltd. The glutamate residue is in pink. Dha and Dhb are 2,3-didehydroalanine and (Z)-2,3didehydrobutyrine, respectively. (b) Cyclization of lanthipeptides catalyzed by LanC. The gray rectangle highlights the final lanthionine (R = H; Ala-S-Ala) or methyl-lanthionine (R = CH3; Abu-S-Ala) units.

subfamilies of RiPPs: class I lanthipeptides115 and thiopeptides.116−119 These RiPPs include dehydroamino acids during their maturation process. These dehydroamino acids are formed in a two-step reaction, from serine and threonine residues (Figure 13a). Current knowledge suggests that the side chains of Ser/Thr residues are first activated by glutamylation. The glutamate is subsequently eliminated to form 2,3-didehydroalanine (Dha) and (Z)-2,3-didehydrobutyrine (Dhb), respectively. This process is catalyzed by a dedicated tRNA-dependent dehydratase generically called LanB (for lanthionine biosynthetic enzyme B51) or by LanB-like enzymes.52 In lanthipeptides, cysteine thiols are added to some Dha or Dhb residues by a Michael-type addition catalyzed by the cyclase LanC to form lanthionine (Ala-S-Ala) or methyl-lanthionine (Abu-S-Ala; Abu stands for L-α-aminobutyric acid) (Figure 13b) (for further details, refer to the review devoted to lanthipeptide biosynthesis in the present thematic issue120). In thiopeptide biosynthesis, dehydration provides the two dehydroalanine precursors for the central six-membered heterocycle via intramolecular [4 + 2] cycloaddition.117,118 Not all the Dha and Dhb residues formed during processing are involved in the cyclization steps, and some are present in the final compounds. The structures of some lanthipeptides and thiopeptides are presented in Figure 14. Figure 14 also shows the structure of goadsporin 14. This RiPP belongs to the family of linear azol(in)e-containing peptides (LAP), defined by the presence of azol(in)e rings on a non-macrocyclized peptide (as opposed to the thiopeptides).51 The distinguishing feature of

3. aa-tRNA-DEPENDENT DEHYDRATASES IN RiPP SYNTHESIS RiPPs are ribosomally synthesized peptides that undergo extensive posttranslational modifications, endowing them with considerable chemical and structural diversity. These features are associated with diverse biological activities, including antibacterial, antifungal, and antiviral activities.111−114 RiPPs can be classified into different subfamilies on the basis of their structural characteristics and biosynthetic machinery. Regardless of the subfamily to which they belong, RiPP synthesis begins with the ribosomal production of a precursor peptide that is longer than 5588


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Figure 14. Representative structures of some of the RiPPs mentioned in this review. Dehydroalanine (Dha) results from the dehydration of serine and is shown in green. Dehydrobutyrine (Dhb) results from the dehydration of threonine and is shown in light blue. In lanthipeptide molecules, lanthionines (Ala-S-Ala) are shown in red and methyl-lanthionines (Abu-S-Ala) are shown in dark blue. In thiopeptides, the central six-membered heterocycle is shown in orange and is formed by cycloaddition between two Dha precursors. Thiopeptides are classified in structural series according to the oxidation state of this nitrogenous ring.

3.1.1. Mechanism and Structure of LanB. Two LanBs have been characterized biochemically and structurally: NisB (117 kDa, from the firmicute Lactococcus lactis)39 and MibB (120 kDa, from the actinobacterium Microbispora sp. 107891)48 (Figure 15). These proteins belong to the two different clades identified by phylogenetic studies, and their sequences are less than 20% identical. Despite their distant phylogenetic relationship, they adopt similar folds and both have been shown to use Glu-tRNAGlu as a cosubstrate to catalyze the dehydration of serine and threonine residues. These findings suggest that a mechanism involving the use of Glu-tRNAGlu is common to class I lanthipeptide dehydratases. The dehydratase NisB is involved in the synthesis of nisin 9 (Figure 14), a lantibiotic that has been used as a food preservative for over 40 years123 and is produced from a ribosomally synthesized precursor peptide called NisA. NisB dehydrates three serine residues and five threonine residues in the NisA core region, yielding Dha and Dhb residues (Figure 16). The cyclase NisC catalyzes the formation of five lanthionine or methyllanthionine rings, through the nucleophilic addition of cysteine thiols to one Dha and four Dhb residues. The fully modified prenisin molecule is then exported by the ABC transporter NisT.

goadsporin 14 regarding other LAP peptides is to contain two Dha residues, which are generated by LanB-like enzymes. 3.1. Full-Length LanB Dehydratases in Lanthipeptides Biosynthesis

Full-length LanBs, referred to hereafter as LanBs, usually have about 1,000 residues and are involved essentially in biosynthesis pathways for class I lanthipeptides. Many of these lanthipeptides have antimicrobial activity and are known as lantibiotics. LanBs display no sequence similarity to the proteins of other families and therefore constitute a unique family. They consist of two domains: an N-terminal lanthipeptide dehydratase domain and a small C-terminal SpaB_C domain. Phylogenetic analyses of LanBs have shown that they form two distinct groups, according to the phyla from which they originate121,122 (Figure 15). The first group consists of proteins from actinobacteria, whereas the second consists mostly of LanBs from firmicutes. The second group also contains the identified enzymes from bacteroidetes and proteobacteria, which cluster together in a small subgroup nested deeply within the firmicutes group.122 Both within a phylogenetic group and between the two different groups, sequence identity between two LanBs can be as low as about 20%. 5589


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Figure 15. Phylogeny of LanB enzymes. Reprinted with permission from ref 122. Copyright 2012 Proceedings of the National Academy of Sciences. The tree was generated using the PhyML program (v 3.1)66 based on the maximum likelihood method. The host organisms are colored according to their taxonomic origin: actinobacteria (blue), firmicutes (brown), bacteroidetes (green), and proteobacteria (red). The characterized LanBs are indicated in black next to the name of the host organism.

dehydrate five serine residues and two threonine residues within the precursor peptide, MibA. The dehydration of serine and threonine residues requires prior activation of their side chains to facilitate β-elimination upon deprotonation at the α carbon. In class II, III, and IV lanthipeptides, dehydration reactions proceed through the ATP-

The extracellular peptidase NisP then removes the leader sequence, yielding the active mature peptide, nisin 9.124 MibB is involved in the synthesis of NAI-107, also termed microbisporicin A1 10 (Figure 14), a lantibiotic in late preclinical trials with potential for use in the treatment of multidrug-resistant Gram-positive bacterial infections.125,126 MibB is predicted to 5590


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Figure 16. Steps in the biosynthesis of the lanthipeptide nisin. Adapted with permission from ref 39. Copyright 2015 Macmillan Publishers, Ltd. Dehydrated residues and their amino acid precursors are colored as in Figure 14. Arrows indicate the formation of thioether bridges between dehydrated residues and cysteines. NisT transports the fully modified precursor outside the cell, where it is cleaved by NisP.

glutamylation active site. The glutamylation domain also contains a small subdomain of about 80 residues (spanning residues 136 through 216 in NisB and 201 through 286 in MibB) responsible for binding the leader peptide. This small structural element was recently shown to be present in numerous unrelated RiPP-modifying enzymes,130−132 requiring the presence of a leader peptide for interaction with their substrate. Structural and biochemical data for NisB and MibB made it possible to propose a global iterative mechanism for the dehydration reaction catalyzed by LanB.39,48 Binding of the leader peptide may bring the core region close to the active site for glutamylation, where the glutamate is transferred from the Glu-tRNAGlu to the side chain of the targeted serine or threonine residue. After each glutamylation reaction, the flexible core peptide then rapidly moves to the elimination domain, where the glutamylated residues are converted into dehydro derivatives upon glutamate elimination. The orientation of the glutamylation and elimination active sites in one monomer suggests that the dimer, observed both in solution and in the crystal, is required for enzymatic activity. Distance constraints would make it necessary for the glutamylation step to occur at the active site for glutamylation of one monomer whereas the elimination step would occur at the active site of the elimination domain of the other monomer (Figure 18). The dehydration of serine or threonine residues within the propeptide (generically called LanA) is not a random process. In the pathways studied, the dehydration events occur in an organized directional scheme that differs between lanthipeptides. In nisin 9 biosynthesis, the dehydration of NisA by NisB occurs in a roughly N- to C-terminal order.133,134 As for MibA, after initial dehydration at the N-terminus, the MibB-catalyzed reaction has been shown to proceed mostly in a C- to Nterminal direction.48 3.1.2. LanBs Are Partners of a Multiprotein Complex. There is substantial evidence to suggest that LanB is part of a multiprotein complex in vivo that also includes the LanC and LanT enzymes and is located at the cytoplasmic membrane.135−137 Extensive genetic and bioengineering studies of nisin biosynthesis have shown that each partner in the

dependent phosphorylation of Ser/Thr hydroxyl groups followed by elimination of the phosphate. The activation strategy observed in class I lanthipeptide synthesis pathways based on the use of glutamate is less common, although a few examples have been reported.127 The glutamate itself must be activated to form an ester bond with the hydroxyl group of a serine or threonine. Biochemical studies on NisB and MibB have shown that dehydratases do not activate glutamate using ATP or another component, instead using the amino acid in the activated GlutRNAGlu form.39,48 They do not catalyze the formation of GlutRNAGlu per se, but hijack the Glu-tRNAGlu generated by GluRS in the cell that is generally used for ribosomal protein synthesis. Resolution of the three-dimensional structures of NisB and MibB showed these proteins to have a multidomain organization (Figure 17). They have dimeric structures, consistent with observations in solution.128,129 Each monomer is curved and consists of two domains separated by a central cleft. Each domain catalyzes a different step of the dehydration reaction. The multidomain N-terminal region, which consists of 700 to 800 amino acids (706 for NisB39 and 809 for MibB48), catalyzes the first glutamylation step. The C-terminal 300-residue domain catalyzes the elimination step. The mechanism involved has not been characterized at the molecular level, but residues essential for the glutamylation and elimination activities have been identified by analyses of sequence conservation patterns within LanBs and mutagenesis studies.39,127 Five residues have been shown to be involved in the elimination reaction (Arg784, Arg786, Glu823, Arg826, and His961; NisB numbering). All are clustered in the same solvent-accessible area of the C-terminal elimination domain, which constitutes the active site for elimination. Six residues have been identified as critical for the glutamylation reaction (Arg83, Arg87, Thr89, Asp121, Asp299, and R464; NisB numbering). All are located within a 10 Å radius, and they form a catalytic pocket in the inner face of the cleft-like structure of the glutamylation domain. A densely basic patch, large enough to accommodate the acceptor stem of the GlutRNAGlu cosubstrate, surrounds this pocket. A docking model for NisB binding to a tRNA has been proposed by Ortega et al.39 and is compatible with the presence of glutamate close to the 5591


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Figure 17. (a) Overall structure of the homodimer NisB in complex with its substrate, NisA (PDB ID, 4WD9). The glutamylation domain of the first monomer is shown in gray, with the peptide-binding site of this domain in blue and the NisA peptide in violet. The elimination domain is shown in light green. The glutamylation domain of the second monomer is shown in yellow, and the elimination domain is in orange. (b) Overall structure of one monomer. For visualization purpose, the overall structure has been rotated compared to that shown in panel a. Glutamylation and elimination active sites are surrounded. Residues essential for each activity appear in zoom-like insets.

cosubstrate Glu-tRNAGlu.39,48 However, for both nisin 9 and microbisporicin 10, the in vitro synthesis of the native pattern of dehydrations requires the simultaneous presence and activity of the related cyclase LanC and the dehydratase LanB. In the absence of MibC, monoglutamylated and partially dehydrated MibA intermediates, corresponding to late-stage glutamylated species, accumulate during the dehydration assay. Addition of MibC to the reaction mixture suppresses these intermediates, consistent with the model, according to which thioether rings must be formed in MibA for the final glutamate elimination reactions to be performed efficiently.48 The formation of the methyl-lanthionine ring of NisA involving residues 25 and 28 prevents the dehydration of Ser29, which remains intact in native nisin. In the absence of NisC, the nine Ser/Thr residues of the core sequence of NisA can be modified by NisB.39,133 The available data support a model in which the formation of thioether rings during lanthipeptide synthesis is a cooperative process involving the iterative alternation of LanB-dependent dehydration and LanC-dependent cyclization.48,133 3.1.3. To Be or Not To Be Substrate. The presence of a leader peptide is essential for propeptide interaction with LanBs.123 In particular, the conserved Phe-Asp/Asn-Leu-Asn/ Asp box within the leader peptide has been shown to be a key

Figure 18. Proposed mechanism for the dehydration of the LanA propeptide catalyzed by LanB.48 (a) Glutamylation reaction involving monomer A. The glutamylation and elimination domains of monomer A (GA and EA, respectively) are in blue and dark blue, respectively. The glutamylation and elimination domains of monomer B (GB and EB, respectively) are in green and dark green, respectively. The leader peptide and core region of LanA propeptide are in purple and red, respectively. The leader peptide-binding domain of GA is in light blue. Active sites are displayed with stars. (b) Subsequent elimination step involving monomer B.

biosynthetic machinery can function independently.39,129,138−140 The recent in vitro biochemical characterization of NisB and MibB conclusively demonstrated that both could perform several dehydration reactions on their respective propeptide substrates in the absence of any partner other than the appropriate 5592


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Figure 19. Gene organization of the goadsporin and thiopeptide clusters. Goadsporin 14 (god, from Streptomyces sp. TP-A0584),156 thiomuracins (tpd, from Nonomurea sp. Bp3714−39, and tbt, from Thermobispora bispora),49,151 GE2270 (pbt, from Planobispora rosea),49,151 thiocillin (tcl, from Bacillus cereus ATCC 14579),148 GE37468 (get, from Streptomyces sp. ATCC 55365),150 lactazole (laz, from Streptomyces lactacystinaeus),146 TP-1161 (tpa, from Streptomyces TFS 65−07),149 cyclothiazomycin (clt, from Streptomyces sp. NR0516),152 nosiheptide 12 (nos, from Streptomyces actuosus),153 nocathiacin (noc, from Nocardia sp. ATCC 202099),154 siomycin (sio, from Streptomyces sioyaensis),147 and thiostrepton 13 (tsr, from Streptomyces laurentii ATCC 31255).147 Candidate genes for the formation of dehydroamino acids, azoles, and central heterocycles are indicated in color. The structural series of the thiopeptide encoded by the cluster is indicated in parentheses.

the first serine or threonine processed.39 Ortega et al. studied the involvement of the leader peptide in the glutamylation and elimination activities of NisB separately. They found that the leader peptide of NisA was essential for glutamylation, but not for elimination,39 which suggests that the local structure of glutamylated nisin is sufficient for binding and processing by the elimination domain. Provided that an appropriate leader peptide is present, LanBs can display a relaxed specificity for their cognate substrates. This property made it possible to use NisB to introduce dehydroamino acids into biological peptides totally unrelated to lanthipeptides, simply by fusing them to the nisin leader sequence.142 The features governing NisB specificity have been studied in detail. The impact of the positions of serine or threonine relative to the leader peptide for dehydration reactions has been investigated, and the relative positions of these residues were found to have no impact on dehydration efficiency.143 NisB can also dehydrate serine or threonine located further away from the leader peptide than position 33, corresponding to the most C-terminal serine residue of nisin.144 Hydrophobic residues flanking dehydratable serine or threonine residues appear to favor dehydration more than hydrophilic residues.144 However, there is no hard and fast rule. NisB, for example, was found to dehydrate multiple flanking threonine residues efficiently.

element for substrate recognition.128,129,141 Consistent with this finding, determination of the structure of the NisB:NisA complex implicated this motif in a strong network of hydrophobic interactions with the peptide leader binding domain of NisB.39 According to this structure, many modifications of the leader sequence away from this motif would be possible without impeding the activity of LanBs. Extensive mutagenesis analyses have been carried out on the nisin leader peptide. Plat et al. engineered different cleavage sites in the C-terminal part of the molecule.141 The specificity of the dedicated protease, NisP, is very narrow, preventing its use for the production of peptides other than nisin. The NisP cleavage site was thus replaced with sites for thrombin, enterokinase, Glu-C, and factor Xa cleavage. A His-tag was also introduced by replacing residues located in the N-terminal part of the leader peptide or just behind the highly conserved box, without impairing the dehydratase activity of NisB.141 Finally, decreasing the length of the leader sequence by more than half (retaining only the 11 N-terminal residues) does not prevent dehydration of the corresponding prenisin variant. However, it almost halved the number of dehydrations of the final product.141 The recent determination of the structure of the NisB:NisA complex provided an explanation for this observation: a minimum distance between the leader peptide-binding domain and the glutamylation active site must be respected for 5593


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two-domain protein, LazF, which has an N-terminal TpdC domain and a C-terminal flavin mononucleotide-dependent dehydrogenase domain. This suggests that LazF may function as a bifunctional enzyme, catalyzing both the glutamylation elimination reaction and the azoline dehydrogenation driven by the C-domain. The various TpdBs characterized generally display between 20 and 40% sequence identity. However, higher levels of identity are observed between TpdBs involved in the synthesis of thiopeptides from the same structural series. Indeed, thiopeptides are classified into five different structural seriesa, b, c, d, and eaccording to the oxidation state of the central nitrogenous heterocycle116 (Figure 14). The a series exhibits a totally reduced central piperidine, whereas the b series is further oxidized and presents a 1,2-dehydopiperidine ring. The d series shows a trisubstituted pyridine ring. As to the e series, it carries an additional hydroxyl group on the central pyridine, now tetrasubstitued. NosE and NocE, both of which are involved in the synthesis of series e thiopeptidesnosiheptide 12 and nocathiacin, respectivelydisplay 55% sequence identity. SioJ and TsrJ, which are involved in the synthesis of series b thiopeptidessiomycin and thiostrepton 13, respectively display 74% sequence identity. However, TbpBs and TpdB1s, from the same clusters, display only moderate sequence identity (about 30%), consistent with a difference in the specificity in the biosynthesis pathway. Levels of sequence identity with the glutamylation domain of LanBs are very low, usually less than 10%. Comparisons of TpdC sequences reveal similar trends. 3.2.2. What We Know about the Mechanism of Split LanBs: TpdB and TpdC. Two different TpdB/TpdC pairs have been characterized in the last year: the TbtB/TbtC pair from Thermobispora bispora, which is involved in the synthesis of thiomuracin 11,49 and the GodF/GodG pair from Streptomyces sp. TP-A0584, which is involved in the synthesis of goadsporin 1450 (Figure 14). The first pair was studied in vitro whereas the second pair was studied in vivo, in an approach combining the analysis of disruptants and the structural characterization of products. Thiomuracin 11 is a series d thiopeptide that blocks bacterial translation by inhibiting the elongation factor EF-Tu.51 The synthesis of its core structure requires six proteins, including TbtB and TbtC, catalyzing 22 chemical transformations.49 Goadsporin 14 is a linear azole-containing peptide with structural similarities with thiopeptides. It has two thiazoles and four (methyl)oxazoles, and it is also decorated with two dehydrolalanines.157,158 It displays antibiotic activity (among other activities), although its precise biological target has yet to be identified.51 Both TbtB and GodF have been shown to catalyze the glutamylation of dehydratable serine residues. In thiomuracin 11 biosynthesis, the glutamylation reaction has been shown to involve Glu-tRNAGlu, originating from the aminoacyl-tRNA pool of the cell, as for LanBs.49 Recent works indicate that the dehydrations installed by the pair TbtB/TbtC are formed in a strict C-to-N-terminal direction.159 For goadsporin 14, the disruption of godG results in the accumulation of a monoglutamylated variant of goadsporin. NMR characterization of this species provided the first demonstration that the glutamyl group was attached to a serine via an ester linkage.50 MS−MS analyses of this species revealed the occurrence of an ordered dehydration process: there were two dehydratable residues, Ser4 and Ser14, but only one of these residues, Ser4, was glutamylated.

Recent studies have highlighted the role of thioether linkages in promoting or preventing the LanB-catalyzed dehydration of neighboring residues (see section 3.1.2 and refs 48 and 133). The only study of the interaction of LanBs with their second substrate Glu-tRNAGlu carried out to date focused on MibB.48 MibB was shown to function with tRNAs obtained by in vitro transcription, indicating that posttranscriptional modifications of tRNA bases are not required for the interaction of tRNAGlu with class I lanthipeptide dehydratases. Ortega et al. showed that MibB specifically recognized nucleotides of the acceptor stem (A73 and U72). The presence of a G in position 73 and a C in position 72 of the tRNAGlu sequence of E. coli prevents the use of this tRNA by MibB. This type of interaction is consistent with observations for other aa-tRNA-dependent enzymes, which interact mostly with the acceptor stems of their tRNA substrates.78,145 However, the specificity of recognition observed for MibB cannot be generalized to all LanBs. Lactococcus lactis contains a unique tRNAGlu isoacceptor. The sequence of the acceptor stem of this tRNA in the region shown to be essential for interaction with MibB is identical to that of the corresponding tRNA from Microbispora. However, NisB is equally able to use the tRNAGlu of E. coli and that of L. lactis,39 suggesting that other types of interaction are involved in the interaction of NisB and its cosubstrate Glu-tRNAGlu and in the possible specificity of NisB. 3.2. Split LanBs in Thiopeptide and Goadsporin Biosynthesis

Protein similarity searches have revealed that LanB-like dehydratases are present in the biosynthesis clusters of thiopeptides and goadsporin 14 (Figure 14).39 In these clusters, the dehydratases are split into two separate polypeptides. The longer polypeptide, which is about 800 residues long, displays sequence similarity to the glutamylation domain of LanBs whereas the smaller polypeptide, which is about 300 residues long, is similar to the elimination domain. Using the standard nomenclature proposed by Arnison et al.,51 the enzyme homologous to the glutamylation domain will generally be referred to here as TpdB (thiopeptide synthesizing enzyme B), whereas that homologous to the elimination domain will be referred to as TpdC. The original names have nevertheless been retained in the characterized clusters. 3.2.1. Characteristics of TpdBs and TpdCs in the Clusters Described. The mature thiopeptide is generated from a 50−60 amino acid preprotein consisting of an N-terminal leader peptide (40−50 amino acids) and the 14−18 amino acid C-terminal core peptide, which undergoes various posttranslational modifications. About 15 different biosynthetic gene clusters had been described by early 2016146−156 (Figure 19). TpdB and TpdC belong to a group of six genes unique to thiopeptide biosynthesis and involved in formation of the central six-membered heterocycle (Figure 14).147 They are generally contiguous, with TpdB preceding TpdC, as found for the organization of the two domains of LanBs. In addition to this standard gene organization, several clusters present unusual features. In three of the clusters characterized (siomycin, thiostrepton 13, and GE2270), there is a gene encoding an additional protein similar to the glutamylation domain of LanBs.147,151 The presence of this additional putative glutamylation protein, generically named TpdB1, suggests that TpdB and TpdB1 may have different patterns of specificity for the synthesis of their cognate thiopeptide, with both able to work with the same partner for the elimination step. Another unusual feature is found in the biosynthesis cluster of lactazole, in which the tpdB and tpdC genes are not contiguous.146 TpdC is part of a 5594


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Analyses of the synthesis intermediates have shown that monoglutamylated species accumulate in the absence of TpdC, for both thiomuracin 11 and goadsporin 14. Diglutamylated species are found in trace amounts. This suggests that the glutamylation and elimination reactions alternate, as observed in reactions catalyzed by LanBs.49,50 The elimination of the glutamyl residues initially formed is required for the next glutamylation step to occur. HHpred searches for enzymes of both the thiomuracin 11 and goadsporin 14 clusters50 revealed that these TpdBs and TpdCs were related to NisB, with a high probability score, allowing the

Figure 21. Proposed pathway for the synthesis of the core scaffold of thiopeptides from ref 49. Dehydration reaction catalyzed by the TpdB/ TpdC pair (dehydrated products are represented by green double bars) requires the prior formation of the thiazoles (represented as blue pentagons) through the combined action of TpdE/TpdF/TpdG. TpdD catalyzes the formation of the central heterocycle (represented as a red hexagon) from two dehydroalanine residues and the concomitant release of the leader peptide.

3.2.3. What We Know about Specificity. Disruptants of godF and godG accumulate a derivative of goadsporin in which the two serine residues are unmodified. Assays of the bioconversion of this derivative into goadsporin by a godA disruptant strain (deficient in the production of the precursor peptide, but containing intact modification enzymes) were unsuccessful.50 A problem with incorporation of the molecule cannot be excluded, but the absence of the leader peptide in the derivative may also explain the lack of dehydration. No threedimensional structures are available for TpdB or TpdC enzymes, but Burkhaert et al.130 identified, by bioinformatics techniques, a leader peptide-binding domain in TpdBs that was localized in the same region as in LanBs. No such binding domain is found in TpdCs. Recent work of van der Donk, Mitchell, and co-workers performed on the thiomuracin pathway showed however that the putative leader peptide-binding domain of TbtB is not functional and that TbtB recognizes instead a specific conformation of the core peptide resulting of posttranslational modifications.159 The specificity of TpdBs for the core peptide begins to be unraveled. The presence of specific functional groups is required. In the case of thiomuracin, TbtB cannot transform the precursor peptide, TbtA. Complete dehydration occurred only after the formation of the six thiazole rings in TbtA49 (Figure 21). Thorough analysis of a panel of TbtA variants has recently revealed that the pair TbtB/TbtC recognizes a specific conformation of the TbtA core peptide induced by the formation of the thiazole at Cys10.159 Similar conclusions were drawn in studies of the enzymes of the goadsporin cluster: the presence of azole groups is probably required for the dehydration of serine residues by GodF/GodG.50 Concerning the specificity of TpdB for its cosubstrate GlutRNAGlu, experiments performed on thiomuracin 11 have indicated a strong similarity to the situation for MibB.49 TbtB does not accept the Glu-tRNAGlu of E. coli as a cosubstrate, and it displays a strong preference for a particular isoacceptor tRNAGlu from T. bispora. Efficiency is highest for this cosubstrate, which differs from other potential cosubstrates by the presence, among others, of an A at position 73 and a U at position 72. However, no systematic study was performed to identify the tRNA bases important for recognition by TbtB. As for MibB, posttranslational modifications to the tRNA are not required for their use by TbtB: all the tRNAs used in the study were produced by in vitro transcription.

Figure 20. Sequence alignment of selected domains of TpdBs (displaying similarity to the glutamylation domain of LanBs) and TpdCs (similar to the elimination domain of LanBs), based on the results of HHpred. The residues essential for the catalytic activity of NisB (red) are labeled above the alignment and conserved in all the sequences, with the exception of Asp299, which may be replaced by a glutamate residue in some TpdBs (green). The biochemically characterized TbdB/TpdC pairs are labeled in bold. TpdB1 enzymes are labeled with a star. Thiocillin (Tcl);148 goadsporin;156 lactazole (laz);146 cyclothiazomycin (clt);151 nocardithiocin (not);155 GE37468 (get);150 GE2270 (pbt);151 thiomuracin (tbt)49 (tpd);151 TP-1161 (tpa);149 nosiheptide (nos);153 nocathiacin (noc);154 thiostrepton (tsr) and siomycin (sio).147

alignment of the two domains of NisB with the TpdBs/TpdCs of the identified clusters (Figure 20). No mutagenesis studies have been performed for TpdBs and TpdCs, but these alignments show that most of the residues identified as forming the active site of the glutamylation domain and all the residues involved in the elimination step are conserved in split LanBs. Taken together, these studies strongly suggest that the dehydration reaction typical of goadsporin 14, thiopeptides, or lanthipeptides I is performed in a similar manner with the twodomain full-length LanBs or with two single-domain split LanB enzymes, TpdBs and TpdCs. 5595


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3.3. “Small LanB” Proteins in Biosynthesis Clusters

Fem transferases required for biosynthesis of the cell wall peptidoglycan.21,166−168 More recently, aaPGSs have been shown to constitute a third family of bacterial aa-tRNAdependent transferases involved in modifying membrane phospholipids for resistance to cationic antibiotics.169 Proteins from these three families use aa-tRNAs of different types (Leu/ Phe-tRNAs for L/F transferases, Gly/Ala/Ser-tRNAs for Fem transferases, and Lys/Ala-tRNAs for aaPGSs) to transfer the aminoacyl moiety to one amino group in the peptide part of their substrate (L/F transferases and Fem transferases) or to one hydroxyl group of the polar head of phosphatidylglycerol (aaPGSs), resulting in the formation of a peptide or amide bond and an ester bond, respectively (Figure 22). The crystal structures of several full-length or truncated members of each family have been determined. They showed that the catalytic domains of these proteins, despite displaying low levels of amino acid sequence identity, adopt the same GCN5-related N-acetyltransferase (GNAT) fold. GNAT foldcontaining proteins constitute a superfamily of more than 10,000 members participating in multiple physiological functions through the acylation of their cognate substrates.41,42,170 The Protein Data Bank (RCSB PDB) currently contains about 200 different proteins harboring the GNAT fold, and the proteins within this database displaying the greatest structural similarity to aaPGSs and L/F transferases are Fem transferases,23,171,172 suggesting the use of a very similar fold for aa-tRNA-dependent transferase activities. The presence of GNAT family annotated

This category includes proteins displaying similarity to the glutamylation domain of LanBs for which no putative elimination protein is encoded by the surrounding genes, contrary to TpdBs. These proteins have been identified in bioinformatics studies, but none has been studied experimentally as yet. These small LanBs are often found in nonribosomal peptide synthesis clusters.39 The absence of an elimination partner suggests that they may add, in a tRNA-dependent manner, an amino acid to a growing peptide synthesized by an NRPS. Most small LanBs are stand-alone polypeptide chains, but Zhang et al.160 identified a gene cluster containing a small LanB within an NRPS module, together with an adenylation domain and a peptidyl carrier protein. This finding supports the hypothesis that small LanBs are involved in the assembly line of certain NRPS biosynthetic pathways. The putative amino acid incorporated by these enzymes and their catalytic mechanism remain to be determined.

4. aa-tRNA-DEPENDENT TRANSFERASES aa-tRNA-dependent transferase activities were first identified in bacteria about 50 years ago.161−165 Intensive biochemical characterizations subsequently identified model enzymes with such activities and the physiological functions they supported, revealing the existence of a family of leucyl/phenylalanyl-tRNA protein transferases (L/F transferases) involved in the N-end rule pathway for protein degradation in bacteria, and the family of

Figure 22. Activity of selected aa-tRNA-dependent transferases. (a) L/F transferase of E. coli. R is a polypeptide chain. (b) FemX and FemA enzymes of S. aureus, FemXSa and FemASa, respectively. R is N-acetylmuramic acid linked to undecaprenyl lipid. (c) Lys-PGS of S. aureus. R is (CH2)7(CH)2(CH2)7CH3. 5596


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However, it was not possible to determine the chemical structure of this compound unequivocally because of its chemical instability, resulting in the generation of N-(L-seryl)-isobutylhydroxylamine 33. NMR experiments suggested that this compound was O-(L-seryl)-isobutylhydroxylamine 32, consistent with the known chemical instability of O-acyl hydroxylamines, resulting in the generation of N-acyl hydroxylamines.33,182 The subsequent steps in the valanimycin biosynthetic pathway are less well documented. The proposed pathway is based on the NMR determination of the chemical structure of valanimycin hydrate 34, an intermediate that accumulates in vlmJ and vlmK mutants and has been shown to be converted into valanimycin in vlmH mutants.183 However, the pathway for valanimycin hydrate biosynthesis remains unknown. Feeding experiments with purified valanimycin hydrate 34 and protein annotation have provided support for the proposed roles of VlmJ and VlmK. Finally, one of the proposed regulatory proteins, VlmI, belongs to the family of Streptomyces antibiotic regulatory proteins and is essential for valanimycin production, due to its role in activating the transcription of vlmA, vlmHORBCD, and vlmJKL.184 The aa-tRNA-dependent transferase VlmA is a 338-residue protein, the function of which was not deduced primarily from its amino acid sequence. Indeed, Parry and co-workers focused on VlmA because of the lack of incorporation of radioactivity from L[U-14C]seryl-tRNA into new compounds in precursor incorporation experiments with vlmA mutants.33 They noted that VlmA displayed low levels of sequence identities with aaPGS and investigated the activity of VlmA in coupled activity assays using E. coli tRNA and the Ser-tRNA synthetase VlmL.33 It is questionable how the activity of VlmA is dependent on the presence of VlmL. Indeed, disruption of the vlmL gene by a single-crossover insertion significantly decreased valanimycin production.33 Furthermore, VlmL displays moderate sequence identity (39%) to the Streptomyces viridifaciens housekeeping SerRS, SvsR, particularly in the N-terminal region involved in tRNA binding.181 Parry and co-workers suggested that this Nterminal region of VlmL might also interact with other proteins involved in valanimycin biosynthesis. The spatial proximity of VlmA and VlmL might favor the efficient use of Ser-tRNA by VlmA. Sequence-based clues to the presence of aa-tRNA-dependent transferases in secondary metabolite biosynthesis were identified later, as described below for phosphonopeptides and uridyl peptide antibiotics. However, we thought it would be interesting to carry out an up-to-date inspection of the VlmA sequence with the tools currently available (June 2016). HHpred searches88,89 performed with the VlmA amino acid sequence retrieved, as best hits, with high probability scores, aaPGS followed by FemXWv and FemASa, consistent with conservation of the GNAT fold. However, searches for conserved domains of VlmA in the NCBI database retrieved only the DUF2156 superfamily (pfam09924 entry), which contains the C-terminal domain of aaPGS. BLAST searches for VlmA homologues in the NCBI database retrieved about 120 entries displaying more than 50% sequence identity to VlmA, including seven entries with identity levels >60%. For the five best hits, the putative VlmA genes were associated with genes encoding proteins very similar to those involved in valanimycin biosynthetic pathways. However, some proteins required for valanimycin biosynthesis were missing, and the entire cluster could give rise to novel azoxy compounds. An original chromosomal organization was found in one species of Frankia, with a 30 kb region bordered by genes encoding VlmL and VlmA

proteins in biosynthetic gene clusters for peptide secondary metabolites recently led several teams to investigate the tRNAdependent biosynthesis of these metabolites. Over the past 10 years, the involvement of aa-tRNAs and their cognate GNAT superfamily transferases was demonstrated in the biosynthesis of four different types of secondary metabolites, the azoxy compound valanimycin, phosphonopeptides, uridyl peptide antibiotics, and compounds related to the antibiotic streptothricin,33,36,37,40 and was postulated for others.38,173 The biochemically characterized tRNA-dependent biosynthetic pathways are described below. They highlight the diversity of means by which the peptide chains of secondary metabolites are synthesized in natural conditions. 4.1. Biosynthesis of the Azoxy Compound Valanimycin: The First Involvement of aa-tRNAs in Secondary Metabolite Biosynthesis To Be Described

Valanimycin 22 belongs to a family of natural products that carry the rare azoxy function but are only poorly related in terms of their chemical structure (Figure 23). Only a few azoxy NPs have been described. Most of these compounds originate from Streptomyces bacteria, with the exception of lyophillin 25, which is produced by the fruiting bodies of the fungus Lyophyllum connatum. Valanimycin 22 was isolated from Streptomyces viridifaciens MG456-hF10 during the studies seeking to identify potential antitumor antibiotics.174 It has weak antibacterial activity and can prolong the life of mice inoculated with carcinoma cells, although it is toxic at high doses. The biosynthesis of valanimycin 22 has been studied for about 30 years. Initial enzymatic and genetic investigations175−179 led to the isolation from Streptomyces viridifaciens MG456-hF10 of a 14-gene cluster, the presence of which in Streptomyces lividans TK24 is sufficient for valanimycin production180 (Figure 24a). These initial investigations and subsequent studies33,181−183 allowed establishing an incompletely deciphered valanimycin biosynthetic pathway (Figure 24b). The presence of the SertRNA synthetase VlmL, whose activity was confirmed in vitro,181 together with the known requirement of L-serine for valanimycin biosynthesis,176 led Parry and co-workers to investigate the role of seryl-tRNA.33 By feeding S. viridifaciens mutants with labeled seryl-tRNA, they identified VlmA as essential for the incorporation of radioactivity into intermediates of valanimycin biosynthesis. In vitro characterization of the activity of the purified recombinant VlmA demonstrated the use of Ser-tRNASer and isobutylhydroxylamine 31 to form a new compound.

Figure 23. Chemical structures of selected naturally occurring azoxy compounds. The azoxy group is shown in magenta. 5597


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Figure 24. Biosynthesis of valanimycin 22. (a) Organization of the valanimycin biosynthetic gene cluster. Genes are drawn according to NCBI entry AY116644.180 They are colored according to the proposed functions of the encoded proteins inferred from bioinformatic analyses and experimental data. (b) Proposed biosynthetic pathway of valanimycin 22. The protein names are shown in color, according to their proposed functions in panel a. The dashed box encloses steps for which catalysis has not been demonstrated in vitro.

Figure 25. Selected synthetic and natural phosphonopeptides. The methyl 2-hydroxy-2-phosphonoacetate (MeHPnA) moiety of fosfazinomycin A and B is highlighted in fuchsia.

attracted considerable attention in attemps to develop bioactive molecules stably mimicking phosphate esters and carboxylic acids inhibiting enzyme activities. Phosphonopeptides combine a peptide backbone with one or several C−P bonds. They generally have a phosphoryl group replacing the C-terminal carboxyl group of a small peptide chain, but some variations may exist (Figure 25). The first phosphonopeptide to be described, alaphosphin 35, was chemically synthesized in the 1970s and was developed during the search for novel inhibitors of bacterial cell wall biosynthesis.186 Phosphonopeptides have since been identified among NPs, but their production in natural conditions is limited and restricted to a few bacteria.185,187 Most known phosphonopeptides were originally isolated during screening for bioactive compounds with diverse activities. The useful proper-

homologues (56% and 63% sequence identity, respectively) and carrying some vlm gene homologues plus genes encoding modules of polyketide synthases. Valanimycin is probably not the only azoxy compound for which biosynthesis is dependent on aatRNAs. 4.2. The tRNA-Dependent Biosynthesis of Phosphonopeptides

Phosphonopeptides belong to the family of phosphinic and phosphonic acid compounds (C−P compounds) which are chemically characterized by the presence of carbon−phosphorus bonds. Since the first identification of a naturally produced C−P compound in 1959, many different C−P compounds have been identified in archaea, bacteria, and eukaryotes, in which they perform diverse functions.185 These compounds have also 5598


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Figure 26. Mode of action and biosynthesis of DHP. (a) DHP metabolism within bacteria. (b) Organization of the DHP biosynthetic gene cluster. Gene organization is drawn in accordance with GenBank accession number GU199252.196 Genes are colored according to the proposed functions of the encoded proteins inferred from bioinformatic analyses and experimental data. DhpH is a two-domain protein. (c) Proposed biosynthesis pathway for DHP. The names of the proteins are colored according to their proposed functions in panel b. The box outlined with a dashed line indicates steps for which no experimental evidence has been obtained that nevertheless appear to be the most plausible reactions on the basis of the predicted features of the proteins and the other identified steps. DhpH-N and Dhp-C correspond to the N- and C-terminal domains of DhpH, respectively (residues 1−356 and 353−698). DhpY is an unidentified protein not encoded in the DHP biosynthetic gene cluster that may be required for the conversion of OP-EP to AP. α-KG, α-ketoglutarate; PLP, pyridoxal 5′-phosphate; SAM, S-adenosylmethionine; PEP, phosphoenolpyruvate; PnPy, phosphonopyruvate; PnAA, phosphonoacetaldehyde; 2-HEP, 2-hydroxyethyl phosphonate; DHEP, 1,2-dihydroxyethyl phosphonate; HP-EP, 1-hydroxy-2-phosphorylethyl phosphonate; OP-EP, 1-oxo-2-phosphorylethyl phosphonate; pSer(P), phosphoserine phosphonate; AP, acetyl phosphonate; L-Ala(P), L-1-aminoethyl phosphonate; SAHC, S-adenosylhomocysteine.

ties of these molecules and the interest aroused by the mechanism of C−P bond formation triggered considerable interest in the biosynthesis of phosphonic and phosphinic acids, including phosphonopeptides. Investigations of these pathways revealed unusual enzymology, not only for C−P bond formation but also for the synthesis of the skeletons of C−P compounds.185,187,188 The unusual enzymes observed include the recently discovered aa-tRNA-dependent transferases in the dehydrophos 37 (DHP) biosynthetic pathway.37 Two different tRNA-dependent-transferase activities are involved in DHP biosynthesis, one of which is performed by a bifunctional enzyme. We present this pathway, the biochemical characterization of which is almost complete, focusing on transferases. We then discuss studies suggesting the involvement of aa-tRNA-

dependent transferases in other phosphonopeptide biosynthetic pathways. 4.2.1. The Biosynthetic Pathway of Dehydrophos. Dehydrophos 37 (DHP) is a phosphonopeptide antibiotic originally isolated from Streptomyces luridus NRRL 15101189 (formerly known as A53868 factor A). The structure of DHP has been revised several times, and is currently considered to consist of a Gly-L-Leu dipeptide, the C-terminus of which is linked by a peptide bond to an O-methylated vinyl phosphonate190 (Figure 25). DHP has broad-spectrum antibacterial activity against Gram-negative and Gram-positive bacteria.189 Structure−activity relationship studies191 and analyses of DHP-resistant bacteria192 have led to a suggested mode of action for DHP. The antibacterial activity of DHP depends on its metabolism within 5599


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that DhpK catalyzes the last step in DHP biosynthesis. It should be noted that DhpI activity was evaluated with the tripeptide GlyL-Leu-L-Ala(P) rather than the dipeptide L-Leu-L-Ala(P) 59. However, this enzyme was shown to have low substrate specificity, with selectivity for phosphonate analogues of alanine. Two proteins, DhpH (698 residues) and DhpK (422 residues), use charged tRNAs to add single amino acids to phosphonate units. A role for these two proteins in amino acid transfer was suspected on the basis of their N-acyltransferase domains, as previously reported for several amino acid transferases using charged tRNAs as substrates. DphH was predicted to be a bifunctional protein, with an N-acyl transferase domain in the C-terminal part of the protein (DhpH-C, residues 353−698). HHpred searches with DhpH-C and DhpK identified FemXWv and FemASa as the best hits, with a high probability score. The purification from E. coli of recombinant DhpH or DhpH-C was accompanied by the copurification of nucleic acids identified in aminoacylation assays as tRNALeu, suggesting that DhpH was responsible for incorporating the leucyl residue into the phosphonate unit. Further experiments established the LeutRNALeu-dependent activity of DhpH and DhpH-C with LAla(P) 58 to generate L-Leu-L-Ala(P) 59. The full-length and Cterminal DhpH molecules copurified with tRNAs and had similar activities, suggesting that the N-terminal domain of DhpH played no role in tRNA binding and transferase activity. Moreover, no nucleic acids were copurified with DhpH-N (DhpH-N; residues 1−355), indicating that only the C-terminal domain of DhpH was able to bind tRNAs. The association, in a single polypeptide, of aa-tRNA-dependent transferase activity and a domain characteristic of PLP-dependent aspartate aminotransferases (DhpH-N) has raised a number of questions. Van der Donk and co-workers suggested that the activities of the two domains may be coupled to ensure the efficient conversion of an unstable intermediate generated by DhpH-N from pSer(P) 56.37 However, their subsequent experiments did not confirm this hypothesis and even demonstrated a role for DhpH-N in acetylphosphonate AP 57 biosynthesis. The nature of the substrate of DhpH-N remains unknown. The reason for the presence of a bifunctional DhpH enzyme in the DHP biosynthesis pathway, if indeed there is one, remains to be determined. Despite the predicted structural similarity between DhpK and DhpH-C, these two proteins display only 16% sequence identity. DhpK was produced in E. coli as an N-terminal maltose binding protein fusion, because the unfused protein was not sufficiently soluble. It was shown to add a Gly residue to the N-terminus of phosphonate units in an aa-tRNA-dependent manner. It is active on L-Leu-L-Ala(P) 59, L-Leu-ΔAla(P), and LLeu-ΔAla(P)-OMe 61. No conversion was observed when DhpK was incubated with L-Ala(P) 58 and the Gly-tRNAGly regeneration system. Blast searches of the NCBI database (June 2016) for DhpH and DhpK homologues identified three Streptomyces strains, NRRL S-118, NRRL S-920, and S. alboniger, with genomes harboring biosynthetic gene clusters encoding DhpA-DhpL homologues displaying 94−99%, 58−89%, and 58−89% sequence identity, respectively, to the query sequences. A partial phosphonopeptide biosynthetic gene cluster was also found in Streptomyces sp. NRRL F-3213, and the encoded DhpA-H and DhpJ homologues displayed 54−76% identity to the query sequences. However, even though the final products were predicted to possess a scaffold similar to DHP, their chemical structures may be slightly different from that of DHP, as the corresponding biosynthetic gene clusters encoded additional

bacteria by peptidases releasing 1-aminovinyl methyl phosphate 46 (Figure 26a). This unstable compound undergoes tautomerization and hydrolysis to generate methyl acetylphosphonate 48 (MAP), an inhibitor of the essential bacterial enzymes pyruvate dehydrogenase193 and 1-deoxy-D-xylulose 5-phosphate synthase.194,195 The Gly-L-Leu N-terminal dipeptide of DHP appears to be essential for antibacterial activity, facilitating uptake into the cell: MAP 48 has no antibacterial activity against E. coli and Bacillus subtilis unlike DHP 37,191 and DHP-resistant mutants of Salmonella enterica had mutations mapping to genes encoding nonspecific oligopeptide permeases.192 Metcalf and co-workers isolated the DHP biosynthetic gene cluster by screening a fosmid library of S. luridus NRRL 15101 for a gene encoding a phosphoenolpyruvate mutase,196 an enzyme previously shown to catalyze the first step required for the formation of the C−P bond of several phosphonate biosynthetic pathways.197−199 They identified a 16-gene cluster, dhpA-P, required for DHP production in a heterologous host.196 This cluster included genes required for DHP biosynthesis (dhpA-K), but also genes encoding membrane proteins DhpL-N thought to play a role in transport and/or resistance, and the dhpO dhpP genes encoding putative proteins involved in transcriptional regulation (Figure 26b). Bioinformatics analyses of the predicted proteins of the DHP biosynthetic cluster, analyses of the metabolic intermediates detected following the knockout of dhp genes in recombinant strains carrying the DHP cluster, and in vitro activity assays with purified Dhp proteins have made it possible to establish most of the reactions catalyzed by DhpA-K in the DHP biosynthetic pathway37,196,200,201 (Figure 26c). The first three steps, catalyzed by DhpE, DhpF, and DhpG, respectively, mediate the synthesis of 2-hydroxyethylphophonate 52 (2-HEP) from phosphoenolpyruvate 49 (PEP) and are common to many phosphonate biosynthetic pathways. The hydroxylation of 2-HEP 52 to generate DHEP 53 has been demonstrated both in vitro and by gene knockout experiments in vivo. A role for DhpB in DHEP 53 phosphorylation was suggested by both the presence of phosphorylated DHEP (HPEP 54) in culture supernatants of dhpC mutants and the presence of only DHEP 53 in dhpB mutants. These findings are consistent with the bioinformatic annotation of DhpB as a glycerate kinase. The next step, catalyzed by DhpC, was inferred from the intermediates accumulating in dhpC mutants and the similarity of DhpC to malate and lactate dehydrogenases. However, the subsequent step has remained more elusive, because of the detection in dhpH mutant culture supernatants of phosphoserine phosphonate pSer(P) 56,196 the biosynthesis of which from OPEP 55 has been shown not to be catalyzed by DhpD,37 a possible candidate for such transformation. However, pSer(P) 56 is a substrate for recombinant DhpH and DhpH-N (the N-terminal domain of DhpH comprising residues 1 to 356), and its incubation with DhpH or DhpH-N results in the appearance of acetylphosphate (AP) 57. The authors envisaged two possibilities.37 Either a protein not encoded in the DHP cluster was involved in pSer(P) 56 synthesis from OP-EP 55, or the DhpH N-terminal domain was able to use L-Ala to form a pyridoxamine phosphate from PLP, further contributing to the elimination of phosphate from OP-EP 55 to generate AP 57. The activities of the enzymes involved in the final steps leading to the production of DHP 37 from AP 57 have been clearly characterized in vitro, in assays with purified recombinant enzymes and chemically synthesized substrates.37,201 Given the selectivity of DhpJ for L-Leu-L-Ala(P)-OMe 60, it has been suggested that the methyl transferase DhpI acts before DhpJ, and 5600


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proteins not found in the dhpA-dhpP cluster of S. luridus NRRL 15101. The almost complete biochemical characterization of the DHP biosynthetic pathway has greatly added to our knowledge of C− P compound biosynthesis. In the past few years, van der Donk, Metcalf, and co-workers have investigated the diversity of the phosphonate biosynthetic pathways existing in natural conditions, by screening a large number of genomes and strains from different collections for phosphonate production. They used a dual approach, consisting of genome mining for PEP mutase genes, as PEP mutase catalyzes the first step of almost all phosphonate biosynthetic pathways and is highly conserved,188 and the detection, by 31P spectroscopy, of phosphonate compound production in cultures of selected strains. The C−P bond has unique chemical properties facilitating its detection even in complex mixtures.202 Two aa-tRNA-dependent biosynthetic pathways for novel phosphonopeptides were predicted and are considered highly likely, despite the lack of detailed biochemical characterization. We present these pathways below. 4.2.2. Putative tRNA-Dependent Biosynthetic Pathways for Other Phosphonopeptides. Fosfazinomycins A 41 and B 42 are antifungal phosphonopeptides containing a hydrazide bond between the carboxyl group of Arg and the phosphonate group of methyl 2-hydroxy-2-phosphonoacetate (Me-HPnA)203,204 (Figure 25). They were first isolated from Streptomyces lavendofoliae 630, and their presence was also recently detected in spent culture medium from Streptomyces sp. XY732, a bacterium with a phosphonate biosynthetic gene cluster.205,206 An almost identical phosphonate biosynthetic cluster is present in Streptomyces sp. WM6372, but attempts to detect fosfazinomycins A and B in culture medium from Streptomyces sp. WM6372 were unsuccessful. Instead, MeHPnA, a substructure of fosfazinomycins A and B, was detected and confirmed after purification and 1H and 13C NMR spectroscopy.206 Based on bioinformatics analysis, the authors described a 19-gene cluster from Streptomyces sp. WM6372 potentially responsible for fosfazinomycin A 41 synthesis. The recombinant expression of this biosynthetic cluster in S. lividans did not result in fosfazinomycin production, but the intermediate Me-HPnA was detected. The authors suggested that regulatory or metabolic limitations were responsible for the absence of fosfazinomycin production, as observed in the parental strain. The activities of the encoded proteins were not experimentally determined. Instead, they were deduced from database annotations and similarities to proteins of known function. The clusters from both WM6372 and XY732 each encoded a protein annotated as GCN5-related N-acetyltransferase with probable aa-tRNA-dependent transferase activity. The two proteins, FzmI from Streptomyces sp. WM6372 (orf10 in Figure S3 of ref 205 and accession number AGZ93904.1) and AGZ93765.1 from Streptomyces sp. XY732 (orf11 in Figure S2 of ref 205), are 426 and 399 residues long, respectively, and their sequences are 90% identical. HHpred searches with each sequence retrieved the FemXWv and FemASa proteins as best hits, with high probability scores. These proteins display 22−27% sequence identity to the transferase PacB involved in pacidamycin biosynthesis (see section 4.3), DhpH-C and DhpK. The role of these proteins in fosfazinomycin biosynthesis is unclear, and the authors suggested that they were involved in the formation of a bond between a nitrogen nucleophile and an acid electrophile. These include the two peptide bonds of fosfazinomycin A and the bond between the hydrazide group and the phosphorus atom (Figure 25). Two other encoded proteins

may be involved in the formation of these bonds: a member of the ATP GRASP enzyme family and a member of the glutamine synthase family. Further investigations are required to determine the roles of these proteins. Mining the genomes of a collection of 10,000 actinomycetes for phosphonate production led to the isolation of Streptomyces monomycini NRRL B-24309, a producer of two novel antibiotic phosphonopeptides called argolaphos A 44 and B 45207 (Figure 25). The structures of the purified argolaphos A and B were determined by 1H and 13C NMR spectroscopy and mass spectrometry, which revealed a common motif composed of an aminoethylphosphonate linked via a peptide bond to N5hydroxyarginine, and an additional peptide bond to Val for argolaphos A (Figure 25). The PEP mutase-encoding cluster of S. monomycini NRRL B-24309 was cloned. Five genes were predicted to encode proteins responsible for aminoethylphosphonate synthesis from PEP on the basis of identity to PhpA-E from the biochemically characterized phosphinothricin biosynthetic pathway.208 A 342 amino acid encoded protein was predicted to be an aa-tRNA-dependent transferase: it displays 34% amino acid sequence identity to DhpH, and HHpred searches retrieved FemXWv and FemASa as the best hits. However, the role of this protein in the formation of peptide bonds in argolaphos and its dependence on aa-tRNA remain to be demonstrated. In this global mining approach on a large collection of strains, the authors retrieved 278 strains carrying a PEP mutase gene indicative of a biosynthetic cluster for phosphonate production.207 Using previously established methods,205 they tried to link the genetic data for each cluster to the phosphonate compounds produced. They characterized 64 discrete groups of phosphonate biosynthetic clusters in terms of (i) the relatedness of the PEP mutase proteins and (ii) the similarity of the genomic environments of PEP mutase genes. The phosphonopeptide biosynthetic pathways described here for DHP, fosfazinomycin, and argolaphos belonged to three different groups. Each of these groups contained several biosynthetic gene clusters in addition to those already described: five for the DHP group, seven for the fosfazinomycin group, and three for the argolaphos group (http://www.igb.illinois.edu/labs/metcalf/gcf/Phosphonates. html). There were differences and similarities between these clusters, particularly as concerns the predicted aa-tRNAdependent transferases. However, although conserved within the cluster, these proteins displayed high levels of amino acid sequence identity to the previously described cognate transferases: 70−96% and 65−94% to DhpH and DhpK, respectively, for clusters of the DHP group; 88−91% to FzmI for clusters of the fosfazinomycin group; 97% to the predicted transferases of the argolaphos group. Furthermore, in most cases, the conservation of aa-tRNA-dependent transferase was accompanied by conservation of the rest of the cluster, suggesting the biosynthesis of highly similar phosphonopeptides. 4.3. Biosynthesis of Uridyl Peptide Antibiotics (UPAs)

Pacidamycins (PACs), napsamycins (NAPs), mureidomycins (MRDs), and sansanmycins (SANs) form a family of UPAs produced by Streptomyces strains.209−213 They are structurally characterized by a 3′-deoxy-4′,5′-enaminouridine or its dihydrouridine derivative connected to an N-methylated 2S,3Saminobutyric acid (DABA) from a tetra- or pentapeptide via an amide bond (Figure 27). The direction of the peptide chain is reversed twice: once at the DABA unit whose β-amino and αamino groups are involved in peptide bonds with aminoacyl 5601


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strains of M. tuberculosis.211 However, the mode of action of SANs against mycobacteria is unknown. In addition to displaying antibacterial activities due to the inhibition of a promising and poorly exploited target in medicinal chemistry,223 UPAs have unusual structural features such as enamide and ureido links and the DABA unit in the peptide chain. However, biosynthetic gene clusters for UPAs were not identified until 2010, more than 20 years after the isolation and characterization of the first UPAs. The discovery in 2007, within the biosynthetic gene cluster for friulimicin,224 a lipopeptide antibiotic, of genes dedicated to DABA synthesis enabled two different teams to identify the biosynthetic gene cluster for PACs.225,226 Since then, the biosynthetic pathway for PAC has been deciphered,36,227,228 revealing, for the first time, an association of NRPS and an aa-tRNA-dependent transferase, PacB, in the synthesis of the polypeptide chain in a NP. 4.3.1. The Transferase PacB in the Biosynthesis of PACs. About 10 different natural variants of PACs differing in the nature of the amino acids found in positions 1, 2, and 5 of the polypeptide chain (Figure 27) have been isolated from Streptomyces coeruleorubidus.209,214,229 Biosynthetic gene clusters for PACs have been isolated from two S. coeruleorubidus producers, the NRRL 18370 and AB1183F-1164 strains, and shown to be identical.225,226,230 We have chosen to use the names suggested by Walsh and co-workers, as they further characterized the enzymatic activities of most of the components of the PAC biosynthetic pathway. The PAC biosynthetic pathway is encoded by 24 genes distributed between two discrete clusters, the 30.3 kb pacA-V cluster and the pacWX two-gene cluster (Figure 28a). Figure 28b shows the proposed biosynthetic scheme for PAC 3 64. A detailed description of the multiple catalytic events is beyond the scope of this review, and we invite the reader to refer to the original articles for details.36,224−226,228,230 In this biosynthetic pathway, the transferase PacB uses Ala-tRNA to introduce the Ala1 amino acid at the N-terminus of the tetrapeptide m-Tyr2-DABA3-Ala4-CO-m-Tyr5 tethered to PacH 92, resulting in 93. Alternatively, during PAC biosynthesis, PacJL may be loaded with L-phenylalanyl or L-tryptophanyl residues rather than with an L-m-tyrosyl residue in 88, resulting in the final uridyl pentapeptides PAC 1 and PAC 2 with a C-terminal L-Trp or L-Phe, respectively (Figure 27). PacB activity was elucidated late in the deciphering of the PAC biosynthetic pathway,36 probably because PacB catalyzes one of the final steps but also because this 350-residue protein does not display sufficient sequence similarity to a protein of known function or a specific sequence signature for the inference of a putative function. A first indication came from HHpred analysis, which predicted that PacB would have a structure similar to those of FemXWv and FemASa. The activities of these enzymes, consisting of the addition of amino acids to a target amino group of the peptidoglycan precursor with aa-tRNAs used as substrates,231 led Walsh and co-workers to hypothesize that PacB would use a similar substrate for the insertion of the first amino acid, Ala1, in PACs. A pacB mutant of S. coeruleorubidus NRRL 18370 produces uridyl tetrapeptides, just like the parental strain, but the pentapeptide forms are not detected in bacterial extracts, suggesting a role for PacB in the N-terminal extension of the polypeptide chain. The in vitro reconstitution of PACs synthesis with purified enzymes highlighted the efficiency of the system for uridyl pentapeptide production in the presence of E. coli tRNAs and aaRSs: tRNA omission decreased the production of uridyl pentapeptides. However, PacB can use alternatively activated alanyl species, such as alanyl-AMP, albeit less efficiently

groups 2 and 4, respectively, and then again at the site of a ureido group between aminoacyl groups 4 and 5. All of these uridyl peptides have antibacterial activity against various Pseudomonas aeruginosa strains.210−212,214,215 Their mode of action has been characterized for MRD A 72 and MRD C 74.216−218 They inhibit peptidoglycan biosynthesis by targeting bacterial translocase I, which catalyzes the essential first step of cell wall biosynthesis by linking undecaprenyl phosphate and UDP-N-acetylmuramyl-pentapeptide to generate lipid intermediate I.219 Surprisingly, these peptides inhibited bacterial translocases I of diverse origins (P. aeruginosa, E. coli, and Staphylococcus aureus) to a similar extent in vitro whereas the antibacterial activity of MRD A 72, MRD C 74, and other uridyl peptides is highly selective for P. aeruginosa, suggesting differences in target accessibility or resistance mechanisms between bacterial strains. Several structure−activity relationship studies have established the importance of the uridine moiety, the 5′ uridine ester bond, and the central N-methyl amide bond common to all of these uridyl peptides for the inhibition of E. coli MraY (Figure 27).220−222 No other antibacterial activities against Gram-negative or -positive bacteria have been described for PACs, NACs, and MRDs, but SANs have been shown to inhibit the growth of M. tuberculosis H37Rv and multidrug-resistant

Figure 27. Chemical structures of selected uridyl peptides. The 3′deoxy-4′,5′-enaminouridine nucleoside is represented in blue. The peptide backbone is shown in black, with the DABA unit in pink. The chemical groups known to be important for antibiotic activity are highlighted in green. 5602


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Figure 28. Biosynthesis of PACs. (a) Map of the pacidamycin gene cluster of S. coeruleorubidicus NRRL 18370. Genes are drawn according to GenBank accession numbers HM855229 (pacA-V) and HQ874646 (pacWX), except for pacB, which has been lengthened by 195 bp at the 5′-end because of start codon misannotation in HM855229.225 (b) Proposed biosynthetic scheme for PAC 3. NRPSs are represented with a color and letter code for the different domains: red, adenylation (A) domain; green, thiolation (T) domain; blue, thioesterase (TE) and condensation (C) domain. A star indicates that the domain is predicted to be inactive. SAM, S-adenyl methionine; PH4, tetrahydropterin; UMP, uridine monophosphate.

CO-m-Tyr5 91 to generate the pentapeptide tethered to PacH 93 ready for uridylation by PacI. 4.3.2. PacB Homologues Encoded in the Biosynthetic Gene Clusters of UPAs. NpsN of Streptomyces sp. DSM 5940 displays 67% amino acid sequence identity to PacB36 and may therefore have similar activities. NpsN is encoded by a biosynthetic gene cluster, the heterologous expression of which in S. coelicolor M1154 led to the production of napsamycins NAP A 68 and NAP C 70, and mureidomycins MRD A 72 and MRD B 73,232,233 suggesting that NAPs and MRDs share the same biosynthetic pathway. NAPs and MRDs were originally isolated from different bacterial strains,212,213 Streptomyces sp. DSM 5940 and S. f lavidovireus, respectively, but a recent reexamination of the uridyl peptides detected in extracts of the genuine producer Streptomyces sp. DSM 5940 confirmed the presence of MRDs.233 The NAP/MRD biosynthetic gene cluster of Streptomyces sp.

than Ala-tRNA, as indicated by the rate of peptidyl-S-PacH 93 formation measured in an in vitro assay, which is at least an order of magnitude lower than that with Ala-tRNA as the substrate. PacB activity is also dependent on the tethering of its peptidyl substrate to the PacH thiolation domain 92 via the pantetheinyl arm. Attempts to form the uridyl pentapeptide from the uridyl tetrapeptide or to aminoacylate L-m-Tyr with PacB, tRNAs, and aaRSs have been unsuccessful. Walsh and co-workers demonstrated that PacB aminoacylated the tetrapeptide tethered to PacH (as shown in Figure 28b), but also the minimal substrate mTyr2-DABA-S-PacH. Thus, the incorporation of Ala in position 1 by PacB may occur as indicated in Figure 28b, or before extension of the polypeptide chain toward the C-terminus from the α-amino group of DABA in 84, yielding Ala1-m-Tyr2-DABAS-PacH. It remains to be determined whether PacD can metabolize this intermediate in the presence of PacN-S-Ala45603


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Figure 29. Chemical structures of streptothrisamines, streptothricins, and ST-related compounds. The carbamoylated D-gulosamine and the streptolidine lactam are shown in pink and blue, respectively. The L-β-Lys unit and the Gly moiety are shown in green.

uridyl tetrapeptides.210,211 The SAN cluster contains 25 genes, 24 of which encode proteins homologous to those encoded by the PAC cluster. The PacB homologue SsaB has a sequence 70% identical to that of PacB, but, as for NpsN and SSEG_02996, there is no evidence of SsaB activity during SAN biosynthesis. The SAN cluster is expressed under the control of SsaA, a DNAbinding protein corresponding to a novel class of transcriptional regulators.236 Indeed, SsaA binding to the promoter region increases transcription, and sansanmycins SAN A 76 and SAN H inhibit this DNA-binding activity. SsaA has homologues in the other uridyl peptide biosynthetic pathwaysthe cryptic NAP/ MRD biosynthesis pathway in S. roseosporus NRRL 15998 was thought to be due to an inactive SsaA homologue235suggesting a similar mode of regulation. PacA and NspM, the SsaA homologues in the PAC and NAP/MRD biosynthetic pathways, respectively, may be involved in the regulation of their genuine clusters. Assuming that PacB and its homologous proteins NpsN, SSEG_02996, and SsaB have similar activities, on the basis of the 67−96% sequence identity between these proteins, differences in uridyl peptide composition at position 1 may reflect differences in transcription regulation mechanisms. A recent BLAST search for PacB homologues (June 2016) identified only five sequences that have not been described elsewhere and display more than 50% identity to PacB (62− 91%). All five sequences originate from Streptomyces strains. Four of these PacB homologues cluster with genes encoding proteins highly similar (more than 70% sequence identity) to proteins from the PAC biosynthetic pathway, including those involved in DABA 81 and 3′-deoxy-4′,5′-enaminouridine 94 biosynthesis. However, these clusters differ from the PAC biosynthetic gene cluster, by either lacking some genes or containing additional genes possibly encoding proteins directing the synthesis of novel pacidamycin-related compounds. The gene of the last PacB homologue to be identified forms a cluster with unusual associations.

DSM 5940 contains 28 genes, and most of the encoded proteins display 67−88% sequence identity to proteins encoded by the PAC biosynthetic gene cluster, suggesting similar biosynthetic pathways despite differences in genetic organization. However, no evidence has been reported of NpsN activity in the NAP/ MRD biosynthetic pathway. NpsN may introduce a Gly residue at position 1 of the pseudopentapeptide, as observed in MRD C 74 and MRD D 75 (Figure 27), but the uridyl peptides detected in bacterial extracts upon heterologous expression of the NAP/ MRD biosynthetic gene cluster have no amino acid at position 1. The production of NAP A−D 68−71 and MRD A−D 72−75 by Streptomyces roseosporus was detected by a highly sensitive method combining MS/MS networking and peptidogenomics genome mining.234 The genome of S. roseosporus NRRL 15998 contains a biosynthetic gene cluster that is almost identical to the NAP/MRD biosynthetic gene cluster of Streptomyces sp. DSM 5940:235 it has an identical genetic organization, and the encoded proteins are 94−99% identical. However, Guoqing Niu and coworkers reported that uridyl peptides were produced by S. roseosporus only upon heterologous expression of the foreign transcriptional regulator SsaA of Streptomyces sp. strain SS,235 suggesting that the cluster of S. roseosporus is cryptic. The detected uridyl peptides belong to the MRD family, and they display variations of the uridyl moiety (hydrogenation) and of the pseudopentapeptides at positions 4 (Leu, MetSO) and 5 (Phe). However, none of these molecules has an amino acid in position 1. Furthermore, they are all modified by N-acetylation of the amino group of the amino acid in position 2. These two studies use the same strain of S. roseosporus NRRL 15998, and differences in the nature of the uridyl peptides detected may reflect differences in growth conditions and in the sensitivity of the detection methods used. Neither study provided any evidence for a role of the PacB homologue encoded by SSEG_02996 in uridyl peptide biosynthesis. The detection of the uridyl pentapeptides MRD C 74 and D 75,234 which carry an N-terminal Gly residue in position 1, may result from the activity of a PacB homologue, but, as for NpsN, the role of the PacB homologue encoded by this cluster remains unclear. The presence of a PacB homologue in the NAP/MRD biosynthesis pathway is not surprising, given the description of MRDs with a Gly residue in position 1 of the pseudopentapeptide backbone (Figure 27). However, the presence of such a homologue was unexpected for the SAN biosynthesis pathway of Streptomyces sp. SS,236 which has been shown to produce only

4.4. Biosynthesis of the Streptothricin Analogue BD-12

Streptothricins 96 (STs) and ST-related compounds 97−101 are broad-spectrum antibiotics produced by Streptomyces strains. The first member of this group of compounds, ST-F, was isolated more than 70 years ago.237 These compounds consist of the amino sugar streptothrisamine 95, composed of streptolidine lactam attached to carbamoylated D-gulosamine, linked to an L-βLys pseudopeptide (one to seven residues) by an amide bond in STs 96, or to a glycyl derivative in ST-related compounds 97− 5604


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Figure 30. Proposed biosynthesis of BD-12 in S. luteocolor NBRC 13826. (a) BD-12 biosynthetic gene cluster of S. luteocolor NBRC 13826. Genes are drawn according to GenBank accession number LC122485. Genes are colored according to the function of the encoded protein in BD-12 biosynthesis. Dotted shading indicates genes encoding proteins for which function was deduced exclusively from protein annotation. (b) Biosynthesis of streptolidine lactam in S. lavendulae BCRC 12163.245 The names of the enzymes are shown in black, and the names of the corresponding homologues identified in S. luteocolor NBRC 13826 are shown in parentheses. (c) Biosynthesis of carbamoylated D-gulosamine in Streptomyces sp. TP-A0356.246 The names of the enzymes are shown in black, and the names of the corresponding homologues identified in S. luteocolor NBRC 13826 are shown in brackets. (d) Biosynthesis of BD-12 from streptothrisamine involving the aa-tRNA-dependent transferase Orf11.

101238−243 (Figure 29). In these two classes of molecules, the Dgulosamine and the streptolidine lactam are conserved, but they can be tailored by various chemical modifications. Hamano and co-workers recently described tRNA-dependent aminoacylation in the biosynthesis of BD-12 99, an ST-related compound produced by Streptomyces luteocolor NBRC 13826.40 They sequenced the genome of the producing organism and identified a gene cluster very similar to the ST-biosynthetic gene cluster from Streptomyces rochei NBRC 12908 they had described a few years earlier244 (Figure 30a). The recombinant expression of the 34 kb fragment carrying the corresponding S. luteocolor NBRC 13826 gene cluster was sufficient to induce BD-12 99 production in S. lividans TK23. The BD-12 biosynthetic gene cluster carries 26 open reading frames (orf), including genes encoding proteins very similar to recently characterized enzymes implicated in streptolidine lactam 106245 and carbamoylated Dgulosamine moiety of ST 95, 110−112246 biosynthesis in S. lavendulae BCRC 12163 and Streptomyces sp. strain TP-A0356, respectively (Figure 30b,c). The authors noted the absence of NRPSs equivalent to those previously shown to incorporate the 244 L-β-Lys pseudopeptide in STs, suggesting that the incorporation of the glycyl residue through amide bond formation with the D-gulosamine unit involves a NRPS-independent mecha-

nism. Using HHpred, they identified the product of orf11 (Orf11; 288 residues) as a structural homologue of FemASa and FemXWv. They purified recombinant Orf11 as a 67 kDa homodimer, and demonstrated that it catalyzed the in vitro attachment of glycyl to streptothrisamine 95 with E. coli GlytRNAGly as the substrate, to generate glycylthricin 113, the chemical structure of which was determined by HPLC−ESI-MS analysis (Figure 30d). The recombinant expression in Streptomyces avermitilis of orf11, together with a mutated ST biosynthetic gene cluster from S. rochei NBRC 12908 directing streptothrisamine production, led to the accumulation of a compound identified by mass spectrometry and NMR as acetylglycylthricin, demonstrating that Orf11 was also active in vivo. The production of acetyl-glycylthricine, an inactive derivative of the antibiotic glycylthricine, was attributed to the presence in the ST biosynthetic gene cluster of a gene encoding an ST acetyl transferase able to acetylate glycylthricine.40 A recent BLAST search for Orf11 homologues in the NCBI database (June 2016) retrieved three original hits displaying 74%, 73%, and 52% sequence identity to the query sequence, originating from Streptomyces rapamycinicus NRRL 5491, Streptomyces violaceusniger NRRL F-8817, and Streptomyces sp. NBRC 110468, respectively. In the two first cases, the orf1− 5605


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Figure 31. GNAT fold of aa-tRNA-dependent transferases involved in primary metabolism. (a) Topological scheme of the GNAT fold. Reproduced with permission from ref 41. Copyright 2005 Elsevier. The N- and C-termini are indicated. The secondary structure elements β-strands and α-helices are colored in blue and red, respectively. (b) aa-tRNA analogues binding and catalytic residues of the GNAT fold 2 of L/F transferase and FemXWv. Parts of the GNAT fold delineating the binding site of aa-tRNA analogues of L/F transferase26,172 and FemXWv252 are indicated in pink and green rectangles, respectively. The names of the catalytic residues of L/F transferase and FemXWv are shown in pink and green, respectively.

orf 22 cluster is well conserved, except that orf6, encoding a putative methyltransferase, is deleted. This cluster would therefore be predicted to direct the synthesis of compound chemically very similar to BD-12 97. In Streptomyces sp. NBRC 110468, the Orf11 homologue-encoding gene is associated with genes encoding homologues of Orf5, Orf7, and Orf9−Orf22 and genes encoding original NRPS and methyl transferase activities. Provided that all the genes for streptolidin lactam biosynthesis are present (the biosynthetic pathway from (4R)-hydroxycapreomycidine 105 to streptolidine lactam 106 is unknown (Figure 30)), this cluster could direct the biosynthesis of derivatives of streptothrisamine with original peptide moieties.

the C-terminal GNAT fold, and cd-Ala-PGSPa and cd-Lys-PGSBl have an additional α-helix between strand β5 and helix α4 of GNAT fold 2. The N-terminal domains of FemXWv, FemASa, cdAla-PGSPa, and cd-Lys-PGSBl also have structures ressembling that of the GNAT fold, although some secondary structure elements might be absent. The N-terminal domain of L/F transferase displays no evident structural similarity to the GNAT fold. However, despite these differences, all these proteins belong to the Dupli-GNAT superfamily predicted to originate from an ancestral enzyme resulting from duplication of the GNAT fold.249 Structural and biochemical data have provided important information about how the L/F transferase of E. coli and FemXWv bind their cognate donor and acceptor substrates and the catalytic mechanisms involved in amino acid transfer. These two enzymes have different preferential donor substrates, leucyl/ phenylalanyl-tRNALeu/Phe and Ala-tRNAAla, respectively, but the binding sites of various aa-tRNA analogues (there are no crystal structures of transferase in complex with a charged or uncharged tRNA) are localized within the same regions of GNAT domain 220,26,172 (Figure 31). However, the cognate donor substrate is recognized in different ways. The L/F transferase has a hydrophobic pocket that can accommodate a leucyl or phenylalanyl moiety of the aa-tRNA,26,172 and part of the acceptor stem is important for efficient binding and catalysis.145 FemXWv specificity involves steric hindrance of the aminoacyl moiety to exclude amino acyl units larger than alanyl,250 and specificity determinants on the acceptor stem distinguishing between Ala-tRNAAla and Gly-tRNAGly.251,250 For both proteins, the binding regions of the acceptor stems are unknown. Furthermore, these two proteins have an α-helix preceding the β1 strand of GNAT domain 2, which exposes positively charged residues at the surface of the protein. In both proteins, this αhelix is in contact with equivalent secondary structure elements of the GNAT fold 2 and the distances between this positively charged region and the substrate binding pocket are almost the same. The replacement of positively charged residues from the

4.5. Similarities and Differences between aa-tRNA-Dependent Transferases

Here, we consider how our knowledge about long-studied transferases involved in primary metabolism could help to increase scientific understanding of the transferases involved in NP biosynthesis. We will first describe the aa-tRNA-dependent transferases involved in primary metabolism, focusing on what we know about the mode of donor substrate binding. The crystal structures of members of the three groups of aatRNA-dependent transferases involved in primary metabolism were determined either for free enzyme or for complexes between enzyme and substrate analogues. Such structures have been determined for the L/F transferase of E. coli (234 residues),26,171,172 the peptidoglycan assembly proteins FemXWv (336 residues)20,247 and FemASa (420 residues),248 and the catalytic domains of aaPGSs from P. aeruginosa (cd-Ala-PGSPa; 339 residues) and B. licheniformis (cd-Lys-PGSBl; 332 residues).23 These proteins have two domains, and the C-terminal domains, despite displaying low levels of sequence identity, have a conserved GNAT fold consisting of seven β-strands and four αhelices arranged in the polypeptide chain in the order β1-α1-α2β2-β3-β4-α3-β5-α4-β6, with the N-terminal domain providing the final β7 strand (Figure 31).42 FemASa has a 60-residue insertion forming helical arms between the β2 and β3 strands of 5606


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Table 1. Amino Acid Sequence Identity between aa-tRNA-Dependent Transferasesa involved in primary metabolism

a

involved in secondary metabolism

transferase

FemXWv

FemASa

L/F transferase

cd-Ala-PGSpa

cd-Lys-PGSBl

VlmA

DhpK

DhpH-Cter

PacB

Orf11

FemXWv FemASa L/F transferase cd-Ala-PGSPa cd-Lys-PGSBl VlmA DhpK DhpH-Cter PacB Orf11

100 25 8 11 13 11 14 7 12 18

25 100 9 12 15 13 15 13 15 15

8 9 100 18 12 10 19 18 17 17

11 12 18 100 34 19 18 14 17 14

13 15 12 34 100 20 16 17 17 10

11 13 10 19 20 100 15 14 11 10

14 15 19 18 16 15 100 24 23 17

7 13 18 14 17 14 24 100 14 28

12 15 17 17 17 11 23 14 100 15

18 15 17 14 10 10 17 28 15 100

Amino acid sequence identities were obtained at the EMBL-EBI Web site using the ClustalΟ free program and are indicated in percentage.

comprising the end of strand β4, the β4−α5 loop, and the beginning of helix α5 (Figure 31b). A similar aminoacyl binding cavity was identified in the GNAT domain 2 of cd-Ala-PGSPa and cd-Lys-PGSBl, by determining the crystal structure of cd-Lys-PGSBl in complex with L-lysine amide.23 This cavity is bordered by elements of the GNAT fold 2, essentially the α1−α2 loop, the C-terminus of strand β4, and the end of the β5−α5 loop. These regions of the GNAT fold correspond, at least partially, to those described for the binding of aa-tRNA analogues by L/F transferase and FemXWv. Mutagenesis and biochemical data revealed that differences in the acceptor stem sequences control the relative specificity of cd-AlaPGSPa for Ala-tRNAAla rather than Lys-tRNALys. Furthermore, the two proteins have an α-helix preceding the GNAT fold 2 and exposing positively charged residues, as observed in the L/F transferase and FemXWv. Their replacement with small polar residues in cd-Ala-PGSPa decreased only transferase activity, whereas charge reversal completely abolished cd-Ala-PGSPa activity. A docking model of tRNA on cd-Ala-PGSPa and cdLys-PGSBl positioned the positively charged residues of this helix at van der Waals distance from the acceptor stem of the tRNA. Finally, the contiguous pair of amino acids (Phe and Lys) essential for activity was found to be present in helix α5 of the GNAT fold 2, as previously shown in FemXWv. Mutagenesis experiments highlighted the essentiality of other residues for activity, but their exact function in binding and/or mechanism remains to be determined. Concerning the aa-tRNA-dependent transferases involved in secondary metabolism, we will focus on the biochemically characterized transferases VlmA, DhpH, DhpK, PacB, and Orf11, as several homologues can be identified in databases from bioinformatics analyses as described above and elsewhere.38,173 Sequence identities between transferases are generally lower than 20% (Table 1), with the highest scores for transferases involved in similar functions, such as FemXWv and FemSa or cd-Ala-PGSSa and cd-Lys-PGSBl. The transferases involved in secondary metabolism display 7−20% sequence identity to the structurally characterized transferases, highlighting the difficulty to infer a function for these enzymes in the biosynthetic pathways. The function of several of these transferases has been hypothesized from secondary structure predictions, highlighting potential structural similarities to Fem proteins (see above). Figure 32 shows a structural alignment of VlmA, DhpH, DhpK, PacB, and Orf11 with FemXWv, one of the best hits identified in HHpred searches with each of the corresponding amino acid sequences. Predicted secondary structures for transferases closely match the secondary structure elements observed for

corresponding α-helix in the L/F transferase of E. coli with alanine residues significantly decreases transferase activity.172 Based on docking models of aa-tRNA binding to L/F transferase, various groups have suggested a role for charged residues in nonspecific binding of the phosphate backbone of the D-stem172 or the acceptor stem.145 To our knowledge, no report has been published on the role of the equivalent positively charged α-helix in FemXWv. Key differences between L/F transferase and FemXWv include the catalytic residues identified and the deduced catalytic mechanism. Initially, structural and biochemical data led Tomita and co-workers to suggest that L/F transferase catalyzes peptide bond formation by activating the α-amino group of the acceptor substrate for nucleophilic attack of the carbonyl carbon of the aminoacyl bond of the donor substrate.26 Activation would then proceed through proton abstraction by Gln188, which, together with Glu186, forms an electron relay essential for this activation. An additional residue, Asn191, was thought to favor nucleophilic attack by increasing the polarity of the carbonyl group of the donor substrate. Fung et al. recently reconsidered the mechanism and suggested, on the basis of biochemical data and discrepancies between existing crystal structures, that peptide bonds were formed through substrate-assisted catalysis involving the 2′-OH of the donor substrate. In this case, the essential residues Glu186 and Gln188 would be involved in the correct positioning of the acceptor substrate for the initial nucleophilic attack and the subsequent proton transfer events. Arthur and co-workers established the catalytic mechanism of FemXWv by solving its crystal structure in complex with a ligand composed of a peptidoglycan-like moiety linked to a tRNA-like moiety via a functional group containing a triazole as a bioisostere of esters.20,253 Inspection of the residues around the triazole group in the crystal structure of the complex revealed the absence of a catalytic base or acid for peptide bond formation, as described in L/F transferase. The authors proposed a substrateassisted mechanism involving the 3′-hydroxy group of terminal adenosine, consistent with its essential role for efficient aminoacyl transfer.252 They also highlighted the role of Phe304 and Lys305, two residues highly conserved in Fem aminoacyl transferases. The replacement of these residues in FemXWv with Ala and Ala, Met, or Arg, respectively, decreased turnover number. Phe304 was observed in stacking interactions with the C75 ribose of the tRNA-like moiety, and it was suggested that Lys305 stabilized the tetrahedral intermediate formed during catalysis. The catalytic residues described for L/F transferase and FemXWv are located in a similar area of the GNAT fold 2 5607


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Figure 32. Structural alignment of secondary metabolism aa-tRNA-dependent transferases with FemXWv. A multiple alignment of FemXWv and transferases was generated with ClustalΟ on the European Bioinformatics Institute (EBI) Web site. This alignment was adjusted manually on the basis of HHpred searches performed on the Max Planck Institute Web site with the sequences of each transferase. Residues are colored according to the secondary structures observed in the crystal structure of FemXWv linked to a substrate analogue (PDB ID, 3GKR) and the predicted secondary structures obtained with HHpred: β-strand, blue; α-helix, red. Secondary structure elements of FemXWv, which participate to the GNAT folds of domains 1 and 2 (noted in gray and green, respectively), are named according to the description of the GNAT fold in Figure 31 and to their belonging to domain 1 (GNAT1 in index) or domain 2 (GNAT2 in index). The helices preceding the β1GNAT2 secondary structure elements (observed in FemXWv and predicted for other proteins) are highlighted in yellow, and their positively charged residues are highlighted in gray.

FemXWv, particularly for the C-terminal GNAT fold 2. VlmA shows a slightly poorer fit, and HHpred searches identified cdAla-PGSPa and cd-Lys-PGSBl as best hits for this protein. All these proteins have a predicted α-helix with positively charged residues preceding the β1 strand of GNAT fold 2 (Figure 32), as observed in all crystal structures for transferases. Furthermore, no residues equivalent to the catalytic residues of L/F transferase were identified, but a Lys residue was observed in the predicted α-helix corresponding to helix α4 of the GNAT fold 2 (Figure 32), reminiscent of the essential Lys of FemXWv (Figure 31b). In DhpH and Orf11, a Phe residue precedes this Lys residue, as observed in FemXWv. Thus, biochemical and structural characterizations of transferases involved in primary metabolism could be of great value to obtain greater insight into the transferases of secondary metabolism, as similarities between members of each of these groups of enzymes are predicted to go beyond the use of aa-tRNAs to transfer amino acyl residues. Finally, aa-tRNA-dependent transferases involved in NP biosynthesis constitute a recently discovered group of enzymes. They appear to have a limited distribution, as characterized and predicted members are scarce and belong to the actinomycetes group, being found mostly in Streptomyces species. Five such enzymes have been biochemically characterized and shown to introduce different aminoacyl residues (Ser, Leu, Gly, or Ala) into four different types of NPs, through ester or amide linkages. Database mining has revealed the presence of homologues within original biosynthetic clusters that may conceal novel NPs and novel activities for these transferases. These enzymes display low levels of amino acid sequence identity to each other and to other well-studied transferases involved in primary metabolism, such as

Fem enzymes, L/F transferases, and aa-PGSs. Despite these low levels of identity, all these transferases share or are predicted to share a common Dupli-GNAT fold supporting the aa-tRNAdependent transfer of an aminoacyl residue. Further studies are required to establish the catalytic mechanisms of transferases involved in secondary metabolism and to determine the importance of these enzymes for NP diversity.

5. THE AVAILABILITY OF aa-tRNAS FOR FUNCTIONS OUTSIDE OF TRANSLATION The past decade has seen the identification of new aa-tRNAdependent enzyme families involved in NP biosynthesis. An increasing number of NPs are being predicted to be synthesized in a tRNA-dependent manner. This leads to questions about the availability of free aa-tRNAs within the cell suitable for use as substrates by these biosynthetic enzymes. Indeed, in growing cells, tRNAs are rapidly aminoacylated by aaRSs (it is estimated that charged tRNAs account for 80% of all tRNAs in exponentially growing cells254) and the resulting aa-tRNAs are bound by elongation factor EF-Tu. This binding prevents the cleavage of the labile ester bond between the aminoacyl and tRNA moieties and promotes the efficient delivery of aa-tRNAs to the A site of the ribosome for protein synthesis.255 Experiments performed essentially with E. coli components have suggested that this association is almost complete in vivo, as indicated by the high-affinity binding constants of the active conformation EF-Tu·GTP for aa-tRNAs (KD of ∼5 nM),256−259 the estimated concentration of EF-Tu (∼100 μM),260 and the concentration of each tRNA isoacceptor (∼1−10 μM).82 This association between EF-Tu and aa-tRNAs may be universal in 5608


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prevented, at least partly because these misacylated tRNAs are not very good ligands of the elongation factor due to a thermodynamic compensation mechanism. Moreover, in several cases, competition of the Adt with EF-Tu is avoided by the direct channeling of Asp-tRNAAsn from AspRS to the AdT through the formation of a complex containing the tRNA, the aaRS, and the AdT.267−269 However, all these strategies are highly specific, and even if similar escape routes can be used in a specific manner (e.g., in the valanimycin biosynthesis pathway), this seems difficult to envisage for CDPS-dependent pathways or in RiPP biosynthesis. Fahlman and co-workers recently suggested a more generalized escape route for the delivery of aa-tRNAs to L/F transferase.145,270 Previous investigations showed that the presence of EF-Tu·GTP significantly decreased L/F transferase activity in vitro.172 Concomitant binding of L/F transferase and EF-Tu to aa-tRNAs was excluded because no ternary complex was detected in cross-linking experiments172 and an overlap including the 3′-aminoacyl adenosine was identified in the parts of the aa-tRNAs involved in binding to these two proteins.145,271,272 Fahlman and co-workers suggested that a stringent response might improve aa-tRNA delivery to L/F transferase. Stringent responses are an adaptative response to various stimuli, mediated by an increase in intracellular levels of pentaphosphate guanosine and tetraphosphate guanosine, (p)ppGpp, due to the consumption of GTP.273 In these conditions, the amount of EF-Tu·GTP available for aa-tRNA binding is decreased, not only by the lower level of GTP but also by the presence of ppGpp, which binds to EF-Tu, preventing formation of the ternary complex GTP·EF-Tu·aa-tRNA. This stringent response has many consequences for bacterial physiology, including the arrest of transcription and translation. This may result in aa-tRNAs becoming available for alternative functions. However, it is difficult to see how this strategy could work for processes requiring large amounts of aa-tRNAs during exponential growth, such as peptidoglycan biosynthesis. How could FemXWv divert Ala-tRNAAla from the translation machinery in such conditions? In studies of aa-tRNA-dependent NP biosynthesis, efforts must be made to determine the amounts of NPs naturally produced by the host and the timing of this production. The biosynthesis of NPs produced in small amounts during stationary phase might benefit from a stress response like mechanism for diverting aa-tRNAs from the protein biosynthesis machinery. Notably, CDPSs are overproduced in E. coli as active proteins, and many produce large amounts of cyclodipeptides during active growth (∼ a few milligrams to a dozen milligrams per liter of culture). It remains unclear whether these enzymes make use of a specific mechanism to utilize EF-Tu sequestered aa-tRNAs.

bacteria, as suggested by the recent work of Shrader and Uhlenbeck, who used a thermodynamic model to predict the ΔG° for the binding of any tRNA to EF-Tu.261 However, if this is the case, it remains unclear how aa-tRNAs escape the translational machinery to supply alternative functions, including NP biosynthesis. Very little is known about aa-tRNA availability in vivo for NP biosynthesis. The only exception is the valanimycin biosynthetic pathway. This pathway includes a dedicated SerRS, VlmL,181 and inactivation of the gene encoding this protein greatly decreases valanimycin production,180 highlighting the difficulty of the transferase, VlmA, using Ser-tRNASer synthesized by the housekeeping SerRS, SvsR,181 and suggesting that idiosyncratic features of VlmA, VlmL, and the tRNASer isoacceptor may be required for efficient valamimycin production. However, several ways to escape the protein biosynthesis machinery have been identified for aa-tRNAs used by enzymes involved in primary metabolism. FemASa preferentially uses a specific aminoacylated isoacceptor, which binds EF-Tu·GTP very poorly, leaving it available for alternative functions.262 This isoacceptor was also identified as a poor EF-Tu binder by the thermodynamic model developed by Shrader and Uhlenbeck.261 Indeed, whereas most of the 5849 tRNA sequences analyzed gave ΔG° values matching those determined for E. coli tRNAs, 19 tRNA sequences, including the S. aureus tRNAGly(UCC), gave ΔG° values suggesting weaker binding to EF-Tu. Based on the experimental evidence for the role of S. aureus tRNAGly(UCC) in peptidogycan biosynthesis,262 the authors suggested that the identified tRNAs with higher ΔG° values might be used for alternative functions in vivo. Another solution was adopted by the Lys-PGS MprF2 of Clostridium perf ringens: this enzyme has a high affinity for Lys-tRNALys, facilitating efficient competition with EF-Tu· GTP for Lys-tRNALys binding in physiological conditions.169 Another well-known escape route is exemplified by the initiator methionyl-tRNA,10 which binds the ribosomal P-site directly to provide the first amino acid for incorporation into a polypeptide.263 The bacterial initiator methionyl-tRNA (MettRNAfMet) is formylated on the amino group of the esterified methionine through the action of a methionyl-tRNAfMet formyltransferase (FMT). This modification prevents undesirable binding to EF-Tu. Before formylation, the affinity of MettRNAfMet for EF-Tu is one-tenth that of elongator aa-tRNAs,257 because initiator tRNAs have a characteristic acceptor stem, with mispairing of the 1−72 bases. This feature decreases competition between FMT and EF-Tu for the uptake of Met-tRNAfMet. Another well-studied escape route involves selenocysteyltRNASec. This rare tRNA incorporates selenocysteine into the polypeptide during ribosomal protein synthesis, in response to unusual UGA codons located in a specific context. This incorporation is achieved with the assistance of a specific form of elongation factor, different from EF-Tu. Again, to avoid the misappropriation of Sec-tRNASec (and its precursor SertRNASec), tRNASec contains specific nucleotides, impairing its binding to canonical EF-Tu.264 Another more recently identified escape route involves the use by many organisms (archaea, organelles, and many bacteria) of indirect routes for the aminoacylation of amidated amino acids (Asn and Gln) on tRNAs. These routes use AspRS or GluRS to load tRNAAsn or tRNAGln with Asp or Glu, respectively. The Asp-tRNAAsn or GlutRNAGln is then converted into Asn-tRNAAsn or Gln-tRNAGln, respectively, through the action of specific tRNA-dependent amidotransferases (AdTs).265,266 The erroneous capture of misacylated Asp-tRNAAsn and Glu-tRNAGln by EF-Tu must be

6. CONCLUSION The aa-tRNA-utilizing enzymes involved in NP biosynthesis could probably best be described as “diverse”. These enzymes catalyze diverse reactions via diverse catalytic mechanisms to produce a diverse set of NPs (Figure 3). They are used in many different ways to produce NPs and, therefore, play a role in the biosynthesis of the ribosomal peptides into which they introduce posttranslational modifications, or in the biosynthesis of nonribosomal peptides and pseudopeptides. So far, these aa-tRNA-utilizing enzymes have been classified into three main groups defined on the basis of catalytic activity and the conserved structural fold: CDPSs, LanB-like dehydratases, and dedicated transferases (Figure 3). They are present in bacteria, rare in eukarya, and absent in archaea, and their 5609


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diversity of aa-tRNA use by the enzymes involved in the biosynthesis of new NPs.

distribution in bacteria is nonuniform (Table 2). All of the dedicated transferases identified to date were found in Streptomyces strains (see section 4), and these enzymes therefore appear to be limited to phylum Actinobacteria. CDPSs and LanB-like enzymes are also found predominantly in this phylum, consistent with the robustness of Actinomycetes as a source of NPs. However, these two groups of enzymes are also well represented among Proteobacteria and Firmicutes (Table 2). The CDPS group, which is currently the largest, is the only group to have a few eukaryotic members, from the Ascomycota and Cnidaria (Table 2). We previously showed that a CDPS from the starlet sea anemone Nematostella vectensis was an active CDPS, and this enzyme is, to date, the only one shown to be involved in nonribosomal peptide synthesis in animals.61 The discovery in 2008 of the first aa-tRNA-utilizing enzyme involved in NP biosynthesis33 was followed by the identification of an increasing number of these enzymes. Less than 10 years later, about 750 putative enzymes have been identified, about 70 of which have been characterized biochemically (Table 2). With the increasing recognition of these enzymes, others are likely to be identified, and new groups will be defined. For example, an enzyme thought to catalyze the transfer of an alanyl moiety in an Ala-tRNAAla-dependent manner during biosynthesis of the nucleoside antibiotic ascamycin has been identified in Streptomyces sp. JCM9888.274 However, this enzyme displays no similarity to any of the enzymes of the three defined groups, and is predicted to have an α/β-hydrolase fold. It may therefore be the first member of a new group of aa-tRNA-utilizing enzymes involved in NP biosynthesis. As knowledge about these aa-tRNAutilizing enzymes increases, we can expect to see a greater

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. ORCID

Muriel Gondry: 0000-0003-0398-3404 Author Contributions †

M.M. and P.B. contributed equally to the writing of this review.

Notes

The authors declare no competing financial interest. Biographies Mireille Moutiez studied Chemistry at the Ecole Nationale Supérieure de Chimie of Paris, France (1989−1992), and received her PhD in Biochemistry in 1995 for research on oxidative stress in the Trypanosomatidae at the Pasteur Institute of Lille, France. She joined the Protein Studies and Engineering Department of the CEA, Saclay (France), where she spent 12 years working on the characterization and engineering of protein disulfide oxidoreductases and the refolding of disulfide-containing proteins. She joined Muriel Gondry’s team in 2007, and her research interests now focus on the characterization and manipulation of CDPS-containing biosynthesis pathways. Pascal Belin graduated from Montpellier SupAgro in 1989, where he studied Agronomy and Applied Microbiology. He then studied at Paris 6 University, France, and received his PhD in Microbiology in 1994. After a two-year postdoctoral position at the CEA, Saclay, he was hired by the CEA to develop new approaches for improving recombinant protein expression. He joined Muriel Gondry’s team in 2004, and has since focused on deciphering CDPS biosynthetic pathways. His current research interests lie in characterizing the tailoring enzymes of these pathways and engineering CDPS-dependent pathways.

Table 2. Genome Distribution of aa-tRNA-Utilizing Enzymes in NP Biosynthesisa

origin Bacteria Actinobacteria Proteobacteria Firmicutes Bacteroidetes Cyanobacteria Chlamydiae Parcubacteria Synergistetes Microgenomates Chloroflexi Eukaryotes Ascomycota Cnidaria Total

no. of avail. genomes in NCBI databaseb

CDPSs

c

1402 3525 1688 778 121 43 208 27 54 126

261 (28) 128 (23) 50 (6) 3 (1) 2 (1) 4 (2) 3 1 1

604 9 8585

8 4 (1) 465 (62)

LanB-like enzymesd 136 (3) 17 59 (1) 10 1

transferases

Muriel Gondry completed her PhD in Biochemistry at Paris-Sud University at Orsay (France) in 1994, under the supervision of Florence Lederer. She joined the group of Roger Genet (1995−1997) at the CEA of Saclay (France) as a postdoctoral worker and has remained at the same institution ever since. She currently directs the “Enzymology and Non-Ribosomal Peptide Biosynthesis” group of the Institute for Integrative Biology of the Cell (CEA, CNRS, Paris-Sud University). Her research interests lie in deciphering natural product biosynthetic pathways and manipulating their gene clusters for the synthesis of new molecules.

e

48 (5)

ACKNOWLEDGMENTS The work carried out in the authors’ laboratory is supported by the CEA, the CNRS, Paris-Sud University, and grants from the French National Research Agency (ANR 2010/Blan 1501 01 and ANR-14-CE09-0021-01). The authors warmly thank Jérôme Seguin, Emmanuel Favry, and Morgan Babin for their continuous support and encouragement throughout the writing of this review.

1

224 (4)

48 (5)

a The number of putative enzymes identified in databases is indicated. The number of biochemically characterized enzymes is indicated in parentheses. bThe number of available genomes in the NCBI database in October 2016 is indicated in order to give some idea of the number of identified enzymes compared to the number of available genomes. c Putative CDPSs were identified by searching the NCBI database in March 2016. dPutative LanB-like enzymes were obtained from Ortega et al.39 ePutative transferases were retrieved from published results and database searches done in June 2016 (see section 4 for details).

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Review

(274) Zhao, C.; Qi, J.; Tao, W.; He, L.; Xu, W.; Chan, J.; Deng, Z. Characterization of biosynthetic genes of ascamycin/dealanylascamycin featuring a 5′-O-sulfonamide moiety in Streptomyces sp. JCM9888. PLoS One 2014, 9, e114722.

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