Cyclic natural product oligomers: diversity and (bio)synthesis of macrocycles

Abstract

Cyclic compounds are generally preferred over linear compounds for functional studies due to their enhanced bioavailability, stability towards metabolic degradation, and selective receptor binding. This has led to a need for effective cyclization strategies for compound synthesis and hence increased interest in macrocyclization mediated by thioesterase (TE) domains, which naturally boost the chemical diversity and bioactivities of cyclic natural products. Many non-ribosomal peptide synthetase (NRPS) and polyketide synthase (PKS) derived natural products are assembled to form cyclodimeric compounds, with these molecules possessing diverse structures and biological activities. There is significant interest in identifying the biosynthetic pathways that produce such molecules given the challenge that cyclodimerization represents from a biosynthetic perspective. In the last decade, many groups have pursued the characterization of TE domains and have provided new insights into this biocatalytic machinery: however, the enzymes involved in formation of cyclodimeric compounds have proven far more elusive. In this review we focus on natural products that involve macrocyclization in their biosynthesis and chemical synthesis, with an emphasis on the function and biosynthetic investigation on the special family of TE domains responsible for forming cyclodimeric natural products. We also introduce additional macrocyclization catalysts, including butelase and the C_T-mediated cyclization of peptides, alongside the formation of cyclodipeptides mediated by cyclodipeptide synthases (CDPS) and single-module NRPSs. Due to the interdisciplinary nature of biosynthetic research, we anticipate that this review will prove valuable to synthetic chemists, drug discovery groups, enzymologists, and the biosynthetic community in general, and inspire further efforts to identify and exploit these biocatalysts for the formation of novel bioactive molecules.

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Songya Zhang

Songya Zhang received his Master of Science degree from Shenyang Pharmaceutical University under the supervision of Prof. Huiming Hua. In 2018, he obtained his PhD degree at Albert-Ludwigs-Universität Freiburg in Germany under the supervision of Prof. Andreas Bechthold. Subsequently, he was selected into the Shenzhen High-Level Talent Plan and joined the lab of Prof. Tong Si at the Shenzhen Institute of Advanced Technology, Chinese Academy of Sciences for postdoctoral research. Later, he became an assistant professor in the lab of Prof. Youming Zhang at the Shenzhen Institute of Advanced Technology, Chinese Academy of Sciences. His research is centered around exploring enzymes possessing special catalytic functions and conducting synthetic biology research based on the non-ribosomal peptide derived from natural products.

Shuai Fan

Shuai Fan obtained his PhD in 2017 from the Chinese Academy of Medical Sciences & Peking Union Medical College. Subsequently, he joined the Institute of Medical Biotechnology at the Chinese Academy of Medical Sciences, where he has held positions as an assistant researcher and an associate researcher. His primary research focuses on enzymes with novel catalytic mechanisms involved in the biosynthetic pathways of natural products, combining theoretical calculations to elucidate catalytic mechanisms and rational enzyme engineering.

Haocheng He

He Haocheng obtained his PhD from Hunan Normal University in 2021, under the supervision of Professor Xia Liqiu. Following his doctoral studies, he joined the laboratory of Professor Zhang Youming at the Shenzhen Institutes of Advanced Technology as a postdoctoral researcher. His research primarily focuses on constructing chassis microorganisms for synthetic biology, aimed at optimizing the production of natural products and enzymes. He Haocheng has accumulated considerable research experience in exploring and regulating the production of natural products, as well as optimizing nitrogenase expression.

Max J. Cryle

Max J Cryle obtained his PhD in chemistry from the University of Queensland in 2006. He then moved to the Max Planck Institute for Medical Research in Heidelberg Germany as an HFSP Cross-Disciplinary Fellow and later as an Emmy Noether group leader funded by the DFG. Since 2016 he is an EMBL Australia group leader based within the Biomedicine Discovery Institute at Monash University, where his team focusses on understanding antibiotic biosynthesis (with particular interest in non-ribosomal peptide synthesis) as well as developing new antibiotics. He is also a Chief Investigator in the Australian Research Council Centre of Excellence for Innovations in Peptide and Protein Science (CIPPS).

Hai-Yan He

Hai-Yan He received his BS in chemistry from Tianjin University. He obtained his PhD in bioorganic chemistry at the Shanghai Institute of Organic Chemistry, Chinese Academy of Sciences, under the supervision of Prof. Gong-Li Tang. Following a post-doctorate with Katherine Ryan at the University of British Columbia, he began his independent career as a Principal Investigator at the Institute of Medicinal Biotechnology, Chinese Academy of Medical Sciences & Peking Union Medical College, China. His research group is devoted to the development of new biocatalysts for production of valuable chemicals and clinical drugs.

Youming Zhang

Youming Zhang received his PhD at Ruprecht-Karls-Universität Heidelberg in 1994. After his postdoctoral research in Heidelberg University and European Molecular Biology Laboratory (EMBL), he founded Gene Bridges GmbH (Germany) in 2000 as the co-founder and Chief Scientific Officer. In 2013 he was appointed as the director of the State Key Laboratory of Microbial Technology, Shandong University. He was elected as a member of Academia Europaea (2019) and German National Academy of Science and Engineering (2020). His research is dedicated to development of the Red/ET recombineering and its application in microbial natural product heterologous expression, humanized animal models and tumor targeting bacteria.

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1. Introduction

Natural products (NPs) have historically provided us with a crucial source of unique molecular scaffolds, enabling the development of new therapeutic agents based on their wide range of biological activites.^1–3 The cyclization of molecules has been an important strategy in the development for new drugs, and macrocycles are widely present in NPs from various sources. Currently, many NP-derived drugs are focusedon a cyclized scaffold, such as the immunosuppressant cyclosporine, the hormone oxytocin, and numerous antibiotics including vancomycin, daptomycin, erythromycin, and amphotericin. To date, around 70 macrocyclic compounds have been developed as FDA-approved medicines.⁴

Cyclization, a crucial chemical reaction, modifies the activity of natural products by forming a ring structure, of which the size and shape can significantly impact the activity of the molecule, particularly in the design of peptides.^5–7 Also, the cyclic nature of the ring is typically essential for the activity, proving highly beneficial in enabling the specific interactions between molecule and its biological target, and improving its pharmacokinetic properties, such as half-life and permeability.⁷ This makes cyclic peptides highly promising for drug development. Cyclic peptides also have low structural Gibbs free energy, increasing binding affinity and selectivity for target molecules. They can selectively bind to unstable protein surfaces and penetrate cell membranes, thus facilitating target drug development.^8,9

The polymerization of molecules as a biosynthetic strategy can bring many advantages,¹⁰ for example by generating compounds with high structural complexity and chemical diversity from a much smaller biosynthetic assembly line. Dimerization (as well as higher order multimerization) reactions can also be used to introduce specific functional groups or structural features, whilst at the same time generating enlarged molecules with improved stability or selectivity, which are crucial factors in target binding.^11,12 Larger molecules are also able to access multiple binding sites, with examples showing how two separate binding sites on a receptor molecule can be targeted by a dimeric drug, creating much tighter binding than would be possible with two isolated monomeric compounds.^13,14 To date, many dimeric natural products have been shown to exhibit enhanced bioactivities when compared to their monomeric form.^15–18 Indeed, in some cases, dimeric compounds have been shown to be is active whilst the monomer remains inactive¹⁹ (e.g. dimeric lipopeptide fusion inhibitors exhibit significantly greater antiviral potency compared to monomeric derivatives,^20,21 and cyclic peptide oligomers display more potent antimicrobial effect than their linear counterparts).²² Due to their broad range of biological activities, some dimeric NPs have been investigated as potential therapeutic agents or have inspired the creation of structural mimics.^23–25

This review will focus primarily on cyclodimeric (as well as higher order multimeric) products formed through cyclisation of a dimeric precursor (referred to as macrocyclization), with a subset formed through direct cyclodimerization (the dimerization of two identical monomers to form a cyclic product). Cyclopeptides can be cyclized in various ways, including head-to-tail, head-to-side chain, side chain-to-tail, and side chain-to-side chain. Both de novo design and in vitro evolution are major strategies utilized for current cyclopeptide development.²⁶ A detailed review of cyclic peptide cyclization strategies has been recently published, and will not be discussed in detail here.^27–29

In natural product biosynthesis, TE domains play a crucial role in enhancing the chemical diversity and bioactivities of these compounds through cyclization. Typically, TE domains control the final chain length after the assembly of monomers from precursors. Macrocycle formation, often mediated by TE domains, is a common strategy in PKS- and NRPS-mediated natural product assembly lines.^30–32 Various types of reactions, including TE-mediated cyclization, cycloaddition, cycloisomerization, esterification, and radical reactions, contribute to the formation of dimeric natural products.^11,33 Understanding the enzymes catalysing these reactions within biosynthetic pathways is essential for drug discovery and the development of new biocatalysts for macrocyclization.

Moreover, research has shown that TE domains can mediate diverse atypical cyclization. For example, in teixobactin biosynthesis, the tandem TEs (TE1 and TE2) within the NRPS terminal module (Txo2) collaborate to create an active site for substrate cyclization, with biochemical experiments confirming their essential roles through serine residue acetylation.³⁴ Similarly, the ObiF1 TE domain in obafluorin biosynthesis catalyzes β-lactone ring closure via transthioesterification, influenced by intramolecular interactions with the C-terminal integrated MbtH-like protein (MLP).³⁵ Additionally, the TE domain within curacin A's polyketide synthase mediates a unique termination reaction within module CurM, preceded by sulfonation catalysed by an integrated sulfotransferase (ST) domain.³⁶ Furthermore, the recombinant TE domain StmC/ACP–TE, discovered by the Ge group, showcases tandem reactions, acting as both a thioesterase and cyclase during streptoseomycin biosynthesis.³⁷ Notably, Kobayashi et al. developed a chemoenzymatic approach to synthesize cyclic peptides by exploiting SurE, a protein homologous to a penicillin-binding protein rather than a traditional TE, thereby expanding the scope of chemoenzymatic synthesis of cyclopeptides.^38,39 YsfF and YsfF_HBT, both with thioesterase activities, are involved in complex biochemical reactions leading to the formation of youssoufene A1, which shows increased inhibition against multidrug-resistant bacteria. It was found that YsfF/YsfF_HBT catalyse various reactions, including 6π-electrocyclic ring closure and dimerization.⁴⁰ These examples collectively highlight the diverse and pivotal roles TE domains play in orchestrating the biosynthesis of natural products.

Currently, many non-ribosomal peptide synthetase (NRPS) and polyketide synthase (PKS) derived cyclodimeric NPs have been identified, with these molecules possessing diverse structures and biological activities. The identification of biosynthetic pathways that produce such molecules is of great interest due to the challenge that cyclization represents: specifically, how do the TE domains in NRPS or PKS assembly lines recognize and accommodate large substrates during catalysis? In the last decade, several groups have pursued identification of such TE domains from NRPS or PKS assembly lines, which has led to new insights to this biocatalytic machinery.^41–43 Whilst these have typically not been involved in cyclodimer formation, our team has been one of those that has sought to overcome this by extensively investigating PKS/NRPS synthetases related to such dimerizing TE domains (including those involved in mohangamide and disorazole biosynthesis). Recently, several excellent reviews about TE-related biosynthesis have been published.^31,32 However, these works only describe a single example of dimer cyclization. This review focuses on the special class of TE domains involved in the cyclization of dimeric (as well as higher order multimeric) natural product intermediates during NRPS and PKS assembly, including discussion concerning their mechanism.

In this review, we examine key examples of cyclic dimeric (as well as higher order multimeric) natural products. We illustrate the synthetic and biosynthetic routes to these cyclodimeric compounds, including the catalytic mechanism of key biosynthetic enzymes. We also strive to emphasize the gaps in both knowledge and methodology that must be addressed to effectively access these cyclic scaffolds. We limit our discussions to focus primarily on works that generate cyclodimeric natural products including novel analogues, thus largely excluding approaches that lead to the production of high polymer products through cyclo-oligomerization. This review covers the period until July 2023.

2. Cyclodimeric natural products

The cyclization and release reaction are the crucial final step for many natural products biosynthesis pathways. In this regard, thioesterase (TE) domains play a central role, where they exhibit impressive stereospecificity and selectivity that makes them highly valuable biocatalysts for industrial applications involving peptide cyclization. TE domains found in PKS or NRPS pathways commonly facilitate crucial steps of oligomerization and macrocyclization in the biosynthesis of many bioactive natural products.^44,45 This section will provide a brief introduction of TE domain structure and the catalytic mechanism, with particular focus on how TE domains involved in cyclodimerization catalyze this transformation during peptide biosynthesis (Fig. 1).

Fig. 1 The general cyclisation mechanism of macrocyclic natural products.

2.1 TE domains

Nonribosomal peptide synthetases (NRPS) and polyketide synthases (PKS) are two major types of microbial biosynthesis pathways. These two machineries generate a range of bioactive molecules, including antibacterial, antifungal, immunomodulatory and antitumor agents, all of which have extensive applications in medicine.^46–48 PKS and NRPS are both representatives of microbial modular biosynthesis assembly lines. These enzymes utilize simple building blocks (such as two-carbon or three-carbon (acetyl-coenzyme A, malonyl-coenzyme A, propionyl-coenzyme A, and butyryl-coenzyme A) or amino acid units, and through a sequence of modules working in concert, assemble them into long-chain products or macrocyclic precursors with molecular weights of up to several kilodaltons.⁴⁹ Thioesterase (TE) domains are then typically located in the final module of these systems, where they are responsible for the release of the final product of the assembly line through hydrolysis or through macrocyclization with intramolecular nucleophilic attack by free amino or hydroxyl groups.⁵⁰ TE domains are responsible for the release of most of the products from both NRPS and PKS pathways. As the last enzyme in the PKS and NRPS assembly lines, the thioesterase controls the recognition and release of the product.³¹ The recognition of substrates and subsequent catalysis by thioesterase domains are therefore crucial for obtaining the expected product in a manner that does not stall the assembly process, whilst also serving as a control element for peptide modification in trans in certain cases.⁵¹

To accurately reflect the scope of our investigation, we have clarified that the phylogenetic analysis specifically covers TEs from reported bacterial PKS or NRPSs pathway. We excluded the TEs from fungal PKS/NRPSs which clearly have different evolutionary histories. In some fungal peptide synthetases, the oligomerization is catalysed by the C_T domain which will be elaborated in the next section. Interestingly, TE domains with reported iterative catalytic function do not gather in a single cluster (Fig. 2), and the mechanism underpinning their catalysis still requires investigation. It is also apparent that a large number of uncharacterized TE domains remain spread across these different clusters and await characterization.

Fig. 2 Phylogenetic analysis of the TE domain family sequence. Colours represent TE domain function and substrate type (type I or II from NRPS or PKS pathway and others). The characterized TE domain with iterative catalytic function mentioned in this paper was labelled by asterisk. This phylogenetic analysis was based on TE proteins from representative phyla involved in reported biosynthetic pathways. This analysis was conducted using protein data sourced from the reference collections of the Boddy group⁵² and the Deng group.⁵³ The detailed protein information can be found in the ESI† (Table S1). The sequence alignment and phylogenetic tree was generated by MEGA, the tree was modified by iTOL.^54,55 For a more detailed phylogenetic analysis, refer to Caswell et al. (2022).⁵⁶

Generally, these thioesterases can be grouped into type I or type II TE domains from NRPS or PKS pathways. Type I TEs are typically integrated into the final module of the biosynthetic assembly line and catalyze product release through hydrolysis or cyclization, whereas type II TEs function independently, often playing a role in proofreading or editing the assembly line intermediates.^52,57 (Fig. 2). For example, the TEs from DptD, EndC, Sas19, GlmI and GrsB group together in one cluster, as they are all domains that are found in the final module of an NRPS and are responsible for peptide cyclization (in daptomycin, enduracidin, WS9326, glidomide and gramicidin biosynthesis, respectively). In contrast, PKS-derived intra-module TE domains exist together in different cluster (e.g. TE domains from pikromycin (pikAIV TE), erythromycin (DEBS TE), amphotericin (AmphK TE) and FR-008 (FscF TE) biosynthesis).

Many PKS and NRPS pathways contain stand-alone enzymes known as type II TEs (TEII), which group into one cluster in the phylogenetic tree (Fig. 2).^52–56 Type II TEs are not covalently bound to the multienzyme, with recent research suggesting that Type II TEs instead are capable of diverse catalytic functions. Type II TEs belong to the α/β hydrolase family, where their role is typically to hydrolyze thioester-bound residues on the 4′-phosphopantetheine (4′-PPant) arms of acyl carrier proteins (ACPs) or peptidyl carrier proteins (PCPs), such as the TEII BarC from barbamide pathway^58,59 or Tcp39 from teicoplanin biosynthesis.⁵¹ Some type II TEs have been characterized with the function of chain translocation, such as the type II TEs WS5 (Cal5 or Sas5) and WS20 (Cal20 or Sas20) from the WS9326 pathway and TEII LgnA from legonindolizidin A pathway, where they work as shuttling enzymes for substrates during peptide assembly.^52,60 In the case of PnG, this TEII enzyme performs multiple functions in the biosynthesis of phoslactomycin PKS (Pn PKS), including release of an ACP-tethered intermediate as well as proofreading.⁶¹

TE domains involved in NRPS biosynthesis. Nonribosomal peptides (NRPs), which are secondary metabolites typically produced by microbes, are synthesized by a class of large multienzyme machineries known as non-ribosomal peptide synthetases (NRPSs).⁶² The peptides produced by these assembly lines include more than 300 building blocks, with key examples seen in compounds such as vancomycin (antibiotic),⁶³ cyclosporine (immunosuppressant),⁶⁴ and bleomycin (antineoplastic drug).⁶⁵ NRPSs are modular and use an assembly-line mechanism for peptide biosynthesis, with each module independently cycling to elongate the peptide chain. Each module consists of several catalytic domains, including adenylation domains (A domains), condensation domains (C domains), and peptidyl carrier protein/thiolation domains (PCP/T domains).⁶⁶ A domains select and activate specific (typically amino acid) substrates through ATP consumption to form aminoacyl adenylates, which are covalently connected to the 4′-phosphopantetheine prosthetic group found at the beginning of the second helix in the neighbouring PCP domain, forming an aminoacyl-S-PCP. Subsequently, C domains catalyse amide bond formation, initially between two aminoacyl-S-PCPs to produce a PCP-bound dipeptide.⁶⁷ Throughout the extension process, the growing peptide intermediates remain covalently linked to different PCP domains. Additionally, there are further auxiliary domains that can also play a role in structural modification of the peptide, including the activity of methyltransferase domains (MT domains), epimerization domains (E domains), and cyclization domains (Cy domains), which further enrich the structural diversity and biological activity of NRPs.⁶⁶ Termination domains, such as the TE thioesterase located at the end of NRPS, are also vital to the function of NRPSs, as they serve to remove the final product from the enzyme, thus allowing multiple rounds of biocatalysis to occur.

The TE domain of NRPS machineries is responsible for catalyzing the hydrolysis or cyclization reactions of the NRPS polypeptide chain. Studies have shown that NRPS type I thioesterases recognize specific linear peptide-PCP substrates (or even peptide-S-CoA/SNAC substrates) and utilize internal nucleophiles to cyclize the peptide via carbon-to-nitrogen intramolecular lactonization or lactamization.⁶⁸ As a crucial NRPS catalytic domain, research into the protein structure and enzymatic catalytic function of TE domains has been steadily increasing.

Structural and mechanistic insights into NRPS TE domain. To date, the structures of the TE domains excised from the enterobactin (EntF TE, PDB: 3TEJ),⁶⁹ valinomycin (Vlm2 TE, PDB: 6ECE),⁷⁰ surfactin (SrfTE, PDB: 1JMK),⁷¹ fengycin (FenTE, PDB: 2CB9)⁷² and skyllamycin B NRPS assembly lines (Skyxy-TE, PDB: 7CRN, 7DXO)⁷³ have been solved, providing insights into the mechanism underlying enzymatic peptide cyclization.

Structural analysis has revealed that TE domains belong to the α/β hydrolase superfamily, which includes enzymes such as lipases, proteases, and esterases.⁷¹ The typical α/β hydrolase fold is comprised of a repeating β/α/β motif that forms a seven- or eight-stranded parallel β-sheet wrapped by α-helices on both sides, while several helices on top form the lid region.^74,75 In general, the first N-terminal β-strand of the hydrolase fold is absent in TE domain. Instead, the second β-strand serves as the only anti-parallel strand among the remaining six or seven strands in the structure. Sequence alignments showed that lid region significantly differs between TE domains. The differences observed in TE structures mainly lie between the sixth and seventh β-strands, with the α-helices linking these strands displaying diversity in structure, including the insertion of an extra structural domain.⁷⁶ It has been hypothesized, based on crystal structures and MD simulations, that the lid region opens to accommodate the presentation of thioester substrates. These studies offer detailed insights into the dynamics of the lid region, which is thought to play a role in substrate loading and release.^77–79

The structures of SrfTE and FenTE confirm that thioesterases belong to the α/β hydrolase superfamily.^71,72 The asymmetric unit of SrfTE reveals the presence of two distinct molecules, each exhibiting closed and open conformations. While SrfTE exists as a dimer in the asymmetric unit, it has been shown to exist as a monomer in a solution environment. In the SrfTE, the lid structure covers three α helices and has two different conformations. When in the “open” conformation, the lid structure folds back, thus exposing the catalytic center. Conversely, in the “closed” conformation, the catalytic center is almost completely covered by the lid. In the Fen TE, the lid structure is the shortest seen for NRPS TE domains, which causes the catalytic center of Fen TE to be exposed, possibly serving to make this TE highly promiscuous. Fully or partly exposed catalytic triads were observed in the related lysosomal palmitoyl protein thioesterases,⁸⁰ whose substrates are similarly sterically demanding when compared to NRPS peptides.

In terms of the structure of TEs, cyclodimerization is performed by EntF TE and Vlm2, making these of particular interest (Fig. 3A and B). The structure of Vlm2 TE adopts the α/β-hydrolase fold typical of type I TE domains, with a canonical Ser-His-Asp catalytic triad covered by the lid structure.⁷⁰ The lid of Vlm2 TE is a mobile component that includes an extended loop, three helices (α4–6), a five-residue helix (α7), a long helix (α8), and another short helix (α9), which is thought to have various functions including substrate positioning and solvent exclusion (Fig. 3E). The structure of the EntF PCP–TE di-domain represents the physiological structure before CoA addition to the PCP domain. The PCP and TE domains have a well-defined relative orientation with a primarily hydrophobic interface. The globular core of the TE domain displays two α-helices (α6–α7) protruding from this core, resembling webbed fingers that form a lid covering the TE active site (Fig. 3F). This allows the 4′-PPant arms, which is tethered to Ser 48 of the PCP domain in the holo state and can span up to ∼20 Å, to reach Ser 180 in the EntF TE domain.

Fig. 3 Overall structure of TEs. Structures of Vlm2 TE (A), EntF TE (B), DEBS TE (C) and PICS TE (D). Secondary-structure elements of Vlm2 TE (E), EntF TE (F), DEBS TE (G) and PICS TE (H).

The typical thioesterase mechanism belongs to the Bi–Bi model, where a covalent intermediate is formed between the substrate and the enzyme during the reaction process.⁵⁶ The first step involves substrate nonribosomal peptide being loaded onto the TE domain by transesterification of the PCP thioester, forming the acetyl–enzyme intermediate.^78,81 The second step involves a histidine residue acting as a base to help deprotonate a nucleophile within the substrate (such as the side chain amine, hydroxy and thiol groups of amino acids) that then can attack the acetyl–enzyme complex, forming the tetrahedral intermediate. The tetrahedral intermediate is stabilized by the oxyanion hole by hydrogen bonding from two backbone amide NH groups. Finally, the cyclic product is released from the TE domain, and as a result, the active site of the TE domain is reactivated (Fig. 4).

Fig. 4 Mechanism of TE-mediated reaction for nonribosomal peptide cyclization. (1) Peptide transfer from PCP to active site serine of TE (2) acyl-TE adduct is attacked by the nucleophilic residue, forming a tetrahedral intermediate that is stabilized by the oxyanion hole. (3) Cyclic product is released and TE domain is reactivated.

The catalytic core of the α/β hydrolase region is highly conserved, usually consisting of a catalytic triad Ser-His-Asp present comprising a nucleophilic serine adjacent to the β5 fold belonging to the GXSXG motif,⁸² an aspartic acid usually located at the loop following the β7 strand according to its canonical site in α/β hydrolase folds, and finally a histidine positioned on the loop after the last β-fold, which is also highly conserved (Fig. 4).^69–71 Interestingly, some TE domains contain a Cys in place of the Ser in catalytic triads.^83,84 The backbone amides of the amino acid following the nucleophilic amino acid, as well as another usually located between the β3 fold and the α-helix, form the negatively charged oxyanion hole, which stabilizes the intermediate during the reaction.⁸⁵

The formation of a linear dimerization intermediate is a prerequisite for cyclodimerization mediated by TE domains. Two possible pathways for the oligomerization of NRPS intermediates by TE domains in valinomycin synthesis have been postulated.⁷⁰ In the first scenario, ‘forward transfer’, the distal hydroxyl group of the peptidyl-O-TE complex attacks the thioester group in the peptidyl-S-PCP enzyme intermediate, directly forming dipeptidyl-O-TE as a product. In the second scenario, ‘reverse transfer’, the distal hydroxyl group of the peptidyl-S-PCP complex attacks the ester group in the peptidyl-O-TE enzyme intermediate, forming dipeptidyl-S-PCP as a product, which would then be transferred onto the serine of TE domain (Fig. 5). The analysis of the synthetic intermediates identified during valinomycin synthesis has revealed that Vlm TE catalyzes oligomerization through the “reverse transfer” pathway.⁷⁰

Fig. 5 (A) The sequence conservation analysis of TE domain; (B) two scenarios for oligomerization.

Thioesterase-mediated cyclisation with stereochemical inversion in NRPS biosynthesis. D-Configured amino acids are characteristic of the products of many NRPSs, and two examples of TE domains have recently been identified that perform stereoinversion prior to hydrolysis. Skyllamycin B, a cyclic peptide molecule containing 11 amino acid residues, contains one D-configuration hydroxyl-substituted leucine. Using a combination of in vivo and in vitro experiments, it has been shown confirmed that Skyxy-TE domain (PDB: 7CRN, 7DXO) performs the isomerization and regioselective cyclization of the C-terminal L-conformation amino acid residue of the linear substrate.⁷³ Site-directed mutagenesis experiments have revealed that Gln125 of Skyxy-TE plays a critical role in the selectivity for isomerization, with the mutant Q125A producing a cyclized product containing an L-leucine residue without isomerization.

Notably, Gln125 of Skyxy-TE is not conserved among other TE enzymes. The catalysis by Skyxy-TE of the triad's His254 might extract an α-proton from the substrate, and Skyxy-TE is capable of utilizing an oxyanion hole to stabilize the enolized intermediate, thereby facilitating the spontaneous epimerization reaction. Nocardicin is a monocyclic β-lactam antibiotic biosynthesised by an NRPS assembly line of five modules, despite only three residues being contained in the final peptide product. In nocardicin biosynthesis, the TE domain NocTE (PDB: 6OJD) has two functions, catalyzing the C-terminal epimerization of an L-Hpg residue to D-Hpg before performing the hydrolytic release of the mature product.^86,87 The peptide binding pocket has been resolved by high-resolution crystal structures of the thioesterase domain in ligand-free form and trapped with a fluorophosphonate mimic of the substrate, which has aided in rationalizing the stereoinversion performed by this TE domain.⁸⁸ The spontaneous epimerization reaction mechanism of NocTE is similar to that of Skyxy-TE. In the substrate of NocTE, the substitution of the hydroxyphenyl moiety stabilizes the carbanion; consequently, the substrate of NocTE can spontaneously epimerize to its stereoisomer (Fig. 6).

Fig. 6 Overall structure of Skyxy-TE (A) and NocTE (B). The catalytic triad is displayed as a green stick model, while the oxyanion hole and key residues are shown as pink stick models.

TE domains involved in PKS biosynthesis. Bacterial and fungal-derived polyketide synthases (PKSs) generate a plethora of intricately structured compounds, which find extensive applications in the field of medicine, such as the antibiotic erythromycin,⁸⁹ the cholesterol-lowering agent lovastatin,⁹⁰ antiparasitic compound nemadectin⁹¹ and the epothilones, which are anticancer agents.⁹² Based on the structural characteristics and functions of PKSs, these can be classified into three major types: type I, type II, and type III PKS.⁹³ Similar to NRPSs, PKSs are modular assembly lines that synthesize complex polyketides and that can be further modified by post-PKS modification enzymes. In type I cis-AT PKSs for example, they contains three basic units in each module: acyltransferase (AT), ketosynthase (KS), and acyl carrier protein (ACP) domains.⁹⁴ The AT domain selects and loads short carbon units onto the ACP. The downstream KS domain receives the polyketide chain synthesized by the previous module and transfers the elongated polyketide chain onto the ACP loaded with the short carbon units, thus forming an elongated chain with typically two additional carbon units. In the Virginiamycin trans-Acyl Transferase Polyketide Synthase Like NRPS systems, all PKS assembly machinery contain a terminal TE domain that is often responsible for the release and cyclization of the polyketide chain.

Structural and mechanistic insights into polyketide thioesterases. In the PKS thioesterase family, the most representative structures are the 6-deoxyerythronolide B synthase (DEBS) TE^94,95 and the picromycin synthase (PICS) TE (Fig. 3C, D, G and H).⁹⁶ The crystal structures of curacin A TE domain (CurM TE, PDB: 3QIT),⁹⁷ aflatoxin TE domain (PksA TE, PDB: 3ILS),⁸⁵ and tautomycetin TE domain (TMC TE, PDB: 3LCR)⁷⁹ have also been sequentially reported. The first crystal structure obtained for macrocycle-forming PKS TE domains was solved for DEBS, which assembles the antibiotic erythromycin.⁹⁸ This work was followed by the structural elucidation of the TE domain from PICS.⁹⁹

Both the DEBS TE and PICS TE are classified within the α/β hydrolase family and exhibit a hydrophobic cavity that traverses the protein, along with a hydrophobic dimer interface enriched in leucine residues. The catalytic triad is located in the center of the active site pocket, indicating that substrates must access this pocket during the reaction. The DEBS TE consists of a central β-sheet composed of seven strands, with β2 oriented in antiparallel manner to the remaining strands. Notably, the length of the β7 strand is shorter than β8, and the C-terminus of β7 is twisted 90° to form part of the substrate channel. The β strands are flanked on either side by α helices, two on one side and four on the other. Additionally, at the N-terminal extremity of the TE, two supplementary α-helices constitute the dimer interface.^95,98

A two-step mechanism has also been proposed for the TE-mediated formation of macrocyclic polyketides. PKS and NRPS release mechanisms the first step involves the acylation of a linear polyketide thioester at a serine residue that is conserved within the catalytic triad of the PKS TE domains. This step generates an acyl–enzyme intermediate which can be stable for an extended period of time.⁵⁶ The second step is a nucleophilic attack of an intramolecular hydroxyl group leading to cyclization (Fig. 7), or hydrolysis of the acyl–enzyme intermediate when no efficient intramolecular nucleophile is available.

Fig. 7 Mechanism of TE-mediated reaction for polyketide cyclization. (1) Peptide transfer from ACP to active site serine of TE (2) acyl-TE adduct is attacked by the nucleophilic residue, forming a tetrahedral intermediate (3) cyclic product is released and TE domain is reactivated.

In the biosynthetic pathway of pikromycin (PIK), PIK-TE-1 demonstrated a higher frequency of “active” conformations, enabling easier transformation into the pre-reaction state necessary for macrocyclization. This transformation was influenced by a hydrogen-bonding interaction with His268 and essential hydrophobic interactions for substrate recognition. Notably, Tyr178 was proposed to widen the exit and facilitate product release in PIK-TE-1. The substrate channel of PIK-TE-1 is crucial for the mechanism and selectivity of macrolactonization, featuring a hydrophobic spacious chamber within the substrate channel, while a hydrophilic barrier at the distal end of the substrate channel induces the long-chain substrate to curl towards the catalytic residue Ser148, thus facilitating macrolactonization. A similar phenomenon exists in Pim TE, where its substrate channel is bifunnel-shaped, with a highly hydrophobic cavity near the catalytic triad, and a hydrophilic region specifically inducing the linear substrate to curl and complete cyclization. Thus, the substrate channel interrupted by a hydrophilic barrier represents a universal mechanism that induces essentially linear hydrophobic substrates to curl into conformations suitable for forming macrolactones.

Computational analysis supported the notion that macrocyclization catalyzed by PIK-TE via re-face nucleophilic attack was thermodynamically and kinetically more favorable than si-face attack.¹⁰⁰ Chen et al. combined MD simulations with QM and QM/MM calculations for complexes of DEBS TE and its substrates. The DEBS TE and substrate undergo induced fit aided by hydrogen bonding and hydrophobic interactions. This transition is accompanied by a conformational change in active site pocket of the enzyme, shifting from a “closed” state to an “open” state. Proton abstraction from the substrate is followed by nucleophilic attack on the carbonyl carbon at C1, resulting in the formation of a charged tetrahedral intermediate. The energy barrier for this reaction step is 9.9 kcal mol⁻¹. Finally, the final macrocyclic product is released with a very low barrier of 0.3 kcal mol⁻¹.¹⁰¹

2.2 Cyclodimeric non-ribosomal peptides

Cyclic peptides are currently used with great success in drug therapy, including applications such as the antibiotics vancomycin, daptomycin, dalbavancin and polymyxin B, the hormone analogues octreotide and vasopressin, the immunosuppressant cyclosporine, and the antitumor agents lanreotide and romidepsin.¹⁰² As drug molecules, cyclooligomer peptides also have a wide range of biological activities such as anticancer, antiviral, antibacterial and enzyme inhibitors. The cyclization of polypeptide structures is important to maintain the properties of peptide drugs.^103–105

For NRPS-derived peptides, cyclodimerization mediated by the TE domain further boosts the chemical diversity of the peptide products of these assembly lines. Beyond this, nature employs diverse strategies in constructing macrocyclic molecular scaffolds. For example, P450 oxidases and radical SAM enzymes also can catalyse macrocyclic formation, such as macrocyclic construction in vancomycin and streptide.^106,107 Diels–Alder enzymes utilize a cycloaddition reaction to construct the macrocyclic scaffold in pyrroindomycin, whilst the bifunctional oxidase LkcE catalyses a unique combination of amide oxidation and a Mannich reaction to assemble the macrocyclic system in lankacidin.¹⁰⁸ Cyclodimerization in NRPS pathway greatly increases the possibilities to access peptides with high structural complexity and bioactivity diversity. Moreover, NRPS modules can be used iteratively to generate larger peptide rings, following the principle of molecular efficiency and atom economy. Understanding how nature generates cyclooligomer peptides from acyclic precursors can provide inspiration for the construction of functional macrocyclic molecules. In this section, we will focus on NRPS-derived NPs that include cyclodimerization mediated by a TE domain in their biosynthesis, and particular focus on the structural characterization and catalytic mechanism of these TE domains.

Gramicidin S. Gramicidin S and tyrocidine belong to cyclic β-sheet decapeptides and were clinically applied as antibiotics early in the 20th century. Gramicidin S was first isolated from Brevibacillus brevis by the Gause group in 1944.^109,110 Gramicidin S (Soviet) demonstrates high antibiotic activity towards Gram-negative and positive bacteria, as well as several strains of pathogenic fungi, with gramicidin S recognized as interacting with the (bacterial) lipid bilayer.^111,112 Like tyrothricin, a mixture of gramicidin and tyrocidine, gramicidin S has seen limited clinic application due to its significant hemolytic side effects.^112,113 Gramicidin S shows distinct structure features in comparison with other gramicidin derivatives such as gramicidin A.^114,115 Gramicidin S is a cyclic decapeptide consisting of a primary dimerized liner peptide backbone (D-Phe₁-L-Pro₂-L-Val₃-L-Orn₄-L-Leu₅-D-Phe₆-L-Pro₇-L-Val₈-L-Orn₉-L-Leu₁₀),¹¹⁶ which then undergoes cyclization to generate a symmetric cyclopeptide cyclo-(Pro-Val-Orn-Leu-D-Phe)₂via conjugation in a head-to-tail fashion. In gramicidin S, the two Val-Orn-Leu tripeptides form an antiparallel β-sheet structure comprising two β-strands, while the two D-Phe-Pro dipeptides act as two type II′ β-turns. Four intramolecular hydrogen bonds stabilize the β-hairpin of its secondary structure (Scheme 1), with the β-sheet conformation playing an important role for the antibacterial bioactivity of this peptide.¹¹⁷

Scheme 1 The gramicidin S biosynthesis pathway. The dotted lines are used to indicate the four intramolecular hydrogen bonds found in gramicidin S.

Tyrocidine is a cyclic decapeptide first isolated from the soil bacteria Bacillus brevis in 1940, and is the major constituent of tyrothricin, an antibiotic mixture also containing gramicidin.¹¹⁸ Tyrothricin was an important milestone in antibiotic development, as it was the first antibiotic commercialized for clinical use. Tyrocidine A is active against a broad spectrum of Gram-positive bacteria and acts by disrupting the synthesis of the peptidoglycan layer of bacterial cell walls.¹¹⁹ Tyrocidine has also been shown to be effective at controlling the fungal pathogens.¹²⁰

Biosynthesis of gramicidin S and tyrocidine. Marahiel and colleagues first reported the gramicidin S BGC (grs) from Bacillus brevis ATCC 9999.¹²¹ This cluster consists of three genes: grsT, grsA, and grsB. The biosynthetic assembly line is comprised of two NRPS proteins, GrsA and GrsB, encoding one and four modules respectively. Thus, the NRPS proteins GrsA and GrsB assemble the basic linear scaffold of the decapeptide. The GrsB TE domain then iteratively catalyses the dimerization of two identical assembled pentapeptides monomer, via head-to-tail macrocyclization, leading to the release of the symmetric cyclo-decapeptide gramicidin S.¹²² Recently, the Kries group showed that the type II thioesterase GrsT plays a proofreading function during this assembly process.^123,124 Hoyer et al. investigated the substrate specificity and cyclisation capacity of the TE domain from the gramicidin S synthase (GrsB TE domain). Protein sequence alignment between GrsB TE and other iterative TE domains (EntF, DhbF and TioS) indicated that the extended linker region could be an important factor for cyclodimerization. A series of biochemical assays confirmed that the truncated GrsB TE domain (250 residues) shows a 5-fold reduction in catalytic activity, supporting this hypothesis.¹²⁴

Further experiments then demonstrated that the ability of the GrsB TE domain to catalyse the cyclodimerization of peptide substrate decreases with increasing peptide length when these are loaded onto PCP–TE didomain constructs. The GrsB TE was shown to efficiently dimerize or even trimerize 6–15 amino acid containing linear peptides, demonstrating significant flexibility and substrate tolerance in the catalytic pocket of this domain.¹²⁴ Additional chemoenzymatic studies demonstrated that the GrsB TE can cyclize immobilized linear decapeptide precursors, suggesting its potential use in chemoenzymatic analogue synthesis.¹²⁵

TycC TE from tyrocidine biosynthesis can catalyze the formation of gramicidin S. Walsh et al. first reported that the TycC TE domain from the tyrocidine producing NRPS also can dimerize the truncated thioester peptide precursor D-Phe-L-Pro-L-Val-L-Orn-L-Leu-SNAC and cyclize the product to generate gramicidin S (Scheme 2).¹²⁶ Thus, describing the TycC TE domain here is also helpful for our understanding of the macrocyclization mechanism mediated by TE domains in NRPS assembly, as this demonstrates the potential versatility of this process by TE domains more generally.

Scheme 2 The biosynthesis pathway of tyrocidine A and gramicidin S.

In 1997, the Marahiel lab reported the biosynthetic gene cluster of tyrocidine in Bacillus brevis ATCC 8185. The tyrocidine NRPS synthetase enzymes TycA, TycB and TycC are responsible for peptide assembly of the linear decapeptide.¹¹⁷ The C-terminal TE domain found in TycC (TycC TE) is then responsible for the macrocyclization of the decapeptide to generate tyrocidine A.¹²⁷ The TycC TE domain was the first excised TE domain (35 kDa) shown to retain cyclization activity. Moreover, it has been demonstrated that the recombinant TycC TE domain is a versatile macrocyclization catalyst, which can generate tyrocidine variants with altered biological activity.¹²⁷ Whilst the TycC TE domain typically catalyses the cyclization of a decapeptide-thioester to form tyrocidine A, it has been shown that the TycC TE can also catalyse the cyclodimerization of the gramicidin S pentapeptide-SNAC (D-Phe-Pro-Val-Orn-Leu-SNAC) to generate gramicidin S (Scheme 2).¹²⁶

The TycC TE can further catalyse the macrocyclization of peptide substrates that differ significantly from the linear precursor of tyrocidine. Tyrocidine and gramicidin share structural similarity in both their N-terminal (D-Phe-Pro) and C-terminal residues (Val-Orn-Leu). Biochemical results suggest that TycC TE can cyclize the gramicidin S precursor decapeptide-SNAC (D-Phe-Pro-Val-Orn-Leu-D-Phe-Pro-Val-Orn-Leu-SNAC) and a tyrocidine A precursor (D-Phe-Pro-Phe-D-Phe-Asn-Gln-Tyr-Val-Orn-Leu-SNAC) at a comparable rate to the wildtype peptides. These results suggest that the TycC TE can cyclize different substrates provided that the necessary “recognition residues” near each end of the substrate are present.¹²⁶ Furthermore, the TycC TE tolerates the replacement of residues 5–8 in peptidyl thioesters, and that it shows comparable catalytic efficiency towards various artificial tyrocidine variants, including glycosylated peptide thioesters.¹²⁸

To further explore the substrate specificity of the TycC TE, Xie et al. established positional-scanning libraries based on SNAC substrate mimics to determine the catalytic activity of TycC TE towards these libraries (K_cat and K_m). In general, TycC TE is less efficient in cyclizing such libraries by a factor of 2- to 82-fold in comparison with the substrate sequence of wild type. Analysis of the catalytic efficiency (K_cat/K_m) of these reactions suggested that the N-terminal D-Phe1 and C-terminal Orn9 residues are essential for substrate recognition by the TycC TE domain,¹²⁹ which is consistent with other reported results.¹³⁰ It was proposed that the efficient occurrence of TycC TE-mediated cyclization requires the formation of key hydrogen bonding partners between the amine group of the Orn side chain and the carbonyl group of the N-terminal aromatic residue.³¹ In addition, the results obtained using sub-library O₃, where the catalytic efficiency showed the greatest reduction (82-fold), suggest that the side chain of the 4th residue significantly influences TE domain activity (Scheme 2).¹³¹

Chemical synthesis of gramicidin S and related congeners. While thioesterase-mediated enzymatic cyclization is crucial in the biosynthesis of natural peptides like gramicidin S, chemical methods also play an important role in the synthesis and modification of these compounds. One notable example is the solid-phase synthesis of gramicidin S and related congeners, which provides an efficient route for producing cyclic peptides in the laboratory.

In 2006, Wadhwani et al. synthesized gramicidin S utilizing solid-phase Fmoc chemistry. The sequential synthesis starts from the 2-chlorotrityl (2CT) resin loaded with Fmoc-protected D-phenylalanine, and proceeds using typical SPPS conditions until the linear decapeptide 1 was cleaved from the solid support using TFA/ⁱPr₃SiH/H₂O. An overall yield of 69% was obtained after peptide cyclisation mediated by PyBOP/HOBT in DCM solution and hydrazine-mediated Dde deprotection (Scheme 3).¹³²

Scheme 3 Total synthesis of gramicidin S and its analogues.

Rivera and colleagues developed an on-resin Ugi reaction system for the synthesis of gramicidin S analogues. 2CT resin was used as the solid support for peptide synthesis by solid phase peptide synthesis (SPPS). Next, the intermediate product resin-bound acyclic decapeptide 2 was directly subjected to Ugi macrocyclization to generate a range of gramicidin S analogues (Scheme 4).^133,134 Grotenbreg et al. designed an alternate synthesis of gramicidin S via stepwise SPPS using HMPB-resin. After the linear nonapeptide 3 was generated using standard Fmoc-based SPPS, cyclization was performed through the incorporated azide functionality (Scheme 5). After deprotection, the cyclopeptide gramicidin S analogue was obtained in 96% yield.¹³⁵

Scheme 4 Total synthesis of gramicidin S and its analogues based on Ugi reaction system.

Scheme 5 Total synthesis of gramicidin S and its analogues via SPPS using HMPB-resin.

TE domains can be combined with SPPS of linear peptide precursors to cyclize the immobilized peptide, which can greatly enrich the structural diversity of the peptides obtained. As a key feature of the enzymatic synthesis of cyclic peptide molecules via NRPS-mediated biosynthesis is the linkage of an activated linear intermediate peptide to a carrier protein domain via a thioester linkage, this has similarities to SPPS approaches.¹³⁶ TE domains can act as isolated cyclization catalysts, and further these TE domains can also act on solid-phase linked peptides that contain biomimetic linkers to complete peptide cyclisation, thus enabling the integration of biocatalysis with combinatorial SPPS.¹³⁷ Kohli and colleagues also synthesized linear peptide SNACs (that mimic the thioester attachment of peptides to the NRPS) to investigate the catalytic function of the TycC TE domain; the results of their study further demonstrate the versatility of this approach for peptide cyclization (Scheme 2).¹³⁷

Mohangamides. The mohangamides are dimeric cyclopeptides characterized by symmetrical dimers with identical peptide backbones and distinct pendant acyl chains.^138,139 Mohangamide A was first isolated from Streptomyces sp. SNM55 together with the cyclic monomer WS9326A, where Mohangamide A displayed strong inhibitory activity against Candida albicans isocitrate lyase.¹⁴⁰ Despite being a pseudodimer of WS9326A, bacterial strains known to produce WS9326A (such as S. calvus and S. asterosporus) do not produce mohangamide A.^141,142

Yoon and colleagues recently investigated the macrocyclization of mohangamide A by the TE domain found in the mohangamide NRPS assembly line (Scheme 6). Sequence alignment revealed few differences between the TE domains from Streptomyces sp. SNM55 (WS19T-TE) and S. calvus (Cal19T-TE) (Fig. 2).¹⁴² Their biochemical experiments suggested that the mohangamide TE domain displays strict specificity for its substrate, and is only able to catalyse peptide cyclization when both native precursors of mohangamide A are present. This suggests that the structure of the unsymmetrical monomeric peptides is restricted in order to lead to a competent reaction for the WS19T-TE/Cal19T-TE. Further structural investigation is needed to illustrate the detailed catalytic mechanism of this unusual pseudo-dimeric scaffold.^143,144

Scheme 6 The biosynthetic pathway of mohangamides.

Recently, it was demonstrated that Tyr dehydrogenation occurs during the WS9326A peptide assembly, with the cytochrome P450 Sas16 catalysing the dehydrogenation of the PCP-dipeptide intermediate (Z)-2-pent-1′-enyl-cinnamoyl-Thr-N-Me-Tyr. The P450_Sas-mediated incorporation of the double bond follows N-methylation of the Tyr by the N-methyl transferase domain found within the NRPS, and P450_Sas appears to be specific for substrates containing the (Z)-2-pent-1′-enyl-cinnamoyl group. The P450_Sas structure reveals differences with other P450s involved in the modification of NRPS-associated substrates, including the substitution of the canonical active site alcohol residue with a phenylalanine (F250), which in turn is critical to P450_Sas activity and WS9326A biosynthesis. This discovery expanded the repertoire of P450 enzymes that can be used to produce biologically active peptides,¹⁴⁵ and raises further questions regarding the specificity of the mohangamide TE domain for the acyl side chains of its substrates.

Enterobactin family siderophores. Siderophores are iron-binding agents secreted by bacteria and fungi that help in the solubilization and transport of Fe(III) in iron-deficient environments.¹⁴⁶ Structurally, siderophores are generally less than 1000 Daltons in size. Depending on the structure of the moieties donating the oxygen ligands for Fe(III) coordination, they are divided into five main classes: catecholate, phenolate, hydroxamate, α-keto-carboxylate and α-hydroxy-carboxylate siderophores. These functional groups all possess negatively charged oxygen atoms that can bind to Fe(III) (Fig. 8). Enterobactin-type siderophores are a representative family dependent on NRPS biosynthesis.¹⁴⁷

Fig. 8 Examples of enterobactin family siderophores and representative siderophore chelating units.

Enterobactin. Enterobactin belong to catecholate-type siderophores that are utilized by many bacteria as iron chelators, and it is one of the strongest natural iron-binding compounds.¹⁴⁸ The linearized enterobactin trimer also can play the role of detoxifying extracellular reactive oxygen species, as it has been shown to be a natural substrate of multidrug efflux pump MacAB.¹⁴⁹ Enterobactin was first identified from Salmonella typhimurium and E. coli cultural supernatants.^150–152 Structurally, enterobactin is a cyclic trimer macrolactone (tris-(N-2,3-dihydroxybenzoyl-L-serine)) that is composed of three 2,3-dihydroxybenzoate (DHB) and L-serine residues connected by an amide and an ester bond. Enterobactin coordinates Fe(III) through its 2,3-dihydroxybenzoyl (DHB) catechol groups.¹⁵¹

Catechol-type siderophores are widely diverse in their structural composition. In contrast to enterobactin, streptobactin has a cyclic triester-bonded macrocycle scaffold (L-Thr-L-Arg-DHBA)₃, which generate a C₃ symmetry axis (Fig. 8). Streptobactin was discovered from the marine actinomycete Streptomyces sp. YM5-799, with the corresponding dimer dibenarthin, trimer tribenarthin and monomer benarthin all discovered from the same strain.¹⁵³ The derivative corynebactin is assembled from the corresponding tri-lactonic scaffold containing the units L-Thr, L-Gly and DHBA (Fig. 8). Corynebactin was discovered from Brevibacterium sp, which are obligate aerobic Gram-positive bacteria found in milk and on human skin.^154,155

The enterobactin biosynthetic pathway in E. coli is a well-researched example of a non-linear NRPS assembly line. The enterobactin gene cluster encodes six functional enzymes EntA-F.^156,157 The formation of 2,3-dihydroxybenzoic acid (DHBA) requires the activities of EntC, EntB and EntA.¹⁵⁶ The three core NRPS enzymes EntE, EntB and EntF comprise the two-module NRPS assembly line, which is responsible for the iterative condensation of three DHBA and three L-serine residues to produce the final iron-chelating product enterobactin.¹⁵⁸

EntE is an AMP ligase that activates DHBA through adenylation. EntB, consisting of two domains (C-PCP), accepts the activated dihydroxybenzoate (DHB) and loads this aryl acid on its PCP domain. EntF catalyses the final step in the pathway and comprises four domains (A-C-PCP–TE). In this protein, the A domain initially adenylates a L-Ser residue and transfers it to the adjacent PCP domain. The subsequent C domain catalyses amide bond formation between the 2,3-DHB bound to the upstream EntB Ar-CP domain and the PCP-bound L-Ser residue to generate DHBA-Ser-S-PCP in EntF. The EntF C domain has been demonstrated to possess specificity for L-Ser residue as the downstream (acceptor) substrate.^159,160

Cyclotrimerization is required to create the trilactone backbone of enterobactin, which is performed by the final two domains of EntF (PCP–TE) via sequentially transferring oligomer intermediates between the 4′-PPant arm of the PCP domain and the active site serine residue of the TE domain (Scheme 7). The EntF TE domain therefore fulfils roles as both a terminal domain for acyl transfer from the PCP domain and an extended catalyst for enterobactin trimerization. Through three cycles of iterative (2,3-dihydroxybenzoyl)-L-serine (DHB-Ser) synthesis, three DHB-Ser moieties are connected whilst remaining linked to the TE domain by ester bond. Finally, the linear DHB-Ser trimer is cyclized and released from TE domain (Scheme 7).^160,161 Structurally, EntF TE is similar to homologous thioesterase domains, such as AB3403-TE, Vlm-TE, FenTE and SrfTE.¹⁶² Using solution phase nuclear magnetic resonance (NMR) spectroscopy, Frueh et al. have also successfully elucidated the structure of the apo-PCP–TE didomain (PDB: 2ROQ, 37 kDa) of the enterobactin NRPS synthetase EntF (vide infra).⁶⁹

Scheme 7 The biosynthetic pathway of enterobactin.

Even though some factors that impact the cyclization and hydrolysis of TE domain mechanism have been investigated for NRPSs, the fundamental structural basis for both iteration and cyclization remain elusive. Whilst multiple sequence alignment of triscatechol siderophore TE domains provides possible clues, the exact mechanism (iteration, hydrolysis, and cyclization) awaits further elucidation.¹⁶³ Structurally, the conserved Ser-His-Asp catalytic triad (Ser1138, His1271, Asp1165) is vital for EntF-TE thioesterase activity. Ser1138 acts as the nucleophile that binds DHB-Ser precursors. Mutation of Ser1138 to Ala results in complete disruption of substrate oligomerization. Mutation of His1271 to Ala can still generate enterobactin but with an approximately 10 000-fold decrease in yield.^158,164 Interestingly, the residue Pro1073 is conserved in the EntF and other PCP–TE homologues, and it was recognized to play important role of stabilizing the oxyanion hole in these structures (Fig. 9).¹⁶⁵

Fig. 9 Structure of the PCP–TE di-domain from EntF. (A) The overall structure of the PCP–TE conjugate. The PCP domain (pink) is connected to the TE domain (light blue) through a short linker. The TE lid region is shown in wheat. (B) Interaction of the 4′-PPant arms with the active site of the EntF-TE domain. Residues of the catalytic triad (S1138, H1271, D1165) and oxyanion hole (A1074 and L1139) are shown as green sticks. The phosphopantetheinyl analog is shown as magenta stick.

Iterative biosynthesis facilitates the repeated use of modules and necessitates a “waiting room” for reaction intermediates.¹⁶⁴ According to solution NMR and protein interaction analysis, the catalytic pocket formed by PCP–TE di-domains is dynamic. The lid region of the EntF holo-PCP–TE (PDB 3TEJ) domain needs to fluctuate to accommodate various intermediates in the hydrophobic pocket and to remain open to the 4′-PPant arm of the corresponding PCP domain bearing the tripeptide.⁶⁹ In addition, the process of cyclotrimerization, which is facilitated by the EntF TE, presumably involves an interaction between the two domains, allowing the structure bound to the PCP domain to be transferred into the substrate pocket of the TE domain.¹⁶⁵

Protein–protein interactions are also hypothesized to influence the catalytic function of TE domains from specific systems. The Cryle group reported the characterization of the TE domain from the teicoplanin NRPS.⁵¹ Based on the results of a series PCP-substrate cleavage activity assays, they found that the activity of the Tcp12-TE domain is dependent on the presence of an unusuallylong N-terminal linker region, which appears to be exclusive to the NRPS machinery seen in GPA biosynthesis. In addition, the longer Tcp12-TE (L-TE) domain of the teicoplanin NRPS has a strong selectivity for the PCP-bound peptide in its optimal cross-linking condition,⁵¹ indicating that the TE in GPA biosynthesis acts as a logic gate for the complex X-domain/P450 mediated peptide cyclisation cascade.^126,166,167

TE domains are relatively flexible and possess significant conformational diversity. To trap a PCP complex with a TE, Bruner and colleagues utilised the chemical probe α-chloroacetyl-amino-coenzyme A, which is a mimic of the natural thioester substrate targeting the Ser residue in the catalytic triad (Ser1138, His1271, and Asp1165) of TE domains (Fig. 9). After bioconjugation with the PCP domain via the activities of a phosphopantetheinyl transferase, the probe allowed a stable complex of the PCP–TE didomain protein to be trapped, with the resultant structure demonstrating the interaction between the TE and PCP domains.^165,168 The Gulick lab revealed that the EntF-TE domain also exhibits significant conformational mobility, as it can adopt multiple positions with respect to other domains based on the analysis of negative-stain electron microscopy.^162,169 Based on these structures, it is postulated that the mature PCP domain-bound peptide chain can be transferred to the TE domain via rotation of the linker attached to PCP.¹⁶²

The total synthesis of enterobactin was first reported in 1977.¹⁷⁰ Shanzer et al. developed a synthetic route to the enterobactin cyclic trilactone backbone (4) using an organotin template. This method required fewsynthetic steps, but delivered a relatively low yield of cyclic trilactone (23%).¹⁷¹ In 1997, Ramirez et al. designed a strategy to improve the synthetic yield of enterobactin. Using methyl N-triphenyl-methyl-L-serinate (5) as starting material and a variation on Shanzer's organotin template, they also synthesized the cyclic trilactone (4) intermediate with a much-improved yield (81%) (Scheme 8).¹⁷²

Scheme 8 Total synthesis of enterobactin.

Amphi-enterobactin. Amphi-enterobactin has been discovered in multiple strains, and its biosynthetic gene cluster was analysed from the strain Vibrio harveyi BAA-116, which is a model bacterium for quorum sensing.¹⁷³ Amphi-enterobactin is a type of triscatecholate siderophore that bears resemblance to enterobactin, but is characterized by an expanded tetralactone core and features a fatty acid attached to the amine of the additional L-Ser, which is in turn decorated with 2,3-dihydroxybenzoate (DHB) (Scheme 9). The fatty acid tail may vary in length (ranging from C₁₀ to C₁₆) and bear additional modifications installed after release from the NRPS.^173–175

Scheme 9 The biosynthetic pathway of amphi-enterobactin.

Through reconstitution and heterologous expression, it was demonstrated that the NRPS enzyme AebF catalyses amide bond formation and cyclization of the tetralactone backbone. The C domain in this single module NRPS performs bifunctional condensation catalytic activity (Scheme 9). The long-chain fatty acid CoA ligase (FACL) AebG is required to activate the fatty acid as a fatty acid CoA thioester. In contrast to the trilactone backbone of enterobactin, the AebF-TE can accommodate the larger 16-member tetralactone backbone of amphi-enterobactin.^164,173 Based on the “waiting room” model, the AebF-TE serves as a temporary attachment site for L-Ser-DHBA during amphi-enterobactin assembly.

Bacillibactin. Bacillibactin (2,3-DHB-Gly-^LThr)₃ is a cyclic catecholic siderophore produced by the soil bacterium Bacillus subtilis (Scheme 10). The enzyme EntF in enterobacteriin biosynthesis in E. coli is a monomodular NRPS, while the analogous enzyme DhbF from bacillibactin biosynthesis is a dimodular NRPS.¹⁷⁶ Paenibactin is a bacillibactin derivative isolated by Paenibacillus elgii B69. Structurally, paenibactin is a cyclic trimeric lactone comprising 2,3-dihydroxybenzoyl-alanine-threonine (2,3-DHB-^LAla-^LThr)₃.¹⁷⁷ Two bacillibactin derivatives (bacillibactin E and bacillibactin F) were discovered from marine Bacillus sp. WMMC1349. In contrast to bacillibactin, one of the DHB residues contained in bacillibactin is replaced by nicotinic acid and benzoic acid, respectively. The bacillibactin BGC of WMMC1349 includes a trimodular NRPS (DhbE, DhbB, DhbF) (Scheme 10). The discovery of bacillibactin E and bacillibactin F suggested that the DhbE and its ortholog are promiscuous in their activation of structurally related benzoic acid derivatives.¹⁷⁸

Scheme 10 The biosynthetic pathway of bacillibactin.

Vicibactin. Vicibactin is a cyclic trihydroxamate siderophore produced by many bean root rhizobia (Scheme 11). These types of siderophores, which acquire iron within the rhizosphere, play a vital role in nodulation and nitrogen fixation.¹⁷⁹ Structurally, vicibactin is composed of three D-3-hydroxybutyric acid moieties and three N²-acetyl-N⁵-hydroxy-D-ornithine residues, which are connected by alternating amide and ester bonds.

Scheme 11 The biosynthetic pathway of vicibactin.

Many studies have reported the genes related to vicibactin biosynthesis and proposed the steps of biosynthetic pathway.^180,181 The biosynthesis of vicibactin produced by the symbiotic bacterium Rhizobium leguminosarum involves a cluster of eight genes: vbsG, vbsS, vbsO, vbsA, vbsD, vbsL, vbsC and vbsP.¹⁶³ Among them, VbsS, a NRPS enzyme (C-A-PCP–TE) plays a crucial role in the assembly of the vicibactin scaffold. VbsS activates L-N⁵-hydroxy-ornithine and attaches it to the PCP domain via covalent attachment as an acyl thioester. Further, the assembly of vicibactin siderophore involves acylation catalyzed by VbsA, followed by epimerization and acetylation mediated by VbsL. In the last step, trimerization and cyclization are mediated by the TE domain of VbsS (Scheme 11). The cyclotrimerization reaction, catalyzed by VbsS, is similar to that catalyzed by EntF in the biosynthesis of enterobactin.¹⁸¹

Bisucaberin, putrebactin. In siderophore biosynthesis, alternative methods of macrocyclization are also possible. Siderophores such as desferrioxamine, bisucaberin and putrebactin are all examples of natural product oligomeric macrocycles (Fig. 10). Recent studies have revealed that they undergo distinct mechanisms of oligomerization.

Fig. 10 Examples of cyclic siderophores.

The siderophore synthetase DesD, a member of the NIS synthetase superfamily, catalyses the biosynthesis of desferrioxamines in Streptomyces coelicolor M145. DesD iteratively catalyses the ATP-dependent condensation of the repeating units of N-hydroxy-N-succinyl-cadaverine to form the dimer or trimer.^182,183 In the biosynthesis of bisucaberin, the BibC multidomain enzyme from Vibrio salmonicida has been identified as catalyzing the macrocyclization of the monomer N-hydroxy-N-succinyl-cadaverine to produce bisucaberin.¹⁸⁴ The closely related analogue PubC catalyzes specific macrocyclization and cyclotrimerization in putrebactin biosynthesis (Scheme 12).¹⁸⁵

Scheme 12 The biosynthetic route to bisucaberin and putrebactin.

Valinomycin and cereulide. Valinomycin, a potassium ionophore cyclodepsipeptide antibiotic, was first identified from Streptomyces fulvissimus in 1955.¹⁸⁶ Valinomycin is a 36-membered cyclododecadepsipeptide composed of a triple repeating unit of tetrapeptide D-α-Hiv-D-Val-L-Lac-L-Val (Fig. 11).^187,188 Valinomycin has been shown to possess antifungal, antiviral, insecticidal, and anticancer activity.¹⁸⁹ The conformation of valinomycin creates a polar cavity and hydrophobic surface that can coordinate a single potassium ion. The hydrophobic surface of valinomycin allows it to enter biological membranes and transport K⁺ ions, disrupting the normal K⁺ gradient and dissipating the membrane potential, ultimately leading to cell death.¹⁹⁰

Fig. 11 The chemical structures of valinomycin and similar symmetric cyclopeptides. D-Hiv = D-α-hydroxyisovaleric acid; D-Hmp = D-alpha-hydroxy-4-methylpentanoic acid; L-Lac = L-lactic acid; L-MeLeu = L-N-methyl-leucine; D-Pla = D-phenyl-lactic acid.

Cheng et al. first reported the complete biosynthetic gene cluster of valinomycin (termed as vlm) in Streptomyces tsusimaensis ATCC 15141.¹⁹¹ The vlm gene cluster encode two NRPS enzymes: Vlm1 and Vlm2. Vlm1 (370 kDa) includes two modules: A₁-KR₁-PCP₁-C₂-A₂-PCP₂-E₂, and Vlm2 (284 kDa) includes another two modules: C₃-A₃-KR₃-PCP₃-C₄-A₄-PCP₄-TE, a total of 16 domains constituting 4 modules, each of which is responsible for the incorporation of the building blocks D-α-hydroxyisovaleric acid (D-Hiv), D-valine (D-Val), L-lactate (L-Lac), L-valine (L-Val). The NRPS domain system is homologous to the assembly of related cyclopeptide cereulide (Scheme 13).¹⁹²

Scheme 13 The biosynthetic pathway of valinomycin.

In the valinomycin biosynthesis pathway, the tetrapeptide precursor is synthesised three times, with each monomer synthesis halting at the TE domain. The Vlm2 TE domain catalyses the final oligomerization, cyclization and the release of valinomycin (Scheme 13).¹⁹³ Montanastatin was discovered as a shunt product of valinomycin. These molecules both comprise the same tetradepsipeptide basic unit, but montanastatin's basic unit only repeats twice, while valinomycin's basic unit repeats three times. Studies have shown that both valinomycin and montanastatin follow the same biosynthetic pathway, which implies that the TE domain loosely controls the basic unit repetition in the valinomycin system.¹⁹¹ It remains unclear how the TE domain can regulate the number of iterations during cyclo-oligomerization.

In 2023, Konno et al. created a substrate peptide-based p-nitrophenyl phosphonate that forms a stable covalent complex with recombinant TycC TE. This method is useful for comprehending substrate identification and substrate-TE interaction during macrocyclization.¹⁹⁴ Similarly, recent work by the groups of Schmeing and Chin has provided further structural insights into TE mechanisms, focusing on the Vlm2 TE. By employing genetic code expansion to install the amine-containing analogue 2,3-diaminopropionic acid (Dap) in the active site, they were able to stabilize and trap acyl-TE intermediates. Using this method, the first and last acyl-TE intermediates in the catalytic cycle as Dap conjugates were trapped and resolved, which provides structural insight into the conformational changes in this TE domain that control the oligomerization and cyclization of linear substrates. Moreover, this Dap substitution strategy can be widely used for protein–ligand related characterization or assays. According to this study, the Vlm TE (PDB: 6ECE) lid region could contribute by changing the shape of the depsipeptide chain to encourage intramolecular cyclization. By comparing the protein structure between tetradepsipeptidyl-TE_DAP (6ECD) and the dodecadepsipeptidyl-TE_DAP (6ECE and 6ECF), significant conformational changes could be observed in the lid region (α4–α8) (Fig. 12). It was proposed that this lid rearrangement is necessary for reorganizing the substrate during the cyclo-oligomerization.⁷⁰ In addition, by capturing the key intermediate (octadepsipeptidyl-SNAC), their investigation confirmed that Vlm TE mediates peptide oligomerization in the “reverse transfer” mode (Scheme 13), and other NRPS-TEs would also be expected to catalyse the peptide ring dimerization in this manner.

Fig. 12 The structural alterations in lid helices α4–α9 among the Vlm TE (A), tetradepsipeptidyl-Vlm TE_DAP (B) and dodecadepsipeptidyl-Vlm TE_DAP (C). The structures of Vlm2 TE and dodecadepsipeptidyl-Vlm TEDAP were overlaid for comparative analysis (D).

Shemyakin et al. first reported the chemical synthesis of valinomycin in 1963. The open-chain depsipeptide was synthesized using SPPS, which was subsequently cyclized into valinomycin using acid chloride after being cleaved from the resin. Many other reported syntheticroutes follow essentially the same strategy.^{188,195–197} In 2020, Li and colleagues developed a new biosynthetic method of valinomycin, in which they used a cell-free protein synthesis (CFPS) system to express the entire valinomycin BGC, giving rise to about 37 g L⁻¹ of valinomycin. This demonstrated for the first time that a single-pot CFPS reaction could enable the complete biosynthesis of a complex nonribosomal peptide.¹⁹⁸

Cereulide. The cereulides are series of valinomycin-like dodecadepsipeptides first isolated from Bacillus cereus.^199,200 Cereulide is an effective ionic carrier with a high affinity for potassium. As an emetic toxin, it can depolarize bacterial and mitochondrial membranes, which also causes the vomiting syndrome of human food poisoning.^201,202Bacillus strains vary in their ability to produce cereulides.²⁰³

Cereulide consists of three monomers each comprising the four amino acids D-O-Leu, D-Ala, L-O-Val and L-Val. The BGC of cereulides and isocereulides have been characterized as the ces NRPS from Bacillus cereus.^204–206 The cereulide synthetase gene cluster (ces, 24 kb) contains 7 genes: cesH, cesP, cesT, cesA, cesB, cesC and cesD. The biosynthesis of cereulide is closely controlled via post-transcriptional and post-translational regulation.²⁰⁷ The NRPSs CesA and CesB complete the assembly of three repeating tetrapeptide motifs (D-O-Leu-D-Ala-L-O-Val-L-Val) as the cereulide backbone. In addition, CesB2 is a termination module containing the TE domain, which releases the mature nonribosomal peptide by cyclization (Scheme 14).^191,208 CesB2 catalyzes the trimerization and macrocyclization of tetrapeptide substrates to finally generate cereulide. Biochemical studies have shown that this TE domain is selective for the nucleophilicity of hydrolyzed deaminopeptidyl-TE intermediates.²⁰⁹

Scheme 14 The biosynthetic pathway of cereulides.

Bisintercalators. Bisintercalator depsipeptides belong to the chromodepsipeptide family and are exemplified by echinomycin (Fig. 13). Such compounds function as DNA intercalators and are widely used in medicine as well as other fields, as bisintercalator derivatives exhibit a variety of biochemical and physical properties, including antibacterial, antiviral, antitumor, and antiparasitic properties.^138,210

Fig. 13 Examples of bisintercalator depsipeptides.

Bisintercalator depsipeptides feature a C₂-symmetric macrocyclic scaffold, which is a cyclic structure formed by various amino acids, including two identical planar bicyclic heteroaromatic chromophore moieties that are found at opposite ends of this symmetric peptide scaffold. These chromophore units are the first moieties incorporated during the assembly of these by NRPSs, and is also the reason why these compounds are known as bisintercalators (Fig. 13).²¹¹ The chromophore subunits employed in bisintercalators are 3-hydroxy-quinaldic acid (3HQA) and quinoxaline-2-carboxylic acid (QXCA).

This general structure enables bisintercalators to position the two chromophores on the peptide scaffold and in such a manner that they interact deeply with the minor groove of DNA.²¹² This structure leads to the unwinding and extension of the DNA helix, causing changes in a variety of cellular processes related to replication and transcription and that ultimately result in the death of the affected cells.²¹³

Many of these compounds (thiocoraline, triostin, SW-163 and echinomycin) share high degrees of similarity in terms of chromophores and the number and type of amino acids, which suggests that they originate from similar biosynthetic pathways.²¹⁴

Quinomycin. Quinomycin derivatives, represented by echinomycin, are an important group of antitumor antibiotics.^138,215,216 Echinomycin was first identified as a non-ribosomal peptide in Streptomyces echinatus.²¹⁵ So far, five quinomycin derivatives have been reported.^216,217 Quinomycin is formed by the polymerization of two identical tetrapeptide units (D-Ser₁-L-Ala₂-N-Me-L-Cys₃-N-Me-L-Val₄). The two peptide chains are connected by ester bonds and contain a unique connection between cysteine residues. The final product is formed by the condensation of two peptide chains with the hydroxyl group of the serine or threonine side chain.

Echinomycin B, C, D and E have been identified as being produced by the same assembly line, and occur as a consequence of the alternate incorporation of a variety of branched amino acids such as isoleucine indicating a relaxed specificity for some A-domains within the NRPS enzymatic assembly line.^218,219 Echinomycin (Quinomycin A) is the most widely studied member of the quinoxaline/quinoline antibiotic group and comprises a C₂-symmetric scaffold containing the cross-bridged cyclic octapeptide dilactone core linked by twin quinoxaline/quinoline 2-carbonyl chromophores. Like other bisintercalators, echinomycins show significant inhibitory against cancer stem cells and hypoxia-inducible factor-1 (HIF-1) by inhibition of DNA replication and RNA synthesis.²²⁰

The quinomycin biosynthetic gene cluster encodes the enzymes that perform its biosynthesis in two key steps. The first is the synthesis of quinoxalin-2-carboxylic acid (QXCA), which involves not only the secondary metabolic pathway but also the ACP protein from fatty acid metabolism.^221–223 Next, the assembly of the echinomycins is performed by an NRPS (Ecm6, Ecm7) that iteratively catalyses the assembly of the linear tetrapeptide peptide precursor. Ecm1 and FabC assist the loading of the chromophore quinoxaline-2-carboxylic acid. The echinomycin TE domain (Ecm TE), located in the final module of Ecm7, is responsible for the dimerization, final cyclization and release of echinomycin (Scheme 15).²²⁴

Scheme 15 The biosynthesis pathway of echinomycin and triostin A.

Echinomycin and triostin A possess an identical cyclic depsipeptide core that contains C₂ symmetry.²²⁵ Genome sequencing has demonstrated that triostin A and echinomycin are produced by the same BGC, with the thioacetal bridge-forming enzyme encoded by Ecm18 being responsible for the conversion of triostin A to echinomycin (Scheme 15).²²²

In the Ecm-TE-mediated cyclization, the reaction has been shown to be limited by the hydrolysis of the product.^226,227 Koketsu and colleagues designed a DNA-binding strategy to increase the catalytic efficiency of Ecm TE-catalyzed cyclization.²²⁸ This was based on the binding preference of tetrapeptide unit products to a specific DNA sequence, which can inhibit unnecessary product hydrolysis and significantly improve the yield of cyclization products.²²⁵

Koketsu et al. synthesized series octapeptide substrates with various substituted amino acid residues and chromophores. Use of these in biochemical assays suggests that the Ecm TE has a relatively broad substrate specificity, and further that the excised Ecm7 TE domain can also catalyse the dimerization of the tetrapeptidyl-SNAC substrates, generating Triostin A.²²⁸ In addition, these assays showed that the Ecm TE mediates the cyclization of pyrazine ring-containing analogues at a slower rate (K_cat/K_m = 0.046 mM⁻¹ min⁻¹) compared to a higher rate (K_cat/K_m = 3.87 mM⁻¹min⁻¹) for the naphthalene ring-containing analogue. This suggests that the ring size of the chromophore has a major influence on substrates recognition by the Ecm TE domain.

Triostin A, by virtue of being a symmetric bicyclic depsipeptide, is an easier synthetic target than echinomycin and a variety of total synthesis methods in solid or liquid phases have been developed to generate triostin A.^229–232 In all of these methods, the amino acids are first linked into linear tetradepsipeptides by SPPS, which is followed by a double cyclisation reaction in which the two tetrapeptide chains are first linked to form a linear octapeptide, and then macrocyclization is carried out with a catalyst to obtain the cyclodimeric peptide skeleton.

Sable et al. reported a different method for triostin A synthesis. Here, allyl and alloc-protecting groups were used in the generation of the linear tetradepsipeptides by SPPS, which were then removed using [Pd(PPh3)₄] and PhSiH₃ in CH₂Cl₂. Cyclisation was then achieved using N,N′-diisopropylcarbodiimide (DIC)/HOAt in a DMSO/DMF (3 : 2) solvent mixture to obtain the cyclized intermediate 6, which was cleaved from the resin by treatment with TFA/TIS/CH₂Cl₂ (10 : 5:85) in 4.0% overall yield (Scheme 16A). In an alternative route, the 2-quinoxaline carboxylic acid was coupled to the peptide after the cyclization. The desired product was then isolated in 9.1% overall yield (Scheme 16B).²³⁰

Scheme 16 Total synthesis of triostin A with different approaches.

Nagasawa et al. developed a 13-step solution-phase synthesis of triostin A, with cyclization achieved through a macrocyclization reaction. Using iodine for oxidative deprotection of the Bam group on the linear octapeptide forms a disulfide bond, and subsequently, when mixed with EDCI/HOAt at high dilution (0.001 M), the anticipated macrolide 7 was formed with a yield of 48%. Triostin A was then synthesised by removing the Cbz groups on 7 with thioanisole/TFA. This was followed by coupling two 2-quinoxaline carboxylic acid residues, which gave a 17.5% yield of the desired product (Scheme 16C).²³¹

The presence of the thioacetal bridge makes the synthesis of echinomycin and quinomycin more challenging and delayed the first complete synthesis of echinomycin until 2020. In their synthesis, the Ichikawa group prepared the ester 8 and sulfide 9 as key precursors based on their retrosythetic analysis. Simultaneous cyclization and two-directional peptide chain elongation were used to create the C₂-symmetrical bicyclic octadecadepsipeptide.²³³ The sulfur of 10 was oxidized to sulfoxide with m-CPBA, and the sulfoxide was converted into the corresponding chloride using AcCl through the Pummerer rearrangement. Finally, using the nucleophile TMSSMe and ZnCl₂ the methylthio group was introduced to generate echinomycin (Scheme 17).²³³

Scheme 17 Total synthesis of echinomycin.

SW-163C-E (Fig. 13) is produced by Streptomyces sp., and its biosynthetic pathway is similar to triostin A, thiocoraline and echinomycin. The SW-163 BGC was first annotated in Streptomyces sp. SNA15896, where fifteen genes were shown to be involved in the biosynthesis of SW-163s.²²³ Amongst these, two NRPSs (Swb16, Swb17) are involved in the assembly of the tetrapeptide chain and the final cyclisation to generate WS-163C. The SAM-dependent methyltransferases Swb8 and radical SAM protein Swb9 are responsible for the conversion of SW-163C to SW-163D and SW-163E-G, respectively.¹³⁸ In contrast with echinomycin, whose chromophore is quinoxaline, quinoline is found in SW-163-type compounds. In addition to the three common amino acids in the tetrapeptide chain of SW-163-type compounds, they also contain the rare amino acid N-Me-norcoronamic acid.²³⁴

Sandramycin. Sandramycin is an antitumor antibiotic produced by a Nocardioides sp.²³⁵ Sandramycin contains no disulfide bonds and is assembled from a C₂-symmetric cyclic decadepsipeptide moiety comprising glycine (Gly), sarcosine (Sar), N-methyl-L-valine (N-Me-Val), D-serine (D-Ser), and L-pipecolic acid (Pip) together with two QXCA molecules as chromophores (Scheme 18).

Scheme 18 Total synthesis of sandramycin.

The solution-phase total synthesis of sandramycin and its analogues was first completed in 1996 and used a strategy similar to the synthesis of other bisintercalators such as quinomycin. The symmetrical pentadepsipeptides were synthesized first, followed by coupling and macrocyclization of a 32-membered decadepsipeptide at a single secondary amide site to afford the target compound.²³⁶

Unlike the sequential peptide coupling approach, Ichikawa et al. recently performed the total synthesis of sandramycin through an Ugi three-component (11, 12, 13) reaction as the key step to obtain a linear pentadepsipeptide 14. Sandramycin was generated by the stepwise coupling of two pentapeptide monomers, with subsequent introduction of the quinaldin chromophores (Scheme 18).²³⁷ Chemical synthesis methods mostly use C₂ symmetry to synthesize the dimerized backbone of sandramycin, which limits derivatives to those containing C₂ symmetry. In 2023, the Ichikawa group further optimized the SPPS approach based on the previous coupling strategy and could synthesize desymmetrized analogues of sandramycin (Scheme 18).²³⁸

Luzopeptins. Luzopeptin A–C and quinoxapeptin A–C are C₂-symmetric cyclic depsidecapeptides bearing two heterocyclic chromophores. The luzopeptins were first isolated from Actinomadura luzonensis, and are structurally similar to sandramycin, comprising two peptide chains of five amino acid residues.²³⁹ The luzopeptins display antibacterial and antitumor activity, with the derivative quinoxapeptin C exhibiting potent HIV-1 reverse transcriptase inhibition activity.²⁴⁰

The Du group identified the biosynthesis gene cluster of luzopeptin A (luz, 48 kb) from Actinomadura luzonensis DSM 43766 and its analogue korkormicins from Micromonospora sp. ATCC 55011 (Scheme 19). Two NRPS enzymes Luz1 and Luz2 are responsible for the assembly of the peptide using the serine, tetrahydropyridazine-3-carboxylic acid (Thp), glycine (2x) and threonine units. The TE domain of Luz2 performs the final release of the product via head-to-tail cyclization (Scheme 19).²⁴¹ After release, a series post-NRPS modifications are required for the generation of mature luzopeptins. The tailoring enzymes Luz25 (cytochrome P450), Luz26 (cytochrome P450) and Luz27 (membrane acyltransferase) are responsible for the modification of the cyclic peptide to generate the non-proteinogenic amino acid residues β-OH-N-methyl-L-Val and the acyl-substituted tetrahydropyridazine-3-carboxylic acid (Thp), respectively.²⁴¹

Scheme 19 The biosynthetic pathway of luzopeptin A.

As with the synthesis of sandramycin, the synthesis of the luzopeptins commences with the preparation of the linear monomer decadepsipeptide, which is achieved through the coupling of pentadepsipeptides 15 and 16, mediated by EDCI-HOAt (CH₂Cl₂, 0 °C, 2 h). By using transfer hydrogenolysis (25% aqueous HCO₂NH₄, 10% Pd/C, EtOH/H₂O), the FMOC group and benzyl ester were cleaved, with macrocyclization (EDCI-HOAt, CH₂Cl₂, 0 °C, 2 h) providing the 32-membered cyclic decadepsipeptide, and finally generating the luzopeptins (Scheme 20).²⁴⁰

Scheme 20 Total synthesis of luzopeptins.

IB-01212. Cyclodepsipeptide IB-01212 was isolated from marine fungus Clonostachys sp. ESNA-A009. It features a C₂-symmetrical cyclic-octapeptide backbone containing two pairs of tetrapeptides containing the residues L-N, N-Me₂Leu, L-Ser, L-N-MeLeu, and L-N-MePhe (Scheme 21). IB-01212 exhibits significant cytotoxicity to different tumor cell lines.²⁴² Several different synthetic strategies towards IB-01212 have been developed, including a linear SPPS route, dimerization of the peptide fragments, and a convergent synthesis. The convergent [4+4] SPPS synthesis proved to be most economical, making it the preferred strategy for the large-scale synthesis of IB-01212 and related peptides (Scheme 21).^243,244

Scheme 21 Convergent [4+4] solid-phase synthesis of IB-01212.

Thiocoraline. Thiocoraline-type bisintercalators comprise a macrothiolactone scaffold and are an octathiodepsipeptide belonging to the family of bisintercalator natural products produced by two strains of marine actinomycetes in the Micromonospora sp. genus (Scheme 22).²⁴⁵ In contrast to other bisintercalators, thiocoraline contains 3-hydroxyquinaldic acid moieties, and its tetrapeptide chain includes N,S-dimethyl-L-Cys.²⁴⁶

Scheme 22 The biosynthetic pathway of thiocoraline.

TioR and TioS are the NRPS enzymes involved in thiocoraline biosynthesis, with TioR comprising two modules (C₁-A₁-PCP₁-E₁-C₂-A₂-PCP₂) and TioS two modules (C₃-A₃-M₃-PCP₃-C₄-A₄-M₄-PCP₄-TE).²⁴⁷ Studies suggest that 3HQA, which is activated by TioJ, is not directly loaded onto the NRPS module but rather onto the fatty acid synthase ACP domain FabC located outside the gene cluster; this phenomenon is similar to that found in the biosynthesis route of echinomycin (Scheme 22).^221,248

The TioS TE domain is the first NRPS-derived thioesterase reported to be capable of catalysing the formation of macrothiolactone and macrolactone functionalities in an iterative fashion, which is typical for the assembly of quinoline or quinoxaline carrier compounds. Robbel et al. used TioS PCP–TE to synthesise thiocoraline analogues and showed that the TioS PCP–TE could catalyse the attachment and subsequent cyclisation of tetrapeptide sulphate substrates. By expressing the TE domain of thiocoraline independently and utilizing SNAC-thiophenol mimics, it was also possible to synthesize a range of thiocoraline analogues in vitro using various activated tetrapeptides. These studies highlighted the importance of D-stereochemistry for specific residues to promote cyclization, as opposed to hydrolysis.²⁴⁹ In addition, substrate specificity studies using the TioS PCP–TE showed that the spatial requirement and polarity of the C-terminal amino acid is essential for substrate recognition, ligation and macrocyclization by this TE domain.

Boger and coworkers have reported the total synthesis of thiocoraline using a convergent strategy to prepare the precursor tetradepsipeptide. After deprotection of the amine and carboxylic acid, the key intermediate octadepsipeptide 19 was generated through the coupling of monomer tetradepsipeptides 17 and 18. The ring was closed through a second round of BOC deprotection, Tce ester deprotection and amide formation. 3-Hydroxyquinoline-2-carboxylic acid residues were attached to the scaffold after removing the Cdz-protecting group using TFA-thioanisole at 25 °C for 4 h (Scheme 23).²⁵⁰ Other thiocoraline analogues have been synthesized using a similar strategy.²⁵¹

Scheme 23 Total synthesis of thiocoraline.

Terrequinone A. Terrequinone A was originally isolated from Aspergillus terreus.²⁵² and belongs to the bisindole alkaloids family including the indolocarbazoles rebeccamycin and staurosporine (Scheme 24).^253,254 Terrequinone A features an asymmetric bisindolylbenzoquinone core, which is also present in other natural products such as cochliodinol, asterriquinone CT5, asterriquinone C1 and asterriquinone B1 (Fig. 14). Most of these compounds exhibit a range of the antifungal, cytotoxic and antitumor bioactivities.^255,256 Biochemical experiments have demonstrated that didemethylasterriquinone D is a key intermediate in terrequinone A biosynthesis.^257,258

Scheme 24 The biosynthetic pathway of terrequinone A.

Fig. 14 Example of terrequinone A and related compounds.

The five genes tdiA–tdiE were identified in Aspergillus nidulans and annotated as the biosynthetic gene cluster of terrequinone A. Walsh and colleagues reconstituted all five proteins (TdiA–TdiE) in E. coli and in doing so achieved the production of terrequinone A.²⁵⁸ The bisindolylbenzoquinone backbone is constructed by indole pyruvic acid (IPA) through dimerization, with the initial substrate IPA derived from tryptophan through the actions of the PLP-dependent transaminase TdiD.

In this pathway, TdiA comprises a single-module NRPSs, which contains three domains responsible for adenylation, substrate tethering and thioester cleavage. The single-module TdiA ends with a thioesterase (TE) domain. Biochemical investigations have suggested that the TdiA TE domain facilitates an intramolecular Claisen condensation reaction, resulting in the symmetric cyclization of two IPA molecules to generate the final product (Scheme 24). The TdiA TE domain can therefore facilitate the formation of carbon–carbon bonds. The mutagenesis of S774A, located within the conserved GXSXGG motif in the TE domain of TdiA, prevented the generation of didemethylasterriquinone D from IPA. This result provides evidence that the TE domain plays a crucial role in facilitating the cyclization of IPA monomers.²⁵⁸

Serratamolide. Serratamolide (serrawettin W1) is a surfactant lipodepsipeptide composed of two symmetric fatty acyl-serinyl (FA-Ser) moieties linked by an ester bond between a β-hydroxyl group in the fatty acid and a carboxyl group of Ser. In the BGC of serratamolide, the NRPS SwrW is proposed to assemble the peptide scaffold, commencing with the selection for Ser by the A domain. The C domain then catalyzes amide bond formation with the activated fatty acyl to give FA-L-Ser-S-PCP, which is then transferred to the TE domain. This TE domain is then responsible for the cyclization and release of the final product (Scheme 25).^259,260

Scheme 25 The biosynthetic pathway of serratamolide.

2.3 Cyclodimeric polyketides

Macrodiolides are effective drug candidates with antibacterial capabilities and have drawn great attention due to their distinctive bioactivity. Polyketides make up an important class of macrocyclic natural products. In the class of polyketides, macrolides are the most common form and are primarily biosynthesised by type I PKSs. The most common type of macrolide is a lactone, which is a cyclic ester formed from the condensation of a carboxylic acid and an alcohol. The structural characteristics of macrolides can have a significant impact on their biological activity and other properties.²⁶¹ Many macrolide-type antibiotics have been used clinically to treat bacterial infections, including erythromycin, clarithromycin, and azithromycin.

Dimeric polyketides exhibit a wide range of distinct structural characteristics and have a significant potential to form several new medicinally relevant molecules.^11,262,263 Their applications include immunosuppressive drugs, antibiotics, herbicides, fungicides, and active medications against cancer, HIV.^264,265 Dimeric polyketides are mainly generated through a dimerization process, including radical reactions, cycloadditions, esterification, and acetal formation.³³

Macrodiolides are compounds primarily synthesized via the PKS or hybrid PKS/NRPS pathway and feature a symmetrical or unsymmetrical polyketide scaffold. In PKSs assembly lines, the TE domain is responsible for iteratively synthesising two monomeric precursors before linking these together in a head-to-tail orientation, thus creating a dimer product which serves as the molecular scaffold for the cyclic diolide structure. In this section, we will mostly examine and review polyketides involving macrocyclization in their biosynthesis, with particular interested in considering how the TE domain functions in these processes.

Elaiophylin. The 16-membered diolide Elaiophylin (azalomycin B) was the first reported C₂ symmetrical macrodiolide and was isolated from Streptomyces melanosporus in 1960 (Fig. 15).²⁶⁶ Azalomycin B, efomycin E, and gopalamycin, obtained from other species of Streptomyces, were determined to be identical to elaiophylin.^267–269 Elaiophylin was reported to display antibacterial, cytotoxic and immunosuppressive activities. Despite of no activity against Gram-negative bacteria, yeast, or fungi, elaiophylin can enhance the activity of rapamycin against Candida albicans.²⁷⁰ Elaiophylin and its analogues also showed activity against drug-resistant pathogens including methicillin-resistant Staphylococcus aureus and vancomycin-resistant enterococci (VRE).²⁷¹ Recently, elaiophylin has been reported as a new type of autophagy inhibitor and anti-cancer candidate, which exerts its anti-cancer effect though inhibiting mitophagy and oxidative stress and targeting SIRT1/Nrf2 signalling.²⁷²

Fig. 15 The chemical structure of elaiophylin and its aglycone derivative elaiolide.

Due to its structural complexity and promising biological activity, several groups have completed the synthesis of elaiophylin or aglycon derivatives.^273–275 The challenges in such syntheses were the low selectivity of stereospecificity during aldol condensation. Evans et al. developed a cyclic protecting group to restrict the conformational flexibility of the ketone and thus gaining high aldol diastereoselectivity. They first synthesized the hydroxy acid 20 in five steps with carboximide as the starting compound, with cyclodimerization of 20 preformed using a modified Yamaguchi's macrocyclization strategy (Scheme 26). The dimer 21 was converted to 23 in two steps ready for crucial aldol coupling. Dialdehyde 23 was treated with the chlorophenylboryl enolate of protected ethyl ketone 24, providing the bis-aldol adduct 25 as the only detectable product. Finally, deprotection and subsequent cyclization of 25 completed the synthesis of the elaiophylin aglycon elaiolide (Scheme 26).²⁷⁶

Scheme 26 Chemical synthesis of elaiolide.

In 2004, Haydock et al. identified the putative biosynthetic gene cluster of elaiophylin from the elaiophylin-producing strain Streptomyces sp. DSM4137, which was found to contain five PKS genes encoding eight modules, consistent with the assembly of the elaiophylin polyketide chain. The C-terminal TE domain in Ela5 (Ela-TE) was shown to catalyse polyketide chain release and intramolecular lactonization to generate the matured C₂ symmetrical macrodiolide (Scheme 27).²⁷⁷

Scheme 27 In vitro activities of Ela-TE.

To study the mechanism and selectivity of macrodiolide formation, Leadlay et al. attempted to reconstitute the in vitro activity of the TE domain. Two alternative mechanisms for the formation of symmetrical diolide was proposed, similar to the cyclization steps seen in nonribosomal peptide synthetases. They synthesized SNAC thioesters of tetraketide 26 and pentaketide 27 as substrate mimics, demonstrating that incubation of pentaketide 27 with Ela-TE formed a homodimerized 16-membered decaketide diolide 28. The intermediate 29, a linear dimer linked with SNAC, and hydrolysis product 30 were also detected in these Ela-TE in vitro assays. Ela-TE was also shown to catalyze the conversion of the purified intermediate 29 into the final product 28. When pentaketide 27 and tetraketide 26 were mixed with Ela-TE, the C₂-synmmetric product 28, together with a novel “hybrid” macrodiolide 31, could be observed, demonstrating that Ela-TE exhibits substrate flexibility. These results further confirm that the elaiophylin TE iteratively catalyzes two acylation and two deacylation reactions to form the diolide (Scheme 28).²⁷⁸

Scheme 28 Alternative mechanisms for TE-catalyzed formation of symmetrical diolide.

Interestingly, a recent study has revealed that the biosynthetic pathway of elaiophylin also generates a non-dimerized product known as pteridic acid. This compound exhibits entirely different biological functions from elaiophylin, as it enhances plants' resistance to abiotic stressors. The biosynthesis of pteridic acid also relies on TE enzymes (Scheme 29), possibly due to the broad substrate specificity and catalytic versatility of TE domain.²⁷⁹

Scheme 29 The proposed biosynthetic pathway of pteridic acids and elaiophylin.

6-Deoxyerythronolide B synthase (DEBS) is a modular PKS that catalyses the biosynthesis of 6-deoxyerythronolide B, the macrocyclic core of the antibiotic erythromycin.²⁸⁰ The DEBS assembly line generate more than 50 erythromycin derivatives using combinatorial biosynthesis, which suggests promiscuity of the corresponding TE domain towards different substrates.^98,281 Boddy et al. reported that the incubation of the DEBS-TE with the non-natural SNAC thioester mimic 32 generated the 14-membered macrolactone and the hydrolysis product as major products but also minor amounts of C₂-symmetrical, head-to-tail dimerized macrodiolide (Scheme 30).⁵³ The observation of linear dimer and the 28-membered macrodiolide is consistent with the reactions catalyzed by Ela-TE.

Scheme 30 In vitro reconstitution of conglobatin macrodiolide formation.

Conglobatin and samroiyotmycins. Conglobatin, another 16-membered symmetrical macrodiolide, was originally isolated from the ionomycin producer Streptomyces conglobatus ATCC 31005 (Fig. 16).²⁸² Recently, the analogues conglobatins B–E were identified in the fermentation extracts of a soil actinomycete, Streptomyces sp. MST-91080.²⁸³ More recently still, a new conglobatin analogue, conglobatin F2, was detected from mutants of Streptomyces pactum ATCC 27456.²⁸⁴ Many studies have demonstrated the critical role of the oxazole moiety in disorazole,^285–287 an analogue of conglobatin. The oxazole moiety has been shown to be essential for its bioactivity. Conglobatin displays activity as a heat-shock protein (Hsp) 90 inhibitor as well as antitumor activity.^288,289 All conglobatins show some toxicity towards mammalian cells using the NS-1 cell line. Interestingly, the conglobatins showed a high degree of selectivity between the tumour cell line (ATCC TIB-18) and the non-tumour cell line.²⁸³ Compared to conglobatin, conglobatin F2 showed enhanced antitumor activity against HeLa and NCI-H460 cancer cell lines.²⁸⁴

Fig. 16 Chemical structure of conglobatin congeners.

In 2015, Leadlay et al. sequenced the conglobatin producer Streptomyces conglobatus and elucidated its biosynthetic gene cluster, suggesting conglobatin was biosynthesized by a PKS-NRPS hybrid. Based on this biosynthetic gene cluster, they reconstituted conglobatin macrodiolide in vitro.²⁶⁴ SNAC thioester 33 was synthesized as a substrate, which when incubated with Cong-TE was converted into the liner dimer 34 and conglobatin. Time course assays showed the production level of conglobatin increased continuously whilst 34 reached a plateau after 2 hours. This suggested that 34 could be an intermediate in diolide formation, which was then demonstrated in the conversion of purified 34 by the Cong-TE into the final product conglobatin (Scheme 31).

Scheme 31 The function of DEBS TE in the biosynthesis of erythromycin.

In an attempt to generate hybrid polyketides with Cong-TE, SNAC thioesters 35 and 36 previously synthesized for use with the elaiophylin TE were also tested as substrates. These experiments demonstrated that when the substrates 34 and 35/36 were co-incubated with Cong-TE, liner dimeric or trimeric products and their corresponding carboxylic acid could be detected. These results reveal significant subtleties exist in the iterative mechanism controlled by macrodiolide TE domains (Scheme 32).

Scheme 32 Hybrid polyketide dimers and trimers generated from mixed substrates.

The soil bacterium Streptomyces pactum ATCC 27456 is the producer of many secondary metabolites, including conglobatin, aromatic polyketide NFAT-133 and pactamycin. Several new NFAT-133 derivatives were identified from a mutant of S. pactum, with structural elucidation confirming two of these (TM-127 and TM-128) as hybrids of NFAT-133 and conglobatin (Scheme 33). In vivo gene inactivation confirmed that both biosynthetic gene clusters of NFAT-133 and conglobatin are necessary to produce the hybrids TM-127 and TM-128. The Cong-TE from S. pactum ATCC 27456 was then purified and assayed with NFAT-133 and the SNAC thioester of the conglobatin monomer; compound TM-127 was successfully formed in this assay. In contrast, the assays using inactivated Cong-TE or without Cong-TE were unable to generate TM-127. When Cong-TE was incubated with NFAT-133 and the conglobatin monomer, TM-127 was not produced, demonstrating that both ACP and TE domains are required for such hybrid formation, and further shows that Cong-TE displays broad substrate scope.²⁹⁰

Scheme 33 Proposed formation of TM-127 and TM-128 catalyzed by Cong-TE.

Recently, Zhou et al. isolated a series of compounds 38–43 from gene-inactivated mutants of the conglobatin producer S. conglobatus ATCC 31005. Two of these compounds (38/39) are new oxazole-containing esters, whilst 40 is a biosynthetic precursor of 38 and 39 (Fig. 17). Inspired by the formation of 38/39, together with TM-127/TM-128 and considering the substrate scope of Cong-TE, 69 alcohol-containing compounds bearing benzylic, allylic, propargyl, primary, secondary, tertiary, and sulfhydryl hydroxyl groups were tested as substrates for Cong-TE, which was able to form 43 new hybrid esters. When the 43 alcohols that had been accepted by the Cong-TE were individually fed to the strain, 36 of these were successfully incorporated in the Cong NRPS-PKS to yield corresponding oxazole containing esters 44in vivo (Fig. 18).²⁹¹

Fig. 17 Structures of the compounds 38–43 produced by S. conglobatus.

Fig. 18 Exotic nucleophilic alcohols accepted by Cong-TE for esterification formation.

Samroiyotmycin A, a C₂-symmetric macro diolide displaying activities against the MDR malarial strain P. falciparum K1 and lung carcinoma cell line NCI-H187, was isolated from crude extracts of Streptomyces sp. BCC33756.²⁹² Despite no investigations into the biosynthesis of the samroiyotmycins to date, the dimerization process appears similar to that found in conglobatin biosynthesis. In 2021, Hulme et al. completed the first total synthesis of samroiyotmycin A, not relying on ester-mediated dimerization but rather a one-pot alkyne cross metathesis–ring-closing metathesis (ACM–RCAM) reaction to generate the desired 20-membered macrocyclic framework (Scheme 34).²⁹³

Scheme 34 Chemical synthesis of samroiyotmycin A.

This reaction employed two molybdenum alkylidynes (47a/b) endowed with a privileged tripodal silanolate ligand. Treatment of monomer 45 with the complex 47a provided the dimer 46 in good yield by raising the temperature. Under these conditions, higher-order oligomers were also found to be formed via concentration-induced polymerization, and the yield was significantly improved when using a more dilute solution of 45. Treatment of 45 with complex 47b gave full conversion to the desired product 39, but purification of 46 was hindered by fragmentation of the catalyst ligands. Hydrostannation of 46 catalyzed by ruthenium catalyst 48 and subsequent protodestannation in the presence of copper(I) salt 49 led to successful synthesis of the desired (5E,21E) isomer samroiyotmycin A as the major stereoisomer.

Disorazoles. Disorazoles, a family of 29 macrodiolides, are intriguing polyketide-derived myxobacterial metabolites isolated from the fermentation broth of the gliding bacterial strain Sorangium cellulosum So ce12.²⁹⁴ Although the disorazoles do not appear to possess antiviral or antibacterial activity, some congeners, like disorazole A1 and disorazole C1, exhibit high activity against established animal and human cancer cell lines (Fig. 19).^295,296

Fig. 19 The structure of disorazoles.

Disorazoles are biosynthesized by a trans-AT PKS-NRPS hybrid pathway, with the trans-acyltransferase encoded by a separate gene, dszD(disD). Analysis of the domain organization of this PKS/NRPS reveals that DszC(DisC) contains one NRPS module, utilizing serine to form the oxazole, and two non-extending PKS modules located between NRPS and the TE domain. It has been proposed that a monomer is transferred to the serine residue of the TE domain from the acyl carrier protein before being dimerized with the other monomer loaded on the adjacent acyl carrier protein. Since both non-extending domains contain ACP domains along with the NRPS module, which contains a PCP domain, it is unclear which carrier proteins are involved in the dimerization of monomers (Scheme 35).²⁹⁷

Scheme 35 Biosynthesis of disorazoles.

In 2004, Wipf and Graham reported the first asymmetric total synthesis of disorazole C1.²⁹⁸ The dimer was separated into four segments and both segments 50 and 51 were utilized twice for the convergent synthesis. Sonogashira cross-coupling of 50 and 51 formed the protected monomer 52 and subsequent acylation and a second Sonogashira coupling afforded seco-disorazole C1. Selective mono-saponification of 54, followed by a Yamaguchi lactonization provided macrocycle 55. Finally, double alkyne reduction with Lindlar catalyst afforded disorazole C1 (Scheme 36).

Scheme 36 Total synthesis of disorazole C₁.

Nicolaou et al. also employed a convergent strategy to synthesize disorazoles A1 and B1.²⁹⁹ Monomers 56, 57 and 58 were constructed by the connection of different building blocks through a Wittig reaction, Suzuki coupling and Stille coupling, with a Sharpless asymmetric epoxidation exploited to synthesize the epoxy vinyl moiety in 57 and 58. Condensation of 56 and 57 by a Yamaguchi esterification provided linear diester 59. Removal of TMS from 59, followed by a Yamaguchi macrolactonization afforded the final product disorazole A1. Using the same strategy, the epoxy monomers 56 and 58 were used to synthesize the C₂-symmetrical disorazole B1 (Scheme 37).

Scheme 37 Total synthesis of disorazole A₁ and disorazole B₁.

Quartromicins. Quartromicins consist of at least six congeners A1-3 and D1-3 and were isolated from extracts of Amycolatopsis orientalis No. Q427-8. They display broad antiviral activity against herpes simplex virus type 1 (HSV-1), the influenza virus, and the human immunodeficiency virus (HIV).^296,300 Structurally, as novel 32-member macrocyclic compounds, the carbocyclic architecture of the quartromicins consists of four spirotetronic acid units that are connected by enone linkers in a head-to-tail fashion, with quartromicin A3 and D3 C₂-symmetric (Fig. 20).

Fig. 20 The structure of quartromicin.

In 2012, He et al. reported the biosynthetic gene cluster of quartromicins from the producer A. orientalis No. Q427-8. The QmnA1-QmnA2-QmnA3 PKS assembly is responsible for the synthesis of two polyketide chains through a ‘module skipping’ pathway, after which 3-oxoacyl-ACP synthase III QmnD5 catalyzes the condensation of a glycerate unit and the mature polyketide chains to form tetronates 60 and 61.³⁰¹ Dehydration of 60–61 relies on QmnD3 and QmnD4 to afford 62 and 63.³⁰² Further biosynthetic studies concerning natural spirotetronate compounds have revealed enzymatic [4+2] reactions are used to form the spirotetronate intermediate.³⁰³ A similar intramolecular [4+2] route is anticipated as the source of the four chiral spirotetronate centers found in the quartromicins (Scheme 38).

Scheme 38 Putative biosynthetic pathway of quartromicins.

Roush et al. completed the diastereoselective synthesis of the endo- and exo-spirotetronate subunits 66 and 67 of the quartromicins through an enantioselective Diels–Alder reaction of an acyclic (Z)-1,3-diene and partially assigned the stereochemical assignment of quartromicins A3 and D3.^304,305 They further synthsised the vertical and horizontal bis-spirotetronate (68) units of quartromicins A3 and D3 (Fig. 21).³⁰⁶ However, due to the structural complexity, the total synthesis of quartromicins has not been completed to date.

Fig. 21 Synthetic spirotetronate subunits for quartromicins.

15G256 polyesters. 15G256 polyesters and the menisporopsins form a group of macrocyclic polylactones produced by fungi. 15G256ι and 15G256ω are symmetric dimers/trimers discovered from the marine fungus Hypoxylon oceanicum,³⁰⁷ whilst menisporopsin A was isolated from a cell extract of the seed fungus Menisporopsis theobromae BCC 4162 (Fig. 22).

Fig. 22 Examples of 15G256 polyesters.

Menisporopsin A, 15G256ι and 15G256ω all contain 3-hydroxybutyric acid and 3,5-dihydroxy-7-(β-hydroxypropyl)-benzoic acid moieties, while menisporopsin A also contains an extra 2,4-dihydroxy-6-(2,4-dihydroxy-n-pentyl)-benzoic acid unit. The menisporopsins exhibit a broad spectrum of cytotoxic, antimycobacterial and antimalarial activity.^308,309

Concerning their biosynthesis, a ¹³C-labeling experiment has demonstrated that the pentalactone scaffold of menisporopsin A is assembled by a polyketide synthase, with all the carbons of menisporopsin A derived from acetate as the sole building block.³¹⁰

Two polyketide synthases, Men1 (HR-PKS) and Men2 (NR-PKS), were identified in Menisporopsis theobromae BCC 4162 and were shown to be responsible for the biosynthesis of menisporopsin derivatives. These two enzymes are involved in the iterative biosynthesis of the polyketide backbone and the subsequent cyclization of the molecule.

The TE domain in Men2 is responsible for the release of the final product from the PKS assembly line (Scheme 39). The esterification and cyclolactonization reactions required for the synthesis of the menisporopsins could therefore reside in this TE domain of the NR-PKS, which is similar to that of the NRPS catalyzing the elongation and cyclization of trilactone in enterobactin biosynthesis and that of modular PKSs catalyzing macrodiolide formation in elaiophylin (from Streptomyces violaceusniger DSM 413721) and conglobatin biosyntheses (from Streptomyces conglobatus).³¹¹

Scheme 39 Putative biosynthetic pathway of the menisporopsin A.

Boromycin. Boromycin, first isolated from a marine strain streptomyces antibioticus, represents a family of boron-containing natural products (Fig. 23).³¹² Boron-containing natural products, such as boromycin and the tartrolons, display broad biological activities.^313–315 Although boromycin is ineffective against Gram negative bacteria due their outer membrane blocking access to the cytoplasm, boromycin inhibits mycobacterial growth with strong bactericidal activity against actively growing and dormant drug tolerant persister bacilli.³¹⁶ Tartrolon E can also target the asexual and early sexual stages of Cryptosporidium parvum, showing high activity against multiple apicomplexan parasites.³¹⁷

Fig. 23 The representative boron-containing natural products.

The boromycins were discovered from the strains Streptomyces antibioticus ETH 28 829 and Streptomyces sp. MA4423 and are notable for being the first natural products found that contain the element boron.^318,319 Boromycin analogues Tartrolon A and B were isolated from the culture broth of Sorangium cellulosum So ce 678, which contain a similar boron binding region as is found in boromycin and aplasmomycin. The tartrolons exhibit significant antibacterial and cytotoxic activity.^314,320

Feeding experiments with ¹³C-labeled precursors have demonstrated that the biosynthesis of boromycin and aplasmomycin utilise similar assembly pathways.^321,322 Recent studies have revealed the biosynthesis of boromycin-related natural products at the genetic level. Elshahawi et al. first identified the tartrolon BGC (trtA–trtJ) from the strain Teredinibacter turnerae T7901.³²³ Among the trt cluster, trtDEF comprise three large genes that encode trans acyltransferase (AT) type I PKSs. These three multimodular PKS ORFs contain 11 modules in addition to the loading module. No detailed research concerning the mechanism of macrocyclization in this pathway has yet been performed, with the intra-module TE domain in TrtF postulated to be involved with release and cyclization (Scheme 40).³²⁴ It has been hypothesized that natural products related to boromycin undergo cyclization catalysed by a TE domain in a process similar to that observed in disorazole biosynthesis; this is due to their similar dimercyclic scaffold.³²⁴

Scheme 40 Putative biosynthetic pathway of the tartrolons.

In 2014, Avery et al. completed the chemical synthesis of the symmetrical 36-membered macrodiolide aplasmomycin A via Mukaiyama lactonization strategies. A base-promoted Chan rearrangement method was used to generate the symmetrical 36-membered diolide intermediate during the synthesis. The final product, aplasmomycin A, was generated by biomimetic modification of desboroaplasmomycin A, followed by boration reaction with trimethyl borate. This synthetic strategy is also relevant for the synthesis of boromycin (Scheme 41).³²⁵

Scheme 41 The synthesis of aplasmomycin A.

Nonactin, pamamycin. Nonactin is a member of the macrotetrolide antibiotics family. Structurally, Nonactin consists of a 32-membered ring assembled from two asymmetrical nonanoate units in a head-to-tail manner. The reported producer of nonactin includes Streptomyces griseus DSM40695, Streptomyces tsukubensis, Streptomyces chrysomallus and Streptomyces werraensis, amongst others.^326,327 Nonactin derivatives exhibit various bioactivities, including antibacterial and anticancer activity, likely due to their function as ionophores, specifically their ability to chelate potassium and sodium ions.³²⁸

The pamamycins are group of macrodiolides with antifungal activity isolated from Streptomyces alboniger.^329–332 Investigation of S. alboniger IFO 12738 revealed that the derivative pamamycin-607 is the main component in the pamamycin mixture.^333,334 The biosynthesis of pamamycin-607 has been investigated by isotopic feeding, which suggested the building blocks of pamamycin-607 were derived from the acetate, propionate, succinate and amino acid (Fig. 24).³³⁵

Fig. 24 The chemical structures of nonactin and its analogues.

Luzhetskyy et al. recently reported the identification of the nonactin and pamamycin BGCs in S. alboniger DSMZ40043 and S. sp. HKI 118.³²⁹ The nonactin BGC contains five KSs-encoding genes, and was characterized from Streptomyces griseus.³²⁹ Two KSs, NonJ and NonK, which are highly homologous to the KSs PamJ and PamK in pamamycin biosynthesis, catalyse the C–O condensation reaction of acyl CoA substrates and result in the closure of the macrodiolide ring (Scheme 42).³²⁶ Interestingly, L-valine supplementation increased pamamycin production and larger derivatives due to increased availability of CoA thioesters, but negatively affected growth and repressed pamamycin formation in the heterologous host S. albus J1074/R2.³³⁴

Scheme 42 The putative key step in the pamamycin biosynthesis.³²⁹

The total syntheses of the macrotetrolide family antibiotics have been realized through a variety of methods. The key macrocyclic reaction can be performed through macro-lactonization reaction using different catalysts (Scheme 43). Based on retrosynthetic analysis, diverse esterification routes have also been developed to realize the formation of the pamamycin dilactone backbone by coupling two precursor “fragments”.^336–342

Scheme 43 The synthetic routes towards pamamycin and derivatives.

3. Alternative approaches to macrocyclization

Natural product biosynthesis contains many strategies to perform the synthesis of carbohydrate-containing macrocycles that extend beyond macrocyclization mediated by TE domains.³⁴³ In this section, we will introduce representative examples of macrocyclization, with a focus on the synthesis of macrocyclic molecules that have not been extensively summarized.

3.1 Macrocyclization via imine formation

NRPS-mediated peptide synthesis can also be terminated through the reduction of the thioester of the terminal module to liberate an aldehyde, which then undergoes macrocyclization. The nostocyclopeptide class of macrocyclic imines, isolated by the Moore group from the terrestrial cyanobacterium Nostoc sp. ATCC53789, are cyclic heptapeptides, not dimers or oligomers, that differ in the residues present in the terminal position of the peptide.³⁴⁴ Marahiel and co-workers were first to demonstrate that the reductase domain present in the final module of the NRPS machinery was able to liberate the aldehyde heptapeptide product that then spontaneously cyclized to form the nostocyclopeptide products.³⁴⁵ Related classes of cyclic imine heptapeptide natural products isolated from cyanobacteria include koranimine³⁴⁶ and scytonemide A (Scheme 44).³⁴⁷ The facile nature of the macrocyclization has been explored via total synthesis for scytonemide A (Scheme 44),³⁴⁸ and has also shown versatility as a tool in the synthesis of peptide macrocycles more generally.³⁴⁹

Scheme 44 The cyclic imine natural products. (A) total synthesis of scytonemide A; (B) the structure of koranimine and scytonemide A.

3.2 Alchivemycin A

Alchivemycin A (74), which was isolated from a plant-derived strain of Streptomyces, displays a novel polycyclic structure containing a rare 2H-tetrahydro-4,6-dioxo-1,2-oxazine moiety.³⁵⁰ Whilst initial biosynthetic postulates for this PKS-NRPS hybrid relied on NRPS-mediated activation of N-hydroxyglycine,³⁵¹ recent work from the Tan and Ge groups³⁵² has shown that this moiety is in fact generated by the activity of the FAD-dependent monooxygenase AvmO1 that performs oxygen insertion into the amide bond of a 4-hydroxy-1,5-dihydro-2H-pyrrol-2-one ring in a cyclic precursor molecule 73. The formation of 73 has been demonstrated to commence with intermediate 70 that is itself is proposed to be formed via a [4+2] cycloaddition of the final NRPS-bound intermediate 69 together with tetramic acid formation catalyzed by the Dieckmann cyclase AvmQ.³⁵³ Formation of 72 is catalyzed by the enzyme AvmM, which is a novel enzyme class, and involves sequential dehydration to intermediate 71, followed by cyclization through Michael addition to form 72 (Scheme 45).³⁵³ The biosynthesis of alchivemycin A (74) therefore represents a fascinating array of novel biosynthetic chemistry.

Scheme 45 The synthesis of alchivemycin A.

3.3 Butelase-mediated cyclodimerization

Butelase 1 and sortase A are powerful cyclases that can mediate macrocyclization in cyclopeptide synthesis.^354,355 The Tam group have reported a biomimetic cyclo-oligomerization synthesis method using butelase 1, which is an Asn/Asp-specific ligase. This method can be used to perform the cyclization of 3–15 amino acids ending with Asn and a His-Val tail (Scheme 46). A one-pot synthesis of dimerized cyclopeptides is also possible through butelase-controlled cyclo-oligomerization. Furthermore, butelase 1 is highly effective at cyclizing macrocycles with more than 200 amino acids and displays a wide substrate recognition range covering L-amino acids and D-amino acids.^356,357

Scheme 46 Butelase-mediated cyclization in cyclopeptide synthesis.

3.4 Beauvericin, bassianolide and enniatin

Filamentous fungi are important sources of cyclopeptide natural products. In fungal NRPS systems, the terminal condensation-like (C_T) domain is often used to synthesize macrocyclic peptidyl products and is responsible for the release of the cyclopeptide product (Fig. 25).

Fig. 25 Examples of macrocyclic peptidyl products mediated by the terminal condensation-like (C_T) domain.

In comparison with TE domains, C_T domains show a different protein fold and different catalytic mechanism. The catalytic triad in the TE domain transfers the peptidyl intermediate to the active site serine, which then attacks the peptidyl-thioester in an intramolecular reaction to complete cyclization. In contrast, C_T-catalyzed cyclization does not generate a covalent adduct with the enzyme. Instead, the catalytic histidine in the C_T domain (conserved “HHxxxDG” motif) deprotonates the amine nucleophile, which attacks the thioester carbonyl bound to the PCP domain to cyclize and liberate the product.³⁵⁸ In addition, the bacterial TE-catalyzed reactions differ from the fungal C_T domain catalysed reactions in that the TE domain can accept peptidyl-S-N-acetylcysteamine (SNAC) mimics, whereas the C_T domain needs specific protein–protein interactions with the upstream PCP domain to be competent for catalysis, requiring a specific PCP domain partner. Gao et al. demonstrated that C_T domains only recognized peptide substrates tethered to the native PCP domains, while peptidyl-SNAC and peptidyl-CoA substrates were not cyclized.³⁵⁹ The interactions between the PCP and C_T domain seems play a “proofreading mechanism” for the correct peptide cyclization and prevent spontaneous hydrolysis reaction.³⁵⁸

Despite sharing the conserved motif with the canonical C domains, C_T domain is placed in a separate evolutionary taxonomy due to functional divergence. The structural basis of the unique macrocyclization activity of C_T domain has been identified through the analysis of the structural elements of C_T domains compared to canonical C domains.³⁵⁹ Specifically, although the protein sequence of C_T domains exhibits low similarity to that of canonical C domains (VibH,³⁶⁰ TycC-C6,³⁶¹ PCP₂-C₃ didomain³⁶² and SrfA-C³⁶³), its overall fold is strikingly similar to these C domains, heterocyclization (Cy) domain and epimerisation (E) domains. C_T domains comprise of two largely separate subdomains, with both N-/C-subdomains arranged in a V-shaped manner and adopting the chloramphenicol-acetyltransferase fold (Fig. 26).

Fig. 26 The overall structure of the C_T domain. The H3766, D3770, and D3773 are drawn as green sticks, while the α2 helix of C_T domain is coloured in red.

However, the main structural feature which distinguishes C_T domains from classic C domains is the replacement of the N-terminal loop in C domains with a α1 helix in C_T domains. The insertion of this α2 helix in the C_T domain results in the compaction of the two subdomains of C_T domains close to the acceptor site, ultimately blocking this substrate access channel (Fig. 26). This structural dissimilarity from canonical C domains is noteworthy, as it enables C_T domain to have distinctive macrocyclization activities.³⁵⁹

In some fungal depsipeptides synthetases, C_T domains are also involved in the catalysis of cyclo-oligomerization. Enniatin (cyclic trimer), beauvericin (cyclic trimer), and bassianolide (cyclic tetramer) are representative cyclo-oligomeric depsipeptides (CODs) that are biosynthesized as dimers, trimers, or tetramers through iterative cyclo-oligomerization by C_T domains. There are numerous impressive reviews on these types of peptides from fungi.^364–367 In this article, we will briefly summarise some of the previously reviewed material and describe some of the defining characteristics of enniatin, beauvericin, bassianolide, and PF1022A. These cyclopeptides are all assembled via the actions of iterative NRPSs (type B), in which individual modules within the NRPS assembly lines are reused for iterative cycles of peptide assembly.³⁶⁸

The enniatins and beauvericin are mycotoxin CODs found in many fungi, such as Fursarium spp., Verticillium hemipterigenum, Halosarpheia sp., and entomopathogen Beuveria bassiana, amongst others, and exhibit a wide range of biological activities, includes antibacterial, insecticidal, and anticancer activity.^369–371 Beauvericin, due to its ionophoric properties, exhibits a similar range of diverse biological activities, including antibiotic, antifungal, insecticidal, and cancer cell antiproliferative and antihaptotactic activity.^372–374 As a broad-spectrum anthelmintic, the semisynthetic PF1022A derivative emodepside (bis-para morpholino-PF1022A) has even been brought to market.³⁷⁵

The assembly of enniatin, beauvericin, and bassianolide is performed by an NRPS in each pathway. These NRPSs resemble each other, consisting of a di-module (C₁-A₁-PCP₁, C₂-A₂-MT-PCP_2a-PCP_2b-C₃) responsible for peptide synthesis (Scheme 47). As an example, the bbBeas encodes a BbBEAS nonribosomal peptide synthetase that was isolated from the strain B. bassiana and confirmed to be responsible for the beauvericin biosynthesis by targeted disruption. BbBEAS utilizes D-2-hydroxyisovalerate (D-Hiv) and L-phenylalanine (Phe) for the iterative synthesis of a predicted N-methyl-dipeptidol intermediate and forms the cyclic trimeric ester beauvericin from this intermediate in an unusual recursive process.³⁷⁶

Scheme 47 (A) The gene cluster organization of bassianolide, beauvericin, PF1022A and enniatin, and (B) their biosynthetic pathway.

Interestingly, these NRPSs uses a specific iterative assembly pathway to form the final cyclic oligomer. It is proposed that A₁ and A₂ domain adenylate the requisite amino acids (D-α-hydroxycarboxylic acids and various L-amino acids, respectively) and transfer these to PCP₁, or PCP_2a/PCP_2b. PCP_2a and PCP_2b play alternate roles as acceptors and donor in the condensation process. The Mt-domain catalyses the methylation of the amino acid residue once bound to a PCP domain. The tandem PCP domains in COD synthetases are presumed to store the depsipeptide during the process of elongation (Scheme 47). In this process, the N-terminal C domain (C₁) is inactive, while the C₂ domain catalyses the condensation of the amino acid acceptor monomers (peptidyl thioester intermediate) attached to PCP_2a/PCP_2b and the elongated intermediate donors attached to PCP₁. The C_T domain controls the length of peptide chain, the formation of the ester bond, as well as the macrocyclization and release of the cyclic peptide from the assembly line. As the peptide chain reaches a certain length, the C_T domain function shifts from elongation to cyclization, catalysing the attack of the terminal hydroxyl on the thioester, thus releasing the mature cyclopeptide. To date, the exact mechanism of this function switching in C_T domains remains enigmatic.¹⁹²

Recently, Steiniger et al. investigated the combinability of these two types of synthetases (iterative NRPSs and linear NRPSs) and compared the exchange unit variants of four hybrid modules. They demonstrated that the selectivity of the C domain influences the assembly line functionality. These experiments suggested that the PCP-C and C-A domain linker regions significantly influence the iterative peptide assembly process and production titres. Furthermore, they also suggest that swapping parts of the C_T domain can be used to uncovered functional aspects of macrocyclization and ring size control in these domains.³⁷⁷ Based on their hybrid synthetase assays, they propose a two-face XU concept for fungal NRPSs engineering, which emphasises the importance of C domain specificity and employs C-A-PCP and A-PCP-C combinatorial engineering methods, which is also potentially transferrable to other NRPS systems from a bioengineering perspective.

PfSYN (350 kDa) is the nonribosomal peptide synthetase that produces PF1022 in an iterative manner.³⁷⁸ PFSYN accepts L-leucine, D-lactate, D-phenyllactate, S-adenosyl-L-methionine (AdoMet) and ATP as substrates to assemble this cyclooctadepsipeptide scaffold.^378,379 The Süssmuth group performed a comprehensive investigation into the substrate tolerance of PfSYN, which exhibited broad substrate tolerance including aromatic, heteroaromatic and aliphatic residues. In contrast to the α-hydroxy activating domain of enniatin synthetase (ESYN), this assembly line can activate hydroxy acids derivatives containing para-halogenated phenyl rings and propargyl sidechains.³⁷⁹

Many enniatins have been identified, demonstrating that these cyclodepsipeptide synthetases possess innate flexibility in their substrate specificities and thus can incorporate a wide range of amino and hydroxy acids.^370,380 As seen with enniatin biosynthesis, the beauvericin depsipeptide synthetase (BbBEAS, 351 kDa) adopts a similar mechanism,³⁷⁶ although the beauvericin synthetase selectively accepts N-methyl-L-phenylalanine and other aliphatic hydrophobic amino acids as substrates.

In the synthesis of these cyclopeptides the main challenge is the cyclization of the linear tetradepsipeptide precursors. Following solution phase or solid phase peptide synthesis, linear tetradepsipeptide 75 is subjected to Mitsunobu esterification using [DIAD (2 equiv.), PPh₃ (3 equiv.), benzene (5 mM)]. These salt-free conditions lead to the isolation of bassianolide, with the yield of tetradepsipeptide 75 macrocyclization increased to 31% through the addition of NaBF₄ (Scheme 48).^365,381

Scheme 48 Total synthesis of bassianolide.

Verticilide is a 24-membered cyclo-oligomeric depsipeptide consisting of α-hydroxy acid and α-amino acid monomers. It was first isolated from fungus Verticillium sp. FKI-1033 in 2006 by Ōmura et al. Whilst verticilide has no antibiotic activity, it is a candidate for pesticide development.³⁸² In addition, verticilide and its derivatives have been shown to function as an inhibitor of insect RyR, ACAT2, as well as ryanodine receptors.^369,383,384 In 2006, Ōmura et al. reported the solution synthesis of verticilide based on Boc/benzyl ester protecting group macrocyclization. After the deprotection of the N- and C-termini and esterification under high dilution, verticilide was generated in a yield of 66% in 13 steps. In this synthesis, PyBrop-mediated N-Me macro-lactamization was employed to construct the acyclic oligomer (Scheme 49).³⁸²

Scheme 49 General chemical synthesis step of verticilide and enniatin B. Ghosez's reagent = 1-chloro-N,N,2-trimethyl-1-propenylamine, DMAP = 4-dimethylamino-pyridine, DIPEA = diisopropylethylamine. PyBroP = Bromo-tris-pyrrolidino-phosphonium hexafluorophosphate.

In 2012, Hu et al. reported an optimized flow-synthesis method for the cyclic hexadepsipeptide enniatin B. In this synthesis, the linear monomer hexadepsipeptide 76 was firstly constructed by the coupling of amine salts, with EDCI and Ghosez's reagent (1-chloro-N,N,2-trimethyl-1-propenylamine) used for amide and ester bond formation. In contrast with the reported verticilide synthesis, the macrocyclization step of enniatin B was catalysed using Ghosez's reagent, providing an overall yield of enniatin B of 36% (Scheme 49).^385,386

3.5 Cyclodipeptides

Cyclodipeptides (CDPs) contain a characteristic diketopiperazine (DKP) cyclic dimer scaffold, present diverse chemical structures, bioactivities, and are widely present in secondary metabolites from bacteria, fungi, plants and animals.³⁸⁷ Their skeleton comprises a stable six-membered diketopiperazine ring, which is derived from two α-amino acids condensed by peptide bonds. This stable six-membered skeleton helps compounds resist protease hydrolysis and facilitates drug passage through the intestinal and blood-brain barriers.³⁸⁸ DKPs are an important pharmacophore in medicinal chemistry and have attracted increasing interest as drug precursors. Cyclodipeptides are currently being developed as potential anticancer, antibiotic and anti-inflammatory agents, such as plinabulin, phenylahistin and Bicyclomycin.^389–391

Cyclodipeptide synthases (CDPSs) are the main biosynthetic pathways for DKP biosynthesis. CDPSs use intracellular aminoacyl-tRNA (aa-tRNAs) as substrates to catalyze the cyclization of two amino acids.³⁹² CDPS protein family are usually composed of 200 to 300 amino acid residues. In 2009, the AlbC protein derived from Streptomyces noursei was shown to catalyse the synthesis of cyclo-(L-Phe-L-Leu)-the skeleton to albonoursin, becoming the first CDPS protein whose function had been characterised.³⁹³ Many symmetrical cyclodipeptides have been reported to be synthesized by CDPS enzymes, such as mycocyclosin biosynthesized (Rv2775),³⁹⁴ pulcherrimin (YvmC),³⁹⁵ and cyclo(L-Trp-L-Trp) (Amir_4627) (Scheme 50).³⁹⁶

Scheme 50 The representative symmetrical cyclodipeptides biosynthesized through CDPS pathway.

Following DKP synthesis, tailoring enzymes encoded in the DKP gene cluster then catalyse modifications of the DKPs to generate diverse structures from the basic cyclic dipeptide skeleton. The post-modified enzymes related to CDPS synthesis that have been reported to include cytochrome P450 monooxygenases (e.g. in mycocyclosin, pulcherrimin and guanitrypmycin),^394,395,397 oxidases (in bicyclomycin)³⁹⁸ and methyltransferases (in drimentines),³⁹⁹ amongst other.⁴⁰⁰ Since there are significant numbers of reviews concerning CDPS biosynthesis, we will not elaborate this pathway further here.^{392,393,401,402}

Beyond the actions of CDPS enzymes, DKPs can also be synthesized via NRPS pathways. Gliovirin, pretrichodermamides, FA2097, aspergillazines, peniciadametizines, acetylaranotin and aspirochlorine are examples of a class of natural products originally derived from the cyclodipeptide cyclo-L-Phe-L-Phe scaffold, which is assembled by an NRPS pathway. The biosynthesis of pretrichodermamide A was recently reported, with this fungal diketopiperazine natural product featuring an α,β-disulfide bridge. Here, the DKP cyclodipeptide backbone of pretrichodermamide A is assembled by the NRPS TdaA (PCP-C-A-PCP-C domain architecture).^403,404 The NRPS AtaP from the acetylaranotin pathway, AclP from the aspirochlorine pathway, and Glv21 from the gliovirin pathway all contains a similar NRPS domain architecture (PCP-C-A-PCP-C). These NRPS synthetases are responsible for the assembly of the dipeptide, with the C-terminal C_T domain responsible for dimerization and cyclization (Scheme 50). The catalytic core of this domain resembles that of the C_T domain in beauvericin biosynthesis.^405–407

Indigoidine is a blue pigment found in different types of bacteria and it is a highly effective radical scavenger. Indigoidine exhibits a broad spectrum of medicinal, industrial potential and is biosynthesized through an NRPS pathway,⁴⁰⁸ where L-Glutamine is the precursor. Indigoidine is assembled by condensing two molecules of L-glutamine by indigoidine synthetase, which is the single-module NPRS BpsA. It comprises of two adenylation (A), one oxidation (Ox), one PCP and one TE domains, with the active holo-BpsA demonstrated to be responsible for the dimerization of two L-glutamine residues to form indigoidine (Scheme 51). Pang et al. reported that the BpsA Ox domains catalyse the key intermediate formation through oxidation.⁴⁰⁹

Scheme 51 The representative symmetrical cyclodipeptides biosynthesized through NRPS pathway.

3.6 Bipentaromycins

Bipentaromycins are dimeric aromatic polyketides featuring the rare 5/6/6/6/5 pentacyclic carbon skeleton. Cyclodimerization in these molecules is achieved via head-to-tail linkage of the two anthraquinone-based monomers. Recently, the heterologous expression of the bipentaromycin BGC bpa was performed in Streptomyces avermitilis SUKA17. From these experiments, it was proposed that cyclodimerization of an ortho-quinone methide intermediate, either spontaneously or via enzyme catalysis, initiates the 1,4-Michael addition step that leads to the formation of bipentaromycins-like dimerized intermediates (Scheme 52).⁴¹⁰

Scheme 52 The putative cyclodimerization mechanism of bipentaromycins during biosynthesis.

3.7 Cylindrocyclophanes

The cylindrocyclophanes are a class of cyanobacterial natural products that were first isolated from the terrestrial blue-green alga Cylindrospermum licheniforme. Members of the cylindrocyclophane family display various bioactivities, including antimicrobial activity, antiproliferative activity and cytotoxicity.^411–414 The cylindrocyclophanes consist of an unusual [7.7] paracyclophane scaffold, which is structurally derived from the head-to-tail dimerization of alkylresorcinol analogues. Based on isotopic feeding experiments and a retrosynthetic biosynthesis hypothesis,⁴¹⁵ the paracyclophane skeleton of the cylindrocyclophanes has been proposed to arise via dimerization of two identical resorcinol fragments, with subsequent electrophilic aromatic alkylation with a nucleophilic ion in the alkyl chain (Scheme 53).⁴¹⁶

Scheme 53 Proposed biosynthetic routes for cylindrocyclophane F.

In the cylindrocyclophane (cyl) cluster from Cylindrospermum licheniforme ATCC 29412, a hemolysin-type calcium-binding protein CylK was biochemically characterized and demonstrated that cyclodimerization reaction uses an alkyl chloride precursor via the S_N2 mechanism. The halogenase CylC mediates the chlorination, which was essential for the subsequent nucleophilic displacement of the chloride ion to generate the aryl-alky linkage. A BLAST search has demonstrated that CylC and CylK homologs are widespread in cyanobacterial genomes, offering a possible route to the discovery of new analogues of this compound class (Scheme 53).⁴¹⁷

In terms of cylindrocyclophane synthesis, the monomeric precursor thioacetate mesylate 77 was prepared before undergoing cyclodimerization using treatment with NaOMe in MeOH at ambient temperature to afford the corresponding macrocyclic bis (thioether). Oxidation of this bis(thioether) with H₂O₂ in the presence of (NH₄)₆Mo₇O₂₄·4H₂O furnished the macrocyclic bis (sulfone) 78 in 51% overall yield. Finally, the intermediate 78 was then modified to generate the cylindrocyclophanes (Scheme 54).⁴¹⁸

Scheme 54 Total synthesis of cylindrocyclophane A.

3.8 Melanthioidine

Tetrahydroisoquinoline alkaloids and dimeric macrocyclic diaryl ethers represent a large family of natural products discovered in plants.^419,420 Structurally, these compounds display dimerization using both head-to-tail and head-to-head routes. Tetrahydroisoquinoline alkaloids have been demonstrated to possess a broad range of biological activities including antitumor, anti-inflammatory, anti-hypertensive, anti-plasmodial and antimalarial activity.⁴²¹

Tubocurarine is a naturally dimeric tetrahydroisoquinoline alkaloid isolated from South American plants such as Chondodendron tomentosum.⁴²² Tubocurarine, as a neuromuscular blocking agent, has previously been widely used as a muscle relaxant during surgical procedures. However, this molecule has largely been replaced by safer and more effective drugs such as vecuronium, rocuronium and atracurium in modern procedures.⁴²³

Tubocurarine possesses a “head-to-tail”, asymmetrical, quaternary ammonium structure with two ether bridges (8-O-12′ and 11-O-7′) linking the coclaurine monomers. It has been proposed that tubocurarine biosynthesis involves a radical coupling of two enantiomeric monocationic tetrahydrobenzylisoquinolines, the two enantiomers of N-methyl-coclaurine.⁴²⁴ Subsequent methylation of the amine and hydroxyl substituents are then enzymatically performed through the consumption of S-adenosyl methionine (SAM) (Scheme 55).⁴²⁴

Scheme 55 Proposed biosynthetic routes for production of tubocurarine.

In 2016, Wang et al. completed the synthesis of the dimeric macrocyclic alkaloid melanthiodine. The enantioselective phenethyltetrahydroisoquinoline 79 was first synthesised as a monomeric precursor, with subsequent cyclodimerization performed using copper-mediated catalysis. After optimization, the yield of macrocyclic dimer O,O-dibenzylmelanthioidine 80 reached 35% when using the combination of the soluble copper salt CuBr·SMe₂ (1 equiv.), Cs₂CO₃ (3 equiv.), pyridine (0.16 M), 150 °C, 48 h to catalyse this reaction (Scheme 56).⁴²¹

Scheme 56 Total synthesis of melanthiodine using copper-mediated cyclodimerization.

3.9 Griffipavixanthone

Griffipavixanthone and garciobioxanthone are group of dimeric xanthones isolated from plants that exhibit a wide range of biological activity, including antitumor and antioxidant properties.^425,426 Structurally, unlike other cyclic dimeric PKS products such as gossypol or cyclic dimeric PKS products such as hypericin (both of which are linked by aryl–aryl or C–O bonds), griffipavixanthone and garciobioxanthone possess a unique cyclodimeric backbone consisting of two monomeric tetrahydroxyxanthone units.

A Diels–Alder cycloaddition reaction was recognized to be the key step during the biosynthesis of these natural products. Using griffipavixanthone as an example, retrosynthetic analysis suggested that the cyclodimeric structure would be derived from the prenylated xanthone monomer 82, and the core cyclohexene ring would be formed through [4+2] cycloaddition (Scheme 57).⁴²⁷ Oxygenated styrenes are the key intermediates for this cyclization, which is initiated by the formation of a benzyl cation that then undergoes Friedel–Crafts cyclization after adding to the C–C double bond of another styrene molecule.

Scheme 57 Total synthesis of griffipavixanthone.

In 2017, Porco and co-workers reported the first biomimetic total synthesis of griffipavixanthone (81). The key synthetic step is the formation of a highly strained cyclohexene ring generated through an intermolecular cationic [4+2] cycloaddition reaction. The monomer intermediates vinyl para-quinone methide 83 and an in situ generated isomeric 1,3-butadiene 84 were afforded through isomerization mediated by Lewis or Brønsted acids (Scheme 56).⁴²⁷ In 2018, Schaus, Porco and co-workers achieved the asymmetric synthesis of (+)-griffipavixanthone employing a chiral phosphoric acid-catalyzed cycloaddition.⁴²⁸

3.10 Cyanolide A

Cyanolides are glycosidic macrodiolides possessing C₂ symmetry discovered in the cyanobacterium Lyungbya bouillonii that exhibit strong molluscicidal activity against Biomphalaria glabrata (Fig. 27).⁴²⁹ Cyanolide A contains a symmetrical dimeric backbone, with the monomeric unit of cyanolide A derived from a polyketide chain containing five acetate units that is modified by SAM-derived methylation.

Fig. 27 Examples of cyanolides, clavosolides, carpaines and azimines.

Many studies concerning the synthesis of cyanolide A have been reported.^430,431 To summarise, two main synthesis routes have been explored, which either perform glycosylation before or after cyclodimerization.

To afford the C₂ symmetric diolide, macrocyclization reactions mainly exploit the Shiina lactonization strategy.^430–432 The Krische group has reported the most direct six-step total synthesis of cyanolide A starting from neopentyl glycol. The synthesis uses no protecting groups and chiral auxiliaries, with the key intermediate C₂-symmetric diol 85 constructed using an asymmetric iridium-catalysed dehydrogenation/allylation strategy (Scheme 58).⁴³³

Scheme 58 Total synthesis route for cyanolide A.

3.11 Clavosolide

Clavosolide A is a C₂-symmetric 20-membered glycosidic macrodiolide discovered from the sponge Myriastra clavosa.⁴³⁴ Clavosolide A and derivatives have been chemically synthesized in a variety of ways.^435–438 Recently, Krische and colleagues reported a 7-step total synthesis route for clavosolide A. Following retrosynthetic analysis, the key intermediate hydroxy acid 86 was firstly prepared, followed by macrolactonization to afford clavosolide A in 52% yield (Scheme 59).⁴³⁸

Scheme 59 Total synthesis route for clavosolide A.

3.12 Azimine and carpaine

Carpaine and azimines are class of alkaloids found in certain species of plants, such as Carica papaya, Uncaria tomentosa and Uncaria guianensis.⁴³⁹ They are structurally classified as alkaloids, a class of naturally occurring compounds that contain a macrocyclic dilactones containing a 2,3,6-trisubstituted piperidine skeleton.

Carpaine has been shown to possess a variety of biological activities, including antimicrobial, antiparasitic, and anti-inflammatory effects. Carpaine has also been investigated for its potential use in the treatment of various diseases, such as cancer, heart disease, and neurodegenerative disorders.^440,441 The effects of carpaine may be related to its macrocyclic dilactone structure, a possible cation chelating structure.⁴⁴² The biosynthesis of carpaine is not well understood, but is hypothesised to begin with the decarboxylation of the amino acid tyrosine to form tyramine, which is then further metabolized to form the alkaloid. The intermediate products of this pathway are believed to be modified and rearranged through additional enzymatic reactions to form the final carpaine molecule.

Taro et al. first reported the retrosynthetic routes of (+)-azimine and (+)-carpaine, where the N-derivatives of azimic acid and carpamic acid were used as starting substrates. Taking the example of (+)-carpaine synthesis, the precursor N-Cbz-carpamic acid was di-lactonized under Yamaguchi macrocyclization condition to generate the cyclodimerized N-Cbz-carpaine in 71% yield. Subsequent deprotection led to the isolation of (+)-carpaine (Scheme 60).⁴⁴³

Scheme 60 Synthetic routes of (+)-azimine and (+)-carpaine.

3.13 Glycolipids

Macrolactone-containing glycolipids are glycosyl derivatives of lipids widely present in microorganisms. Glycolipids possess C₂-symmetrical core structures that are formed through cyclodimerization, with some derivatives containing a C₃-symmetrical scaffold.^444,445 Representative glycolipids include the glucolipsins, cycloviracins, macroviracin, fattiviracins and arthrobacilin (Fig. 28). To date, few of these compounds have been studied in terms of their biosynthesis.

Fig. 28 Examples of macrolactone-containing glycolipids.

3.14 Glucolipsins and cycloviracin

Glucolipsin A and B are macrocyclic dilactones produced by Streptomyces purpurogeniscleroticus WC71634 and Nocardia vaccinii WC65712.⁴⁴⁶ The glucolipsins demonstrate activity in relieving the inhibition of glucokinase by stearoyl-CoA.⁴⁴⁷ In the glycolipid family, the cycloviracins are glycoside derivatives of the glucolipsins and thus contain a related aglycone. Cycloviracin B1 was discovered to be produced by the actinomycete Kibdelosporangium albatum so. nov. (R761-7) and exhibits significant antiviral activity against influenza A virus, varicella-zoster virus, and human immunodeficiency virus, amongst others.⁴⁴⁸

A synthesis route towards glucolipsin A exploited the cyclodimerization of monomer 87, with the key cyclic dimer intermediate 88 obtained through macrodilactonization, with the reaction using the activating agent 2-chloro-1,3-dimethylimidazolinium chloride (89). The cyclodimerized product 88 was obtained in 54% yield using optimized conditions (Scheme 61).^446,449

Scheme 61 Total synthesis of glucolipsin A.

The synthesis of cycloviracin B was also enabled using 2-chloro-1,3-dimethylimidazolinium chloride (89) in the macrodilactonization step.^449–451 Interestingly, it was observed that the addition of KH significantly increases the yield of 90 to 71% (Scheme 62), and from this it was postulated that the presence of potassium cations may structure the precursor molecules in a way that favours cyclodimerization.

Scheme 62 Total synthesis of cycloviracin B and macroviracin A.

3.15 Macroviracin

The macroviracins are a class of C₂-symmetric macrocyclic compounds, which were isolated from the mycelium extracts of Streptomyces sp. BA-2836 and classified into eight types of congeners (A–D) by the length (C₂₂ or C₂₄) of the constitutive fatty acids. These cyclic glycolipids exhibit a powerful antiviral activity against HSV-1 and VZV, with their potency is 10 times higher than acyclovir.⁴⁵² Takahashi and co-workers first reported the synthesis of macroviracin A, which was constructed using the precursor C₂₂-hydroxy carboxylic acid 91 (glucosyl fatty acid) and subsequent intermolecular cyclodimerization. Under optimized conditions (2-chloro-1,3-dimethylimidazolinium chloride, DMC/DMAP, sodium hydride), the dimeric product was obtained in 35% yield, whilst also avoiding the unwanted side products caused by lactonization of the glucosyl fatty acid 91 (Scheme 62).³⁴³

3.16 Fattiviracins

The fattiviracins are a group of antiviral antibiotics discovered from strain Streptomyces microflavus.⁴⁵³ They exhibit strong antiviral effects against several viruses, such as herpes simplex virus, influenza A virus, and human immunodeficiency virus.⁴⁵⁴ The fattiviracins consist of macrocyclic diesters, which are created by the linkage of two trihydroxy fatty acids and two D-glucose monomers.⁴⁵⁵ The water solubility of fattiviracin A1 is facilitated by the presence of four sugar moieties in its structure. This characteristic has the potential to make fattiviracin A1 a promising new antiviral drug with a broad spectrum of activity.⁴⁵³

3.17 Arthrobacilin

The arthrobacilins are cyclic glycolipids containing a 27-membered ring scaffold bearing multiple hydroxyl groups. These compounds were discovered from the strain Arthrobactor sp. NR2967,⁴⁵⁶ and exhibit weak cytotoxic and antibiotic activity.⁴⁵⁷ In 1994, Garcia et al. reported the synthesis of the arthrobacilins, with analogues synthesized using the monomer ω-hydroxylic acid 92 as the substrate. Treatment of key intermediate 93 with 2-chloro-1,3-dimethylimidazolinium chloride (DCC)/DMAP catalysed the key cyclisation, followed by benzyl deprotection afforded the natural product arthrobacilin A. (Scheme 63).⁴⁵⁸

Scheme 63 Total synthesis of arthrobacilin A.

4. Chemical routes to macrocyclic scaffolds

Despite recent advances in natural product synthesis, accessing complex cyclodimeric compounds through cyclodimerization and macrocyclization with high synthetic efficiency remains challenging. The specific methods used for cyclodimerization depend on the structure of the target compound and the availability of precursor molecules and reagents. General cyclodimerization approaches were typically developed from classic macrocyclization mechanisms, which mainly include ring-closing metathesis (RCM), multicomponent reactions (MCRs), cycloaddition reactions, macrolactonization, macrolactamization, click reactions and cross-coupling reactions. Thus, diverse macrocyclization methodologies have been developed, with such reactions having been extensively reviewed.^459,460

Here we instead seek to emphasize the chemical synthesis reactions developed to generate cyclodimeric scaffolds. As most of the synthetic strategies for cyclodimerization and macrocyclization have been described in the above sections on individual cyclodimerized natural products, here we will restrict ourselves to the introduction of representative examples of cyclodimerization or cyclo-oligomerization related to NPs or NP-like small molecules. The chemical synthesis of cyclic supramolecular polymers will not be discussed.

4.1 Lactonization/lactamization

The generation of amide, ester, thioester, disulfide, olefin, or C–C bonds is a common route for achieving peptide cyclization. In cyclopeptide synthesis, the final ring-closing reaction is usually accomplished through lactamization, lactonization, or the generation of a disulfide bridge.^27,459

Macrocyclic structures with one or more ester connections are known as macrolides. Lactonization often represents an efficient approach for the synthesis of macrolides.⁴⁶¹ Porco and co-workers reported distannoxane-catalyzed cyclodimerization, which is mediated by distannoxane transesterification catalysts that facilitate the production of dimeric macrodiolides from various hydroxy esters substrates. They reported that substrate enantioenriched hydroxy esters are readily able to generate the 14- to 22-membered cyclodimerization reaction when using distannoxane transesterification catalysts (Scheme 64).⁴⁶² Studies have shown that this is also an effective strategy to create stereochemically enriched homodimers from a variety of hydroxy esters monomer pairs. The effectiveness of cyclodimerization reactions has been shown to depend significantly on both microwave power and reaction temperature employed for such reactions.⁴⁶³

Scheme 64 Distannoxane-catalyzed cyclodimerization.

Distannoxane catalysts effectively promote the macrolactonization of seco-ester precursors. Collins and co-workers reported a reaction protocol for macrodiolide synthesis based on seco-acids using Lewis acid Hf (OTf)₄ as catalyst, with the only by-product generated in this reaction being water. After optimization, this method was shown to be suitable for the synthesis of the 22-membered macrodiolide 95, and macrocycles 94 and 96 through cyclodimerization (Scheme 65).⁴⁶⁴

Scheme 65 The macrodiolide synthesis based on seco-acids using Lewis acid Hf (OTf)₄.

Zhang et al. reported a facile synthesis method for cyclic polyamides through cyclodimerization, with monomer pentafluorophenol esters prepared in advance. After deprotection, the two active groups (NH₂ and CO₂C₆F₅) were exposed on both termini, which promoted macro-lactamization by monomers cyclodimerization in an alkaline environment. (Scheme 66).⁴⁶⁵

Scheme 66 Total synthesis of cyclic polyamides.

Miao et al. investigated the autocatalytic polymerization reaction of tripeptides linked by disulfide bond using (Cys-Xxx-Gly-SEt)₂ as the “monomer” starting material. The resulting macrocyclic peptides, containing up to 69 amino acids, displayed autocatalytic kinetics and were formed via native chemical ligation; thiol–thioester exchange “hopping” was likely responsible for this behaviour. This method allows for the straightforward production of thiol-rich macrocyclic oligopeptides, which can be used as a tool for studying functional cyclopeptides (Scheme 67).⁴⁶⁶

Scheme 67 Autocatalytic polymerization reaction for the synthesis of macrocyclic peptides.

4.2 Multicomponent reactions

Multicomponent reactions (MCRs) have proven to be powerful macrocyclization tools for cyclopeptide synthesis. This method has been developed to synthesize heterocycles, polymers, and natural products, with numerous reviews written about this topic.^134,467 Well-known MCRs strategies for cyclodimerization include azide–alkyne cycloadditions such as the “click” reaction, Ugi reaction and Ugi-azide reaction. These methods are especially suitable for the synthesis of cyclic peptide structures, with Ugi multicomponent stapling approaches especially suitable for cyclizing helical peptides (e.g. gramicidin S and sandramycin) as described in the Section 2.2 and Scheme 68.

Scheme 68 The generation of macrocyclic rings by copper-catalysed azide–alkyne cycloaddition.

Click chemistry has been powerful strategy for chemical backbone synthesis and biomimetic applications.⁴⁶⁸ Many click reactions have proven to be efficient peptide macrocyclization methods. The azide–alkyne cycloaddition methodology is a valuable technique for generating macrocyclic rings.⁴⁶⁹ Based on the 1,3-dipolar cycloaddition of azides and alkynes, triazole-containing macrocycles can be readily synthesized. The Liskamp group developed the synthetic route of the ABC ring system of vancomycin through ruthenium-based azide–alkyne cycloaddition (RuAAC). This macrocyclization method exhibited high intramolecular selectivity to mimic the bicyclic formation.⁴⁷⁰

Jagasia et al. reported a copper-catalysed azide–alkyne cycloaddition reaction (CuAAC) that combined the methods of SPPS and click chemistry. This method can also be applied to perform the head-to-tail cyclodimerization of resin-bound oligopeptides. The advantage of this method is that this reaction was independent of peptide sequence (Scheme 69).^471,472

Scheme 69 The generation of macrocyclic rings by copper-catalysed azide–alkyne cycloaddition.

The Ghadiri lab has reported a solution-phase synthesis of C₂ symmetric cyclic peptide scaffolds through 1,3-dipolar azide–alkyne cycloaddition reactions. This technique utilised a tandem dimerization-macrocyclization approach enabled by facile click reactions of an azido-dipeptide alkyne.⁴⁷³

The azido-alkyne “click reaction” strategy also can be used in the synthesis of cyclodextrin analogues, with the cyclodimerization of an azido-alkyne saccharides performed via Cu-catalyzed dipolar cycloaddition of alkyne and azide functional groups (Scheme 70).^474,475 Menand et al. developed an efficient domino Staudinger aza-Wittig reaction to synthesis sugar-aza-crown ethers through the cyclodimerization of C-glycosyl azido aldehydes (Scheme 70).^476,477

Scheme 70 The synthesis of cyclodextrin analogues through azido-alkyne “click reaction” strategy.

The exploration of active precursors for efficient cyclodimerization synthesis has been a longstanding objective of organic chemists. Recently, vinylethylene carbonates (VECs) were discovered as promising dipole precursors to iridium-catalysed cascade allylation-macrolactonization reactions.⁴⁷⁸ Wang and co-workers developed a wide range of C₂-symmetric chiral macrodiolides based on VEC substrates and isatoic anhydride analogues (Scheme 71). Using palladium-mediated catalysis, VECs were converted into the key zwitterionic p-allyl palladium intermediate. Subsequently, this intermediate was used to perform diverse reactions including cyclodimerization, which also maintains high diastereo- and enantioselectivity in the final products.⁴⁷⁹

Scheme 71 The iridium-catalysed cascade allylation-macrolactonization based on vinylethylene carbonates precursors.

4.3 Coupling reactions

Pd-catalysed cross-coupling reactions have been widely applied in many fields related to pharmaceutical development and natural products synthesis.⁴⁸⁰ Mendive-Tapia et al. reported a cyclodimerization method involving the coupling between tryptophan and L-phenylalanine units to yield the aryl-indole coupled peptide through the Pd-catalysed C–H activation reaction. In addition, the residue number between para-L-Phe and Trp effects the catalytic efficiency of this palladium-catalyzed C–H arylation, with molecules containing short linkers (i, i + 1-meta/para; i + 2-para) tending to afford cyclodimerization. Pd-catalysed C–H activation has also been used to cyclise monomers 97 to generate DKPs 98 with yields of 24% (meta-regiochemistry) and 28% (para-regiochemistry), respectively (Scheme 72).⁴⁸¹

Scheme 72 Pd-catalysed cross-coupling reactions to cyclopeptide formation.

4.4 Cycloaddition reactions

Cycloaddition reactions are an important way to catalyse the cyclodimerization of unsaturated substrates.^482,483 In natural product biosynthesis, the most representative cycloaddition reaction is the Diels–Alder reaction, which catalyses the concerted [4+2] cycloaddition of a precursor diene and a dienophile to form a new 6-membered ring, containing two new C–C bonds. In the case that a heteroatom is involved, this is known as a hetero-Diels–Alder reaction.^484–486 Cycloaddition reactions are especially useful for the formation of small-to-medium size cycles and are present in the biosynthesis of many natural products. As a result, many enzymes have been reported as cyclases catalysing [4+2], [2+2] or [6+4] cycloaddition reactions.^37,487–489

In organic synthesis, cycloaddition reactions are regarded as one of the most effective strategies for constructing cyclic structures. Cycloaddition dimerization obviously differs from the cyclodimerization reaction because cycloaddition involves the interaction between two distinct reactants, usually a conjugated diene and a dienophile, resulting in the creation of a new cyclic molecule. In contrast to cyclodimerization, cycloaddition reaction forms a single cyclic product, not a dimer made of identical or similar reactants. Cycloaddition reaction have also been extensively reviewed.^490–493

5. Conclusion and perspectives

With the increasing number of cyclic peptides being developed for use as therapeutic agents, there is great need for efficient and cost-effective synthetic methods for the large-scale synthesis of cyclic molecules. The efficient synthesis of large libraries of cyclic peptides or cyclic peptide-like molecules containing non-natural amino acids (e.g.D- or non-natural amino acids for increased stability and diversity) is also of significant importance.⁴⁹⁴ However, the efficient cyclisation of peptides using traditional chemical synthesis currently remains a challenge. Many studies have shown that enzymatic domains found in the modular biosynthetic pathways of PKSs and NRPSs can carry out catalytic cyclisation reactions in vitro with a wide range of substrates.^4,495 This can enable cyclisation reactions that are challenging to carry out using conventional chemical synthesis.

In this review, we summarised the structural traits, distribution, biological functions, and uses of natural products that feature cyclodimeric/cyclomultimeric scaffolds, along with a summary of what is known regarding their biosynthesis and origins. We sought to pay particular attention to the role played by the TE domain of PKS and NRPS machinery in the macrocyclization process. Among these compounds, the most representative examples are gramicidin S and tyrocidine, in which the catalytic mechanism of these TE domain have been relatively well-studied. In addition, we also introduced examples of alternate catalytic cyclization reactions, such as the butelase- and C_T-mediated macrocyclization of peptides, as well as the formation of cyclodipeptides mediated by CDPSs and single-module NRPSs. Whilst there are several alternate mechanisms found in nature that perform cyclisation within biosynthetic assembly lines, arguably the most relevant for cyclodimerization is the role of TE domains, given their potential application as biocatalysts. These domains have been widely investigated for their biosynthetic and biocatalytic potential, and whilst we now know significant details concerning the mechanism of cyclisation or dimerization performed by TE domains, there is far less known about the aspects of these domains that control the substrate specificity and positioning that gives rise to their specific products.

Examples of this phenomena can be seen in the activity of the TE domains from gramicidin S (GrsB), tyrocidine (TycC) and WS9326A/mohangamide biosynthesis. The GrsB TE efficiently catalyses the dimerization of assembled pentapeptides to form the cyclic decapeptide gramicidin S, with additional potential for chemoenzymatic analogue synthesis. The TycC TE has been widely studied and cyclises the tyrocidine precursor and a range of modified peptides and is also able to dimerize the gramicidin S pentapeptide, thus emphasizing its versatility. The ability of both TycC TE and GrsB TE to accommodate variations in substrate sequences while maintaining catalytic efficiency suggests a degree of plasticity in their active sites. This plasticity enables these enzymes to recognise and cyclise substrates with similar structural features, even if they are derived from different biosynthetic pathways or synthetically derived.^124,125 An excellent example of this substrate promiscuity is the use of the TycC TE domain to generate libraries of tyrocidine derivatives with modified amino acid residues at position 4 of the peptide.¹³¹ Kohli et al. noted that the structure of tyrocidine—an amphipathic β-sheet—can also be used to predict the propensity of the TE domain to accept modified residues at P4 in terms of how these might disrupt the structure of the peptide, thus suggesting preorganization of the peptide within the TE must be important for cyclisation. This is in addition to constraints placed on the activity of TE domains (and related PBP-type TE domains) that is seen in the amino acid specificities of such enzymes at the N- and C-termini of the peptide substrates.^126,128 The ability of the TycC TE domain to generate the dimeric gramicidin S-structure is also highly significant in this regard, although again there are restraints placed in terms of the amino acids that are accepted at the peptide termini and the structure adopted by the final cyclic peptide in this case. This raises a more general question concerning the importance of preorganization of the substrate for cyclisation by TE domains—something that has largely been investigated in peptide biosynthesis only—and could well call into question the ability to apply TE domains to the cyclisation of substrates that differ significantly from the natural products produced by these enzymes.

Nonetheless, the impressive tolerance of the PBP-type TE SurE for different ring sizes shows that this limitation can be overcome, at least for some systems.³⁹ The TE domain from WS9326A/mohangamide biosynthesis is also a fascinating example of how we are yet to understand the substrate positioning governed by these domains.^142,143 In this case, the results of in vitro cyclisation assays clearly show that the differentiating feature in whether the TE domain catalyses a standard cyclisation (to afford WS9326A) or a pseudo-dimerization (to generate mohangamide) is the structure of the acyl unit present on the peptide monomer. Whilst it appears as though mohangamide is likely an unusual minor product of this pathway due to the loading of an unusual dihydropyridine-containing acyl unit, this raises the question of how this seemingly minor alteration to a single building block can completely alter the specificity of the reaction towards dimerization. This requirement for the presence of two different monomers to afford mohangamide is also tantalizing for the generation of controlled dimers via the use of TE domains, as the dihydropyridine-containing acylated peptide appears unable to undergo cyclisation without the presence of the alternate monomer.¹⁴⁰ Taken together, these results clearly show that the field requires far more insights into the binding and dynamics of peptides within TE domains in order to understand and exploit such catalytic behaviour.

The interplay of the substrates with the TE domain makes the value of isolated crystal structure without substrate bound of somewhat reduced value, and even the pioneering work of Chin and Schmeing demonstrated the resistance of TE domain complexes to crystallographic investigation, at least from the perspective of explaining the noted cyclisation behaviour of the TE domain studied in this case.⁷⁰ This strongly suggests that a greater application of computational approaches coupled with in solution biophysical approaches (as have been demonstrated recently for NRPS systems in pioneering work from Mootz and co-workers) is needed to understand these phenomena more completely.⁴⁹⁶

Despite their challenges, TE domains do offer the major advantage over C_T domains in their ability to accept soluble SNACs as substrates, which is a major advantage when exploring the potential for cyclodimerization by such enzymes. Additional examples of TE domains accepting ester substrates provides further support for the value of TE-domains as biocatalysts, which is due to the synthesis of such substrates is significantly simplified because of the improved stability of ester linkages during solid phase peptide synthesis. An important area of future investigation—particularly in regard to pseudo cyclodimerization—is to understand how acyl moieties can be used to control the specific dimerization of two substrates. This would be a very powerful tool for synthetic biology and would have great potential in generating libraries of larger cyclic peptides from a smaller pool of appropriately protected linear intermediates. These findings collectively underscore the versatility and substrate tolerance of these TE domains observed in the NRPS assembly in nature, shedding light on their potential applications in peptide engineering and synthesis.

Data availability

Data availability is not applicable to this article as no new data were created or analyzed in this review article.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

This work was supported by National Natural Science Foundation of China (no. 82104044 to Songya Zhang, no. T2250710184 to Youming Zhang), Shenzhen Science and Technology Innovation Commission (no. ZDSYS20220303153551001 to Youming Zhang), CAMS Innovation Fund for Medical Sciences (CIFMS, no. 2021-I2M-1-029 to Ran Shi, no. 2022-I2M-2-002 to Hai-Yan He), the Non-profit Central Research Institute Fund of Chinese Academy of Medical Sciences (no. 2021-RC350-004 to Hai-Yan He), National Natural Science Foundation of China (no. 22277146 to Hai-Yan He) and National Science Foundation of Beijing Municipality (7244468 to Shuai Fan); This research was also conducted by the Australian Research Council Centre (CE200100012) and funded by the Australian Government.

References

1. A. G. Atanasov, S. B. Zotchev, V. M. Dirsch, The International Natural Product Sciences and C. T. Supuran, Nat. Rev. Drug Discovery, 2021, 20, 200–216.
2. F. Kopp and M. A. Marahiel, Nat. Prod. Rep., 2007, 24, 735–749.
3. D. J. Newman and G. M. Cragg, J. Nat. Prod., 2020, 83, 770–803.
4. D. G. Jimenez, V. Poongavanam and J. Kihlberg, J. Med. Chem., 2023, 66, 5377–5396.
5. K. Tang, S. Wang, W. S. Gao, Y. H. Song and B. Yu, Acta Pharm. Sin. B, 2022, 12, 4309–4326.
6. H. Zhang and S. Chen, RSC Chem. Biol., 2022, 3, 18–31.
7. E. M. Driggers, S. P. Hale, J. Lee and N. K. Terrett, Nat. Rev. Drug Discovery, 2008, 7, 608–624.
8. Y. U. Kwon and T. Kodadek, Chem. Biol., 2007, 14, 671–677.
9. J. Samanen, F. Ali, T. Romoff, R. Calvo, E. Sorenson, J. Vasko, B. Storer, D. Berry, D. Bennett and M. Strohsacker, et al. , J. Med. Chem., 1991, 34, 3114–3125.
10. J. Sun, H. Yang and W. Tang, Chem. Soc. Rev., 2021, 50, 2320–2336.
11. J. Liu, A. Liu and Y. Hu, Nat. Prod. Rep., 2021, 38, 1469–1505.
12. T. Wezeman, S. Brase and K. S. Masters, Nat. Prod. Rep., 2015, 32, 6–28.
13. A. Paquin, C. Reyes-Moreno and G. Berube, Molecules, 2021, 26, 2340.
14. A. Nudelman, Curr. Med. Chem., 2022, 29, 2751–2845.
15. Y. C. Park, G. Rimbach, C. Saliou, G. Valacchi and L. Packer, FEBS Lett., 2000, 465, 93–97.
16. N. G. M. Gomes, R. B. Pereira, P. B. Andrade and P. Valentao, Mar. Drugs, 2019, 17, 551.
17. K. S. Lam, G. A. Hesler, J. M. Mattei, S. W. Mamber, S. Forenza and K. Tomita, J. Antibiot., 1990, 43, 956–960.
18. G. Berube, Curr. Med. Chem., 2006, 13, 131–154.
19. Z. E. Perlman, J. E. Bock, J. R. Peterson and R. S. Lokey, Bioorg. Med. Chem. Lett., 2005, 15, 5329–5334.
20. R. D. de Vries, K. S. Schmitz, F. T. Bovier, C. Predella, J. Khao, D. Noack, B. L. Haagmans, S. Herfst, K. N. Stearns, J. Drew-Bear, S. Biswas, B. Rockx, G. McGill, N. V. Dorrello, S. H. Gellman, C. A. Alabi, R. L. de Swart, A. Moscona and M. Porotto, Science, 2021, 371, 1379–1382.
21. P. W. Schiller, T. M. Nguyen, C. Lemieux and L. A. Maziak, FEBS Lett., 1985, 191, 231–234.
22. M. L. Huang, S. B. Y. Shin, M. A. Benson, V. J. Torres and K. Kirshenbaum, ChemMedChem, 2012, 7, 114–122.
23. V. Razakantoanina, N. K. Phi Phung and G. Jaureguiberry, Parasitol. Res., 2000, 86, 665–668.
24. I. Takahashi, S. Nakanishi, E. Kobayashi, H. Nakano, K. Suzuki and T. Tamaoki, Biochem. Biophys. Res. Commun., 1989, 165, 1207–1212.
25. J. Skubnik, V. S. Pavlickova, T. Ruml and S. Rimpelova, Biology, 2021, 10, 849.
26. Y. Huang, M. M. Wiedmann and H. Suga, Chem. Rev., 2019, 119, 10360–10391.
27. C. Bechtler and C. Lamers, RSC Med. Chem., 2021, 12, 1325–1351.
28. V. Marti-Centelles, M. D. Pandey, M. I. Burguete and S. V. Luis, Chem. Rev., 2015, 115, 8736–8834.
29. C. J. White and A. K. Yudin, Nat. Chem., 2011, 3, 509–524.
30. M. A. Fischbach and C. T. Walsh, Chem. Rev., 2006, 106, 3468–3496.
31. M. E. Horsman, T. P. Hari and C. N. Boddy, Nat. Prod. Rep., 2016, 33, 183–202.
32. R. F. Little and C. Hertweck, Nat. Prod. Rep., 2022, 39, 163–205.
33. Y. Fan, J. Shen, Z. Liu, K. Xia, W. Zhu and P. Fu, Nat. Prod. Rep., 2022, 39, 1305–1324.
34. D. Mandalapu, X. Ji, J. Chen, C. Guo, W. Q. Liu, W. Ding, J. Zhou and Q. Zhang, J. Org. Chem., 2018, 83, 7271–7275.
35. D. F. Kreitler, E. M. Gemmell, J. E. Schaffer, T. A. Wencewicz and A. M. Gulick, Nat. Commun., 2019, 10, 3432.
36. L. Gu, B. Wang, A. Kulkarni, J. J. Gehret, K. R. Lloyd, L. Gerwick, W. H. Gerwick, P. Wipf, K. Hakansson, J. L. Smith and D. H. Sherman, J. Am. Chem. Soc., 2009, 131, 16033–16035.
37. B. Zhang, K. B. Wang, W. Wang, X. Wang, F. Liu, J. Zhu, J. Shi, L. Y. Li, H. Han, K. Xu, H. Y. Qiao, X. Zhang, R. H. Jiao, K. N. Houk, Y. Liang, R. X. Tan and H. M. Ge, Nature, 2019, 568, 122–126.
38. M. Kobayashi, K. Fujita, K. Matsuda and T. Wakimoto, J. Am. Chem. Soc., 2023, 145, 3270–3275.
39. T. Kuranaga, K. Matsuda, A. Sano, M. Kobayashi, A. Ninomiya, K. Takada, S. Matsunaga and T. Wakimoto, Angew. Chem., Int. Ed., 2018, 57, 9447–9451.
40. Z. Deng, C. Liu, F. Wang, N. Song, J. Liu, H. Li, S. Liu, T. Li, Z. Liu, F. Xiao and W. Li, Angew. Chem., Int. Ed., 2024, e202402010,  DOI:10.1002/anie.202402010.
41. M. A. Fischbach and C. T. Walsh, Chem. Rev., 2006, 106, 3468–3496.
42. K. A. Bozhuyuk, J. Micklefield and B. Wilkinson, Curr. Opin. Microbiol., 2019, 51, 88–96.
43. A. Koglin and C. T. Walsh, Nat. Prod. Rep., 2009, 26, 987–1000.
44. S. L. Wenski, S. Thiengmag and E. J. N. Helfrich, Synth. Syst. Biotechnol., 2022, 7, 631–647.
45. T. W. Giessen and M. A. Marahiel, FEBS Lett., 2012, 586, 2065–2075.
46. C. T. Walsh, Acc. Chem. Res., 2008, 41, 4–10.
47. C. Hertweck, Angew. Chem., Int. Ed., 2009, 48, 4688–4716.
48. J. Staunton and K. J. Weissman, Nat. Prod. Rep., 2001, 18, 380–416.
49. K. J. Weissman, Nat. Chem. Biol., 2015, 11, 660–670.
50. S. A. Sieber and M. A. Marahiel, J. Bacteriol., 2003, 185, 7036–7043.
51. M. Peschke, C. Brieke, M. Heimes and M. J. Cryle, ACS Chem. Biol., 2018, 13, 110–120.
52. S. Wang, W. D. G. Brittain, Q. Zhang, Z. Lu, M. H. Tong, K. Wu, K. Kyeremeh, M. Jenner, Y. Yu, S. L. Cobb and H. Deng, Nat. Commun., 2022, 13, 62.
53. T. P. Hari, P. Labana, M. Boileau and C. N. Boddy, ChemBioChem, 2014, 15, 2656–2661.
54. K. Tamura, G. Stecher and S. Kumar, Mol. Biol. Evol., 2021, 38, 3022–3027.
55. I. Letunic and P. Bork, Nucleic Acids Res., 2024, 52, W78–W82.
56. B. T. Caswell, C. C. de Carvalho, H. Nguyen, M. Roy, T. Nguyen and D. C. Cantu, Protein Sci., 2022, 31, 652–676.
57. T. P. Hari, P. Labana, M. Boileau and C. N. Boddy, ChemBioChem, 2014, 15, 2656–2661.
58. Z. Chang, P. Flatt, W. H. Gerwick, V. A. Nguyen, C. L. Willis and D. H. Sherman, Gene, 2002, 296, 235–247.
59. P. M. Flatt, S. J. O'Connell, K. L. McPhail, G. Zeller, C. L. Willis, D. H. Sherman and W. H. Gerwick, J. Nat. Prod., 2006, 69, 938–944.
60. M. S. Kim, M. Bae, Y. E. Jung, J. M. Kim, S. Hwang, M. C. Song, Y. H. Ban, E. S. Bae, S. Hong, S. K. Lee, S. S. Cha, D. C. Oh and Y. J. Yoon, Angew. Chem., Int. Ed., 2021, 60, 19766–19773.
61. K. Geyer, S. Hartmann, R. R. Singh and T. J. Erb, Biochemistry, 2022, 61, 2662–2671.
62. R. D. Sussmuth and A. Mainz, Angew. Chem., Int. Ed., 2017, 56, 3770–3821.
63. A. M. van Wageningen, P. N. Kirkpatrick, D. H. Williams, B. R. Harris, J. K. Kershaw, N. J. Lennard, M. Jones, S. J. Jones and P. J. Solenberg, Chem. Biol., 1998, 5, 155–162.
64. L. Xu, Y. Li, J. B. Biggins, B. R. Bowman, G. L. Verdine, J. B. Gloer, J. A. Alspaugh and G. F. Bills, Appl. Microbiol. Biotechnol., 2018, 102, 2337–2350.
65. B. Shen, L. Du, C. Sanchez, D. J. Edwards, M. Chen and J. M. Murrell, J. Nat. Prod., 2002, 65, 422–431.
66. M. A. Marahiel, Nat. Prod. Rep., 2016, 33, 136–140.
67. T. Izore and M. J. Cryle, Nat. Prod. Rep., 2018, 35, 1120–1139.
68. S. Konno, L. Misson and M. D. Burkart, in Cyclic Peptides: From Bioorganic Synthesis to Applications, ed. J. Koehnke, J. Naismith and W. A. van der Donk, The Royal Society of Chemistry, 2017 10.1039/9781788010153-00033.
69. D. P. Frueh, H. Arthanari, A. Koglin, D. A. Vosburg, A. E. Bennett, C. T. Walsh and G. Wagner, Nature, 2008, 454, 903–906.
70. N. Huguenin-Dezot, D. A. Alonzo, G. W. Heberlig, M. Mahesh, D. P. Nguyen, M. H. Dornan, C. N. Boddy, T. M. Schmeing and J. W. Chin, Nature, 2019, 565, 112–117.
71. S. D. Bruner, T. Weber, R. M. Kohli, D. Schwarzer, M. A. Marahiel, C. T. Walsh and M. T. Stubbs, Structure, 2002, 10, 301–310.
72. S. A. Samel, B. Wagner, M. A. Marahiel and L. O. Essen, J. Mol. Biol., 2006, 359, 876–889.
73. J. H. Yu, J. Song, C. B. Chi, T. Liu, T. T. Geng, Z. H. Cai, W. D. Dong, C. Shi, X. Y. Ma, Z. Y. Zhang, X. J. Ma, B. Y. Xing, H. W. Jin, L. R. Zhang, S. W. Dong, D. H. Yang and M. Ma, ACS Catal., 2021, 11, 11733–11741.
74. T. L. Bauer, P. C. F. Buchholz and J. Pleiss, FEBS J., 2020, 287, 1035–1053.
75. A. Rauwerdink and R. J. Kazlauskas, ACS Catal., 2015, 5, 6153–6176.
76. D. C. Cantu, Y. Chen and P. J. Reilly, Protein Sci., 2010, 19, 1281–1295.
77. A. Koglin, F. Lohr, F. Bernhard, V. V. Rogov, D. P. Frueh, E. R. Strieter, M. R. Mofid, P. Guntert, G. Wagner, C. T. Walsh, M. A. Marahiel and V. Dotsch, Nature, 2008, 454, 907–911.
78. K. D. Patel, M. R. MacDonald, S. F. Ahmed, J. Singh and A. M. Gulick, Nat. Prod. Rep., 2023, 40, 1550–1582.
79. J. B. Scaglione, D. L. Akey, R. Sullivan, J. D. Kittendorf, C. M. Rath, E. S. Kim, J. L. Smith and D. H. Sherman, Angew. Chem., Int. Ed., 2010, 49, 5726–5730.
80. G. Calero, P. Gupta, M. C. Nonato, S. Tandel, E. R. Biehl, S. L. Hofmann and J. Clardy, J. Biol. Chem., 2003, 278, 37957–37964.
81. B. R. Miller and A. M. Gulick, Methods Mol. Biol., 2016, 1401, 3–29.
82. H. Nakamura, J. X. Wang and E. P. Balskus, Chem. Sci., 2015, 6, 3816–3822.
83. L. Serino, C. Reimmann, P. Visca, M. Beyeler, V. D. Chiesa and D. Haas, J. Bacteriol., 1997, 179, 248–257.
84. S. T. Pullan, G. Chandra, M. J. Bibb and M. Merrick, BMC Genomics, 2011, 12, 175.
85. T. P. Korman, J. M. Crawford, J. W. Labonte, A. G. Newman, J. Wong, C. A. Townsend and S. C. Tsai, Proc. Natl. Acad. Sci. U. S. A., 2010, 107, 6246–6251.
86. N. M. Gaudelli and C. A. Townsend, Nat. Chem. Biol., 2014, 10, 251–258.
87. N. M. Gaudelli, D. H. Long and C. A. Townsend, Nature, 2015, 520, 383–387.
88. K. D. Patel, F. B. d'Andrea, N. M. Gaudelli, A. R. Buller, C. A. Townsend and A. M. Gulick, Nat. Commun., 2019, 10, 3868.
89. C. Khosla, Y. Tang, A. Y. Chen, N. A. Schnarr and D. E. Cane, Annu. Rev. Biochem., 2007, 76, 195–221.
90. C. D. Campbell and J. C. Vederas, Biopolymers, 2010, 93, 755–763.
91. H. Wang, C. Li, B. Zhang, H. He, P. Jin, J. Wang, J. Zhang, X. Wang and W. Xiang, J. Biotechnol., 2015, 204, 1–2.
92. B. Julien, S. Shah, R. Ziermann, R. Goldman, L. Katz and C. Khosla, Gene, 2000, 249, 153–160.
93. B. Shen, Curr. Opin. Chem. Biol., 2003, 7, 285–295.
94. K. J. Weissman, Methods Enzymol., 2009, 459, 3–16.
95. P. Argyropoulos, F. Bergeret, C. Pardin, J. M. Reimer, A. Pinto, C. N. Boddy and T. M. Schmeing, Biochim. Biophys. Acta, 2016, 1860, 486–497.
96. D. L. Akey, J. D. Kittendorf, J. W. Giraldes, R. A. Fecik, D. H. Sherman and J. L. Smith, Nat. Chem. Biol., 2006, 2, 537–542.
97. J. J. Gehret, L. Gu, W. H. Gerwick, P. Wipf, D. H. Sherman and J. L. Smith, J. Biol. Chem., 2011, 286, 14445–14454.
98. S. C. Tsai, L. J. Miercke, J. Krucinski, R. Gokhale, J. C. Chen, P. G. Foster, D. E. Cane, C. Khosla and R. M. Stroud, Proc. Natl. Acad. Sci. U. S. A., 2001, 98, 14808–14813.
99. S. C. Tsai, H. X. Lu, D. E. Cane, C. Khosla and R. M. Stroud, Biochemistry, 2002, 41, 12598–12606.
100. T. Shi, L. Liu, W. Tao, S. Luo, S. Fan, X.-L. Wang, L. Bai and Y.-L. Zhao, ACS Catal., 2018, 8, 4323–4332.
101. X. P. Chen, T. Shi, X. L. Wang, J. T. Wang, Q. H. Chen, L. Q. Bai and Y. L. Zhao, ACS Catal., 2016, 6, 4369–4378.
102. D. J. Craik, D. P. Fairlie, S. Liras and D. Price, Chem. Biol. Drug Des., 2013, 81, 136–147.
103. D. Yu, F. Xu, J. Zi, S. Wang, D. Gage, J. Zeng and J. Zhan, Metab. Eng., 2013, 18, 60–68.
104. G. Li Petri, S. Di Martino and M. De Rosa, J. Med. Chem., 2022, 65, 7438–7475.
105. L. Y. Chia, P. V. Kumar, M. A. A. Maki, G. Ravichandran and S. Thilagar, Int. J. Pept. Res. Ther., 2023, 29, 7.
106. K. R. Schramma, L. B. Bushin and M. R. Seyedsayamdost, Nat. Chem., 2015, 7, 431–437.
107. A. Greule, T. Izore, D. Iftime, J. Tailhades, M. Schoppet, Y. Zhao, M. Peschke, I. Ahmed, A. Kulik, M. Adamek, R. J. A. Goode, R. B. Schittenhelm, J. A. Kaczmarski, C. J. Jackson, N. Ziemert, E. H. Krenske, J. J. De Voss, E. Stegmann and M. J. Cryle, Nat. Commun., 2019, 10, 2613.
108. L. C. Cai, Y. M. Yao, S. K. Yeon and I. B. Seiple, J. Am. Chem. Soc., 2020, 142, 15116–15126.
109. G. Gause and M. Brazhnikova, Nature, 1944, 154, 703.
110. W. W. Umbreit, The Search for New Antibiotics. G. F. Gause. Yale University Press, New Haven, Conn, 1960. 97 pp. Illus. $4.75, 1960.
111. L. H. Kondejewski, S. W. Farmer, D. S. Wishart, R. E. W. Hancock and R. S. Hodges, Int. J. Pept. Protein Res., 1996, 47, 460–466.
112. M. Wenzel, M. Rautenbach, J. A. Vosloo, T. Siersma, C. H. M. Aisenbrey, E. Zaitseva, W. E. Laubscher, W. van Rensburg, J. C. Behrends, B. Bechinger and L. W. Hamoen, mBio, 2018, 9, e00802–e00818.
113. K. P. Dimick, Proc. Soc. Exp. Biol. Med., 1951, 78, 782–784.
114. R. Sarges and B. Witkop, J. Am. Chem. Soc., 1965, 87, 2011–2020.
115. K. O. Pedersen and R. L. M. Synge, Acta Chem. Scand., 1948, 2, 408–413.
116. B. F. Erlnager and L. Goode, Nature, 1954, 174, 840–841.
117. H. D. Mootz and M. A. Marahiel, J. Bacteriol., 1997, 179, 6843–6850.
118. R. J. Dubos and R. D. Hotchkiss, J. Exp. Med., 1941, 73, 629–640.
119. P. J. Loll, E. C. Upton, V. Nahoum, N. J. Economou and S. Cocklin, Biochim. Biophys. Acta, 2014, 1838, 1199–1207.
120. M. Rautenbach, A. M. Troskie, J. A. Vosloo and M. E. Dathe, Biochimie, 2016, 130, 122–131.
121. M. Krause and M. A. Marahiel, J. Bacteriol., 1988, 170, 4669–4674.
122. J. Kratzschmar, M. Krause and M. A. Marahiel, J. Bacteriol., 1989, 171, 5422–5429.
123. F. Pourmasoumi, S. Y. T. De, H. Y. Peng, F. Trottmann, C. Hertweck and H. Kries, ACS Chem. Biol., 2022, 17, 2382–2388.
124. K. M. Hoyer, C. Mahlert and M. A. Marahiel, Chem. Biol., 2007, 14, 13–22.
125. X. M. Wu, X. Z. Bu, K. M. Wong, W. L. Yan and Z. H. Guo, Org. Lett., 2003, 5, 1749–1752.
126. J. W. Trauger, R. M. Kohli, H. D. Mootz, M. A. Marahiel and C. T. Walsh, Nature, 2000, 407, 215–218.
127. R. M. Kohli, J. Takagi and C. T. Walsh, Proc. Natl. Acad. Sci. U. S. A., 2002, 99, 1247–1252.
128. H. Lin, D. A. Thayer, C. H. Wong and C. T. Walsh, Chem. Biol., 2004, 11, 1635–1642.
129. R. M. Kohli, J. W. Trauger, D. Schwarzer, M. A. Marahiel and C. T. Walsh, Biochemistry, 2001, 40, 7099–7108.
130. J. W. Trauger, R. M. Kohli and C. T. Walsh, Biochemistry, 2001, 40, 7092–7098.
131. G. Xie, M. Uttamchandani, G. Y. Chen, X. Bu, S. S. Lin, K. M. Wong, W. Yan, S. Q. Yao and Z. Guo, Bioorg. Med. Chem. Lett., 2002, 12, 989–992.
132. P. Wadhwani, S. Afonin, M. Ieronimo, J. Buerck and A. S. Ulrich, J. Org. Chem., 2006, 71, 55–61.
133. F. E. Morales, H. E. Garay, D. F. Munoz, Y. E. Augusto, A. J. Otero-Gonzalez, O. Reyes Acosta and D. G. Rivera, Org. Lett., 2015, 17, 2728–2731.
134. L. Reguera and D. G. Rivera, Chem. Rev., 2019, 119, 9836–9860.
135. G. M. Grotenbreg, M. S. Timmer, A. L. Llamas-Saiz, M. Verdoes, G. A. van der Marel, M. J. van Raaij, H. S. Overkleeft and M. Overhand, J. Am. Chem. Soc., 2004, 126, 3444–3446.
136. L. Du and L. Lou, Nat. Prod. Rep., 2010, 27, 255–278.
137. R. M. Kohli, C. T. Walsh and M. D. Burkart, Nature, 2002, 418, 658–661.
138. O. E. Zolova, A. S. Mady and S. Garneau-Tsodikova, Biopolymers, 2010, 93, 777–790.
139. S. Zhang, Y. Chen, J. Zhu, Q. Lu, M. J. Cryle, Y. Zhang and F. Yan, Nat. Prod. Rep., 2023, 40, 557–594.
140. M. Bae, H. Kim, K. Moon, S. J. Nam, J. Shin, K. B. Oh and D. C. Oh, Org. Lett., 2015, 17, 712–715.
141. C. W. Johnston, M. A. Skinnider, M. A. Wyatt, X. Li, M. R. Ranieri, L. Yang, D. L. Zechel, B. Ma and N. A. Magarvey, Nat. Commun., 2015, 6, 8421.
142. S. Zhang, J. Zhu, D. L. Zechel, C. Jessen-Trefzer, R. T. Eastman, T. Paululat and A. Bechthold, ChemBioChem, 2018, 19, 272–279.
143. M. S. Kim, M. Bae, M. C. Song, S. Hwang, D. C. Oh and Y. J. Yoon, Org. Lett., 2022, 24, 4444–4448.
144. Z. Yu, S. Vodanovic-Jankovic, M. Kron and B. Shen, Org. Lett., 2012, 14, 4946–4949.
145. S. Zhang, L. Zhang, A. Greule, J. Tailhades, E. Marschall, P. Prasongpholchai, D. J. Leng, J. Zhang, J. Zhu, J. A. Kaczmarski, R. B. Schittenhelm, O. Einsle, C. J. Jackson, F. Alberti, A. Bechthold, Y. Zhang, M. Tosin, T. Si and M. J. Cryle, Acta Pharm. Sin. B, 2023, 13, 3561–3574.
146. M. L. Guerinot, Annu. Rev. Microbiol., 1994, 48, 743–772.
147. M. Miethke and M. A. Marahiel, Microbiol. Mol. Biol. Rev., 2007, 71, 413–451.
148. L. D. Loomis and Kenneth N. Raymond, Inorg. Chem., 1991, 30, 906–911.
149. L. M. Bogomolnaya, R. Tilvawala, J. R. Elfenbein, J. D. Cirillo and H. L. Andrews-Polymenis, mBio, 2020, 11, e00528–e00520.
150. J. R. Pollack and J. B. Neilands, Biochem. Biophys. Res. Commun., 1970, 38, 989–992.
151. I. G. O'Brien and F. Gibson, Biochim. Biophys. Acta, 1970, 215, 393–402.
152. I. G. O'Brien, G. B. Cox and F. Gibson, Biochim. Biophys. Acta, 1971, 237, 537–549.
153. Y. Matsuo, K. Kanoh, J. H. Jang, K. Adachi, S. Matsuda, O. Miki, T. Kato and Y. Shizuri, J. Nat. Prod., 2011, 74, 2371–2376.
154. S. Zajdowicz, J. C. Haller, A. E. Krafft, S. W. Hunsucker, C. T. Mant, M. W. Duncan, R. S. Hodges, D. N. Jones and R. K. Holmes, PLoS One, 2012, 7, e34591.
155. H. Budzikiewicz, A. Bossenkamp, K. Taraz, A. Pandey and J. M. Meyer, Z. Naturforsch., C: J. Biosci., 1997, 52, 551–554.
156. I. G. Young, L. Langman, R. K. Luke and F. Gibson, J. Bacteriol., 1971, 106, 51–57.
157. R. K. Luke and F. Gibson, J. Bacteriol., 1971, 107, 557–562.
158. A. M. Gehring, I. Mori and C. T. Walsh, Biochemistry, 1998, 37, 2648–2659.
159. D. E. Ehmann, J. W. Trauger, T. Stachelhaus and C. T. Walsh, Chem. Biol., 2000, 7, 765–772.
160. E. D. Roche and C. T. Walsh, Biochemistry, 2003, 42, 1334–1344.
161. A. M. Gehring, K. A. Bradley and C. T. Walsh, Biochemistry, 1997, 36, 8495–8503.
162. E. J. Drake, B. R. Miller, C. Shi, J. T. Tarrasch, J. A. Sundlov, C. L. Allen, G. Skiniotis, C. C. Aldrich and A. M. Gulick, Nature, 2016, 529, 235–238.
163. Z. L. Reitz, M. Sandy and A. Butler, Metallomics, 2017, 9, 824–839.
164. C. A. Shaw-Reid, N. L. Kelleher, H. C. Losey, A. M. Gehring, C. Berg and C. T. Walsh, Chem. Biol., 1999, 6, 385–400.
165. Y. Liu, T. Zheng and S. D. Bruner, Chem. Biol., 2011, 18, 1482–1488.
166. K. Haslinger, M. Peschke, C. Brieke, E. Maximowitsch and M. J. Cryle, Nature, 2015, 521, 105–109.
167. M. Peschke, M. Gonsior, R. D. Sussmuth and M. J. Cryle, Curr. Opin. Struct. Biol., 2016, 41, 46–53.
168. A. M. Gulick and C. C. Aldrich, Nat. Prod. Rep., 2018, 35, 1156–1184.
169. B. R. Miller, E. J. Drake, C. Shi, C. C. Aldrich and A. M. Gulick, J. Biol. Chem., 2016, 291, 22559–22571.
170. E. J. Corey and S. Bhattacharyya, Tetrahedron Lett., 1977, 3919–3922.
171. A. Shanzer and J. Libman, J. Chem. Soc., Chem. Commun., 1983, 846–847,  10.1039/c39830000846.
172. R. J. A. Ramirez, L. Karamanukyan, S. Ortiz and C. G. Gutierrez, Tetrahedron Lett., 1997, 38, 749–752.
173. H. K. Zane, H. Naka, F. Rosconi, M. Sandy, M. G. Haygood and A. Butler, J. Am. Chem. Soc., 2014, 136, 5615–5618.
174. D. L. McRose, O. Baars, M. R. Seyedsayamdost and F. M. M. Morel, Proc. Natl. Acad. Sci. U. S. A., 2018, 115, 7581–7586.
175. A. M. Jelowicki and A. Butler, J. Biol. Inorg. Chem., 2022, 27, 565–572.
176. J. J. May, T. M. Wendrich and M. A. Marahiel, J. Biol. Chem., 2001, 276, 7209–7217.
177. Y. P. Wen, X. C. Wu, Y. Teng, C. D. Qian, Z. J. Zhan, Y. H. Zhao and O. Li, Environ. Microbiol., 2011, 13, 2726–2737.
178. Q. H. Wu, K. Throckmorton, M. Maity, M. G. Chevrette, D. R. Braun, S. R. Rajski, C. R. Currie, M. G. Thomas and T. S. Bugni, J. Nat. Prod., 2021, 84, 136–141.
179. H. N. Patel, R. N. Chakraborty and S. B. Desai, FEMS Microbiol. Lett., 1988, 56, 131–134.
180. J. R. Heemstra, C. T. Walsh and E. S. Sattely, J. Am. Chem. Soc., 2009, 131, 15317–15329.
181. R. A. Carter, P. S. Worsley, G. Sawers, G. L. Challis, M. J. Dilworth, K. C. Carson, J. A. Lawrence, M. Wexler, A. W. Johnston and K. H. Yeoman, Mol. Microbiol., 2002, 44, 1153–1166.
182. N. Kadi, D. Oves-Costales, F. Barona-Gomez and G. L. Challis, Nat. Chem. Biol., 2007, 3, 652–656.
183. J. Yang, V. S. Banas, K. D. Patel, G. S. M. Rivera, L. S. Mydy, A. M. Gulick and T. A. Wencewicz, J. Biol. Chem., 2022, 298, 102166.
184. N. Kadi, L. Song and G. L. Challis, Chem. Commun., 2008, 5119–5121,  10.1039/b813029a.
185. N. Kadi, S. Arbache, L. J. Song, D. Oves-Costales and G. L. Challis, J. Am. Chem. Soc., 2008, 130, 10458–10459.
186. H. Brockmann and G. Schmidt-Kastner, Chem. Ber., 1955, 88, 57–61.
187. H. Brockmann, M. Springor, G. Traxler and I. Hofer, Naturwissenschaften, 1963, 50, 689.
188. M. M. Shemyakin, N. A. Aldanova, E. I. Vinogradova and M. Y. Feigina, Tetrahedron Lett., 1963, 1921–1925.
189. S. H. Huang, Y. S. Liu, W. Q. Liu, P. Neubauer and J. Li, Microorganisms, 2021, 9, 780.
190. Z. Su, X. Ran, J. J. Leitch, A. L. Schwan, R. Faragher and J. Lipkowski, Langmuir, 2019, 35, 16935–16943.
191. Y. Q. Cheng, ChemBioChem, 2006, 7, 471–477.
192. D. A. Alonzo and T. M. Schmeing, Protein Sci., 2020, 29, 2316–2347.
193. J. Jaitzig, J. Li, R. D. Sussmuth and P. Neubauer, ACS Synth. Biol., 2014, 3, 432–438.
194. S. Konno, M. Tanaka, T. Mizuguchi, H. Toyokai, A. Taguchi, A. Taniguchi and Y. Hayashi, J. Pept. Sci., 2023, 30, e3532.
195. B. F. Gisin, R. B. Merrifield and D. C. Tosteson, J. Am. Chem. Soc., 1969, 91, 2691–2695.
196. Y. L. Dory, J. M. Mellor and J. F. Mcaleer, Tetrahedron Lett., 1989, 30, 1695–1698.
197. J. R. Dawson, Y. L. Dory, J. M. Mellor and J. F. McAleer, Tetrahedron, 1996, 52, 1361–1378.
198. L. Zhuang, S. H. Huang, W. Q. Liu, A. S. Karim, M. C. Jewett and J. Li, Metab. Eng., 2020, 60, 37–44.
199. N. Agata, M. Mori, M. Ohta, S. Suwan, I. Ohtani and M. Isobe, FEMS Microbiol. Lett., 1994, 121, 31–34.
200. V. Walser, M. Kranzler, C. Dawid, M. Ehling-Schulz, T. D. Stark and T. F. Hofmann, Molecules, 2022, 27, 872.
201. M. A. Andersson, P. Hakulinen, U. Honkalampi-Hamalainen, D. Hoornstra, J. C. Lhuguenot, J. Maki-Paakkanen, M. Savolainen, I. Severin, A. L. Stammati, L. Turco, A. Weber, A. von Wright, F. Zucco and M. Salkinoja-Salonen, Toxicon, 2007, 49, 351–367.
202. N. Agata, M. Ohta and M. Mori, Curr. Microbiol., 1996, 33, 67–69.
203. F. Carlin, M. Fricker, A. Pielaat, S. Heisterkamp, R. Shaheen, M. S. Salonen, B. Svensson, C. Nguyen-The and M. Ehling-Schulz, Int. J. Food Microbiol., 2006, 109, 132–138.
204. S. Marxen, T. D. Stark, A. Rutschle, G. Lucking, E. Frenzel, S. Scherer, M. Ehling-Schulz and T. Hofmann, Sci. Rep., 2015, 5, 10637.
205. M. Ehling-Schulz, N. Vukov, A. Schulz, R. Shaheen, M. Andersson, E. Martlbauer and S. Scherer, Appl. Environ. Microbiol., 2005, 71, 105–113.
206. M. Ehling-Schulz, M. Fricker, H. Grallert, P. Rieck, M. Wagner and S. Scherer, BMC Microbiol., 2006, 6, 20.
207. G. Lucking, E. Frenzel, A. Rutschle, S. Marxen, T. D. Stark, T. Hofmann, S. Scherer and M. Ehling-Schulz, Front. Microbiol., 2015, 6, 1101.
208. D. A. Alonzo, N. A. Magarvey and T. M. Schmeing, PLoS One, 2015, 10, e0128569.
209. G. W. Heberlig and C. N. Boddy, J. Nat. Prod., 2020, 83, 1990–1997.
210. G. J. Quigley, G. Ughetto, G. A. van der Marel, J. H. van Boom, A. H. Wang and A. Rich, Science, 1986, 232, 1255–1258.
211. U. Keller and F. Schauwecker, Prog. Nucleic Acid Res. Mol. Biol., 2001, 70, 233–289.
212. H. Chen and D. J. Patel, J. Mol. Biol., 1995, 246, 164–179.
213. J. Y. Park, Y. S. Ryang, K. Y. Shim, J. I. Lee, H. S. Kim, Y. H. Kim and S. K. Kim, Int. J. Biochem. Cell Biol., 2006, 38, 244–254.
214. J. Fernandez, L. Marin, R. Alvarez-Alonso, S. Redondo, J. Carvajal, G. Villamizar, C. J. Villar and F. Lombo, Mar. Drugs, 2014, 12, 2668–2699.
215. R. Corbaz, L. Ettlinger, E. Gaumann, W. Kellerschierlein, F. Kradolfer, L. Neipp, V. Prelog, P. Reusser and H. Zahner, Helv. Chim. Acta, 1957, 40, 199–204.
216. T. Yoshida, K. Katagiri and S. Yokozawa, J. Antibiot., 1961, 14, 330–334.
217. C. Zhang, L. X. Kong, Q. Liu, X. Lei, T. Zhu, J. Yin, B. R. Lin, Z. X. Deng and D. L. You, PLoS One, 2013, 8, e56772.
218. J. Shoji, R. Konaka, K. Kawano, N. Higuchi and Y. Kyogoku, J. Antibiot., 1976, 29, 1246–1248.
219. T. Yoshida and K. Katagiri, J. Bacteriol., 1967, 93, 1327–1331.
220. L. G. May, M. A. Madine and M. J. Waring, Nucleic Acids Res., 2004, 32, 65–72.
221. G. Schmoock, F. Pfennig, J. Jewiarz, W. Schlumbohm, W. Laubinger, F. Schauwecker and U. Keller, J. Biol. Chem., 2005, 280, 4339–4349.
222. A. P. Praseuth, C. C. C. Wang, K. Watanabe, K. Hotta, H. Oguri and H. Oikawa, Biotechnol. Prog., 2008, 24, 1226–1231.
223. K. Watanabe, K. Hotta, M. Nakaya, A. P. Praseuth, C. C. C. Wang, D. Inada, K. Takahashi, E. Fukushi, H. Oguri and H. Oikawa, J. Am. Chem. Soc., 2009, 131, 9347–9353.
224. M. Sato, T. Nakazawa, Y. Tsunematsu, K. Hotta and K. Watanabe, Curr. Opin. Chem. Biol., 2013, 17, 537–545.
225. K. Watanabe, K. Hotta, A. P. Praseuth, M. Searcey, C. C. C. Wang, H. Oguri and H. Oikawa, ChemBioChem, 2009, 10, 1965–1968.
226. J. Grunewald, S. A. Sieber and M. A. Marahiel, Biochemistry, 2004, 43, 2915–2925.
227. C. C. Tseng, S. D. Bruner, R. M. Kohli, M. A. Marahiel, C. T. Walsh and S. A. Sieber, Biochemistry, 2002, 41, 13350–13359.
228. K. Koketsu, H. Oguri, K. Watanabe and H. Oikawa, Chem. Biol., 2008, 15, 818–828.
229. J. Tulla-Puche, E. Marcucci, M. Fermin, N. Bayo-Puxan and F. Albericio, Chem. – Eur. J., 2008, 14, 4475–4478.
230. G. A. Sable and D. Lim, J. Org. Chem., 2015, 80, 7486–7494.
231. K. Hattori, K. Koike, K. Okuda, T. Hirayama, M. Ebihara, M. Takenaka and H. Nagasawa, Org. Biomol. Chem., 2016, 14, 2090–2111.
232. P. K. Chakravarty and R. K. Olsen, Tetrahedron Lett., 1978, 19, 1613–1616.
233. K. Kojima, F. Yakushiji, A. Katsuyama and S. Ichikawa, Org. Lett., 2020, 22, 4217–4221.
234. C. Maruyama, Y. Chinone, S. Sato, F. Kudo, K. Ohsawa, J. Kubota, J. Hashimoto, I. Kozone, T. Doi, K. Shin-Ya, T. Eguchi and Y. Hamano, Biomolecules, 2020, 10, 210.
235. J. A. Matson, K. L. Colson, G. N. Belofsky and B. B. Bleiberg, J. Antibiot., 1993, 46, 162–166.
236. D. L. Boger, J. H. Chen and K. W. Saionz, J. Am. Chem. Soc., 1996, 118, 1629–1644.
237. K. Katayama, K. Nakagawa, H. Takeda, A. Matsuda and S. Ichikawa, Org. Lett., 2014, 16, 428–431.
238. Y. Komatani, K. Momosaki, A. Katsuyama, K. Yamamoto, R. Kaguchi and S. Ichikawa, Org. Lett., 2023, 25, 543–548.
239. M. Konishi, H. Ohkuma, F. Sakai, T. Tsuno, H. Koshiyama, T. Naito and H. Kawaguchi, J. Antibiot., 1981, 34, 148–159.
240. D. L. Boger, M. W. Ledeboer, M. Kume, M. Searcey and Q. Jin, J. Am. Chem. Soc., 1999, 121, 11375–11383.
241. X. J. Shi, L. M. Huang, K. H. Song, G. Y. Zhao, Y. Liu, L. X. Lv and Y. L. Du, Angew. Chem., Int. Ed., 2021, 60, 19821–19828.
242. L. J. Cruz, M. M. Insua, J. P. Baz, M. Trujillo, R. A. Rodriguez-Mias, E. Oliveira, E. Giralt, F. Albericio and L. M. Canedo, J. Org. Chem., 2006, 71, 3335–3338.
243. L. J. Cruz, C. Cuevas, L. M. Canedo, E. Giralt and F. Albericio, J. Org. Chem., 2006, 71, 3339–3344.
244. A. V. Mayorov, M. Y. Cai, E. S. Palmer, Z. H. Liu, J. P. Cain, J. Vagner, D. Trivedi and V. J. Hruby, Peptides, 2010, 31, 1894–1905.
245. J. P. Baz, L. M. Canedo, J. L. F. Puentes and M. V. S. Elipe, J. Antibiot., 1997, 50, 738–741.
246. F. Romero, F. Espliego, J. P. Baz, T. G. DeQuesada, D. Gravalos, F. DelaCalle and J. L. FernadezPuertes, J. Antibiot., 1997, 50, 734–737.
247. F. Lombo, A. Velasco, A. Castro, F. de la Calle, A. F. Brana, J. M. Sanchez-Puelles, C. Mendez and J. A. Salas, ChemBioChem, 2006, 7, 366–376.
248. S. Mori, S. K. Shrestha, J. Fernandez, M. A. San Milian, A. Garzan, A. H. Al-Mestarihi, F. Lombo and S. Garneau-Tsodikova, Biochemistry, 2017, 56, 4457–4467.
249. L. Robbel, K. M. Hoyer and M. A. Marahiel, FEBS J., 2009, 276, 1641–1653.
250. D. L. Boger, S. Ichikawa, W. C. Tse, M. P. Hedrick and Q. Jin, J. Am. Chem. Soc., 2001, 123, 561–568.
251. N. Bayo-Puxan, A. Fernandez, J. Tulla-Puche, E. Riego, C. Cuevas, M. Alvarez and F. Albericio, Chem. – Eur. J., 2006, 12, 9001–9009.
252. J. He, E. M. K. Wijeratne, B. P. Bashyal, J. X. Zhan, C. J. Seliga, M. P. X. Liu, E. E. Pierson, L. S. Pierson, H. D. VanEtten and A. A. L. Gunatilaka, J. Nat. Prod., 2004, 67, 1985–1991.
253. C. Sanchez, I. A. Butovich, A. F. Brana, J. Rohr, C. Mendez and J. A. Salas, Chem. Biol., 2002, 9, 519–531.
254. H. Onaka, S. Taniguchi, Y. Igarashi and T. Furumai, Biosci., Biotechnol., Biochem., 2003, 67, 127–138.
255. S. Shimizu, Y. Yamamoto, J. Inagaki and S. Koshimura, Gan, 1982, 73, 642–648.
256. A. Kaji, R. Saito, M. Nomura, K. Miyamoto and N. Kiriyama, Anticancer Res., 1997, 17, 3675–3679.
257. P. Schneider, M. Weber, K. Rosenberger and D. Hoffmeister, Chem. Biol., 2007, 14, 635–644.
258. C. J. Balibar, A. R. Howard and C. T. Walsh, Nat. Chem. Biol., 2007, 3, 584–592.
259. S. Thies, B. Santiago-Schubel, F. Kovacic, F. Rosenau, R. Hausmann and K. E. Jaeger, J. Biotechnol., 2014, 181, 27–30.
260. H. Li, T. Tanikawa, Y. Sato, Y. Nakagawa and T. Matsuyama, Microbiol. Immunol., 2005, 49, 303–310.
261. C. T. Walsh, Science, 2004, 303, 1805–1810.
262. H. Liu, R. N. Ottosen, K. M. Jennet, E. B. Svenningsen, T. F. Kristensen, M. Biltoft, M. R. Jakobsen and T. B. Poulsen, Angew. Chem., Int. Ed., 2021, 60, 18734–18741.
263. P. Steib and B. Breit, Angew. Chem., Int. Ed., 2018, 57, 6572–6576.
264. Y. J. Zhou, A. C. Murphy, M. Samborskyy, P. Prediger, L. C. Dias and P. F. Leadlay, Chem. Biol., 2015, 22, 745–754.
265. J. Ma, Z. Wang, H. Huang, M. Luo, D. Zuo, B. Wang, A. Sun, Y. Q. Cheng, C. Zhang and J. Ju, Angew. Chem., Int. Ed., 2011, 50, 7797–7802.
266. M. Arai, J. Antibiot., 1960, 13, 46–50.
267. M. Arai, J. Antibiot., 1960, 13, 51–56.
268. M. Gui, M. X. Zhang, W. H. Wu and P. Sun, Molecules, 2019, 24, 3840.
269. M. G. Nair, A. Chandra, D. L. Thorogood, E. Ammermann, N. Walker and K. Kiehs, J. Agric. Food Chem., 1994, 42, 2308–2310.
270. A. Fang, G. K. Wong and A. L. Demain, J. Antibiot., 2000, 53, 158–162.
271. C. Wu, Y. Tan, M. Gan, Y. Wang, Y. Guan, X. Hu, H. Zhou, X. Shang, X. You, Z. Yang and C. Xiao, J. Nat. Prod., 2013, 76, 2153–2157.
272. J. Ji, K. Wang, X. Meng, H. Zhong, X. Li, H. Zhao, G. Xie, Y. Xie, X. Wang and X. Zhu, Cancers, 2022, 14, 5812.
273. K. Toshima, K. Tatsuta and M. Kinoshita, Tetrahedron Lett., 1986, 27, 4741–4744.
274. D. Seebach, H. F. Chow, R. Jackson, K. Lawson, M. Sutter, S. Thaisrivongs and J. Zimmermann, J. Am. Chem. Soc., 1985, 107, 5292–5293.
275. H. Nakamura, K. Arata, T. Wakamatsu, Y. Ban and M. Shibasaki, Chem. Pharm. Bull., 1990, 38, 2435–2441.
276. D. A. Evans and D. M. Fitch, J. Org. Chem., 1997, 62, 454–455.
277. S. F. Haydock, T. Mironenko, H. I. Ghoorahoo and P. F. Leadlay, J. Biotechnol., 2004, 113, 55–68.
278. Y. Zhou, P. Prediger, L. C. Dias, A. C. Murphy and P. F. Leadlay, Angew. Chem., 2015, 127, 5321–5324.
279. Z. Yang, Y. Qiao, N. C. Konakalla, E. Strobech, P. Harris, G. Peschel, M. Agler-Rosenbaum, T. Weber, E. Andreasson and L. Ding, Nat. Commun., 2023, 14, 7398.
280. H. Zhang, Y. Wang, J. Wu, K. Skalina and B. A. Pfeifer, Chem. Biol., 2010, 17, 1232–1240.
281. Q. Xue, G. Ashley, C. R. Hutchinson and D. V. Santi, Proc. Natl. Acad. Sci. U. S. A., 1999, 96, 11740–11745.
282. J. W. Westley, C. M. Liu, R. H. Evans and J. F. Blount, J. Antibiot., 1979, 32, 874–877.
283. H. J. Lacey, T. J. Booth, D. Vuong, P. J. Rutledge, E. Lacey, Y. H. Chooi and A. M. Piggott, J. Antibiot., 2020, 73, 756–765.
284. Z. Ju, W. Zhou, H. A. Alharbi, D. C. Howell and T. Mahmud, ACS Chem. Biol., 2021, 16, 2641–2650.
285. C. P. Bold and K.-H. Altmann, Tetrahedron, 2024, 156, 133908.
286. L. Lizzadro, O. Spiess, S. Reinecke, M. Stadler and D. Schinzer, Molecules, 2024, 29, 1123.
287. L. Lizzadro, O. Spiess, W. Collisi, M. Stadler and D. Schinzer, ChemBioChem, 2022, 23, e202200458.
288. W. Huang, M. Ye, L. R. Zhang, Q. D. Wu, M. Zhang, J. H. Xu and W. Zheng, Mol. Cancer, 2014, 13, 150.
289. W. Huang, Q. D. Wu, M. Zhang, Y. L. Kong, P. R. Cao, W. Zheng, J. H. Xu and M. Ye, Cancer Lett., 2015, 356, 862–871.
290. W. Zhou, P. Posri and T. Mahmud, ACS Chem. Biol., 2021, 16, 270–276.
291. W. Zhang, M. Yang, W. Li, L. Zhou, Y. Shen, S. P. Wang, J. M. Gao, H. W. Lin, J. Qi and Y. Zhou, J. Agric. Food Chem., 2023, 71, 7459–7467.
292. A. Dramae, S. Nithithanasilp, W. Choowong, P. Rachtawee, S. Prabpai, P. Kongsaeree and P. Pittayakhajonwut, Tetrahedron, 2013, 69, 8205–8208.
293. E. Yiannakas, M. I. Grimes, J. T. Whitelegge, A. Furstner and A. N. Hulme, Angew. Chem., Int. Ed., 2021, 60, 18504–18508.
294. R. Jansen, H. Irschik, H. Reichenbach, V. Wray and G. Höfle, Liebigs Ann. Chem., 1994, 1994, 759–773.
295. Y. A. Elnakady, F. Sasse, H. Lunsdorf and H. Reichenbach, Biochem. Pharmacol., 2004, 67, 927–935.
296. C. D. Hopkins and P. Wipf, Nat. Prod. Rep., 2009, 26, 585–601.
297. R. Carvalho, R. Reid, N. Viswanathan, H. Gramajo and B. Julien, Gene, 2005, 359, 91–98.
298. P. Wipf and T. H. Graham, J. Am. Chem. Soc., 2004, 126, 15346–15347.
299. K. C. Nicolaou, G. Bellavance, M. Buchman and K. K. Pulukuri, J. Am. Chem. Soc., 2017, 139, 15636–15639.
300. M. Tsunakawa, O. Tenmyo, K. Tomita, N. Naruse, C. Kotake, T. Miyaki, M. Konishi and T. Oki, J. Antibiot., 1992, 45, 180–188.
301. H. Y. He, H. X. Pan, L. F. Wu, B. B. Zhang, H. B. Chai, W. Liu and G. L. Tang, Chem. Biol., 2012, 19, 1313–1323.
302. L. F. Wu, H. Y. He, H. X. Pan, L. Han, R. Wang and G. L. Tang, Org. Lett., 2014, 16, 1578–1581.
303. Z. Tian, P. Sun, Y. Yan, Z. Wu, Q. Zheng, S. Zhou, H. Zhang, F. Yu, X. Jia, D. Chen, A. Mandi, T. Kurtan and W. Liu, Nat. Chem. Biol., 2015, 11, 259–265.
304. W. R. Roush and D. A. Barda, Org. Lett., 2002, 4, 1539–1542.
305. W. R. Roush, C. Limberakis, R. K. Kunz and D. A. Barda, Org. Lett., 2002, 4, 1543–1546.
306. T. K. Trullinger, J. Qi and W. R. Roush, J. Org. Chem., 2006, 71, 6915–6922.
307. G. Schlingmann, L. Milne and G. T. Carter, Tetrahedron, 2002, 58, 6825–6835.
308. V. Ruanglek, S. Chokpaiboon, N. Rattanaphan, S. Madla, P. Auncharoen, T. Bunyapaiboonsri and M. Isaka, J. Antibiot., 2007, 60, 748–751.
309. M. Chinworrungsee, P. Kittakoop, M. Isaka, P. Maithip, S. Supothina and Y. Thebtaranonth, J. Nat. Prod., 2004, 67, 689–692.
310. P. Wattana-amorn, P. Juthaphan, M. Sirikamonsil, A. Sriboonlert, T. J. Simpson and N. Kongkathip, J. Nat. Prod., 2013, 76, 1235–1237.
311. W. Bunnak, P. Wonnapinij, A. Sriboonlert, C. M. Lazarus and P. Wattana-Amorn, Org. Biomol. Chem., 2019, 17, 374–379.
312. W. Pache and H. Zähner, Arch. Mikrobiol., 1969, 67, 156–165.
313. J. Kohno, T. Kawahata, T. Otake, M. Morimoto, H. Mori, N. Ueba, M. Nishio, A. Kinumaki, S. Komatsubara and K. Kawashima, Biosci., Biotechnol., Biochem., 1996, 60, 1036–1037.
314. M. Perez, C. Crespo, C. Schleissner, P. Rodriguez, P. Zuniga and F. Reyes, J. Nat. Prod., 2009, 72, 2192–2194.
315. Y. Shimizu, Y. Ogasawara, A. Matsumoto and T. Dairi, J. Antibiot., 2018, 71, 968–970.
316. W. Moreira, D. B. Aziz and T. Dick, Front. Microbiol., 2016, 7, 199.
317. A. Cotto-Rosario, E. Y. D. Miller, F. G. Fumuso, J. A. Clement, M. J. Todd and R. M. O’Connor, Microorganisms, 2022, 10, 2260.
318. S. I. Elshahawi, A. E. Trindade-Silva, A. Hanora, A. W. Han, M. S. Flores, V. Vizzoni, C. G. Schrago, C. A. Soares, G. P. Concepcion, D. L. Distel, E. W. Schmidt and M. G. Haygood, Proc. Natl. Acad. Sci. U. S. A., 2013, 110, E295–304.
319. F. Surup, D. Chauhan, J. Niggemann, E. Bartok, J. Herrmann, M. Keck, W. Zander, M. Stadler, V. Hornung and R. Muller, ACS Chem. Biol., 2018, 13, 2981–2988.
320. H. Irschik, D. Schummer, K. Gerth, G. Hofle and H. Reichenbach, J. Antibiot., 1995, 48, 26–30.
321. T. S. S. Chen, C.-J. Chang and H. G. Floss, J. Am. Chem. Soc., 1981, 103, 4565–4568.
322. T. S. S. Chen, C. J. Chang and H. G. Floss, J. Org. Chem., 1981, 46, 2661–2665.
323. J. C. Yang, R. Madupu, A. S. Durkin, N. A. Ekborg, C. S. Pedamallu, J. B. Hostetler, D. Radune, B. S. Toms, B. Henrissat, P. M. Coutinho, S. Schwarz, L. Field, A. E. Trindade-Silva, C. A. Soares, S. Elshahawi, A. Hanora, E. W. Schmidt, M. G. Haygood, J. Posfai, J. Benner, C. Madinger, J. Nove, B. Anton, K. Chaudhary, J. Foster, A. Holman, S. Kumar, P. A. Lessard, Y. A. Luyten, B. Slatko, N. Wood, B. Wu, M. Teplitski, J. D. Mougous, N. Ward, J. A. Eisen, J. H. Badger and D. L. Distel, PLoS One, 2009, 4, e6085.
324. S. I. Elshahawi, A. E. Trindade-Silva, A. Hanora, A. W. Han, M. S. Flores, V. Vizzoni, C. G. Schrago, C. A. Soares, G. P. Concepcion, D. L. Distel, E. W. Schmidt and M. G. Haygood, Proc. Natl. Acad. Sci. U. S. A., 2013, 110, E295–E304.
325. M. A. Avery, S. C. Choudhry, O. P. Dhingra, B. D. Gray, M. C. Kang, S. C. Kuo, T. R. Vedananda, J. D. White and A. J. Whittle, Org. Biomol. Chem., 2014, 12, 9116–9132.
326. H. J. Kwon, W. C. Smith, A. J. Scharon, S. H. Hwang, M. J. Kurth and B. Shen, Science, 2002, 297, 1327–1330.
327. H. J. Kwon, W. C. Smith, L. Xiang and B. Shen, J. Am. Chem. Soc., 2001, 123, 3385–3386.
328. Y. Wu and Y. P. Sun, Org. Lett., 2006, 8, 2831–2834.
329. Y. Rebets, E. Brotz, N. Manderscheid, B. Tokovenko, M. Myronovskyi, P. Metz, L. Petzke and A. Luzhetskyy, Angew. Chem., Int. Ed., 2015, 54, 2280–2284.
330. P. A. McCann and B. M. Pogell, J. Antibiot., 1979, 32, 673–678.
331. M. Natsume, J. Tazawa, K. Yagi, H. Abe, S. Kondo and S. Marumo, J. Antibiot., 1995, 48, 1159–1164.
332. B. M. Pogell, Cell. Mol. Biol., 1998, 44, 461–463.
333. M. Natsume, S. Kondo and S. Marumo, J. Chem. Soc., Chem. Commun., 1989, 1911–1913,  10.1039/c39890001911.
334. L. Glaser, M. Kuhl, J. Stegmuller, C. Ruckert, M. Myronovskyi, J. Kalinowski, A. Luzhetskyy and C. Wittmann, Microb. Cell Fact., 2021, 20, 111.
335. M. Hashimoto, H. Komatsu, I. Kozone, H. Kawaide, H. Abe and M. Natsume, Biosci., Biotechnol., Biochem., 2005, 69, 315–320.
336. G. B. Ren and Y. Wu, Org. Lett., 2009, 11, 5638–5641.
337. O. Germay, N. Kumar and E. J. Thomas, Tetrahedron Lett., 2001, 42, 4969–4974.
338. E. J. Jeong, E. J. Kang, L. T. Sung, S. K. Hong and E. Lee, J. Am. Chem. Soc., 2002, 124, 14655–14662.
339. P. Fischer, A. B. Garcia Segovia, M. Gruner and P. Metz, Angew. Chem., Int. Ed., 2005, 44, 6231–6234.
340. G. Hanquet, X. Salom-Roig and S. Lanners, Chimia, 2016, 70, 20–28.
341. P. Fischer, M. Gruner, A. Jager, O. Kataeva and P. Metz, Chemistry, 2011, 17, 13334–13340.
342. P. A. Bartlett, J. D. Meadows and E. Ottow, J. Am. Chem. Soc., 1984, 106, 5304–5311.
343. J. Xie and N. Bogliotti, Chem. Rev., 2014, 114, 7678–7739.
344. T. Golakoti, W. Y. Yoshida, S. Chaganty and R. E. Moore, J. Nat. Prod., 2001, 64, 54–59.
345. F. Kopp, C. Mahlert, J. Grunewald and M. A. Marahiel, J. Am. Chem. Soc., 2006, 128, 16478–16479.
346. B. S. Evans, I. Ntai, Y. Chen, S. J. Robinson and N. L. Kelleher, J. Am. Chem. Soc., 2011, 133, 7316–7319.
347. A. Krunic, A. Vallat, S. Mo, D. D. Lantvit, S. M. Swanson and J. Orjala, J. Nat. Prod., 2010, 73, 1927–1932.
348. T. A. Wilson, R. J. Tokarski, 2nd, P. Sullivan, R. M. Demoret, J. Orjala, L. H. Rakotondraibe and J. R. Fuchs, J. Nat. Prod., 2018, 81, 534–542.
349. L. R. Malins, J. N. deGruyter, K. J. Robbins, P. M. Scola, M. D. Eastgate, M. R. Ghadiri and P. S. Baran, J. Am. Chem. Soc., 2017, 139, 5233–5241.
350. Y. Igarashi, Y. Kim, Y. In, T. Ishida, Y. Kan, T. Fujita, T. Iwashita, H. Tabata, H. Onaka and T. Furumai, Org. Lett., 2010, 12, 3402–3405.
351. H. Komaki, N. Ichikawa, A. Oguchi, M. Hamada, E. Harunari, S. Kodani, N. Fujita and Y. Igarashi, Stand. Genomic Sci., 2016, 11, 85.
352. H. J. Zhu, B. Zhang, L. Wang, W. Wang, S. H. Liu, Y. Igarashi, G. Bashiri, R. X. Tan and H. M. Ge, J. Am. Chem. Soc., 2021, 143, 4751–4757.
353. H. J. Zhu, B. Zhang, W. Wei, S. H. Liu, L. Xiang, J. Zhu, R. H. Jiao, Y. Igarashi, G. Bashiri, Y. Liang, R. X. Tan and H. M. Ge, Nat. Commun., 2022, 13, 4499.
354. G. K. Nguyen, A. Kam, S. Loo, A. E. Jansson, L. X. Pan and J. P. Tam, J. Am. Chem. Soc., 2015, 137, 15398–15401.
355. C. J. Podracky, C. An, A. DeSousa, B. M. Dorr, D. M. Walsh and D. R. Liu, Nat. Chem. Biol., 2021, 17, 317–325.
356. X. Y. Hemu, X. H. Zhang and J. P. Tam, Org. Lett., 2019, 21, 2029–2032.
357. G. K. T. Nguyen, X. Hemu, J. P. Quek and J. P. Tam, Angew. Chem., Int. Ed., 2016, 55, 12802–12806.
358. X. Gao, S. W. Haynes, B. D. Ames, P. Wang, L. P. Vien, C. T. Walsh and Y. Tang, Nat. Chem. Biol., 2012, 8, 823–830.
359. J. Zhang, N. Liu, R. A. Cacho, Z. Gong, Z. Liu, W. Qin, C. Tang, Y. Tang and J. Zhou, Nat. Chem. Biol., 2016, 12, 1001–1003.
360. T. A. Keating, C. G. Marshall, C. T. Walsh and A. E. Keating, Nat. Struct. Biol., 2002, 9, 522–526.
361. S. A. Samel, G. Schoenafinger, T. A. Knappe, M. A. Marahiel and L. O. Essen, Structure, 2007, 15, 781–792.
362. T. Izore, Y. T. Candace Ho, J. A. Kaczmarski, A. Gavriilidou, K. H. Chow, D. L. Steer, R. J. A. Goode, R. B. Schittenhelm, J. Tailhades, M. Tosin, G. L. Challis, E. H. Krenske, N. Ziemert, C. J. Jackson and M. J. Cryle, Nat. Commun., 2021, 12, 2511.
363. A. Tanovic, S. A. Samel, L. O. Essen and M. A. Marahiel, Science, 2008, 321, 659–663.
364. L. Zhang, C. Wang, K. Chen, W. Zhong, Y. Xu and I. Molnar, Nat. Prod. Rep., 2023, 40, 62–88.
365. S. Sivanathan and J. Scherkenbeck, Molecules, 2014, 19, 12368–12420.
366. F. Kopp and M. A. Marahiel, Nat. Prod. Rep., 2007, 24, 735–749.
367. R. Sussmuth, J. Muller, H. von Dohren and I. Molnar, Nat. Prod. Rep., 2011, 28, 99–124.
368. H. D. Mootz, D. Schwarzer and M. A. Marahiel, ChemBioChem, 2002, 3, 490–504.
369. S. M. Batiste, D. J. Blackwell, K. Kim, D. O. Kryshtal, N. Gomez-Hurtado, R. T. Rebbeck, R. L. Cornea, J. N. Johnston and B. C. Knollmann, Proc. Natl. Acad. Sci. U. S. A., 2019, 116, 4810–4815.
370. L. Richter, F. Wanka, S. Boecker, D. Storm, T. Kurt, O. Vural, R. Sussmuth and V. Meyer, Fungal Biol. Biotechnol., 2014, 1, 4.
371. M. Urbaniak, A. Waskiewicz and L. Stepien, Toxins, 2020, 12, 765.
372. E. Dorschner and H. Lardy, Antimicrob. Agents Chemother., 1968, 8, 11–14.
373. R. L. Hamill, C. E. Higgens, H. E. Boaz and M. Gorman, Tetrahedron Lett., 1969, 10, 4255–4258.
374. A. Logrieco, A. Moretti, G. Castella, M. Kostecki, P. Golinski, A. Ritieni and J. Chelkowski, Appl. Environ. Microbiol., 1998, 64, 3084–3088.
375. A. Harder, L. Holden-Dye, R. Walker and F. Wunderlich, Parasitol. Res., 2005, 97(Suppl 1), S1–S10.
376. Y. Xu, R. Orozco, E. M. Wijeratne, A. A. Gunatilaka, S. P. Stock and I. Molnar, Chem. Biol., 2008, 15, 898–907.
377. C. Steiniger, S. Hoffmann and R. D. Sussmuth, Cell Chem. Biol., 2019, 26, 1526–1534 e1522.
378. W. Weckwerth, K. Miyamoto, K. Iinuma, M. Krause, M. Glinski, T. Storm, G. Bonse, H. Kleinkauf and R. Zocher, J. Biol. Chem., 2000, 275, 17909–17915.
379. J. Muller, S. C. Feifel, T. Schmiederer, R. Zocher and R. D. Sussmuth, ChemBioChem, 2009, 10, 323–328.
380. A. Prosperini, H. Berrada, M. J. Ruiz, F. Caloni, T. Coccini, L. J. Spicer, M. C. Perego and A. Lafranconi, Front. Public Health, 2017, 5, 304.
381. S. M. Batiste and J. N. Johnston, Proc. Natl. Acad. Sci. U. S. A., 2016, 113, 14893–14897.
382. S. Monma, T. Sunazuka, K. Nagai, T. Arai, K. Shiomi, R. Matsui and S. Omura, Org. Lett., 2006, 8, 5601–5604.
383. T. Ohshiro, D. Matsuda, T. Kazuhiro, R. Uchida, K. Nonaka, R. Masuma and H. Tomoda, J. Antibiot., 2012, 65, 255–262.
384. K. Shiomi, R. Matsui, A. Kakei, Y. Yamaguchi, R. Masuma, H. Hatano, N. Arai, M. Isozaki, H. Tanaka, S. Kobayashi, A. Turberg and S. Omura, J. Antibiot., 2010, 63, 77–82.
385. D. X. Hu, M. Bielitza, P. Koos and S. V. Ley, Tetrahedron Lett., 2012, 53, 4077–4079.
386. M. M. Shemyakin, Y. A. Ovchinnikov, A. A. Kiryushkin and V. T. Ivanov, Tetrahedron Lett., 1963, 4, 885–890.
387. A. D. Borthwick, Chem. Rev., 2012, 112, 3641–3716.
388. M. Teixidó, E. Zurita, M. Malakoutikhah, T. Tarragó and E. Giralt, J. Am. Chem. Soc., 2007, 129, 11802–11813.
389. M. Millward, P. Mainwaring, A. Mita, K. Federico, G. K. Lloyd, N. Reddinger, S. Nawrocki, M. Mita and M. A. Spear, Invest. New Drugs, 2012, 30, 1065–1073.
390. Y. Yamazaki, K. Tanaka, B. Nicholson, G. Deyanat-Yazdi, B. Potts, T. Yoshida, A. Oda, T. Kitagawa, S. Orikasa, Y. Kiso, H. Yasui, M. Akamatsu, T. Chinen, T. Usui, Y. Shinozaki, F. Yakushiji, B. R. Miller, S. Neuteboom, M. Palladino, K. Kanoh, G. K. Lloyd and Y. Hayashi, J. Med. Chem., 2012, 55, 1056–1071.
391. M. Nishida, Y. Mine, T. Matsubara, S. Goto and S. Kuwahara, J. Antibiot., 1972, 25, 582–593.
392. P. Belin, M. Moutiez, S. Lautru, J. Seguin, J. L. Pernodet and M. Gondry, Nat. Prod. Rep., 2012, 29, 961–979.
393. M. Gondry, L. Sauguet, P. Belin, R. Thai, R. Amouroux, C. Tellier, K. Tuphile, M. Jacquet, S. Braud, M. Courcon, C. Masson, S. Dubois, S. Lautru, A. Lecoq, S. Hashimoto, R. Genet and J. L. Pernodet, Nat. Chem. Biol., 2009, 5, 414–420.
394. M. W. Vetting, S. S. Hegde and J. S. Blanchard, Nat. Chem. Biol., 2010, 6, 797–799.
395. M. J. Cryle, S. G. Bell and I. Schlichting, Biochemistry, 2010, 49, 7282–7296.
396. T. W. Giessen, A. M. von Tesmar and M. A. Marahiel, Biochemistry, 2013, 52, 4274–4283.
397. J. Liu, X. Xie and S. M. Li, Angew. Chem., Int. Ed., 2019, 58, 11534–11540.
398. N. M. Vior, R. Lacret, G. Chandra, S. Dorai-Raj, M. Trick and A. W. Truman, Appl. Environ. Microbiol., 2018, 84, e02828–e02817.
399. T. Yao, J. Liu, E. Jin, Z. Liu, H. Li, Q. Che, T. Zhu, D. Li and W. Li, iScience, 2020, 23, 101323.
400. M. Moutiez, E. Schmitt, J. Seguin, R. Thai, E. Favry, P. Belin, Y. Mechulam and M. Gondry, Nat. Commun., 2014, 5, 5141.
401. Y. Tingting, Z. Jingxing, L. Jing, L. Huayue and L. Wenli, Chin. J. Org. Chem., 2019, 39, 328–338.
402. J. F. Gonzalez, I. Ortin, E. de la Cuesta and J. C. Menendez, Chem. Soc. Rev., 2012, 41, 6902–6915.
403. J. Fan, H. Ran, P. L. Wei, Y. Li, H. Liu, S. M. Li, Y. Hu and W. B. Yin, Angew. Chem., Int. Ed., 2023, 62, e202217212.
404. H. Liu, J. Fan, P. Zhang, Y. Hu, X. Liu, S. M. Li and W. B. Yin, Chem. Sci., 2021, 12, 4132–4138.
405. Y. Tsunematsu, N. Maeda, M. Yokoyama, P. Chankhamjon, K. Watanabe, K. Scherlach and C. Hertweck, Angew. Chem., Int. Ed., 2018, 57, 14051–14054.
406. C. J. Guo, H. H. Yeh, Y. M. Chiang, J. F. Sanchez, S. L. Chang, K. S. Bruno and C. C. Wang, J. Am. Chem. Soc., 2013, 135, 7205–7213.
407. P. D. Sherkhane, R. Bansal, K. Banerjee, S. Chatterjee, D. Oulkar, P. Jain, L. Rosenfelder, S. Elgavish, B. A. Horwitz and P. K. Mukherjee, ChemistrySelect, 2017, 2, 3347–3352.
408. M. R. Ghiffary, C. P. S. Prabowo, K. Sharma, Y. Yan, S. Y. Lee and H. U. Kim, ACS Sustainable Chem. Eng., 2021, 9, 6613–6622.
409. B. Pang, Y. Chen, F. Gan, C. Yan, L. Jin, J. W. Gin, C. J. Petzold and J. D. Keasling, J. Am. Chem. Soc., 2020, 142, 10931–10935.
410. C. Huang, H. Cui, H. Ren and H. Zhao, JACS Au, 2023, 3, 195–203.
411. B. S. Moore, J. L. Chen, G. M. L. Patterson, R. E. Moore, L. S. Brinen, Y. Kato and J. Clardy, J. Am. Chem. Soc., 1990, 112, 4061–4063.
412. B. S. Moore, J.-L. Chen, G. M. L. Patterson and R. E. Moore, Tetrahedron, 1992, 48, 3001–3006.
413. S. Luo, H. S. Kang, A. Krunic, G. E. Chlipala, G. Cai, W. L. Chen, S. G. Franzblau, S. M. Swanson and J. Orjala, Tetrahedron Lett., 2014, 55, 686–689.
414. H. T. Bui, R. Jansen, H. T. Pham and S. Mundt, J. Nat. Prod., 2007, 70, 499–503.
415. S. C. Bobzin and R. E. Moore, Tetrahedron, 1993, 49, 7615–7626.
416. T. P. Martins, C. Rouger, N. R. Glasser, S. Freitas, N. B. de Fraissinette, E. P. Balskus, D. Tasdemir and P. N. Leao, Nat. Prod. Rep., 2019, 36, 1437–1461.
417. H. Nakamura, E. E. Schultz and E. P. Balskus, Nat. Chem. Biol., 2017, 13, 916–921.
418. K. C. Nicolaou, Y. P. Sun, H. Korman and D. Sarlah, Angew. Chem., Int. Ed., 2010, 49, 5875–5878.
419. A. N. Kim, A. Ngamnithiporn, E. Du and B. M. Stoltz, Chem. Rev., 2023, 123, 9447–9496.
420. A. R. Battersby, R. B. Herbert and F. Šantavý, Chem. Commun., 1965, 415–416.
421. J. Wang and G. Evano, Org. Lett., 2016, 18, 3542–3545.
422. O. Wintersteiner and J. D. Dutcher, Science, 1943, 97, 467–470.
423. W. C. Bowman, Br. J. Pharmacol., 2006, 147(Suppl 1), S277–S286.
424. S. Panjikar, J. Stoeckigt, S. O'Connor and H. Warzecha, Nat. Prod. Rep., 2012, 29, 1176–1200.
425. Y.-J. Xu, S.-G. Cao, X.-H. Wu, Y.-H. Lai, B. Tan, J. Pereira, S. Goh, G. Venkatraman, L. J. Harrison and K.-Y. Sim, Tetrahedron Lett., 1998, 39, 9103–9106.
426. S. Feng, Y. Jiang, J. Li, S. Qiu and T. Chen, Nat. Prod. Res., 2014, 28, 81–85.
427. K. D. Reichl, M. J. Smith, M. K. Song, R. P. Johnson and J. A. Porco, Jr., J. Am. Chem. Soc., 2017, 139, 14053–14056.
428. M. J. Smith, K. D. Reichl, R. A. Escobar, T. J. Heavey, D. F. Coker, S. E. Schaus and J. A. Porco, Jr., J. Am. Chem. Soc., 2019, 141, 148–153.
429. A. R. Pereira, C. F. McCue and W. H. Gerwick, J. Nat. Prod., 2010, 73, 217–220.
430. G. C. Tay, M. R. Gesinski and S. D. Rychnovsky, Org. Lett., 2013, 15, 4536–4539.
431. H. Kim and J. Hong, Org. Lett., 2010, 12, 2880–2883.
432. W. Che, Y. Z. Li, J. C. Liu, S. F. Zhu, J. H. Xie and Q. L. Zhou, Org. Lett., 2019, 21, 2369–2373.
433. A. R. Waldeck and M. J. Krische, Angew. Chem., Int. Ed., 2013, 52, 4470–4473.
434. M. R. Rao and D. J. Faulkner, J. Nat. Prod., 2002, 65, 386–388.
435. J. B. Baker, H. Kim and J. Hong, Tetrahedron Lett., 2015, 56, 3120–3122.
436. G. Peh and P. E. Floreancig, Org. Lett., 2012, 14, 5614–5617.
437. C. S. Barry, J. D. Elsworth, P. T. Seden, N. Bushby, J. R. Harding, R. W. Alder and C. L. Willis, Org. Lett., 2006, 8, 3319–3322.
438. J. M. Cabrera and M. J. Krische, Angew. Chem., Int. Ed., 2019, 58, 10718–10722.
439. K. Jadhav, K. Ahir, S. Desai, S. Desai and S. Acharya, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci., 2022, 1213, 123533.
440. G. Rall, T. Smalberger, H. De Waal and R. Arndt, Tetrahedron Lett., 1967, 8, 3465–3469.
441. H. Rapoport and H. D. Baldridge, Jr., J. Am. Chem. Soc., 1951, 73, 343–346.
442. V. Zunjar, R. P. Dash, M. Jivrajani, B. Trivedi and M. Nivsarkar, J. Ethnopharmacol., 2016, 181, 20–25.
443. T. Sato, S. Aoyagi and C. Kibayashi, Org. Lett., 2003, 5, 3839–3842.
444. R. C. R. Jala, S. Vudhgiri and C. G. Kumar, Carbohydr. Res., 2022, 516, 108556.
445. A. M. Abdel-Mawgoud and G. Stephanopoulos, Synth. Syst. Biotechnol., 2018, 3, 3–19.
446. A. Furstner, J. Ruiz-Caro, H. Prinz and H. Waldmann, J. Org. Chem., 2004, 69, 459–467.
447. J. Qian-Cutrone, T. Ueki, S. Huang, K. A. Mookhtiar, R. Ezekiel, S. S. Kalinowski, K. S. Brown, J. Golik, S. Lowe, D. M. Pirnik, R. Hugill, J. A. Veitch, S. E. Klohr, J. L. Whitney and S. P. Manly, J. Antibiot., 1999, 52, 245–255.
448. M. Tsunakawa, N. Komiyama, O. Tenmyo, K. Tomita, K. Kawano, C. Kotake, M. Konishi and T. Oki, J. Antibiot., 1992, 45, 1467–1471.
449. A. Furstner, J. Mlynarski and M. Albert, J. Am. Chem. Soc., 2002, 124, 10274–10275.
450. A. Furstner, M. Albert, J. Mlynarski, M. Matheu and E. DeClercq, J. Am. Chem. Soc., 2003, 125, 13132–13142.
451. A. Furstner, M. Albert, J. Mlynarski and M. Matheu, J. Am. Chem. Soc., 2002, 124, 1168–1169.
452. S. Takahashi, M. Hosoya, H. Koshino and T. Nakata, Org. Lett., 2003, 5, 1555–1558.
453. K. Yokomizo, Y. Miyamoto, K. Nagao, E. Kumagae, E. S. Habib, K. Suzuki, S. Harada and M. Uyeda, J. Antibiot., 1998, 51, 1035–1039.
454. M. Uyeda, Actinomycetologica, 2003, 17, 57–66.
455. E. S. Habib, K. Yokomizo, K. Murata and M. Uyeda, J. Antibiot., 2000, 53, 1420–1423.
456. T. Ohtsuka, Y. Itezono, N. Nakayama, M. Kurano, N. Nakada, H. Tanaka, S. Sawairi, K. Yokose and H. Seto, Tetrahedron Lett., 1992, 33, 2705–2708.
457. M. Wietz, M. Mansson, J. S. Bowman, N. Blom, Y. Ng and L. Gram, Appl. Environ. Microbiol., 2012, 78, 2039–2042.
458. D. M. Garcia, H. Yamada, S. Hatakeyama and M. Nishizawa, Tetrahedron Lett., 1994, 35, 3325–3328.
459. V. Marti-Centelles, M. D. Pandey, M. I. Burguete and S. V. Luis, Chem. Rev., 2015, 115, 8736–8834.
460. A. Gradillas and J. Perez-Castells, Angew. Chem., Int. Ed., 2006, 45, 6086–6101.
461. G. P. Dinos, Br. J. Pharmacol., 2017, 174, 2967–2983.
462. Q. Su, A. B. Beeler, E. Lobkovsky, J. A. Porco, Jr. and J. S. Panek, Org. Lett., 2003, 5, 2149–2152.
463. A. B. Beeler, D. E. Acquilano, Q. Su, F. Yan, B. L. Roth, J. S. Panek and J. A. Porco, Jr., J. Comb. Chem., 2005, 7, 673–681.
464. M. de Leseleuc and S. K. Collins, Chem. Commun., 2015, 51, 10471–10474.
465. Q. Zhang, X. Cui, S. Lin, J. Zhou and G. Yuan, Org. Lett., 2012, 14, 6126–6129.
466. X. Miao, A. Paikar, B. Lerner, Y. Diskin-Posner, G. Shmul and S. N. Semenov, Angew. Chem., Int. Ed., 2021, 60, 20366–20375.
467. L. A. Wessjohann, D. G. Rivera and O. E. Vercillo, Chem. Rev., 2009, 109, 796–814.
468. C. D. Hein, X. M. Liu and D. Wang, Pharm. Res., 2008, 25, 2216–2230.
469. G. Chouhan and K. James, Org. Lett., 2011, 13, 2754–2757.
470. X. Yang, L. P. Beroske, J. Kemmink, D. T. Rijkers and R. M. Liskamp, Tetrahedron Lett., 2017, 58, 4542–4546.
471. R. Jagasia, J. M. Holub, M. Bollinger, K. Kirshenbaum and M. G. Finn, J. Org. Chem., 2009, 74, 2964–2974.
472. S. Punna, J. Kuzelka, Q. Wang and M. G. Finn, Angew. Chem., Int. Ed., 2005, 44, 2215–2220.
473. J. H. van Maarseveen, W. S. Horne and M. R. Ghadiri, Org. Lett., 2005, 7, 4503–4506.
474. K. D. Bodine, D. Y. Gin and M. S. Gin, J. Am. Chem. Soc., 2004, 126, 1638–1639.
475. N. Pietrzik, D. Schmollinger and T. Ziegler, Beilstein J. Org. Chem., 2008, 4, 30.
476. M. Menand, J. C. Blais, J. M. Valery and J. Xie, J. Org. Chem., 2006, 71, 3295–3298.
477. A. Bordes, A. Poveda, N. Fontelle, A. Arda, J. Guillard, Y. B. Ruan, J. Marrot, S. Imaeda, A. Kato, J. Desire, J. Xie, J. Jimenez-Barbero and Y. Bleriot, Carbohydr. Res., 2021, 501, 108258.
478. W. Fu, L. Wang, Z. Yang, J. S. Shen, F. Tang, J. Zhang and X. Cui, Chem. Commun., 2020, 56, 960–963.
479. Q. Xiong, L. Xiao, X. Q. Dong and C. J. Wang, Org. Lett., 2022, 24, 2579–2584.
480. R. Emadi, A. Bahrami Nekoo, F. Molaverdi, Z. Khorsandi, R. Sheibani and H. Sadeghi-Aliabadi, RSC Adv., 2023, 13, 18715–18733.
481. L. Mendive-Tapia, A. Bertran, J. García, G. Acosta, F. Albericio and R. Lavilla, Chem. – Eur. J., 2016, 22, 13114–13119.
482. J. Garcia-Lacuna, G. Dominguez and J. Perez-Castells, ChemSusChem, 2020, 13, 5138–5163.
483. K. E. Ylijoki and J. M. Stryker, Chem. Rev., 2013, 113, 2244–2266.
484. S. B. Needleman and M. Chang Kuo, Chem. Rev., 1962, 62, 405–431.
485. H. Oikawa and T. Tokiwano, Nat. Prod. Rep., 2004, 21, 321–352.
486. K. Klas, S. Tsukamoto, D. H. Sherman and R. M. Williams, J. Org. Chem., 2015, 80, 11672–11685.
487. H. Wang, Y. Zou, M. Li, Z. Tang, J. Wang, Z. Tian, N. Strassner, Q. Yang, Q. Zheng, Y. Guo, W. Liu, L. Pan and K. N. Houk, Nat. Chem., 2023, 15, 177–184.
488. Q. Zheng, Y. Gong, Y. Guo, Z. Zhao, Z. Wu, Z. Zhou, D. Chen, L. Pan and W. Liu, Cell Chem. Biol., 2018, 25, 718–727 e713.
489. Z. Zhang, C. S. Jamieson, Y. L. Zhao, D. Li, M. Ohashi, K. N. Houk and Y. Tang, J. Am. Chem. Soc., 2019, 141, 5659–5663.
490. B. Briou, B. Ameduri and B. Boutevin, Chem. Soc. Rev., 2021, 50, 11055–11097.
491. M. H. Cao, N. J. Green and S. Z. Xu, Org. Biomol. Chem., 2017, 15, 3105–3129.
492. K. Takao, R. Munakata and K. Tadano, Chem. Rev., 2005, 105, 4779–4807.
493. G. Dominguez and J. Perez-Castells, Chem. Soc. Rev., 2011, 40, 3430–3444.
494. P. J. Salveson, A. P. Moyer, M. Y. Said, G. Gökce, X. Li, A. Kang, H. Nguyen, A. K. Bera, P. M. Levine, G. Bhardwaj and D. Baker, Science, 2024, 384, 420–428.
495. V. Sarojini, A. J. Cameron, K. G. Varnava, W. A. Denny and G. Sanjayan, Chem. Rev., 2019, 119, 10318–10359.
496. A. L. Feldberg, F. Mayerthaler, J. Ruschenbaum, J. Kroger and H. D. Mootz, Angew. Chem., Int. Ed., 2024, e202317753,  DOI:10.1002/anie.202317753.

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Footnotes

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d2cs00909a

‡ These authors contributed equally to this work.

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