University of Groningen
Chemical Modification of Peptide Antibiotics
de Vries, Reinder
DOI:
10.33612/diss.171585325
IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite from it. Please check the document version below.
Document Version
Publisher's PDF, also known as Version of record
Publication date: 2021
Link to publication in University of Groningen/UMCG research database
Citation for published version (APA):
de Vries, R. (2021). Chemical Modification of Peptide Antibiotics. University of Groningen. https://doi.org/10.33612/diss.171585325
Copyright
Other than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons).
Take-down policy
If you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediately and investigate your claim.
Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons the number of authors shown on this cover page is limited to 10 maximum.
Chapter 1
Introduction
In this chapter, the biosynthesis, structure and properties of the classes of antimicrobial peptides that were studied in this thesis will be discussed. An introduction to the chemical modification of such peptides via their dehydroamino acids is given, followed by the current state-of-the-art in the field. Finally, the current challenges in the field are highlighted and the outline of this thesis is presented.
Chapter 1
2
Antimicrobial resistance is a major threat to modern healthcare, and is estimated to cause 50 million deaths annually by the year 2050.[1] Due to overuse in both humans and livestock, bacteria have become resistant to many conventional antibiotics via various sophisticated biological pathways.[2] Moreover, the curative nature of antibiotics and the fact that patients are treated with antibiotics for a relatively short period of time make the development of new antibiotics a very poor investment for the pharmaceutical industry.[3] Consequently, the number of novel treatments released to the market has stalled significantly in the last decades,[3] while the health concerns and economical costs to individuals and society are becoming more and more evident.[4,5] Therefore, the development of new types of treatments that are less susceptible to the antimicrobial resistance associated with conventional antibiotics is crucial.[6]
Antimicrobial peptides isolated from microorganisms have a high activity against a broad range of bacteria and often show lower development of resistance compared to conventional antibiotics,[7,8] which makes them promising candidates in the search for new antibiotics. However, their clinical application is hampered due to poor pharmacokinetics and, in some cases, low water solubility. In order to improve these properties, the late-stage chemical editing to make semi-synthetic analogs has become an increasingly popular approach (Figure 1.1).[9] Nevertheless, achieving chemo- and site-selective modifications on these structurally diverse and complex peptides remains a major synthetic challenge.
In this thesis, the late-stage modification of antimicrobial peptides via their uniquely reactive dehydroalanine residues (Dha) (Figure 1.1) is described. The first part of this introduction will cover several classes of antimicrobial peptides that are studied in this thesis. In the second part dehydroalanine will be introduced as a chemical handle for the selective editing of peptides and proteins and an overview of existing methods for the modification of such residues will be given. Finally, current challenges in the field will be evaluated, followed by the objectives and outline of this thesis.
Introduction
3
Figure 1.1: Schematic representation of the late-stage editing of natural antimicrobial
peptides.
1.1 Antimicrobial peptides
Over the years, bacteria have proven to be a rich source of peptides that have antimicrobial activity.[9] Organisms have evolved these antimicrobial peptides (AMPs) as part of their innate immune system and as a first line of defense against pathogens.[10,11] Bacteria also produce AMPs to eliminate other microorganisms that compete for the same biological environment.[12]
AMPs can be divided into two categories based on their biosynthesis. Ribosomally synthesized AMPs are produced by all life via translation of mRNA.[13] On the other hand, the nonribosomal peptides (NRPs) are mostly produced by bacteria using multifunctional enzymatic “assembly line-like” machineries called nonribosomal peptide synthetases (NRPS).[14] In both ribosomal and nonribosomal AMPs modifications can take place during the final stages of their biosynthesis.[14,15] These are a variety of chemical, mostly enzymatic transformations that occur on specific amino acid residues, either before or after their incorporation into the peptide (Figure 1.2).[14,15] Modifications that affect the overall peptide structure, such as (macro)cyclizations, are also common in AMPs (Figure 1.2).[14,15]
Figure 1.2: Schematic representation of a selection of modifications that can be found
in AMPs (Dha = dehydroalanine, Orn = ornithine). HS
Microorganism
Biosynthesis chemical modificationLate-stage
High activity
Low resistance development
Poor pharmacologcial properties
Isolation
Variant with improved properties, targeting group or fluorescent label Uniquely reactive handle (Dha)
Natural antimicrobial peptide
HO NH2 HS HO NH2 COOH COOH O O N H O N H O S O Lipidation Dehydration Oxidation Incorporation of non-canonical/ D-amino acids Macrocyclization Acetylation
Chapter 1
4
Due to the large variety of modifications that can occur during their biosynthesis, AMPs are a class of natural products that is structurally highly diverse. However, most AMPs have in common that they are short in length (10-50 amino acids), consist for a large part (>50 %) of hydrophobic amino acids and bear a net positive charge.[7] This amphiphilic nature enables AMPs to have strong interactions with bacterial membranes, which is a key factor in their antibacterial activity.[16] Once bound, many AMPs are able to kill their hosts via membrane disruption, forming pores that cause leakage of cytoplasmic contents of the cell.[16] Others are able to translocate over membranes into the cytoplasm and kill the cell by acting on intracellular targets, such as the inhibition of DNA, RNA or protein biosynthesis.[17]
Although resistance mechanisms of bacteria against AMPs are known and have been studied extensively,[18] the development of resistance against AMPs is generally lower compared to conventional antibiotics. The interactions of AMPs with bacterial membranes are highly favorable and it is difficult for bacteria to defend themselves against membrane disruption without disturbing the structural integrity of their membranes.[19] Moreover, many AMPs act moderately on multiple cellular targets rather than binding strongly with one specific target, as is the case with conventional antibiotics, making it more challenging for bacteria to develop resistance via a single mechanism.[20]
Besides low resistance development, AMPs also have an exceptionally broad scope of activity. In addition to high activities against Gram-positive and Gram-negative bacteria the spectrum of AMPs even extends into other domains of life, such as funghi.[21] Moreover, killing of bacteria by AMPs can be extremely rapid compared to conventional antibiotics.[22] Because of their potential as new antibiotics, investigating the use of AMPs for the treatment of infections in human patients is becoming an increasingly popular field of research.[23] Moreover, some AMPs have been used as food preservatives for several years.[24]
In this thesis, several classes of NRPs and ribosomally synthesized and post-translationally modified peptides (RiPPs) have been studied and they will be highlighted in the next sections.
1.2 Non-ribosomal peptides
Nonribosomal peptides (NRPs) are a class of secondary metabolites produced by microorganisms. In contrast to ribosomally produced peptides and proteins, they are synthesized by large, multifunctional enzymes called nonribosomal peptide
Introduction
5
synthetases (NRPSs) that are completely independent of ribosomal translation. These NRPSs work as assembly line-like machineries, building up the peptide chain one amino acid at a time (Figure 1.3). Genes that encode for the peptides can be divided into sequential modules that are each responsible for the incorporation of one amino acid.[25] In turn, each module consists of a set of so-called domains that carry out the different chemical transformations necessary for the incorporation of each amino acid.[25] Each module has a peptidyl carrier protein (PCP)-domain, which carries the growing peptide chain, and an adenylation (A)-domain that recognizes the correct amino acid, activates it and loads the amino acid on the PCP-domain.[26] All modules except the first one also have a condensation (C)-domain, which catalyzes the peptide bond formation of the loaded amino acid with the growing peptide chain.[26] The final module contains a thioesterase (TE)-domain that cleaves the finished peptide from the last PCP-domain.[26] Modules can also contain additional modification domains, allowing for the incorporation of modified or non-canonical amino acids.
Figure 1.3: Schematic representation of the assembly line-like machinery of NRPSs. In
this example, the tripeptide Tyr-Ile-Lys is synthesized, where the Tyr is incorporated as a D-amino acid via an additional epimerization (E) domain.
The adenylation (A)-domain performs the selection of the amino acid and thus effectively controls the amino acid sequence in the final product.[25] This domain uses ATP to activate the amino acid, which results in an aminoacyl adenylate (Scheme 1.1).[27] Next, the A-domain transfers this activated amino acid to the carrier domain (Scheme 1.1).[27] These carrier domains have a conserved serine residue that has a phosphopantetheine cofactor attached, which binds covalently to the amino acid via a thioester linkage (Scheme 1.1).[28]
S O NH2 OH S O NH O NH2 HO S O NH H2N HN O O NH2 OH H2N H N N H O O OH O HO NH2
Chapter 1
6
Scheme 1.1: Activation of the amino acid by the A-domain and transfer to the carrier
domain (top) and chemical structure of the phosphopantetheine cofactor attached to the PCP-domain (bottom).
After the amino acid has been loaded on the PCP-domain, the C-domain catalyzes the peptide bond formation between the amino group of the amino acid on the downstream module with the peptidyl chain on the upstream module (Scheme 1.2).[29,30] Via this cycle of adenylation, loading on the carrier and condensation the peptide is built up one amino acid at a time. Finally, the TE-domain in the last module cleaves the finished peptide from the PCP-domain, which can happen via hydrolysis, resulting in a linear peptide (Scheme 1.2).[26] However, cleavage via macrolactonization is also common in NRPs and gives rise to macrocyclic peptides.[26]
C-domain PCP1 S O H2N R1 PCP2 S O H2N R2 PCP2 S O HN R2 NH2 O R1 PCPn S O HN Rn O TE-domain peptide O H2N R1 O HN Rn O peptide O H2N R1 HO
Scheme 1.2: C-domain catalyzed peptide bond formation between two adjacent
loaded amino acids and cleavage of the finished peptide by the TE-domain.
In addition to the assembly of the peptide chain, a wide variety of modifications can occur during several stages of the biosynthesis of NRPs. First of all, the direct incorporation of non-canonical or modified amino acids can occur during peptide elongation by recognition of such amino acids by specific A-domains.[31] Side chain modifications can also happen during the assembly of the peptide via additional domains within a module, such as epimerization domains for the incorporation of D-amino acids (Figure 1.3).[26] Finally, external tailoring enzymes can modify side chains via enzymatic transformations. These tailoring enzymes can act both during peptide chain elongation or after the peptide assembly is complete, leading to so-called peptide maturation.[26,31] The extent and variety of modifications, combined
H2N OH O R1 A-domain ATP PPi Amino acid H2N O O R1 P O O N N N N NH2 O OH OH O Aminoacyl adenylate A-domain AMP PCP SH PCP S O H2N R1 PCP SH HN O OPO O O OH O H N HN O SH
Introduction
7
with frequently occurring macrocyclizations, make NRPs one of the most structurally diverse classes of secondary metabolites produced by microorganisms.
Scheme 1.3: Structures of bogorol A, brevicidine and laterocidine.
Cationic peptide antibiotics are attractive candidates as antibiotics due to their high efficacy, rapid killing of bacteria and low development of resistance.[32,33] Cationic nonribosomal peptides (CNRPs), such as bogorols, brevicidine and laterocidine (Scheme 1.3), have in common that they are short to medium in length, consist of multiple cationic and apolar residues and little to no anionic residues.[34–37] In addition, several types of CNRPs have a fatty acid tail that is introduced via N-terminal lipidation during biosynthesis and are often also referred to as lipopeptides.[37,38] Their amphiphilic nature enables CNRPs to effectively penetrate and disrupt bacterial membranes and in some cases interact strongly with multiple anionic intracellular targets, which gives them a high activity against both Gram-positive and Gram-negative bacteria.[13,36] However, elucidation of the exact mechanism of action of CNRPs is often challenging because of the rapid killing of bacteria and interaction with multiple targets.[13,36]
OH H N O N H O H N O N H H2N O H N O N H O H N O NH2 O N H H N O N H O H N NH2 N H O OH H N O O OH Bogorol A N H O H N N H H N N H H N N H H N N H O O O O O O O O CONH2 OH NH NH2 NH2 NH2 NH O NH O N H O HN O N H O H N N H H N N H H N N H H N N H O O O O O O O O OH OH NH NH2 NH2 NH2 NH O NH O N H O HN O N H O O O OH CONH2 Brevicidine Laterocidine
Chapter 1
8
Although several CNRPs have made it to clinical trials and even therapeutic application,[23] the investigation of these antibiotic candidates is difficult due to their structural diversity and complexity. Because of this, the discovery of new CNRPs is hampered, since many promising candidates that are genetically encoded are being overlooked.[36] To overcome these challenges, research recently focused towards the development of methods for the systematic discovery, isolation and characterization of new candidates from producing organisms.[36,38]
1.3 RiPPs
Ribosomally synthesized and post-translationally modified peptides, or RiPPs, is a class of AMPs that is growing fast in number and attention as a source of new antibiotics.[15] These peptides are initially produced ribosomally as linear precursor peptides, which then undergo multiple side chain modifications after translation.[39] In general, such a linear precursor consists of a core peptide, which will be the final RiPP product, an N-terminal leader peptide and, in some cases, a C-terminal recognition sequence (Figure 1.4).[39] The latter two are necessary for recognition and guidance of the modifying enzymes. After modifications have taken place, the matured core peptide is cleaved from the rest of the peptide, releasing the RiPP natural product (Figure 1.4).[39]
Figure 1.4: Schematic representation of the ribosomal translation and
post-translational modification machinery involved in the biosynthesis of RiPPs.
As a result of this post-translational machinery, RiPPs have highly modified structures which often contain one or more macrocycles and many structural features that are uncommon in other peptides and proteins.[15] Due to this structural diversity, RiPPs have been divided into families based on similarities in their biosynthesis and the resulting motifs that they have in common in their final structure.[15] The research in this thesis is primarily focused on the thiopeptide and lanthipeptide families of RiPPs, which will be highlighted in the next section.
Introduction
9
1.3.1 Thiopeptides
The thiopeptides are a large family of sulphur-rich RiPPs that are produced by Gram-positive bacteria.[40–42] Characteristic are their heavily modified, macrocyclic structures that contain multiple azole moieties and dehydrated amino acids, such as dehydroalanine (Dha) and dehydrobutyrine (Dhb) (Figure 1.5).[15,40,42] A 6-membered nitrogenous ring is the central scaffold for the macrocycles and a tail region (Figure 1.5). Thiopeptides can be divided into five series (series a-e) based on the oxidation state and substitution pattern of these 6-membered rings (Figure 1.5).[40] On top of these common structural features of thiopeptides, there are additional motifs that are characteristic for each series, or even for individual peptides.
Figure 1.5: Schematic overview of the biosynthesis of thiopeptides.
OH OH HO HO OH OH HO HS HS OH OH N H O Dehydroalanine (Dha) N NH N H O Dehydrobutyrine (Dhb) N X N N N S O Azoles: X=S: thiazole X=O: oxazole OH O OH Series a Series e Series d Series c Series b 6-membered azacycles
Chapter 1
10
The remarkable transformation from linear peptides to these polycyclic structures is the result of a post-translational modification machinery of which the steps have been elucidated only quite recently.[43–46]
The first modification that takes place after ribosomal translation is the dehydrocyclization of Cys, Ser and Thr residues to form thiazoles and oxazoles, respectively (Figure 1.5).[39,47,48] This happens in a two-step sequence of dehydrocyclization by an adenosine triphosphate (ATP)-dependent enzyme to form azoline derivatives, which are subsequently aromatized by a flavin mononucleotide (FMN)-dependent dehydrogenase (Scheme 1.4).[48–52] ATP-mediated phosphorylation is necessary for the activation of alkoxide groups during the cyclodehydration, while FMN is used as an oxidant in the dehydrogenation of the azoline intermediates (Scheme 1.4).[48] N H X O O H R Cys: X=S, R=H Ser: X=O, R=H Thr: X=O, R=Me B: HN X O O R ATP ADP N X O O R P OH O O H B: N X O R FMN N X O R FMNH2 HPO4 2-Azoline Azole
Scheme 1.4: Mechanism of enzymatic azole formation during the biosynthesis of
thiopeptides.
The dehydration of Ser and Thr residues to Dha and Dhb is the next step in the post-translational modification of thiopeptides (Figure 1.5). Enzymatic elimination of a hydroxyl functionality to make a carbon-carbon double bond requires conversion of the hydroxyl to a good leaving group, which typically happens via phosphorylation or acetylation. However, the biosynthesis of Dha and Dhb in thiopeptides involves a unique glutamylation-elimination pathway, where the donor of the glutamyl moiety is glutamyl-tRNAGlu (Scheme 1.5).[47] The glutamylation of Ser and Thr proceeds via a glutamyl-tRNAGlu-dependent enzyme, while the elimination is carried out by a separate enzyme.[47]
Introduction 11 N H R O O O Ser: R=H Thr: R=Me B: Glutamate H N N N N NH2 O OH O O O H3N O O P O O O tRNAGlu N H R O O O B: O NH 3 O O H N H O O R Dha: R=H Dhb: R=Me Scheme 1.5: Mechanism of the enzymatic dehydration of Ser and Thr in thiopeptides.
Finally, the 6-membered azacycle in thiopeptides is formed in a [4+2] aza-Diels-Alder cycloaddition between two dehydroalanines (Figure 1.5). This is done by multifunctional Diels-Alderase enzymes, which perform most of the steps involved in this reaction, including isomerization, cycloaddition, dehydration and subsequent reductions or aromatizations (Scheme 1.6).[53] A tautomerized Dha reacts with another Dha residue in a [4+2] cycloaddition followed by dehydration.[53] The resulting dihydropyridine is either aromatized to a pyridine derivative (series d and e)[54–57] or reduced to a dehydropiperidine (series b) (Scheme 1.6).[46] The dehydropiperidine can be further reduced to a piperidine (series a)[58] or rearranged to form an imidazopiperidine (series c)[59] (Scheme 1.6). After final peptide-specific modifications and cleavage of the leader peptide the natural product is finished.
H2O N H O O N O HO N H O O Tautomerization Aza-Diels-Alder [4+2] N HO H H B: Dehydration N Dihydropyridine Aromatization N Reduction N Pyridine Dehydropiperidine Reduction NH Piperidine N N S OH Rearrangement Imidazopiperidine
Scheme 1.6: Biosynthetic pathway of the 6-membered azacycle formation in
thiopeptides.
Thiopeptides are most well-known for their high antimicrobial activity against Gram-positive bacteria[42], however they also possess antifungal[60] and antitumor[61] properties. Their potent activity stems from their ability to bind to ribosomes very strongly, blocking ribosomal protein synthesis and thereby bacterial growth. However, thiopeptides have little to no activity against Gram-negative bacteria due to their
Chapter 1
12
inability to cross the outer membrane of these species.[62,63] Despite their potential as antibiotics, their poor solubility in aqueous media is a major drawback in their clinical application.[62,63]
1.3.1.1 Thiostrepton
Scheme 1.7: Structure of thiostrepton with its characteristic features highlighted (red:
Dha, orange: Dhb, green: thiazoles, pink: thiazoline, purple: quinaldic acid, brown: oxidized isoleucine, blue: dehydropiperidine).
Thiostrepton (Scheme 1.7) is one of the most well-known and extensively studied members of the thiopeptide family of RiPPs. It was first isolated from the Gram-positive soil bacterium Streptomyces azureus in 1954 and its structure was elucidated in 1970.[64,65] A few years later it was found that thiostrepton is also produced by Streptomyces laurentii.[66] Its full biosynthesis was reported in 2009, showing that thiostrepton was indeed ribosomally synthesized and required posttranslational modification for maturation.[44,46]
Thiostrepton is one of the series b thiopeptides, which have a dehydropiperidine as the central azacycle.[15,65] Similar to the series a and c, the series
b are known to have a second macrocycle, which contains a quinaldic acid residue,
attached to the central 6-membered core (Scheme 1.7).[58,59,65] Besides the other structural motifs that are common in thiopeptides, thiostrepton also contains a thiazoline moiety, which is an azole moiety that did not undergo aromatization during the biosynthesis, and an oxidized isoleucine in its main macrocycle (Scheme 1.7).
The antimicrobial activity of thiostrepton against Gram-positive bacteria is very high. By binding to the bacterial ribosome/L11 protein complex, translocation of
N H N O S N HN O S N H N HO O HN N S H N O S N OH NH NH O O H N O N H O S N NH H N O O NH2 O N OH OH O HO H H O thiostrepton quinaldic acid thiazoline oxidized isoleucine
Introduction
13
the growing peptide-tRNA complex within the ribosome is blocked.[67–69] This halts ribosomal protein synthesis and therefore bacterial growth.[70] In vitro studies have shown that both Gram-positive and Gram-negative bacteria are sensitive to this inhibition of protein synthesis, however the latter are unaffected by thiostrepton because it is not able to cross the outer membrane of Gram-negative bacteria.[62,67] Apart from its antimicrobial activity, thiostrepton also displays high anticancer activity, selectively killing breast cancer cells without affecting healthy mammalian cells.[71] Despite this promising bioactivity profile, its pharmacokinetics and poor water solubility hamper its therapeutic potential, which is currently limited to the veterinary use in salves to treat skin infections.
1.3.1.2 Nosiheptide
Scheme 1.8: Structure of nosiheptide with its characteristic features highlighted (red:
Dha, orange: Dhb, green: thiazoles, purple: indolic acid, blue: 5-hydroxypyridine).
Another well-known member of the thiopeptide antibiotics is nosiheptide, a relatively small thiopeptide produced by Streptomyces actuosus.[72,73] First discovered under the name multhiomycin, its structure was later solved and found to be identical with that of nosiheptide.[74,75] Its biosynthesis has been studied extensively over the years, elucidating the post-translational machinery behind its highly modified structure.[57,76– 78]
Nosiheptide is a series e thiopeptide, having a 5-hydroxypyridine as the central scaffold (Scheme 1.8).[15,72] After the aza-Diels-Alder and aromatization, the pyridine of series e thiopeptides is oxidized further at the 5-position.[57] This gives rise to the tetrasubstituted pyridine, which is the type of azacycle with the highest oxidation state in all thiopeptides. Similar to the series a-c, the series e also contain a second macrocycle, but with an indolic acid moiety incorporated instead of a quinaldic acid (Scheme 1.8).[57,72] N S N N S N S OH NH S O N H O O S N O OH HN O S N NH O NH HO O HN O O NH2 nosiheptide indolic acid
Chapter 1
14
The spectrum of activity against Gram-positive bacteria and the mechanism of action of nosiheptide are similar to that of thiostrepton.[68,79,80] Nosiheptide has a very high potency, which is in some cases even superior to that of thiostrepton.[80,81] However, nosiheptide also suffers from low water solubility and oral bioavailability, limiting its use in medicine. Currently, nosiheptide is used as a feed additive due to its positive effect on the growth of chickens and pigs, while it is not detected in significant amounts in meat.[82]
1.3.2 Lanthipeptides
One of the oldest known families of RiPPs is that of the lanthipeptides.[15] The term lanthipeptides stems from their characteristic lanthionines (Lan) and methyllanthionines (MeLan) which are thioether-bridged amino acids formed during post-translational modifications (Figure 1.6).[15,83] Apart from these thioether crosslinkages lanthipeptides also contain Dha and Dhb. In general, lanthipeptides are less heavily modified than thiopeptides. Lanthipeptides that possess antimicrobial activity are often called lantibiotics.[84]
Figure 1.6: Schematic overview of the biosynthesis of lanthipeptides.
HO SH SH SH HO OH HO HO N H NH S O O N H NH S O O N H O N H O SH SH SH S S S S S Dehydroalanine
Introduction
15
After ribosomal translation, the linear lanthipeptide precursors are modified to form Lan and Melan moieties (Figure 1.6). First, Ser and Thr are dehydrated to Dha and Dhb (Figure 1.6).[39,85,86] Next, the thioether bridges are formed via Michael addition of Cys thiols to Dha or Dhb, giving rise to Lan or MeLan, respectively (Figure 1.6).[39,85,86] These transformations occur enzymatically and lanthipeptides can be divided into classes (I-IV) based on the biosynthetic enzymes involved in these post-translational modifications.[85] In class I lanthipeptides the dehydrations and cyclizations are carried out separately by a dehydratase (LanB) and a cyclase (LanC).[87,88] In classes II-IV one enzyme is responsible for both the dehydrations and cyclizations (LanM, LanKC and LanL, respectively).[89–91] Upon maturation of the core peptide the leader sequence is cleaved off by a separate protease (LanP).[86] While LanX is the generic term for these modifying enzymes, each lanthipeptide has its own particular designation.
Lanthipeptides get their antimicrobial activity from their ability to bind and disrupt membranes. Most lantibiotics bind strongly to lipid II, after which the host is killed via inhibition of cell wall synthesis, formation of membrane pores that allow leakage of cytoplasmic contents or a combination of those two.[92] This dual mode of action makes it very difficult for bacteria to develop resistance against these peptides. 1.3.2.1 Nisin
Scheme 1.9: Structure of nisin Z with its characteristic features highlighted (red: Dha,
orange: Dhb, blue: lanthionine, purple: methyllanthionine, gray: hinge region).
The oldest known lanthipeptide is the lantibiotic nisin (Scheme 1.9), produced by
Lactococcus lactis.[93] The discovery of nisin in 1928 even predates the discovery of penicillin by Alexander Fleming,[93] however it took until 1971 to completely solve its
H N O O NH S NH O N H O H N O O NH H N O S N O O H N HN O O NH H2N O HN NH OO NH HN O H N O S O HN O NH O N H S NH O H2N O O HN S O NH NH2 S O HN NH O N H O H N O NH O S O HN OH O NH O NH HN O HN N O N H O HN O OH NH2 NH2 O H2N A B C D E nisin Z hinge region
Chapter 1
16
structure.[94] The structure of nisin consists of four methyllanthionines and one lanthionine, which together make up five thioether-bridged macrocycles labeled A-E from N to C-terminus (Scheme 1.9). Moreover, there are also two residual dehydroalanines and one dehydrobutyrine that did not end up forming thioether bridges.
Nisin is a class I lanthipeptide, which means that the formation of the Lan and MeLan moieties is carried out by two separate enzymes. The first enzyme, NisB, is responsible for the dehydration to form Dha and Dhb residues from Ser and Thr, respectively.[87] NisB is a Glu-tRNAGlu-dependent enzyme, which glutamylates the hydroxyl group of Ser and Thr, followed by elimination of glutamate to form the double bond.[95] This happens via a mechanism similar to that of the biosynthesis of Dha and Dhb in thiopeptides (vide supra).[95] However, NisB performs the complete dehydration, while in the biosynthesis of thiopeptides glutamylation and elimination are carried out by two separate enzymes. Next, the thioether bridges are formed via thia-Michael additions of cysteine residues on the dehydroamino acids, which is catalyzed by a second enzyme called NisC.[88] Finally, the mature peptide is cleaved from the leader peptide by NisP, a serine protease, to release the finished natural product.[96]
The mode of action of nisin involves binding of its N-terminal region containing the A, B and C rings to lipid II in the cell membrane, inhibiting cell wall synthesis.[97] In addition, the flexible hinge region allows the C-terminal fragment containing the D and E rings to form pores, resulting in membrane disruption and leakage of intracellular contents.[97,98] Due to its high efficacy and low toxicity, nisin has been used as a food preservative for a variety of products, including dairy, meat and fish, for decades.[99]
1.4 Chemical modification of RiPPs
Despite the promising bioactivity profile of RiPPs as antibiotics, their therapeutic use is hampered by their pharmacological and chemical properties. Post-translational modifications such as cyclizations improve proteolytic and chemical stability of RiPPs by limiting conformational flexibility, however many still suffer from poor metabolic stability in vivo.[15] Some families of RiPPs, such as thiopeptides, also have a low aqueous solubility, which further limits their oral bioavailability.[62,63] Moreover, a precise understanding of their mechanism of action is necessary for clinical application, yet frequently lacking. Mitigating these challenges in order to make
Introduction
17
pharmaceutical use possible in the future requires fine-tuning of these antibiotic candidates via chemical modification and labeling.
The synthetic modification of these complex natural products has proven to be a major challenge. Bottom-up reconstruction of RiPPs via solid-phase peptide synthesis to make synthetic analogs is challenging, since it is difficult to precisely mimic the remarkable post-translational machinery that is behind the maturation of these peptides in nature.[100–102] Solution-phase total syntheses of RiPPs have also been described, however they are greatly demanding in terms of stereoselectivity, are labor intensive, require many steps and have taken up to several years to complete.[103–106]
The chemical editing of naturally produced RiPPs to make semi-synthetic derivatives is becoming an increasingly popular approach for the modification of these peptides. By obtaining the finished, pure natural products first and then chemically modifying them in a late stage the number of steps required is greatly reduced and most successful modifications can be performed in one or two steps (vide infra). Moreover, the production and isolation of common RiPPs such as thiostrepton and nisin has advanced to an extent that they are commercially available in high purity for a reasonable price. Over the years, this late-stage chemical editing of RiPPs has led to biologically active, semi-synthetic analogs with improved pharmacological properties.[107] Nevertheless, achieving chemo- and site-selective modification of these structurally diverse and complex natural products remains a daunting challenge. 1.4.1 Selective modification of dehydroalanine residues
The dehydroamino acids that occur naturally in RiPPs have proven to be excellent chemical handles for the selective modification of these peptides. Dehydroalanine residues (Dha) are α,β-unsaturated amino acids, which gives them a distinct reactivity as carbon electrophiles, whereas most reactive residues in peptides and proteins are nucleophilic.[108] This orthogonal reactivity, combined with their natural occurrence in RiPPs, has made Dha an attractive target for the selective chemical editing of these peptide antibiotics.[109–111] Additionally, Davis et al. developed a method to chemically incorporate Dha onto peptides and proteins via bis-alkylation and elimination of cysteine residues (Scheme 1.10).[112] These studies enabled the use of Dha as a chemical handle also for the selective modification of peptides and proteins that do not naturally possess this residue, greatly broadening the scope of this approach.[109–111]
Chapter 1
18
Scheme 1.10: Chemical introduction of Dha onto peptides and proteins via
bis-alkylation of Cys with dibromohexanediamide followed by base-catalyzed elimination.
The electrophilic nature of Dha residues has enabled the selective derivatization of peptides and proteins via a wide variety of reaction classes.[109–111] First of all, they can be used as Michael acceptors in Michael-type additions (Scheme 1.11). Thiols have been shown to react in a thio-Michael addition similar to the addition of Cys thiols to Dha that occurs naturally in the biosynthesis of lanthipeptides.[81,113] This method has been used effectively in the preparation of semi-synthetic derivatives of thiostrepton[81] and in the mimicking of post-translational modifications in histones.[113] In a more recent study, Bernardes employed the aza-Michael addition of amines to Dha as a method to chemoselectively install amine bonds on proteins.[114] Nucleophilic addition of amines to Dha was found to be compatible with disulfide bonds present in native proteins and results in the formation of stable, cropped lysine isosteres.[114] Michael additions on Dha are straightforward, effective and chemoselective, yet often lack regioselectivity, especially with highly reactive thiol substrates.
Scheme 1.11: Derivatization of Dha residues in peptides and proteins via Michael
additions.[81,113,114]
Dha can also act as a somophile in radical additions, where it reacts as an acceptor for nucleophilic radicals. This reactivity was initially described by Davis et al. and Park et al. using carbon-centered radicals that were generated from alkyl iodides via reduction using NaBH4 or Cu(II)/Zn, respectively (Scheme 1.12).[115,116] In these studies, it was demonstrated that radical additions on Dha are a powerful tool for the chemical mutagenesis of proteins.[115,116]
N H O RNH2 RSH N H O NH R N H O S R Bernardes Arndt/Davis Thia-Michael Aza-Michael N H O SH Cys Br H2N O Br NH2 O N H O S NH2 O O H2N bis-alkylation H B: elimination NH O Dha
Introduction
19
Scheme 1.12: Radical addition of alkyl iodides via reduction with NaBH4[115] or a combination of Cu(II) and zinc.[116]
More recently, Roelfes et al. developed a method for the activation of trifluoroborate salts using an iridium photocatalyst to generate radicals that can react with Dha residues (Scheme 1.13).[117] Photoredox catalysis allows for a more steady generation of a limited amount of radicals over time. This lower concentration of radicals hampers the formation of side products due to termination reactions, which makes it a promising strategy for achieving chemoselective modifications. However, the photocatalyst used in this study is not soluble in water and requires the use of co-solvents.[117] To expand the scope of the approach, Roelfes et al. later synthesized a water-soluble iridium photocatalyst and used this to successfully perform radical additions on Dha in RiPPs and proteins using zinc sulfinates as precursors under purely aqueous conditions (Scheme 1.13).[118] Around the same time, Davis et al. reported the functionalization of Dha in proteins using ruthenium-based photoredox catalysts and boronate catechol esters and pyridyl-sulfone-fluorides as radical precursors (Scheme 1.13).[119] The water-soluble Ru(II) catalysts allowed for the installation of a variety of reactive functional groups into diverse protein scaffolds.[119] Radical additions on Dha have proven to be a powerful method for selective C-C bond formations on peptides and proteins, and they have been employed successfully in the selective editing of RiPPs. However, since the radicals are generated in situ, this approach requires strict exclusion of oxygen via degassing or the use of a glovebox.
Scheme 1.13: Overview of radical-based approaches for the modification of Dha via
photoredox catalysis.[117–119] N H O R I N H O R Davis/Park NaBH4or Cu(II)/Zn(0) N H O Blue LED R Ir-cat. SO O Zn 2 Zinc sulfinate or R BF3K trifluoroborate salt N H O R Roelfes Ru-cat. Blue LED B O O R HO or Boronate catechol ester
N S R F F O O pyridyl-sulfone-fluoride N H O RX X X=H or X=F Davis
Chapter 1
20
Transition metal-catalyzed cross-coupling reactions have also been performed on Dha, where it acts as the electron-poor reaction partner (Schemes 1.14). Rhodium catalysis has been shown by Miller et al. to lead to 1,4-addition of aryl boronic acids to Dha residues in thiostrepton (Scheme 1.14A).[120] This site- and diastereoselective editing of thiostrepton enabled the isolation of a variety of new derivatives of thiostrepton, while high antimicrobial activities were maintained.[120] Roelfes and co-workers showed that palladium catalyzed cross-couplings with boronic acids under completely aqueous conditions gives a mixture of the 1,4-addition- and Heck products on Dha-containing peptides and proteins (Scheme 1.14B).[121] The formation of Heck product was the first example of a Dha modification where the sp2 hybridization of the α-carbon was retained. Recently, Miller demonstrated the Co(III)-catalyzed amidation of Dha in thiostrepton using dioxazolone substrates, leading to variants of thiostrepton with increased aqueous solubility (Scheme 1.14A).[122] In this approach, the Dha double bond is also preserved with complete Z-selectivity.[122] Moreover, the reaction showed a high selectivity for the terminal Dha residue in the tail region of thiostrepton (Scheme 1.14A).[122] This site-selectivity is complementary to that of other existing methods, since in most cases modification of the more electron poor sub-terminal Dha residue in thiostrepton is predominant.
Scheme 1.14: A) Rhodium-catalyzed cross-coupling of arylboronic acids and
Co-catalyzed amidation of Dha residues in thiostrepton.[120,122] B) Palladium-catalyzed
cross-coupling of arylboronic acids with Dha in peptides and proteins.[121]
N H O N H O Roelfes Pd-cat. N H O
Conjugate addition product Heck product
R R B(OH)2 R Rh-cat. Co-cat. O N O O R Arylboronic acid Dioxazolone N H N O S N HN O S N H N HO O HN N S H N O S N OH NH NH O O H N O N H O S N NH H N O O NH2 O N OH OH O HO H H O thiostrepton S N NH H N O O NH2 O thiostrepton tail region S N NH H N O O NH2 O thiostrepton HN R O R (Z) amidation product
Conjugate addition product B(OH)2 R Arylboronic acid Miller Miller A) B)
Introduction
21
Finally, diazo reagents can be used in both cycloadditions and cyclopropanations on Dha in RiPPs (Scheme 1.15). Raines demonstrated that diazo reagents can undergo 1,3-dipolar cycloadditions on Dha substrates without any additives.[123] On small α,β-unsaturated alkene substrates the reaction was shown to be completely chemoselective for the diazo reagents, even in the presence of azides.[123] The method was then used to modify the lantibiotic nisin, where the same high chemoselectivity was observed.[123] Bowers and co-workers showed that cytochrome P450 enzymes can catalyze cyclopropanations of Dha in thiopeptides, also using diazo reagents.[124] The latter is the first example of a non-natural modification of Dha in RiPPs using biocatalysis. The enzymes used are involved in post-translational modifications occurring during the biosynthesis of thiopeptides in nature, and this study showed that they can also be repurposed to install abiological functionalities into a range of complex natural products that are similar to the natural substrates of these enzymes.[124]
Scheme 1.15: Overview of chemical reactions that can be used for the modification of
dehydroalanine residues in peptides and proteins.[81,113-124]
N H O X R X = SH, NH2 N H O X R Nucleophilic Addition
Arndt, Davis, Bernardes
R
Radical Addition
N H O R
Davis, Bernardes, Roelfes Pd, Rh Cross-coupling N H O Miller, Roelfes Cyclopropanation P450 N H O R O Bowers N H O N HN O R 1,3-dipolar Cycloaddition Raines N2 R O Amidation Co(III) O NO O R N H O HN R O Miller
Chapter 1
22
In summary, the methods described above show that Dha is a highly versatile chemical handle for selective modification, not only for RiPPs but also for peptides and proteins in general. The current state-of-the-art shows significant progress towards the development of a wide variety of chemical reactions that can be performed on Dha residues (Scheme 1.15). However, every individual method has its limitations and also several challenges that are central to Dha modifications remain, which are currently hampering the advancement of the field towards the semi-synthesis of clinically viable antimicrobial peptides. The research presented in this thesis aims to address several of these challenges, which are highlighted in the next section.
1.5 Aim and outline of this thesis
The objective of this thesis was to develop new methods for the late-stage chemical editing of RiPPs via their dehydroalanine residues. The goal was to also apply these methods in the bioconjugation of these peptides and to synthesize analogs with improved pharmacological properties. In addition to the modification of RiPPs, the characterization of newly discovered antimicrobial NRPs was also studied. The studies presented here aim to address the following challenges currently in the field of Dha modification:
• Catalytic modification of Dha residues often requires the use of heavy metal catalysts, which can be difficult to remove during the purification of the modified peptides. This in turn can pose a problem when evaluating the biological activity of these antibiotics and their clinical application in the future, due to the inherent toxicity of these metals. Therefore, the aim was to develop methods that are transition metal free or only require less toxic, cheap and environmentally benign 1st-row transition metal catalysts.
• The antimicrobial activity of RiPPs is naturally very high, yet improving their generally poor pharmacological properties is critical for enabling clinical application in the future. Surprisingly, for most existing modifications of RiPPs the effect on their antimicrobial activity is evaluated, while the effect on their pharmacological properties is often overlooked. The studies presented here were aimed at improving and evaluating these properties and demonstrating efficient labeling, while also keeping the antimicrobial activity as high as possible.
Introduction
23
• Existing methods allow for the construction of C-C, C-N and C-S bonds on Dha. In this thesis, the aim was to expand this ´toolbox´ of Dha modifications by developing methods for the direct formation of non-natural bonds (e.g. C-B, C-Si) on Dha residues. This should then enable the exploration of the vast chemical space of such abiological functionalities in RiPPs and their effect on the pharmacological properties, activity and labeling of these peptides.
• Although Dha modification is a powerful tool, research has focused primarily on finding new methods, and currently no major progress has been made yet towards the actual application of these methods in bioconjugation. The research presented here was aimed at also applying the developed methods in the semi-synthesis of useful conjugates, or hybrids, for the selective targeting of RiPPs.
In chapter 2 the transition metal free Diels-Alder reaction of Dha in RiPPs with cyclopentadiene is discussed. Several RiPPs were modified efficiently under microwave conditions and the site-selectivity was established using 2D NMR spectroscopy. The norbornylated peptides were subjected to the Inverse Electron Demand Diels-Alder (IEDDA) reaction with a variety of functionalized tetrazines. Chapter 3 describes how the Diels-Alder – IEDDA conjugation method established in chapter 2 is employed in the synthesis of thiostrepton-siderophore hybrids for the targeting of gram-negative bacteria. The synthesis of several tetrazine-functionalized siderophores is described.
Chapter 4 describes the Cu(II)-catalyzed β-borylation of Dha in RiPPs. The reaction is straightforward, fast and highly chemo and site-selective. The borylated peptides were used as intermediates for chemical mutagenesis and in the reversible labeling via boronate ester formation with diols and triols. Also, the effects of borylation on the aqueous solubility and antimicrobial activity of thiostrepton and nosiheptide are discussed.
In chapter 5 the β-silylation of Dha residues in peptides and proteins using the copper catalysis discussed in chapter 4 is described. The reaction shows a broad scope, since not only RiPPs, but also a Dha-functionalized protein was silylated efficiently.
Chapter 6 describes the characterization of newly isolated NRPs via 2D NMR spectroscopy. Three mutants of brevibacillin were investigated, including two mutants
Chapter 1
24
and one succinylated variant. One relacidine, a new class of lipopeptides with high antimicrobial activity against Gram-negative bacteria, was also characterized using 2D NMR techniques.
In chapter 7 the conclusions and perspectives of this research are presented. References
[1] IACG, No Time to Wait: Securing the Future from Drug-Resistant Infections, 2019. [2] A. H. Holmes, L. S. P. Moore, A. Sundsfjord, M. Steinbakk, S. Regmi, A. Karkey, P. J.
Guerin, L. J. V. Piddock, Lancet 2016, 387, 176–187. [3] C. L. Ventola, Pharm. Ther. 2015, 40, 277–283.
[4] R. Sugden, R. Kelly, S. Davies, Nat. Microbiol. 2016, 1, 16187.
[5] N. Mobarki, B. Almerabi, A. Hattan, Int. J. Med. Dev. Ctries. 2019, 3, 561–564. [6] C. Ghosh, P. Sarkar, R. Issa, J. Haldar, Trends Microbiol. 2019, 27, 323–338.
[7] M. Mahlapuu, J. Håkansson, L. Ringstad, C. Björn, Front. Cell. Infect. Microbiol. 2016, 6, 1–12.
[8] R. Spohn, L. Daruka, V. Lázár, A. Martins, F. Vidovics, G. Grézal, O. Méhi, B. Kintses, M. Számel, P. K. Jangir, B. Csörgő, Á. Györkei, Z. Bódi, A. Faragó, L. Bodai, I. Földesi, D. Kata, G. Maróti, B. Pap, R. Wirth, B. Papp, C. Pál, Nat. Commun. 2019, 10, 1–13.
[9] D. J. Newman, G. M. Cragg, J. Nat. Prod. 2016, 79, 629–661. [10] M. Zasloff, Nature 2002, 415, 389–395.
[11] R. E. W. Hancock, H. G. Sahl, Nat. Biotechnol. 2006, 24, 1551–1557.
[12] M. Hassan, M. Kjos, I. F. Nes, D. B. Diep, F. Lotfipour, J. Appl. Microbiol. 2012, 113, 723– 736.
[13] R. E. W. Hancock, D. S. Chapple, Antimicrob. Agents Chemother. 1999, 43, 1317–1323. [14] G. H. Hur, C. R. Vickery, M. D. Burkart, Nat. Prod. Rep. 2012, 29, 1074–1098.
[15] P. G. Arnison, M. J. Bibb, G. Bierbaum, A. A. Bowers, T. S. Bugni, G. Bulaj, J. A. Camarero, D. J. Campopiano, G. L. Challis, J. Clardy, P. D. Cotter, D. J. Craik, M. Dawson, E. Dittmann, S. Donadio, P. C. Dorrestein, K. D. Entian, M. A. Fischbach, J. S. Garavelli, U. Göransson, C. W. Gruber, D. H. Haft, T. K. Hemscheidt, C. Hertweck, C. Hill, A. R. Horswill, M. Jaspars, W. L. Kelly, J. P. Klinman, O. P. Kuipers, A. J. Link, W. Liu, M. A. Marahiel, D. A. Mitchell, G. N. Moll, B. S. Moore, R. Müller, S. K. Nair, I. F. Nes, G. E. Norris, B. M. Olivera, H. Onaka, M. L. Patchett, J. Piel, M. J. T. Reaney, S. Rebuffat, R. P. Ross, H. G. Sahl, E. W. Schmidt, M. E. Selsted, K. Severinov, B. Shen, K. Sivonen, L. Smith, T. Stein, R. D. Süssmuth, J. R. Tagg, G. L. Tang, A. W. Truman, J. C. Vederas, C. T. Walsh, J. D. Walton, S. C. Wenzel, J. M. Willey, W. A. Van Der Donk, Nat. Prod. Rep. 2013, 30, 108–160.
[16] J. P. S. Powers, R. E. W. Hancock, Peptides 2003, 24, 1681–1691.
Introduction
25 [18] D. I. Andersson, D. Hughes, J. Z. Kubicek-Sutherland, Drug Resist. Updat. 2016, 26, 43–
57.
[19] Y. Lai, R. L. Gallo, Trends Immunol. 2009, 30, 131–141.
[20] C. D. Fjell, J. A. Hiss, R. E. W. Hancock, G. Schneider, Nat. Rev. Drug Discov. 2012, 11, 37– 51.
[21] A. K. Marr, W. J. Gooderham, R. E. Hancock, Curr. Opin. Pharmacol. 2006, 6, 468–472. [22] K. A. Brogden, Nat. Rev. Microbiol. 2005, 3, 238–250.
[23] H. B. Koo, J. Seo, Pept. Sci. 2019, 111, e24122.
[24] M. Rai, R. Pandit, S. Gaikwad, G. Kövics, J. Food Sci. Technol. 2016, 53, 3381–3394. [25] R. Finking, M. A. Marahiel, Annu. Rev. Microbiol. 2004, 58, 453–488.
[26] B. R. Miller, A. M. Gulick, Methods Mol. Biol. 2016, 1401, 3–29. [27] A. M. Gulick, ACS Chem. Biol. 2009, 4, 811–827.
[28] A. C. Mercer, M. D. Burkart, Nat. Prod. Rep. 2007, 24, 750–773.
[29] V. Bergendahl, U. Linne, M. A. Marahiel, Eur. J. Biochem. 2002, 269, 620–629.
[30] T. Stachelhaus, H. D. Mootz, V. Bergendahl, M. A. Marahiel, J. Biol. Chem. 1998, 273, 22773–22781.
[31] C. T. Walsh, H. Chen, T. A. Keating, B. K. Hubbard, H. C. Losey, L. Luo, C. G. Marshall, D. A. Miller, H. M. Patel, Curr. Opin. Chem. Biol. 2001, 5, 525–534.
[32] R. E. W. Hancock, R. Lehrer, Trends Biotechnol. 1998, 16, 82–88.
[33] A. T. Y. Yeung, S. L. Gellatly, R. E. W. Hancock, Cell. Mol. Life Sci. 2011, 68, 2161–2176. [34] T. Barsby, M. T. Kelly, S. M. Gagné, R. J. Andersen, Org. Lett. 2001, 3, 437–440.
[35] T. Barsby, K. Warabi, D. Sørensen, W. T. Zimmerman, M. T. Kelly, R. J. Andersen, J. Org.
Chem. 2006, 71, 6031–6037.
[36] Y. X. Li, Z. Zhong, W. P. Zhang, P. Y. Qian, Nat. Commun. 2018, 9, 2–10. [37] X. Yang, A. E. Yousef, World J. Microbiol. Biotechnol. 2018, 34, 1–10.
[38] X. Yang, E. Huang, C. Yuan, L. Zhang, A. E. Yousef, Appl. Environ. Microbiol. 2016, 82, 2763–2772.
[39] T. J. Oman, W. A. Van Der Donk, Nat. Chem. Biol. 2010, 6, 9–18.
[40] M. C. Bagley, J. W. Dale, E. A. Merritt, X. Xiong, Chem. Rev. 2005, 105, 685–714. [41] R. A. Hughes, C. J. Moody, Angew. Chem. Int. Ed. 2007, 46, 7930–7954. [42] X. Just-Baringo, F. Albericio, M. Álvarez, Mar. Drugs 2014, 12, 317–351.
[43] L. C. W. Brown, M. G. Acker, J. Clardy, C. T. Walsh, M. A. Fischbach, PNAS 2009, 106, 2549–2553.
[44] R. Liao, L. Duan, C. Lei, H. Pan, Y. Ding, Q. Zhang, D. Chen, B. Shen, Y. Yu, W. Liu, Chem.
Biol. 2009, 16, 141–147.
[45] R. P. Morris, J. A. Leeds, H. U. Naegeli, L. Oberer, K. Memmert, E. Weber, M. J. LaMarche, C. N. Parker, N. Burrer, S. Esterow, A. E. Hein, E. K. Schmitt, P. Krastel, J. Am. Chem. Soc. 2009, 131, 5946–5955.
Chapter 1
26
[47] G. A. Hudson, Z. Zhang, J. I. Tietz, D. A. Mitchell, W. A. Van Der Donk, J. Am. Chem. Soc. 2015, 137, 16012–16015.
[48] Q. Zheng, H. Fang, W. Liu, Org. Biomol. Chem. 2017, 15, 3376–3390.
[49] E. W. Schmidt, J. T. Nelson, D. A. Rasko, S. Sudek, J. A. Eisen, M. G. Haygood, J. Ravel,
PNAS 2005, 102, 7315–7320.
[50] J. A. Mcintosh, M. S. Donia, E. W. Schmidt, J. Am. Chem. Soc. 2010, 132, 4089–4091. [51] K. L. Dunbar, J. O. Melby, D. A. Mitchell, Nat. Chem. Biol. 2012, 8, 569–575.
[52] K. L. Dunbar, J. R. Chekan, C. L. Cox, B. J. Burkhart, S. K. Nair, D. A. Mitchell, Nat. Chem.
Biol. 2014, 10, 823–829.
[53] D. P. Cogan, G. A. Hudson, Z. Zhang, T. V. Pogorelov, W. A. Van Der Donk, D. A. Mitchell, S. K. Nair, PNAS 2017, 114, 12928–12933.
[54] A. A. Bowers, C. T. Walsh, M. G. Acker, J. Am. Chem. Soc. 2010, 132, 12182–12184. [55] A. A. Bowers, M. G. Acker, A. Koglin, C. T. Walsh, J. Am. Chem. Soc. 2010, 132, 7519–
7527.
[56] W. J. Wever, J. W. Bogart, J. A. Baccile, A. N. Chan, F. C. Schroeder, A. A. Bowers, J. Am.
Chem. Soc. 2015, 137, 3494–3497.
[57] Y. Yu, L. Duan, Q. Zhang, R. Liao, Y. Ding, H. Pan, E. Wendt-Pienkowski, G. Tang, B. Shen, W. Liu, ACS Chem. Biol. 2009, 4, 855–864.
[58] O. D. Hensens, G. Albers-Schönberg, Tetrahedron Lett. 1978, 39, 3649–3652.
[59] M. S. Puar, T. M. Chan, V. Hegde, M. Patel, P. Bartner, K. J. Ng, B. N. Pramanik, R. D. MacFarlane, J. Antibiot. 1998, 51, 221–224.
[60] N. Mizuhara, M. Kuroda, A. Ogita, T. Tanaka, Y. Usuki, K. I. Fujita, Bioorg. Med. Chem. 2011, 19, 5300–5310.
[61] N. S. Hegde, D. A. Sanders, R. Rodriguez, S. Balasubramanian, Nat. Chem. 2011, 3, 725– 731.
[62] C. L. Myers, P. C. Hang, G. Ng, J. Yuen, J. F. Honek, Bioorg. Med. Chem. 2010, 18, 4231– 4237.
[63] X. Just-Baringo, F. Albericio, M. Álvarez, Angew. Chem. Int. Ed. 2014, 53, 6602–6616. [64] R. Donovick, J. F. Pagano, H. A. Stout, M. J. Weinstein, Antibiot. Annu. 1955, 3, 554–559. [65] B. Anderson, D. C. Hodgkin, M. A. Viswamitra, Nature 1970, 225, 233–235.
[66] W. H. Trejo, L. D. Dean, J. Pluscec, E. Meyers, W. E. Brown, J. Antibiot. 1977, 30, 639– 643.
[67] J. Thompson, E. Cundliffe, Biochimie 1991, 73, 1131–1135.
[68] G. Lentzen, R. Klinck, N. Matassova, F. Aboul-ela, A. I. H. Murchie, Chem. Biol. 2003, 10, 769–778.
[69] H. R. A. Jonker, S. Ilin, S. K. Grimm, J. Wöhnert, H. Schwalbe, Nucleic Acids Res. 2007, 35, 441–454.
[70] J. M. Harms, D. N. Wilson, F. Schluenzen, S. R. Connell, T. Stachelhaus, Z. Zaborowska, C. M. T. Spahn, P. Fucini, Mol. Cell 2008, 30, 26–38.
Introduction
27 [71] J. M. M. Kwok, S. S. Myatt, C. M. Marson, R. C. Coombes, D. Constantinidou, E. W. F.
Lam, Mol. Cancer Ther. 2008, 7, 2022–2032.
[72] C. Pascard, A. Ducruix, J. Lunel, T. Prangé, J. Am. Chem. Soc. 1977, 99, 6418–6423. [73] K. A. Acott, T. P. Labuza, A. Minra, J. W. Drake, S. Y. Aguyagi, F. Furkawa, H. Zawadska, O.
S. Bhanet, D. Dufuy, K. H. Erismann, M. E. Grant, R. H. Hall, S. Biolo-, J. Mccann, E. Yamasaki, V. J. Veron, A. P. De Groot, F. Strong, E. Collins, F. Benazet, M. Cartier, J. Florent, C. Godard, G. Jung, J. Lunel, D. Mancy, C. Pascal, J. Renaut, P. Tarridec, J. Theilleux, R. Tissier, M. Dubost, L. Ninet, Experientia 1980, 36, 414–416.
[74] T. Tanaka, T. Endō, A. Shimazu, R. Yoshida, Y. Suzuki, N. Ōtake, H. Yonehara, J. Antibiot. 1970, 23, 231–237.
[75] T. Endō, H. Yonehara, J. Antibiot. 1978, 31, 623–625.
[76] U. Mocek, A. R. Knaggs, R. Tsuchiya, T. Nguyen, J. M. Beale, H. G. Floss, J. Am. Chem. Soc. 1993, 115, 7557–7568.
[77] Y. Yu, H. Guo, Q. Zhang, L. Duan, Y. Ding, R. Liao, C. Lei, B. Shen, W. Liu, J. Am. Chem. Soc. 2010, 132, 16324–16326.
[78] H. Guo, J. Wang, Y. Li, Y. Yu, Q. Zheng, J. Wu, W. Liu, Chem. Sci. 2014, 5, 240–246. [79] E. Cundliffe, J. Thompson, J. Gen. Microbiol. 1981, 126, 185–192.
[80] N. M. Haste, W. Thienphrapa, D. N. Tran, S. Loesgen, P. Sun, S. J. Nam, P. R. Jensen, W. Fenical, G. Sakoulas, V. Nizet, M. E. Hensler, J. Antibiot. 2012, 65, 593–598.
[81] S. Schoof, S. Baumann, B. Ellinger, H. D. Arndt, ChemBioChem 2009, 10, 242–245. [82] F. Benazet, J. R. Cartier, Poult. Sci. 1980, 59, 1405–1415.
[83] M. A. Ortega, W. A. Van Der Donk, Cell Chem. Biol. 2016, 23, 31–44.
[84] N. Schnell, K.-D. Entian, U. Schneider, F. Götz, H. Zähner, R. Kellner, G. Jung, Nature 1988, 333, 276–278.
[85] P. J. Knerr, W. A. van der Donk, Annu. Rev. Biochem. 2012, 81, 479–505.
[86] L. M. Repka, J. R. Chekan, S. K. Nair, W. A. Van Der Donk, Chem. Rev. 2017, 117, 5457– 5520.
[87] L. D. Kluskens, A. Kuipers, R. Rink, E. De Boef, S. Fekken, A. J. M. Driessen, O. P. Kuipers, G. N. Moll, Biochemistry 2005, 44, 12827–12834.
[88] B. Li, J. P. J. Yu, J. S. Brunzelle, G. N. Moll, W. A. Van Der Donk, S. K. Nair, Science 2006,
311, 1464–1467.
[89] C. Chatterjee, L. M. Miller, Y. L. Leung, L. Xie, M. Yi, N. L. Kelleher, W. A. Van Der Donk, J.
Am. Chem. Soc. 2005, 127, 15332–15333.
[90] W. M. Müller, T. Schmiederer, P. Ensle, R. D. Süssmuth, Angew. Chem. Int. Ed. 2010, 49, 2436–2440.
[91] Y. Goto, B. Li, J. Claesen, Y. Shi, M. J. Bibb, W. A. van der Donk, PLoS Biol. 2010, 8, 4–13. [92] N. I. Martin, E. Breukink, Future Microbiol. 2007, 2, 513–525.
[93] L. A. Rogers, J. Bacteriol. 1928, 16, 321–325.
Chapter 1
28
[95] M. A. Ortega, Y. Hao, Q. Zhang, M. C. Walker, W. A. Van Der Donk, S. K. Nair, Nature 2015, 517, 509–512.
[96] J. R. Van der Meer, J. Polman, M. M. Beerthuyzen, R. J. Siezen, O. P. Kuipers, W. M. De Vos, J. Bacteriol. 1993, 175, 2578–2588.
[97] E. Breukink, B. de Kruijff, Nat. Rev. Drug Discov. 2006, 5, 321–323. [98] G. Bierbaum, H.-G. Sahl, Curr. Pharm. Biotechnol. 2009, 10, 2–18.
[99] J. Lubelski, R. Rink, R. Khusainov, G. N. Moll, O. P. Kuipers, Cell. Mol. Life Sci. 2008, 65, 455–476.
[100] J. C. Slootweg, J. Kemmink, R. M. J. Liskamp, D. T. S. Rijkers, Org. Biomol. Chem. 2013,
11, 7486–7496.
[101] J. C. Slootweg, N. Peters, H. C. Quarles Van Ufford, E. Breukink, R. M. J. Liskamp, D. T. S. Rijkers, Bioorg. Med. Chem. 2014, 22, 5345–5353.
[102] W. J. Wever, J. W. Bogart, A. A. Bowers, J. Am. Chem. Soc. 2016, 138, 13461–13464. [103] K. Fukase, M. Kitazawa, A. Sano, K. Shimbo, H. Fujita, S. Horimoto, T. Wakamiya, T. Shiba,
Tetrahedron Lett. 1988, 29, 795–798.
[104] K. C. Nicolaou, B. S. Safina, M. Zak, S. H. Lee, M. Nevalainen, M. Bella, A. A. Estrada, C. Funke, F. J. Zécri, S. Bulat, J. Am. Chem. Soc. 2005, 127, 11159–11175.
[105] K. C. Nicolaou, M. Zak, B. S. Safina, A. A. Estrada, S. H. Lee, M. Nevalainen, J. Am. Chem.
Soc. 2005, 127, 11176–11183.
[106] K. P. Wojtas, M. Riedrich, J. Y. Lu, P. Winter, T. Winkler, S. Walter, H. D. Arndt, Angew.
Chem. Int. Ed. 2016, 55, 9772–9776.
[107] T. Koopmans, T. M. Wood, P. ’T Hart, L. H. J. Kleijn, A. P. A. Hendrickx, R. J. L. Willems, E. Breukink, N. I. Martin, J. Am. Chem. Soc. 2015, 137, 9382–9389.
[108] D. Siodłak, Amino Acids 2015, 47, 1–17.
[109] J. Dadová, S. R. Galan, B. G. Davis, Curr. Opin. Chem. Biol. 2018, 46, 71–81. [110] J. W. Bogart, A. A. Bowers, Org. Biomol. Chem. 2019, 17, 3653–3669.
[111] E. A. Hoyt, P. M. S. D. Cal, B. L. Oliveira, G. J. L. Bernardes, Nat. Rev. Chem. 2019, 3, 147– 171.
[112] J. M. Chalker, S. B. Gunnoo, O. Boutureira, S. C. Gerstberger, M. Fernández-González, G. J. L. Bernardes, L. Griffin, H. Hailu, C. J. Schofield, B. G. Davis, Chem. Sci. 2011, 2, 1666– 1676.
[113] J. M. Chalker, L. Lercher, N. R. Rose, C. J. Schofield, B. G. Davis, Angew. Chem. Int. Ed. 2012, 51, 1835–1839.
[114] A. M. Freedy, M. J. Matos, O. Boutureira, F. Corzana, A. Guerreiro, P. Akkapeddi, V. J. Somovilla, T. Rodrigues, K. Nicholls, B. Xie, G. Jiménez-Osés, K. M. Brindle, A. A. Neves, G. J. L. Bernardes, J. Am. Chem. Soc. 2017, 139, 18365–18375.
[115] T. H. Wright, B. J. Bower, J. M. Chalker, G. J. L. Bernardes, R. Wiewiora, W. L. Ng, R. Raj, S. Faulkner, M. R. J. Vallée, A. Phanumartwiwath, O. D. Coleman, M. L. Thézénas, M. Khan, S. R. G. Galan, L. Lercher, M. W. Schombs, S. Gerstberger, M. E. Palm-Espling, A. J.
Introduction
29 Baldwin, B. M. Kessler, T. D. W. Claridge, S. Mohammed, B. G. Davis, Science 2016, 354, aag1465.
[116] A. Yang, S. Ha, J. Ahn, R. Kim, S. Kim, Y. Lee, J. Kim, D. Söll, H. Y. Lee, H. S. Park, Science 2016, 354, 623–626.
[117] A. D. De Bruijn, G. Roelfes, Chem. Eur. J. 2018, 24, 11314–11318.
[118] R. C. W. van Lier, A. D. de Bruijn, G. Roelfes, Chem. Eur. J. 2020, DOI 10.1002/chem.202002599.
[119] B. Josephson, C. Fehl, P. G. Isenegger, S. Nadal, T. H. Wright, A. W. J. Poh, B. J. Bower, A. M. Giltrap, L. Chen, C. Batchelor-McAuley, G. Roper, O. Arisa, J. B. I. Sap, A. Kawamura, A. J. Baldwin, S. Mohammed, R. G. Compton, V. Gouverneur, B. G. Davis, Nature 2020,
585, 530–537.
[120] H. M. Key, S. J. Miller, J. Am. Chem. Soc. 2017, 139, 15460–15466. [121] A. D. de Bruijn, G. Roelfes, Chem. Eur. J. 2018, 24, 12728–12733.
[122] R. J. Scamp, E. deRamon, E. K. Paulson, S. J. Miller, J. A. Ellman, Angew. Chem. Int. Ed. 2020, 59, 890–895.
[123] M. R. Aronoff, B. Gold, R. T. Raines, Org. Lett. 2016, 18, 1538–1541.
[124] J. G. Gober, S. V. Ghodge, J. W. Bogart, W. J. Wever, R. R. Watkins, E. M. Brustad, A. A. Bowers, ACS Chem. Biol. 2017, 12, 1726–1731.
Chapter 1
