Enzymatic Fluorination and Biotechnological Developments of the

Review pubs.acs.org/CR

Enzymatic Fluorination and Biotechnological Developments of the Fluorinase David O’Hagan*,† and Hai Deng*,‡ †

EaStChem School of Chemistry, University of St Andrews, North Haugh, St Andrews KY169ST, United Kingdom UK Marine Biodiscovery Centre, Department of Chemistry, Meston Walk, University of Aberdeen, Aberdeen AB24 3UE, United Kingdom

‡

Acknowledgments References

1. INTRODUCTION Despite a long and distinguished history focused on the isolation and structural elucidation of secondary metabolites from all classes of organisms, organic chemists have only securely identified five unique fluorine-containing metabolites1 from an estimated 130 0002 structurally characterized natural products (see Figure 1). This absence of fluorometabolites in our natural product inventory arises for several reasons. Fluoride ion has very low abundance (F− = 1.3 ppm) in the oceans relative to chloride (Cl− = 20 000 ppm) and bromide (Br− = 70 ppm),3 and thus its bioavailability is low. It also has the highest heat of hydration (∼120 kcal mol−1); therefore, to achieve nucleophilic catalysis from water, an enzyme has to evolve a desolvation strategy. Much of the biochemistry of the other halogens involves the oxidation of halide ions (X−) to halonium ions (X+) or halide radicals (X·),4 but the high electronegativity of fluorine mitigates against an oxidation approach. It is interesting that iodide, which is even less abundant than fluoride in surface water (F− = 1.3 ppm vs I− = 0.02 ppm),5 has given rise to ∼120 iodine-containing natural products.4b This is because iodide, unlike fluoride, is readily oxidized by haloperoxidases, and it finds an entry into enzymology through the relative ease of iodonium ion (I+) generation. The haloperoxidases do not oxidize F− because the oxidation potential of hydrogen peroxide (−1.8 eV) is above that of fluoride (F− = −2.87 eV) but lower than that of the other halogens (Cl− = −1.36 eV; Br− = −1.07 ev; I− = −0.54 ev).1b Thus, the physical properties of fluoride appear to have limited the evolution of fluorine biochemistry. However, nature has found a way, and a few rare fluorometabolites have been identified, as illustrated in Figure 1. The most ubiquitious metabolite is fluoroacetate 1, which was first identified from Dichapetalum cymosum in South Africa but has been subsequently found in a variety of plants across the globe, particularly in southern and tropical regions of Africa, Australia, and Brazil.6 Fluoroacetate is readily metabolized, via fluoroacetyl-CoA, to the potent toxin (2R,3R)-fluorocitrate 3,7 which has been identified as a cometabolite in some fluoroacetate-producing

CONTENTS 1. Introduction 1.1. Biosynthesis of Fluorometabolites of Streptomyces cattleya 2. Enzymatic C−F Bond Formation 2.1. C−F Bond Formation in the Chemical Rescue of Mutant Glycosyl Transferases 2.2. Bacterial Fluorinases of Secondary Metabolism 2.2.1. Fluorinase from Streptomyces cattleya 2.2.2. Recently Identified Bacterial Fluorinases 2.3. Chlorinase and Hydrolases Related to the Bacterial Fluorinases 3. Biotechnological Potential for Enzymatic Fluorination 3.1. Fluorometabolite Engineering 3.1.1. Engineering Fluorosalinosporamide Production in Salinospora tropica 3.1.2. Incorporation of Fluoroacetyl-CoA and Fluoromalonyl-CoA into Polyketide Frameworks 3.2. Fluorinase As a Catalyst for 18F−C Bond Formation in Positron Emission Tomography 3.2.1. [18F]-Fluoronucleosides 3.2.2. “Last-Step” Labeling of Peptides Using the Fluorinase. 3.2.3. [18F]-Fluoroacetate 3.2.4. [18F]-Fluororibose 3.3. [18F]-Labeling of Peptides with [18F]-FDR 4. Conclusions Author Information Corresponding Authors Notes Biographies © XXXX American Chemical Society

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Special Issue: 2015 Fluorine Chemistry Received: April 14, 2014

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dx.doi.org/10.1021/cr500209t | Chem. Rev. XXXX, XXX, XXX−XXX

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Figure 1. Known fluorinated natural products. The compounds in the box are almost certainly misidentified natural products.

Scheme 1. Intermediates and Enzymes on the Biosynthetic Pathway to Fluoroacetate 1 and 4-Fluorothreonine 2 in Streptomyces cattleya

plants.6 This transformation is responsible for the high toxicity of fluoroacetate 1, as (2R,3R)-fluorocitrate 3 is a potent inhibitor of aconitase, a key enzyme of the citric acid cycle, central to cellular respiration.8 To date the biochemical basis of plant fluoroacetate biosynthesis is unknown. Other metabolites, such as the ω-fluorofatty acids from the West African plant Dichapetalum toxicarium, presumably also originate from fluoroacetate metabolism, where fluoroacetate 1 (or fluoroacetyl-CoA) serves as a starter unit in fatty acid biosynthesis in this organism.9,10 Analysis of the seed lipids of D. toxicarium has revealed that the major lipid is ω-fluorooleic acid 4 and that there is a large diversity of minor ω-fluorinated lipids that appear to be metabolites of the major lipid.11,12 Nucleocidin 5

is an antibiotic produced by the bacterium Streptomyces calvus. It was recognized to contain fluorine in 1969;13 however, subsequently it has proven difficult to produce this metabolite in culture, particularly from the available strains of S. calvus in public culture collections.14 This has frustrated the necessary biosynthesis studies required to understand the origin of the antibiotic, insights which would be revealing because a casual analysis of the structure suggests that this organism possesses a unique process for C−F bond formation, distinct from that found in those that can elaborate fluoroacetate. The Actinomycete soil bacterium Streptomyces cattleya can secrete fluoroacetate 1 and 4-fluorothreonine 2 as part of its metabolic profile.15 4-Fluorothreonine 2 is an antibiotic and presumably B


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Scheme 2. Enzymatic C−F Bond Formation Occurs in Mutant Glycosyl Transferase Enzymes25

fluorothreonine 2 is illustrated in Scheme 1. The “fluorinase”, which catalyzes the first reaction on the biosynthetic pathway, was isolated in 2002, and it remained a unique enzyme for over a decade until recently23,24 four additional fluorinases were reported by genome mining in four different bacterial strains. These isolates are discussed in more detail in section 2.2.2. Given the increasing number of bacterial genomes that are now available, the occurrence of fluorinase genes remains rather few. They appear to have a limited distribution, although we can reasonably expect additional f lA genes to appear as more bacterial genomes are sequenced and the data are deposited in public repositories.

this confers a selective advantage to the organism. The origin of these metabolites from S. cattleya will be discussed in more detail later. There are several probable misidentifications of fluorometabolites in the literature. Fluoroacetone 6 was claimed from a plant extract in the early 1970s, isolated as a dinitrophenylhydrazone derivative (DNP); however, the DNP derivative of fluoroacetone is unstable to fluoride ion elimination,16,17 so this is probably an incorrect isolation or at least the isolation requires revalidation. A range of 5-fluorouracils including 7 were isolated from a sponge (Phakellia f usca Schmidt) from the South China Sea in 2003;18 however, given the direct relationship of these isolates to the well-known pharmaceutical, and the less than obvious enzymatic strategy for fluorine incorporation into such a heterocycle, it is very probable that the sponge has accumulated these compounds from an industrial effluent. Most recently the fluoroarylpropionic acid 8 was reported as an isolate from Streptomyces sp. TC1.19 This observation is intriguing particularly because an enzymatic strategy for aryl fluorination would be an exciting discovery; however, the analytical characterization reported in the isolation paper19 is inconsistent with the claimed product, and it is unlikely that the isolated compound contains a fluorine atom.

2. ENZYMATIC C−F BOND FORMATION 2.1. C−F Bond Formation in the Chemical Rescue of Mutant Glycosyl Transferases

The first examples of enzymatic fluorination (C−F bond formation) reactions were demonstrated by Zechel, Withers, and co-workers25 when exploring mutagenesis and chemical rescue of glycosyl transferase enzymes (Scheme 2). For example, they demonstrated that, when the catalytically important glutamate-358 in a β-glucosidase from an Agrobacterium sp. or the glutamate-429 residue in a β-mannosidase from Cellulomonas f imi was mutated to serine or alanine, respectively, then the enzymes became dysfunctional. However, when fluoride (or azide) anion was added to the assay at relatively high concentrations (1−2 M), the enzymes recovered their glycosyl transferase activities. This was shown by 19FNMR to be due to the intermediacy of glycosyl fluoride 14 in the case of the Agrobacterium glycosyl transferase (Scheme 2a) or a β-D-mannosyl fluoride 15 in the case of the β-mannosidase (Scheme 2b), which presumably arose by anion trapping of intermediate oxocarbenium ions, which would have been otherwise stabilized by the glutamate carboxylates in the wildtype enzymes. Thus, the mutant enzymes accommodated C−F bond formation from fluoride ion in solution. The resultant glycosyl fluorides 14 and 15 are sufficiently reactive to act as electrophiles for nucleophilic displacement by incoming sugars,

1.1. Biosynthesis of Fluorometabolites of Streptomyces cattleya

S. cattleya has served as the most tractable system available for evaluating fluorometabolite biosynthesis and exploring fluorination enzymology.20 The organism was reported in 1986 to produce fluoroacetate 1, as well as the amino acid 4fluorothreonine 2.15 4-Fluorothreonine 2 has antibiotic activity, although its mode of action is unknown. The biosynthesis and enzymology for the assembly of these metabolites has been studied. The first enzyme committed to fluorometabolite production catalyzes a reaction between S-adenosyl-L-methionine (SAM) 9 and fluoride ion to generate 5′-fluorodeoxyadenosine (5′-FDA) 10 and L-methionine.21,22 The biosynthetic pathway from 5′-FDA 10 to fluoroacetate 1 and 4C


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Figure 2. Two representations of the fluoride ion/SAM 9 prereaction complex at the active site of the fluorinase immediately prior to SN2 substitution. (a) Fluoride ion is modeled into the SAM−fluorinase cocomplex, which was solved by X-ray crystallography. (b) Drawing of the same cocomplex shown in (a), with selected hydrogen bonds to the ribose shown.28

recently, an experimental study by Lohman et al.32 explored the second-order rate constants of fluoride (and other halide ions) substitution on sulfonium ylids in water at elevated temperatures (65−200 °C). The study explored the reaction kinetics of Me3S+:BF4− and KF heated in a sealed system to generate fluoromethane, a reaction modeling that of the fluorinase. Extrapolation of the rates of these fluoride ion/sulfonium substitution reactions to 25 °C, and then by correlation to the enzymatic rates, estimated that the fluorinase accelerates the substitution reaction over a noncatalyzed process by up to a 1017-fold rate enhancement!32 2.2.2. Recently Identified Bacterial Fluorinases. Recently four new fluorinases have been identified,23,24 and three of them have been characterized.23 A decade passed without any apparent fluorinase gene (f lA) homologues in the genome databases, and then in the short period between 2011 and 2013, four candidate f lA genes appeared. All have >80% homology to the original fluorinase. The first was identified from Streptomyces sp. MA37, a strain isolated in 2011 from Ghana.23 Full genome sequencing of the organism revealed a gene encoding a putative fluorinase with 87% sequence identity to that of S. cattleya. In culture the organism had the capacity to biosynthesize fluoroacetate 1 and 4-fluorothreonine 2, as well as a range of other minor (so far unidentified) fluorometabolites. Overexpression in E. coli of a codon-optimized synthetic flA gene gave rise to a soluble protein that assayed as a fluorinase. Full genome sequences of the South American hospital pathogen, Nocardia brasiliensis,23a,b and Actinoplanes sp. N902109 were deposited into the public domain in 2012 and 2013, respectively. They also contained flA genes. Synthetic codon optimized genes designed from these sequences were overexpressed in E. coli, and they too assayed as active fluorinases. These three new fluorinases all have similar kinetic profiles to the original S. cattleya enzyme. As stated, Streptomyces sp. MA37 does elaborate fluorometabolites in culture; however, attempts to culture the N. brasiliensis strain have failed to detect the presence of any fluorometabolites despite good cell growth, suggesting that the organism may only possess a latent ability to biosynthesize fluorometabolites. The Actinoplanes sp. N902-109 strain is not publically available, and it remains to be determined if it will produce fluorometabolites in culture. The draft genome of the marine bacterium Streptomyces xinghaiensis NRRL B-24674 was deposited in the public domain in 2011. More recently, a new f lA gene has been identified in S. xinghaiensis genome.24 S. xinghaiensis produces fluoroacetate in a fluoride-containing medium, and the

reenabling enzymatic turnover. An analogous process of oxocarbenium ion quenching by fluoride ion may have a role to play in the ribosyl-4′-fluoride moiety in nucleocidin 5 (Figure 1) biosynthesis; however, that remains to be verified. 2.2. Bacterial Fluorinases of Secondary Metabolism

2.2.1. Fluorinase from Streptomyces cattleya. A fluorination enzyme has been identified in bacteria, the first of which was isolated from the soil bacterium Streptomyces cattelya.21 The enzyme is involved in secondary metabolism, delivering bioactive fluorinated natural products when the bacterium is grown in a medium containing fluoride.15 This fluorinase catalyzes an SN2 substitution reaction where fluoride ion mediates a nucleophilic attack to the 5′-carbon of the ribose ring of SAM 9 to give 10 (Scheme 1). Experiments using deuterium-labeled precursors in in vivo biosynthesis experiments,26 and in in vitro assays using stereospecifically labeled [2H1-5′]-SAM 9,27 have demonstrated that the reaction proceeds with an inversion of configuration at the C-5′ carbon. Studies exploring the kinetics and order of binding have indicated that during turnover fluoride ion, which has a low affinity for the enzyme (high Km ≈ 10 mM), binds first. Structural,28 kinetic,29 and quantum mechanics/molecular mechanics (QM/MM) theoretical studies30 have indicated that fluoride locates in the active site, exchanging two of four water molecules for hydrogen bonding contacts to Ser-158, retaining four hydrogen bonding contacts, and minimizing the penalty of desolvation as illustrated in Figure 2. SAM 9 then binds into the active site with 103 higher affinity (Km ≈ 10 μM) than fluoride ion, and during that process drives full desolvation. The pre-reaction complex makes further hydrogen bonding contacts to the surface of the enzyme (via OH of Thr80) to compensate for the loss of one water molecule, and the fourth coordination site for fluoride locates to the electropositive 5′C-carbon of the positively charged sulfonium species. Thus, there is also electrostatic stabilization in the pre-reaction complex. This coordination then progresses to a nucleophilic attack and C−F bond formation. Another theoretical study modeling the enzyme reaction also concluded that this partially solvated/coordinated fluoride remains a good nucleophile.31 The QM/MM computational study30 explored alternative reaction modes such as sulfonium ylid formation using fluoride ion as a base for deprotonation; however alternative pathways were excluded as they progress through higher-energy intermediates. The study estimated that the enzyme catalyzed the reaction by 108 over the noncatalyzed reaction. Most D


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Figure 3. Comparison of the genes surrounding the fluorinase ( f lA) gene from the genomes of S. cattleya, Streptomyces sp. MA37, Norcardia brasiliensis, and Actinoplanes sp. N902-10923 and the marine microorganism Streptomyces xinghaiensis.24

production of fluoroacetate is dependent on sea salt. Although the fluorinase in S. xinghaiensis was not isolated, the production of fluoroacetate in culture suggests a functioning fluorinase. This is the first instance of a fluorometabolite isolated from a marine organism.24 The availability now of five full genomes of organisms containing the f lA genes allows a comparison of the surrounding genes and gene organization as illustrated in Figure 3. It is intriguing that four organisms (Streptomyces cattleya, Streptomyces sp MA37, N. brasiliensis, and Actinoplates sp. N902-109) appear to have the genetic capacity to biosynthesis 4-fluorothreonine (4-FT) 2, because they all contain a 4-FT transaldolase gene ( f lFT), encoding a pyridoxal phosphate (PLP) enzyme that is responsible for the last step in 4-fluorothreonine biosynthesis (Scheme 1).33,34 Inactivation of f lFT in S. cattleya resulted in the sole production of fluoroacetate, confirming its role in 4-FT biosynthesis.37 4-FT transaldoses (4-FTase) appear to be a hybrid protein containing two catalytic domains, the larger one sharing high homology to a PLP-dependent serine hydroxymethyl transferase (SHMT) and the smaller one to an epimerase/aldose.33 This is not the case for S. xinghaiensis. In S. xinghaiensis, there is a gene, adjacent to the f lA gene, that encodes an open reading frame (ORF) with only 96 amino acids in length. This ORF shares a high sequence identity (70%) only with the epimerase motif of the other 4-FTases, and the majority SHMT domain (ca. 540 aa in length) seems to be missing, clearly suggesting that this truncated 4-FTase could not carry out the biotransformation of 4-FT. This is consistent with the

experimental observation that only FAc was observed in culture.24 The arrangement of the five biosynthetic gene clusters across the individual genomes also differs significantly. In S. cattleya, f lA is located next to the second catalytic gene f lB encoding a purine nucleotide phosphorylase (PNP), which is responsible for converting 5′-FDA 10 into 5′-FDRP 11 (see Scheme 1).35 In all cases (Streptomyces sp MA37, Streptomyces xinghaiensis, N. brasiliensis, and Actinoplates sp. N902-109) the f lA gene is located in close proximity to the corresponding f lB gene. This is also the case for the chlorinase gene (salA) in the biosynthetic gene cluster for salinisporamide in S. tropica.36 Although it is common that genes of many natural product biosynthetic pathways in Streptomyces are clustered, it is not the case for fluorometabolite biosynthesis in S. cattleya.37 The remaining four catalytic genes are scattered across the S. cattleya genome and remote from the Spencer cluster.23,24,37 The 4-FT transaldolase gene (f lFT) encoding the last step of 4-FT biosynthesis is located on the megaplasmid pSCATT (1.8 Mbp in length) rather than the chromosome of S. cattleya. For the three newly sequenced actinomycete strains, the organization of the biosynthetic genes appears much more clustered and in line with expectation. For example, all of the f lFT homologues are in close proximity to the corresponding f lA homologues in the four new sequences.23 The gene f lIso encoding the 5′-FDRP isomerase that converts 5′-FDRP 11 to 5-FDRulP 1238 (see Scheme 1 and Figure 3) is located on the chromosome, remote from the f lA gene in S. cattleya. However, the homologue genes are found to be physically close to their respective f lA genes in N. brasiliensis and Actinoplates sp. N902-109.23 The f lK gene, E


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Scheme 3. Reactions of the Fluorinase, the Chlorinase, and the duf-62 Enzyme Classes; All Catalyse Nucleophilic Attack to C-5′ of SAM but with Different Nucleophiles

Scheme 4. Chlorinase Enzyme Catalyses the First Step in the Biosynthesis of Salinosporamide A from the Marine Organism Salinispora tropica34

a potent inhibitor of the fluorinase, and thus this hydrolase appears to play a role in reducing SAH levels to maximize fluorinase activity. The f lH homologues encode Na+/H+ antiporters. Despite their presence close to f lA in all four genomes, a recent study has demonstrated that the f lH is not essential for fluorometabolite biosynthesis in S. cattleya, and thus the role of this conserved gene is not clear.37 The f lG gene is a slightly unusual member of the LuxR-type subfamily of response regulators. FlG has moderate sequence identity (34%) with the regulatory protein SalR2 found in the biosynthetic pathway of salinosporamide A 19 in S. tropica. A recent report indicated that SalR2 is a pathway-specific regulator in the biosynthesis of salinisporamide A, and its overexpression led to a significant increase in the production of salinosporamide A.42,43 In this context it has been shown that raised fluoride levels amplify the expression of both the f lG and f lA genes by 2.7-fold and 2.8-fold, respectively, indicating that the fluorometabolite pathway is upregulated by increased fluoride ion levels, perhaps due to upregulation of f lG.44

which is thought to confer resistance on the organism to fluoroacetate toxicity, encodes a fluoroacetyl-CoA specific thioesterase in S. cattleya.39,40 This enzyme displays a high selectivity (106-fold) for the hydrolysis of fluoroacetyl-CoA over acetyl-CoA41 and does not allow the organism to accumulate fluoroacetyl-CoA, which would otherwise get converted to the cellular toxin (2R,3R)-fluorocitrate 3. Inactivation of f lK abolished fluorometabolite production in S. cattleya in an otherwise healthy organism,37 so there appears to be a fail-safe mechanism, turning off fluoroacetate biosynthesis, if f lK is inactive. Reintroduction of the f lK4 gene, the f lK homologue gene, into the f lK4 knockout mutant of S. xinghaiensis resulted in the restored production of fluoroacetate.24 The physical location of the fluoroacetate resistance (f lK) gene is very close to the f lA gene in S. cattleya and S. xinghaiensis; however, this is not the case in the three new genomes where the respective f lK homologues are distant from their f lA genes (Figure 3). Interestingly, four genes encoding putative auxiliary functions are highly conserved across the five genomes, including DNA regulation ( f lF, G, and I homologues), and transporter ( flH homologues) genes. The f lI homologues encode S-adenosyl-L-homocysteine (SAH) hydrolases that catalyze the degradation of SAH. SAH, which is directly derived from and structurally analogous to SAM 9, is

2.3. Chlorinase and Hydrolases Related to the Bacterial Fluorinases

The f lA gene was first sequenced in Streptomyces cattleya, located at the center of a 10 kb gene cluster (Spencer cluster).45 F


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Scheme 5. Genetic Engineering of S. tropica, Inserting the Fluorinase ( flA) Gene in Place of the Chlorinase Gene (salL) Resulted in an Organism with the Capacity to Biosynthesize Fluorosalinosporamide-A 2053

containing medicines, in heterologous bacterial strains, by fermentation technology. The first steps in this direction have been achieved and are discussed below. 3.1.1. Engineering Fluorosalinosporamide Production in Salinospora tropica. In view of the similarity between the fluorinase and the chlorinase36 genes/enzymes in S. tropica responsible for salinosporamide-A 19 biosynthesis, there was an opportunity for metabolite engineering. The chlorinase carries out the first step in salinosporamide A 19 biosynthesis to generate 16. Salinosporamide A 19 is cytotoxic51,52 and has progressed to phase II clinical trials for cancer treatment. A summarized biosynthetic pathway to 19 is shown in Scheme 4. The salL (chlorinase) gene is located in the middle of this salinosporamide A cluster. The chlorinase does not accept fluoride ion as a substrate; therefore, it became attractive to try to engineer S. tropica to induce fluorosalinosporamide 20 production by replacing the chlorinase (salL) by the fluorinase ( f lA) gene. This was accomplished by inserting the f lA gene into the middle of the salL gene, to deactivate the latter and position f lA such that it was located close to the other biosynthetic and regulatory genes controlling salinosporamide A biosynthesis.53 Scheme 5 illustrates the event in which the engineered strain had the capacity to biosynthesize 20 in culture. Although a positive development, some problems emerged. S. tropica, which normally grows in the marine environment, proved to be extremely sensitive to fluoride ion in culture, and this limited the prospect for a high titer from large-scale fermentation. Successful production required that cell growth was pre-established in the absence of fluoride, and then after several days, fluoride ion was introduced to mature cells. Thus, a biotechnologically useful organism is required to be engineered, such that fluoride ion toxicity is eliminated. Nonetheless, the ability to engineer the fluorinase into a foreign host, and demonstrate the production of a pharmaceutically relevant bioactive such as 20, offers encouraging prospects for this approach. 3.1.2. Incorporation of Fluoroacetyl-CoA and Fluoromalonyl-CoA into Polyketide Frameworks. Fluoroacetyl-CoA 21 can be generated readily in vivo from fluoroacetate 1. This is the basis for fluoroacetate toxicity where fluoroacetylCoA 21 acts as a substrate for citrate synthase. Fluorooleic acid 4 (Figure 1) almost certainly arises by an in vivo fatty acid assembly process using fluoroacetyl-CoA 21 as a starter unit in the seeds of Dichapetalum toxicarium.10 In an effort to engineer a similar strategy for polyketide synthesis, Spencer and coworkers54 incubated fluoroacetyl-CoA 21 with an overexpressed minimal polyketide synthase (PKS), involved in the biosynthesis of the antibiotic actinorhodin. Incubation of fluoroacetyl-CoA, malonyl-CoA, and the PKS successfully

The gene is 900 bp in length and codes for 299 amino acids. The nearest relative to the fluorinase is a chlorinase from the marine bacterium Salinispora tropica.46 It has a relatively low ∼35% amino acid homology. This enzyme carries out a very similar reaction where it combines SAM and chloride ion to catalyze a nucleophile substitution at C-5′ of SAM to generate 5′-chlorodeoxyadenosine (5′-ClDA) 16 and L-methionine (Scheme 3). The reaction is the first step in the biosynthesis of the marine metabolite salinosporamide A 19 (Scheme 4).42 The chlorinase cannot utilize fluoride ion as a nucleophile, although the fluorinase can utilize both fluoride ion and to a lesser extent chloride ion.47 Crystallography has revealed different active site residues between the fluorinase and the chlorinase, and notably there is no equivalent to Ser-158 (Figure 2a) in the chlorinase, with a different organization for halide binding.48 The next most closely related protein to the fluorinase is a large class expressed by the duf-62 (domains of unknown function-62) genes, which have ∼30% homology.49,50 In view of the relatively close sequence similarity between the fluorinase and the duf-62 family, a representative duf-62 enzyme, from the hyperthermophilic archaeon Pyrococcus horikoshii, was overexpressed. Assays of this enzyme showed that it mediated a nucleophilic attack of hydroxide ion (from water) to C-5′ of SAM, to generate adenosine 17 and Lmethionine (Scheme 3). 49,50 Thus, the fluorinase, the chlorinase, and the duf-62’s utilize different nucleophiles to attack C-5′ of SAM 9. The duf-62 enzyme is unable to utilize fluoride ion as a nucleophile. An inspection of the active site of the duf-62 from P. horikoshi reveals a very different organization of amino acid side chains when compared to the fluorinase. The duf-62 family (∼200 members) has a highly conserved His·· Arg··Asp triad, which is absent in the fluorinase. Although these enzymes (fluorinase, chlorinase, and duf-62’s) have some sequence similarity, as well as a similar quarternary structure, they use different nucleophiles for substitution reactions at C-5′ of SAM, and they have very different amino acid side chains in their active sites. A comparison of the fluorinase with the chlorinase and the duf-62 genes reveals a 21 amino acid insert unique to the fluorinase, which translates to a distinct loop in the monomeric structure found only in the fluorinase and not the chlorinase or duf-62’s.48 This loop does not form part of the active site, and it does not have an obvious role in the catalysis, but it is a signature of the fluorination enzyme.

3. BIOTECHNOLOGICAL POTENTIAL FOR ENZYMATIC FLUORINATION 3.1. Fluorometabolite Engineering

It is an exciting prospect that f lA genes and their associated biosynthetic and regulatory genes could be used to engineer the biosynthesis of novel fluorometabolites and perhaps fluorineG


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incubations of fluoromalonyl-CoA or fluoromalonate with appropriate activating enzymes, including methylmalonyl-CoA 27 and additional cofactors, partially activated SNAC diketide 25, and an assembly module from the erythromycin PKS, generated partially assembled tri- 25 and tetraketides 28 and 29, respectively. The fluorines located to midchain sites. In an in vivo experiment using an engineered E. coli, fluoromalonate was converted to 26 when 25 was added to the incubation as a primer unit. Polyketide metabolites56 contribute many of the important antibiotics used clinically, and these biochemical reactions point the way to preparing fluorinated analogues of more complex structural architectures by fermentation technology.

generated the octaketide 22 carrying a fluoromethyl group originating from fluoroacetyl-CoA 21 (Scheme 6). Scheme 6. Incorporation of Fluoroacetyl-CoA As a Starter Unit in Aromatic Polyketide Biosynthesis54

3.2. Fluorinase As a Catalyst for Positron Emission Tomography

18

F−C Bond Formation in

The fluorinase has proven to be an excellent catalyst for radiolabeling of organic substrates using [18F]-fluoride. The fluorine-18 isotope is widely used in clinical imaging as one of the longer-lived isotopes (t1/2 = 109 min) among those employed in positron emission tomography (PET), and innovative methodologies for its incorporation have been an intense focus of research over the past decade.57 There are limitations associated with using [18F]-fluoride in radiochemistry.58 The [18F]-fluoride ion is generated on a cyclotron by bombardment of a vial (1−2 mL) of [18O]-H2O with a beam of high-energy protons. A nuclear reaction occurs in the water target, converting a few water molecules to [18F]-fluoride. The molar conversion to [18F]-fluoride is low (picomolar, 10−12 M), although the specific activity can be very high (Giga-Mega bequerels, GBq-MBq). The usual procedure involves removing [18F]-fluoride ion from the aqueous solution in which it is generated, by ion-exchange chromatography, for formulation in a Kryptofix [2.2.2]-30 matrix, as its potassium salt. It is this form of [18F]-fluoride that is used for nucleophilic chemistry to prepare radiolabeled ligands for clinical imaging. In this context 2-fluoro-2-deoxyglucose ([18F]-FDG) 33 is the most widely used imaging ligand in the clinic and accounts for >95% of all clinical imaging studies worldwide.59 [18F]-FDG 33 is prepared

One can envisage rather elaborate architectures incorporating fluorine if such pathways could be engineered into host organisms for in vivo fermentations. In this case fluorine results at the carbon chain terminus; however, incorporation into midchain positions requires fluoromalonyl-CoA 21 as a building block. It has recently been demonstrated55 in engineered E. coli that malonyl-CoA synthetase, which normally converts malonate to malonyl-CoA, will convert fluoromalonate 23 to fluoromalonyl-CoA 24, albeit at a reduced rate. Also fluoroacetyl-CoA was converted to fluoromalonyl-CoA by the action of acetyl-CoA carboxylase, at only a moderately reduced rate (4.5 times slower). Therefore, several biochemical strategies for coaxing fluoroacetate to fluoromalonate or fluoromalonyl-CoA exist for future engineering of biosynthesis pathways. If fluoromalonyl-CoA is also accepted by PKS’s, then this opens up possibilities of incorporating fluorine into midchain positions in metabolites from a fluoroacetate/ fluoromalonyl feed stock. The study demonstrated this in principle, and results are summarized in Scheme 7. In vitro

Scheme 7. Biochemical Incorporation of Fluoromalonates Has Been Demonstrated into Midchain Locations of Polyketide Fragments55

H


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Scheme 8. Radiochemical Synthesis of [18F]-2-Fluoro-2-deoxyglucose ([18F]-FDG)

Scheme 9. Enzymatic Synthesis of [18F]-10 Utilizing the Fluorinase and an Amino Acid Oxidase61

Scheme 10. Synthesis of a Range of [18F]-5′-Fluorodeoxy Nucleosides Can Be Prepared after Fluorinase Mediated Fluorination, By Base-Swap Reactions Using Appropriate Purine/Pyrimidine Nucleotide Phosphorylase Enzymes62

h) time periods. The fluorinase has been applied to the synthesis of a range of novel [18F]-radiolabeled products, examples of which are described below. 3.2.1. [18F]-Fluoronucleosides. The first synthesis that was explored using the fluorinase involved a radiolabeled synthesis of [18F]-FDA 10.60,61 This was accomplished most efficiently by the addition of an L-amino acid oxidase (L-AAO) to the reaction cocktail, which contained SAM, fluorinase, and [18F]-fluoride. The reaction is summarized in Scheme 9. The L-AAO acts to oxidize the L-methionine that is formed in the reaction and to prevent the reverse reaction from occurring. It should be noted that SAM 10 from commercial suppliers also contains low levels of L-methionine, and because turnover is at the 10−12 M level, even low levels of L-methionine are in excess for the reverse reaction and reduce the efficiency of the radiochemical synthesis. However, with added L -AAO, satisfactory conversions (∼95%) of [18F]-fluoride to [18F]-10

by nucleophilic displacement of triflate 31 on a protected sugar by Kryptofix secured [18F]-fluoride 30, to generate 32, which, followed by acid catalyzed hydrolysis, gives 33 (Scheme 8). Enzymes operate in water, and therefore an immediate advantage of enzymatic fluorination is that the fluorinase can utilize [18F]-fluoride directly as a solution in the [18O]-H2O target, without recourse to drying the fluoride ion and preparing 30. It emerges that the fluorinase is a very satisfactory catalyst for 18F−C bond formation for PET radiochemistry. The [18F]-fluoride ion concentrations that are generated in [18O]-H2O are very low (10−12 M), but the enzyme concentration is prepared at a μM (mg/mL) level; thus, there is ∼106 molar excess of enzyme over [18F]-fluoride ion. This is a very unusual form of enzyme conversion in that it is hardly catalytic, but it favors 18F−C bond synthesis, and experimentally radiochemical conversions to [18F]-organofluorine products are common within relatively short (0.5−1 I


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Scheme 11. Dimethyl-aza-SAM 36 Is Competent to Act As a Substrate for Fluorinase Catalysed Formation of [18F]-1063

It followed that suitable substituents attached to C-2 of adenine might project out of the active site and have little influence on binding or catalysis. The acetylene moiety was explored because it fulfilled obvious criteria in that it is linear and sterically compact. In the event the fluorinase was able to catalyze a trans-halogenation reaction of acetyleneic ClDA 37 to acetylenic FDA 38 as illustrated in Scheme 12. The reaction followed from an earlier discovery that the fluorinase is reversible and that L-Met or L-selenomethionine (L-SeMet) will displace the chloride of ClDA 16 to generate SAM, and then the SAM is converted to FDA in a one-pot reaction.47 The conversion of acetyleneic ClDA 37 to 38 proceeded at ∼60% of the efficiency of the transhalogenation of ClDA 16 to 5′-FDA 10. The approach was then extended by attaching a pegylated chain to the terminus of the acetylene with substrate 39, as illustrated in Scheme 12. In this substrate the pegylated chain links to a cyclic Arg-Gly-Asp (RGD) peptide, a class of peptides relevant to cancer cell identification and clinical imaging.65 The fluorinase accepted substrate 39 and was able to catalyze its conversion to [18F]-41 in a biotransformation that proceeds via intermediate 40. The radiochemical conversion was excellent, generating [18F]-41 in a 90% radiochemical conversion from [18F]-fluoride in 30 min. The method achieves “last-step” labeling of a peptide in buffer at neutral pH and ambient temperature, conditions previously unavailable for fluorine-18 labeling of bioactive molecules. This approach is currently being developed toward higher molecular weight peptides and proteins for clinical imaging. 3.2.3. [18F]-Fluoroacetate. [18F]-Fluoroacetate 1 has been used as a radiotracer to explore the relative activity of glial neurons in the brain,66 and it is also being developed as a cancer marker,67 where it identifies regions of high levels of acetate metabolism in rapidly metabolizing cells. A simple modification of the enzymatic synthesis of [18F]-10 allowed the chemo-enzymatic preparation of [18F]-1.68 When fluorinase generated [18F]-10 is treated with an oxidizing solution of chromium trioxide (CrO3) and sulfuric acid, in a Kuhn−Roth oxidation, then the C-4′-carbon of the sugar with the attached fluoromethyl group is oxidized to carve out [18F]-1. The oxidation is extremely efficient (∼10 min), and the radiolabeled [18F]-1 is extracted directly into diethyl ether and then neutralized to give sodium [18F]-1 (Scheme 13). 3.2.4. [18F]-Fluororibose. Probably the most usefully applied radiolabeled compound to have been prepared so far using fluorinase technology is [18F]-5-fluoro-5-deoxyribose 42. Ribose [18F]-42 has some similar characteristics to [18F]-FDG 33 in that they are both [18F]-labeled monosaccharides. It has been explored as a tracer for tumor imaging and also as a bioconjugation agent for the radiolabeling of bioactive peptides. Initially a three-enzyme strategy (fluorinase, PNP, and phytase) was used for the preparation of [18F]-42 as illustrated in Scheme 14.61

can be achieved as the enzyme oxidises endogenous Lmethionine.61 The synthesis of a range of additional [18F]nucleosides was prepared via base-swap reactions.62 A purine nucleotide phosphorylase (PNP) enzyme was used in a phosphate buffer to depurinate [18F]-FDA and to generate ribose-phosphate [18F]-11 in the reaction. This is an activated sugar, and by addition of alternative purine bases in an excess, then the PNP was able to catalyze the reverse reaction to generate the purine nucleotides such as [18F]-34. A range of [18F]-fluoropyrimidines could also be prepared by adding a pyrimidine nucleotide phosphorylase to the reaction cocktail. For example fluorouracil [18F]-35 was prepared in this way (Scheme 10). All of these base-swap reactions can be achieved in modest-to-good radiochemical conversions within the 1−2 h time scale. Sergeev et al.63 have recently reported the conversion of the dimethyl-aza-analogue 36 of SAM to generate [18F]-10 (Scheme 11). In this reaction it would appear that 36 is acting as a substrate replacement for SAM. The reaction requires LAAO to move the equilibrium in the direction of [18F]-10 production and arrest any reverse reaction with accumulating coproduced L-Met. Only a radiolabeled reaction was reported, and presumably the turnover is so low that it can only be detected by radioactivity, e.g., rather than by high-performance liquid chromatography (HPLC) assay. The enzyme has also been immobilized to provide a potentially more efficient and rapid preparation of [18F]-10.63 3.2.2. “Last-Step” Labeling of Peptides Using the Fluorinase.64 A significant breakthrough was made recently in terms of expanding the substrate specificity of the fluorinase and therefore improving its potential as a catalyst for the introduction of fluorine-18 into peptides for PET. X-ray structure analysis of the fluorinase cocrystallized with 5′-FDA 10 revealed that the C-2 position of the adenine ring is close to the surface of the enzyme and that C(2)-H is pointing into the solvent void (see Figure 4).

Figure 4. Comparison of X-ray cocrystal structures of the fluorinase active site with 5′-FDA 10 and acetylene 38 overlaid, revealing a weakness in the substrate specificity of the enzyme. The acetylene moiety of 38 projects from the active site into the solvent void. J


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Scheme 12. Efficient Fluorinase Catalyzed Transhalogenation (Organic-Chlorine to [18F]-Organic-Fluorine) Reactions Demonstrating a New Strategy for 18F-Radiolabelling of RGD Peptides for PET under Ambient Aqueous Conditions64

Scheme 13. Chemo-enzymatic Route to Na[18F]-1 Fluoroacetate68

overexpression of this hydrolase IAG-NH allowed the radiochemical synthesis of [18F]-42 in 80% conversion. A chemical hydrolysis protocol replaced the second hydrolytic enzyme, and thus a fluorinase/AAO preparation of [18F]-FDA followed by acidic hydrolysis provides an efficient method for the synthesis of [18F]-42 (Scheme 14). Finally, a totally synthetic protocol for the preparation of [18F]-42 was developed by nucleophilic displacement of tosyl 43 or mesyl 44 protected ribose to generate [18F]-45. Hydrolysis of acetonide [18F]-45 generated [18F]-42 (Scheme 15),71 completely independent of the requirement for the fluorinase enzyme, and similar to the general strategy used for [18F]-FDG preparation (Scheme 4). A study was conducted on the relative abilities of ribose [18F]-42 and glucose [18F]-33 to image tumors.72 This was explored in a subcutaneous tumor (A431 human epithelial carcinoma) in a mouse model. In the event [18F]-42 proved to be metabolically stable, it did not metabolize to [18F]-fluoride. It also was successful in its ability to image tumors over a relatively short time period as illustrated in Figure 5. However, it was noted that with time the efflux rate of the radiotracer from the tumor began to reduce image contrast quality. Conversely, [18F]-33 steadily accumulates in the tumors and gave the more persistent images. This is understood to be due to the ability of [18F]-FDG to become phosphorylated within cells. This study indicated that there was no particular advantage in using [18F]-42 over [18F]-33 as a tumor imaging

Scheme 14. Enzymatic and Chemo-enzymatic Routes to [18F]-4261,68

Although this proved successful, the protocol was first superseded by a two-enzyme strategy that involved direct enzymatic hydrolysis of [18F]-10, rather than proceeding via the intermediate fluororibose phosphate [18F]-11.69 Direct hydrolysis required a less common purine hydrolase activity, which was sourced from the parasite Trypanosoma vivax.70 Thus, K


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Scheme 15. Chemical Synthesis of [18F]-4270,71

Figure 5. Longitudinal sectional PET imaging of a mouse model. Arrows in the central image indicate subcutaneous tumor detection after injection of [18F]-42 in the mouse model.72

Figure 6. Examples of a variety of reactive small molecules that have been used to radiolabel peptides with [18F]-fluoride for positron emission tomography.

47 and 48,74 or fluorinated maleimides 49 for thiol reactions, or acetylenes such as 50 for “click” condensations.75 RGD peptides recognize and bind to integrins (conserved peptide motifs) on the surface of cells.76 Integrins such as the αvβ3 motif are upregulated in many tumor cells. For example, the RGD peptide [18F]-fluciclatide has proven clinically effective as an agent for monitoring tumor regression.77 For diagnostic purposes, [18F]-fluciclatide is labeled in a very classical manner by conjugation to benzaldehyde [18F]-46 through an aminooxy ether functionality to generate a stable oxime (Figure 6).78

agent, although it demonstrated the metabolism-resistant qualities of the radiotracer (Figure 5). 3.3. [18F]-Labeling of Peptides with [18F]-FDR

Ribose [18F]-42 has proven to be an excellent bioconjugation agent for [18F]-tagging bioactive peptides. The field of radiolabeling peptides for targeted clinical imaging by PET is very active. A multitude of small reactive [18F]-labeled molecules have been explored in this context, although [18F]benzaldehyde 46 is among the most common.73 Others are illustrated in Figure 6 and involve activated [18F]-benzoic acids L


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Scheme 16. Ribose [18F]-42 Is an Efficient Small Molecular Tag for the Preparation of [18F]-Radiolabelled Bioactive Peptides70,78,80,81

Ribose [18F]-42 has proven be an excellent conjugation agent in this context.71,79,80 It is predisposed to ring opening as a 5membered ring carbohydrate. Also the electronegative fluorine situated at C-5 promotes ring opening to the reactive aldehyde form. In this way the RGD peptides 51 and 53 illustrated in Scheme 16 have been successfully tagged with [18F]-42 to give [18F]-FDR-RGD adducts 52 and 5479 and the SLIGKV peptide, which is an agonist of the cancer upregulated human protease activated receptor 2 (PAR-2) and has also been successfully labeled to generate [18F]-55 by this strategy.71 The conjugations occur extremely rapidly, at ambient temperature and only under moderately acidic conditions (pH ≈ 4.6). Also peptide Siglec-9 adduct 56, which has been developed as an inflammation marker, was successfully tagged with [18F]-42 to image inflammation in a mouse model PET imaging study.80,81

4. CONCLUSIONS It has been over a decade since the fluorinase enzyme was isolated and characterized, and in that time its mechanism has been evaluated and the enzyme has been applied in several different directions in terms of developing fluorinase-based technologies. There has been some exciting success in cloning the f lA gene to induce fluorometabolite production in a host organism. As described, this was demonstrated in S. tropica where the marine organism was engineered to produce fluorosalinosporamide. The study points the way to preparing other fluorinated bioactives by microbial engineering, although it highlights the necessity for good pathway regulation and to engineer a host organisms that is tolerant to fluoride ion. The enzyme has proven to be a particularly useful catalyst for securing the fluorine-18 isotope from [18F]-fluoride ion, to prepare a range of molecules for positron emission tomogM


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raphy. For example [18F]-FDR 42 is a particularly reactive fluorinated ribose that can efficiently tag PET relevant peptides via oxime ligation. Also the fluorinase offers a strategy to access a range of [18F]-nucleosides and [18F]-fluoroacetate as tracers for PET, as well as a “last-step” [18F]-labeling approach to peptides under ambient conditions. Other exciting prospects involve the isolation of novel fluorination enzymes such as that involved in nucelocidin production from Streptomyces calvus. Nothing is known regarding the biosynthesis of this metabolite. Also the genes and enzymes that are responsible for fluoroacetate production in higher plants from South Africa and Australia remain to be identified and characterized. Dr Hai Deng studied chemistry for his Bachelor’s and Masters’ degrees in China. In 1999, he came to the UK to pursue his Ph.D. in the field of biochemistry and biotransformations at the University of Wales, Swansea. In 2002 he joined Professor David O’Hagan’s group as a postdoctoral research fellow in the Centre for Biomolecular Sciences at the University of St Andrews. He was appointed as a Lecturer at the Department of Chemistry, University of Aberdeen in 2008. His research team is situated in the Marine Biodiscovery Centre (MBC), Department of Chemistry, University of Aberdeen. His research is mainly focused on discovering novel bioactive natural products from various resources, tracing biosynthesis pathways, identifying novel enzyme activities in secondary metabolism, and enzyme mechanism. He is a Member of Royal Society of Chemistry.

AUTHOR INFORMATION Corresponding Authors

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

The authors declare no competing financial interest. Biographies

ACKNOWLEDGMENTS We thank all of our colleagues at the University of St. Andrews who worked on this project over the years, particularly Jim Naismith’s lab for structural biology support. It has been a pleasure to engage in fluorinase collaborations with many scientists in other laboratories including Walter Thiel and Hans Martin Senn of the Max Planck in Mulheim; Bradley Moore and Alessandra Eustáquio of the Scripps Institution of Ocenaography; Matteo Zanda, Sergio D’Allangelo, Lutz Schweiger, Juozas Domarkas, and Ian Fleming of the University of Aberdeen; Wim Versées, Jan Steyaert, and John N. Barlow of Vrije Universiteit Brussel; Jan Passchier and Mayca Onega at Imanova in London; Xiang-Gu-Li of the University of Turku; Bert Windhorst and Danielle Vugts of the Free University, Amsterdam; Kwaku Kyeremeh of the University of Ghana; Yi Yu of Wuhan University; and Hong-Yu Oh of Jiaotong University. D.OH acknowledges the Royal Society for a Wolfson Research Merit Award.

Professor David O’Hagan was born in Glasgow and studied chemistry at the University of Glasgow, graduating in 1982. He carried out a Ph.D (1985) in polyketide antibiotic biosynthesis at the University of Southampton with Professor John A Robinson and then spent a postdoctoral year at the Ohio State University with Professor Heinz G Floss, investigating peptide antibiotic biosynthesis. In 1986 he was appointed to the University of Durham where he continued to explore natural product biosynthesis but also developed a strong interest in organo-fluorine chemistry. He remained at Durham until 2000 before moving to his current position as Professor and Head of Organic Chemistry at the University of St Andrews. His research interests extend from the synthesis and properties of organo-fluorine compounds, fluorination enzymology, fluorine-18 chemistry for positron emission tomography (PET) and through to fluorinated organic materials. He was awarded the RSC Malcolm Campbell Memorial Prize in Medicinal Chemistry in 2005, was a recipient of the RSC Tilden Medal in 2006/was the RSC ’Natural Product Reports Award’ Lecturer in 2009, and was awarded the Americal Chemical Society (ACS) Award for ’Creative Work in Fluorine Chemistry’ in 2012. He received a Royal Society Wolfson Research Merit Award in 2013.

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