Phosphonate Biochemistry - American Chemical Society

Review pubs.acs.org/CR

Phosphonate Biochemistry Geoff P. Horsman*,† and David L. Zechel*,‡ †

Department of Chemistry and Biochemistry, Wilfrid Laurier University, Waterloo, Ontario N2L 3C5, Canada Department of Chemistry, Queen’s University, Kingston, Ontario K7L 3N6, Canada

‡

ABSTRACT: Organophosphonic acids are unique as natural products in terms of stability and mimicry. The C−P bond that defines these compounds resists hydrolytic cleavage, while the phosphonyl group is a versatile mimic of transition-states, intermediates, and primary metabolites. This versatility may explain why a variety of organisms have extensively explored the use organophosphonic acids as bioactive secondary metabolites. Several of these compounds, such as fosfomycin and bialaphos, figure prominently in human health and agriculture. The enzyme reactions that create these molecules are an interesting mix of chemistry that has been adopted from primary metabolism as well as those with no chemical precedent. Additionally, the phosphonate moiety represents a source of inorganic phosphate to microorganisms that live in environments that lack this nutrient; thus, unusual enzyme reactions have also evolved to cleave the C−P bond. This review is a comprehensive summary of the occurrence and function of organophosphonic acids natural products along with the mechanisms of the enzymes that synthesize and catabolize these molecules.

CONTENTS 1. Introduction 2. Occurrence of Organophosphonates in Nature 2.1. Why Nature Chose Phosphonates 2.2. Abiotic Synthesis of Pn 2.3. Phosphonolipids and Phosphonoglycans 2.4. Bioactive Pn Secondary Metabolites 3. Biosynthesis of Pn Building Blocks 3.1. Phosphonopyruvate (1) 3.2. Phosphonoacetaldehyde (2) 3.3. 2-Aminoethylphosphonic Acid (3) 3.4. 2-Hydroxyethylphosphonic Acid (4) 3.5. 1-Hydroxy-2-Aminoethylphosphonic Acid (5) 3.6. Phosphonoalanine (6) 3.7. Phosphonoacetate (7) 3.8. Methylphosphonic Acid (8) 3.9. Phosphonomethylmalic Acid (56) 3.10. 2-Keto-4-hydroxy-5-phosphonopentanoic Acid (57) 3.11. Natural Product Pn of Unknown Provenance 4. Pathways for Synthesizing Complex Pn Natural Products 4.1. Fosfomycin (9) 4.1.1. Mechanism of Action 4.1.2. Biosynthesis of Fosfomycin 4.1.3. Cobalamin-Dependent Methyltransferase Fom3 4.1.4. 2-Hydroxypropylphosphonate Epoxidase Fom4 4.1.5. Biosynthesis of Fosfomycin in Pseudomonas sp. © 2016 American Chemical Society

4.1.6. Mechanisms of Resistance to Fosfomycin 4.2. Phosphinothricin (10a) 4.2.1. Mechanism of Action 4.2.2. Biosynthesis of Phosphinothricin and Derivatives (10a−d) 4.2.3. Hydroxyethylphosphonate Dioxygenase (HEPD) Encoded by PhpD 4.2.4. PhpE, PhpJ, and PhpF Catalyze Phosphonoformate (30) Synthesis and Activation 4.2.5. Glycolytic Enzyme Homologues PhpG and PhpH Generate CPEP (95) 4.2.6. Carboxyphosphonoenolpyruvate (CPEP) Mutase 4.2.7. Similarities to the Citric Acid Cycle en Route to Demethylphosphinothricin (101) 4.2.8. Phosphinothricin Acetyltransferase (PAT): From Self-Protection to Crop Protection 4.2.9. Breaking the NRPS Specificity Code: PhsA, PhsB, and PhsC 4.2.10. Putative Transacylase PhpL and Putative Thioesterase PhpM 4.2.11. Cobalamin-Dependent P-Methyltransferase PhpK 4.2.12. Deacetylase

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Special Issue: Unusual Enzymology in Natural Products Synthesis

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Received: August 11, 2016 Published: October 27, 2016 5704

DOI: 10.1021/acs.chemrev.6b00536 Chem. Rev. 2017, 117, 5704−5783

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Figure 1. Overview of Pn natural products. Pn within the circle, with the exception of 4, are distinguished by their dual roles in biosynthesis and sources of Pi. Pn outside the circle are bioactive secondary metabolites.

4.2.13. Resistance Mechanisms 4.3. Dehydrophos (11) 4.3.1. Mechanism of Action 4.3.2. Biosynthesis of 11 4.3.3. Dioxygenase DhpA 4.3.4. Kinase DhpB, Dehydrogenase DhpC, and Aminotransferase DhpD 4.3.5. DhpH: a tRNA-Dependent Peptide Ligase also Catalyzing PLP-Dependent Elimination 4.3.6. Phosphonate O-Methyltransferase DhpI 4.3.7. Desaturase DhpJ Installs the Vinyl Moiety 4.3.8. Peptide Ligase DhpK 4.4. FR-900098 (12a), Fosmidomycin (12b), and FR-32863 (12c) 4.5. Fosfazinomycins (13a−c) 4.6. Rhizocticins, Plumbemycins, and Phosacetamycin (14b−h) 4.7. Emerging Pn Pathways 4.7.1. Argolaphos A and B (15a,b) 4.7.2. Phosphonothrixin (17) and Valinophos (18) 4.7.3. Nitrilalophos, Hydroxynitrilaphos, and Phosphonocystoximic Acids (19a−d) 5. Pathways for Catabolizing Pn as a Source of Pi 5.1. Hydrolytic C−P Bond Cleavage 5.1.1. Phosphonopyruvate Hydrolase 5.1.2. Phosphonacetaldehyde Hydrolase 5.1.3. Phosphonoacetate Hydrolase

5.2. Radical C−P Bond Cleavage by Carbon− Phosphorus Lyase 5.2.1. Substrate Specificity and Reaction Products 5.2.2. Structural Diversity of the phn Operon 5.2.3. Regulation of the phn Operon 5.2.4. CP-Lyase Pathway 5.2.5. PhnG2H2I2J2K Complex 5.3. Oxidative C−P Bond Cleavage by PhnY*/ PhnZ 6. Summary and Outlook Author Information Corresponding Authors Notes Biographies Acknowledgments References

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1. INTRODUCTION As a class of natural product, organophosphonic acids (Pn) are unique for their Janus-faced relationship with the microbial world. On the one hand the carbon−phosphorus (C−P) bond that defines these molecules is a key driver of their potent bioactivities toward microbes (and other higher order organisms) as well as their stability against enzymatic degradation. On the other hand, the phosphorus atom represents a source of inorganic phosphate (Pi) for microorganisms that, more often than not, are starved for this lifelimiting nutrient, one that Karl has aptly called “the staff of

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Figure 2. Examples of synthetic organophosphonic acids. The biological target or function of each phosphonate is given in parentheses, while the natural substrate or ligand of a biological target is shown in blue.

life”.1 As a result the biochemistry of Pn has evolved into two worlds, with one dedicated to synthesizing the C−P bond and incorporating this motif into complex bioactive molecules, while the other seeks to cleave the C−P bond to release Pi. At the heart of Pn chemical space are eight relatively simple Pn that either form “building blocks” to create more complex natural products, or are substrates for catabolic pathways that allow microorganisms to extract Pi (Figure 1, inner circle). These include phosphonopyruvate 1, phosphonacetaldehyde 2, 2-aminoethylphosphonic acid 3, 2-hydroxyethylphosphonic acid 4, 1-hydroxy-2-aminoethylphosphonic acid 5, phosphonoalanine 6, phosphonoacetate 7, and methylphosphonic acid 8. Some of these simple Pn exhibit surprising distributions and bioactivities. For example, 3 is concentrated in the gills of male blue crabs, but not females,2 while 6 is a potent inhibitor of a neuronal excitatory amino acid receptor associated with phosphoinositide hydrolysis.3 From this center, Nature derives an inspiring array of Pn structures that encompass a wide range of bioactivities. The majority of these compounds are antibiotics, the most famous example being fosfomycin 9, which has a long history as a treatment for urinary tract infections. However, a number of Pn are herbicidal including phosphinothricin 10a, also known as glufosinate, and phosphonothrixin 17. 10a is used commercially in combination with crops that are genetically modified to be resistant to this herbicide. More exotic activities are exhibited by phosphoiodyn A 16,4,5 which acts as an agonist of the human transcription factor peroxisome proliferator-activated receptor gamma (hPPARγ), and the related compounds K-26 21a and 15B2 21b, which are inhibitors of the angiotensin-converting enzyme (ACE). Pn are often thought to be synthetic creations, and indeed over many decades they have been created to serve a variety of functions. As with the natural Pn, the utility of synthetic Pn stems from the stability of the carbon−phosphorus bond as well as the ability of the phosphonic acid moiety to mimic the polar functional groups of enzyme substrates. Some prominent examples illustrate these themes (Figure 2). The most widely

used herbicide on Earth is glyphosate 23, more commonly known as Roundup, which inhibits 5-enoylpyruvylshikimate-3phosphate (EPSP) synthase as a mimic of the substrate phosphoenolpyruvate (PEP) 24. Sarin 25 and VX 26 have earned notoriety as chemical warfare agents. These compounds are analogs of the neurotransmitter acetylcholine 27 and achieve their toxicity by inactivating acetylcholine esterase. Tenofivir 28 is listed by the World Health Organization as an essential medicine for the treatment of HIV and hepatitis B.6 Upon diphosphorylation in the cell, 28 acts as an inhibitor of viral reverse transcriptases by mimicking deoxynucleotide triphosphates. Zoledronate 29 and phosphonoformate (Foscarnet) 30 are analogs of pyrophosphate and are used to treat osteoporosis and herpes, respectively. 29 also has anticancer activity due to its ability to target the dimethylallyl pyrophosphate binding site of human farnesyl pyrophosphate synthase.7 Perzinfotel 31 is an analog of N-methyl-D-aspartate 32 and thus acts as an antagonist toward NMDA receptors. More prosaic applications are fulfilled by multivalent Pn such as EDTMP 33, which are frequently used as metal ion chelating agents. While Pn have been recognized as natural products for nearly 60 years, the enzymology that is involved in the biosynthesis and catabolism of these molecules has, to a large degree, only begun to come to light in the last 20 years. Like other areas of natural product chemistry, the discovery of new Pn pathways and enzymes has greatly benefited from advances in genome sequencing, manipulation of genomic and metagenomic DNA, and improved analytical techniques. The purpose of this review is to comprehensively examine our current understanding of Pn biosynthesis and catabolism while highlighting the unusual enzymology that is often found within these pathways. While the emphasis is on literature from the last 20 years, much older and often underappreciated literature is included for context as well as to highlight some of the remarkable mechanistic puzzles that have required decades of effort to solve. This review will first examine the occurrence of Pn in Nature, followed by the biosynthesis of the Pn “building-blocks” shown in Figure 1. The 5706


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Figure 3. Pn mimicry and stability. (A) Alkyl phosphonate mimicry of a phosphate ester or carboxylate. (B) Mimicry of a transition state for acyl transfer. (C) Calculated bond dissociation energy values (kJ mol−1) for selected O−P and C−P bonds.41 (D) Partial reaction coordinate for competing hydrolysis reactions of O−P and C−P bonds.43 Calculated Gibbs free energy values are shown.

Scheme 1

enzyme structures;27,28 enzyme mechanisms;10,29−32 and the total synthesis of Pn natural products.33

building blocks are a useful organizing principle in Pn biochemistry as these compounds feature prominently in both the biosynthetic and catabolic pathways described later in the review. In addition to reviewing the better-characterized biosynthetic pathways such as fosfomycin, phosphinothricin, and dehydrophos, emerging pathways are also presented, which are known primarily at a genetic level but are predicted to contain exciting enzymology. The resistance mechanisms to bioactive Pn such as fosfomycin and phosphinothricin are also discussed due to the importance of such chemistry to therapeutic and agricultural applications. Finally, the reader is encouraged to supplement this review with others that have been written on various aspects of Pn biochemistry, including biosynthesis;8−10 catabolism;10−14 the global distribution and impact of catabolic pathways on the Pi cycle;15−19 the discovery of new pathways and molecules;20,21 phosphonolipids and phosphonoglycans;22−25 pathway regulation and genetics;14,26

2. OCCURRENCE OF ORGANOPHOSPHONATES IN NATURE 2.1. Why Nature Chose Phosphonates

As a functional group the phosphonate in its ionized form (pKa2 ≈ 7−8)34 can mimic a phosphate monoester or carboxylate of a metabolic intermediate (Figure 3A). It can also mimic the tetrahedral geometry and negative charge development found in the transition state (or intermediate) for an associative acyl transfer reaction (Figure 3B).35−40 Moreover, the C−P bond of a phosphonate is a chemically and thermally stable analog of a P−O bond. In fact, this stability led to the discovery of the first Pn natural product, 2-aminoethylphosphonic acid 3: unlike phosphate esters, 3 stood out for its resistance to hydrolysis by strong acids and even combustion during elemental analysis.22 It is important to note that based on bond dissociation energies 5707


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Figure 4. Structures of common Pn decorated macromolecules and metabolites.

(BDEs) a C−P bond of a phosphonate is 70 to 100 kJ mol−1 weaker than the corresponding O−P bond of a phosphate ester (Figure 3C),41,42 reflecting that the former bond is less polarized than the latter. However, BDEs reflect homolytic bond cleavage, and if one considers heterolytic cleavage the C− P bond is considerably more stable. There are three reasons that contribute to the stability of Pn to heterolytic cleavage arising from nucleophilic attack at the phosphorus center (Figure 3D):43 (1) a preference for a less electronegative carbon-based substituent to occupy the equatorial position of a pentacoordinate phosphoranyl intermediate (or transition state) upon attack by a nucleophile, rather than the required apical position for a leaving group44 (as shown in Figure 3D there is a 8 kcal mol−1 preference for a methoxy group to assume an apical position over a methyl group43); (2) the poor leaving group ability of a carbanion relative to an alkoxide; and (3) the lack of lone pair electrons on the phosphorus bound carbon atom that prevents Lewis or general acid stabilization of a carbanion leaving group by an enzyme. For these reasons

chemists have long used Pn in the design of enzyme inhibitors and mechanistic probes,45−54 with some of these highlighted in Figure 2. 2.2. Abiotic Synthesis of Pn

While Nature has devised biological strategies for the biosynthesis of Pn, it is important to note that simple Pn can be produced spontaneously. Significant yields of hydroxymethylphosphonic acid 35 (20−30%) and 1-hydroxyethylphosphonic acid 36 (1−3%) are obtained by simple UV photolysis (185 nm) of formaldehyde in the presence of phosphite 34 (Scheme 1A).55 The analogous reaction with acetone yields methylphosphonic acid 8 (17%) (Scheme 1B). Such products are proposed to arise by the generation of phosphonyl and carbon centered radicals. 34 is a reduced and reactive form of phosphorus that can be obtained by the hydrolysis of nickelphosphides, such as Schreibersite [(Fe,Ni)3P] 37 (Scheme 1C), which can be found in meteorites and the Hatutruim Formation near the Dead Sea.56 Such chemistry may explain 5708


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Bacteroides and Treponema.72 Genome scanning led to the identification of methylphosphonic acid 8 in the exopolysaccharide of the marine bacterium Nitrosopumilus maritimus.62 The function of 8 is not known, but it is ultimately a major source of methane production by the world’s oceans through the action of carbon−phosphorus lyase. Genome scanning also led to the identification of 4 as a component of the exopolysaccharides in Stackebrandtia nassauensis, Glycomyces sp.,67 and possibly Bacteroides sp.72 3 is also frequently found in O- and N-linked glycoproteins.83 Early research observed that the sea anemone Metridium dianthus contained a large quantity of a glycoprotein that was decorated with a derivative of 3.84 The freshly laid eggs of the freshwater snail Helisoma contain 3 as a component of phosphonoglycoproteins, representing 85% of the total phosphorus.85 A mucin isolated from the jellyfish Aurelia aurita was shown to contain O-glycosylated peptide 46,86 while Nlinked oligosaccharides like 47 are present in the common locust.87 The marine snail Volvarina rubella contains several complex N- and O-linked glycans where 3 and its N-methylated derivative are observed to be attached to galactosamine, mannosamine, and N-acetylglucosamine, often multiple times on the same polysaccharide, and differing in methylation pattern.88 Glycomics workflows involving glycosidase treatment, chromatography, and tandem MS/MS analysis have accelerated the process of solving these phosphonoglycan structures.83 The distribution and abundance of Pn macromolecules among organisms varies widely with species, tissue, or cellular location. While vertebrates have sphingomyelin as the primary sphingosylphospholipid, invertebrates frequently contain high levels of 40.24 Similarly, phosphonoglycocerebrosides like 42 tend to be localized in the nervous tissue of the sea slug Aplysia kurodai.76−78 The parasitic protozoan Trypanosoma cruzi, the causative agent of Chagas’ disease and found primarily in South America, contains phosphonolipids, while Trypanosoma brucei, found primarily in Africa, does not.89 The degree of Nmethylation can vary: mollusks and coelenterata contain primarily 40a, while the marine snail Turbo cornutus is rich in 40b.24,63 Even sex can play a role: the gills of the male blue crab contain 3, while the female crab does not.2 Despite the number of Pn containing macromolecules that have been identified, little is known about their function or how they are biosynthesized. In principle phosphonolipids like 39 would resist the action of mammalian phospholipase D, perhaps explaining their prominence in microorganisms like T. pyriformis that inhabit the rumen of cattle. Although the genes and enzymes necessary for synthesizing 3,90,91 4,67 and 862 are known (and will be discussed later in this review), the enzymes that attach these Pn to lipids and glycans in a regiospecific manner, or control the N-methylation state of 3, are not. The addition of phosphocholine 38 to glycans in bacteria requires activation of the phosphoryl group by the cytidyltransferase LicC followed by transfer of 38 to a specific glycan hydroxyl group by the phosphocholine phosphotransferase LicD.82 S. pneumoniae has two LicD homologues that likely modify different GlcNAc units in the exopolysaccharide sequence. Biosynthetic loci for Pn decorated bacterial exopolysaccharides have been identified,62,67,72,92 but the functions of the genes encoding Pn attachment and modification remain to be determined. It is not clear if bacteria that synthesize phosphonoglycans simply rely on the

the presence of methylphosphonic acid 8, as well as ethyl, propyl, and butylphosphonic acids in the Murchison meteorite.57 Remarkably, even nucleotide monophosphates, which comprise the building blocks of genetic material, can be synthesized by heating aqueous mixtures of nucleosides with 37.58 It has been proposed that such abiotically derived Pn may have served as a form of soluble and accessible phosphorus for early life forms on Earth.59,60 2.3. Phosphonolipids and Phosphonoglycans

Several of the simple Pn shown in Figure 1, inner circle, are known to be incorporated into lipids and polysaccharides (Figure 4). These Pn decorated macromolecules have been isolated from a startling array of organisms, including humans, ruminants (e.g., cows and goats), sea anemones, freshwater and marine bivalves, snails, cephalopods, oysters, abalone, protozoans, bacteria, and fungi61 (for reviews see refs 22−25). The best analytically characterized examples include methylphosphonic acid 8,62 2-aminoethylphosphonic acid 363,64 and its 1hydroxy derivative 5,65,66 and 2-hydroxyethylphosphonic acid 4.67 3 is the most commonly observed Pn due to its structural similarity to phosphocholine 38 (Figure 4) and thus occurs as a polar headgroup in lipids to form phosphonoglycerides 39 and sphingosylphosphonolipids 40. Variation in the degree of methylation of the 2-amino group in 39 and 40 is often observed. Some organisms can accumulate astonishingly high levels of phosphonolipids. For example, 3 was discovered in a rumen protozoan.68 This was likely Tetrahymena pyriformis, where up to 30% of the total cellular lipid is in the form of 39a,69 40a, and the N-methyl derivative 40b.70 The Pn level rises to 60% in the membrane sheath of the cilia, for which 3 was given that name ’ciliatine’.71 5 appears in 41 as a component of the membranes of the Bdellovibrio stolpii, a Gram-negative bacterium that preys upon other bacteria.66 Gene clusters predicted to encode phosphonolipids containing 5 have been identified in several bacterial species, particularly Burkholderia.72 Interestingly, the freshwater cyanobacterium Aphanizomenon f los-aquae, which forms blooms in lochs of Scotland, produces a surfactant 48 where the 2-hydroxyl group of 4 is esterified with a fatty acid.73 The the reef building coral Acropora digitifera may also produce 3 as genes encoding its biosynthesis have been identified from genomic data.74 Pn are also frequently used to decorate the glycans of phosphonoglycocerbrosides and bacterial exopolysaccharides.24 Such Pn macromolecules are common in marine organisms. This was recognized early on in the common amoeba Acanthamoeba castellanii where up to 70% of the phosphoglycocerebroside phosphorus arises from 3 and its 1-hydroxy derivative 5.65 The sea slug Aplysia kurodai is a source of several complex phosphonoglycocerebrosides such as 42 that display multiple copies of 3.75−79 Phosphonoglycocerbrosides such as 43 are also found in sea snails, but in this organism 3 is Nmethylated.24,63 A novel N-hydroxymethylated derivative of 3 was identified in the exopolysaccharide 45 of the ruminal bacterium Fibrobacter succinogenes S85.80 The exopolysaccharide 44 is the virulence factor for the human pathogenic bacterium Bacteroides f ragilis.81 Although it is not known if 3 is essential for virulence, it is well-known that functionalization of glycans with 38 is critical for several pathogens including Haemophilus inf luenzae and Streptococcus pneumoniae.82 Genome sequencing has accelerated the identification of new Pn decorated macromolecules. Glycans esterified with 3 are predicted to be encoded by 27 gene clusters found in 5709


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includes the alkylphosphonate esters dehydrophos 11 and phosphoiodyn A 16. The methyl ester is important for the bioactivity of 1195,96 while the Pn bond of 16 is essential for its activity against the hPPARγ transcription factor.4,5 The phosphonoamide bond has also been sampled in the fosfazinomycins 13a/b in the form of a hydrazide linkage to phosphorus. Finally, there are phosphinic acid secondary metabolites with C−P−C bonds in the form of phosphinothricin 10a and its amino acid derivatives. While many of these compounds bind noncovalently to their targets, 9 is notable for covalently inactivating EPSP synthase by virtue of the epoxide ring, while potential for covalent modification of a biological target is present in the α-haloketone of fosfonochlorin 20. A common theme in natural product chemistry is the creation of “hybrid” molecules through assembly of different building blocks.97 In this way it is possible to have, for example, nonribosomal peptides combined with polyketides and glycosides. The Pn natural products also follow this theme. A common modification for several of Pn secondary metabolites is incorporation of the phosphonate moiety into amino acids and peptides through nonribosomal peptide bond synthesis (Figure 1).95,98 As Pn are inherently polar molecules, this enhances uptake of the molecule by the targeted cell. The peptide moiety can also serve as a Trojan horse whereby peptide bond cleavage inside the cell releases the active molecule. This is the case for 10b-d, which release phosphinothricin 10a; 14b−h, which release APPA 14a; and 11, which is a source of the methyl acetylphosphonic acid 52, an inhibitor of pyruvate dehydrogenase, pyruvate oxidase, and 1-deoxy-D-xylulose 5-phosphate (DXP) synthase.95,96,99 The structure of 16, isolated from a marine sponge, also indicates that Pn have been combined with polyketides.4,5 Like synthetic Pn, the bioactivity of natural product Pn can often be traced to the mimicry of the compound to an enzyme substrate (Figure 5). 9 targets the active site of EPSP synthase as a mimic of phosphoenolpyruvate 34. The phosphinate moiety of 10a mimics the tetrahedral intermediate 51 formed by glutamine synthase. Methylacetylphosphonate 52 is a mimic of pyruvate 53; the former is spontaneously generated following peptide bond cleavage of 11. This accounts for the inhibition of the thiamin dependent enzymes that use pyruvate as a substrate, including pyruvate dehydrogenase, pyruvate

promiscuity of LicC or LicD homologues, substituting 3 for 38 (Scheme 2), or if there are enzymes that are specific for Pn. Scheme 2

Enzyme promiscuity certainly explains the occurrence of Pn macromolecules in eukaryotes. Such organisms do not have the capacity to synthesize 3 but instead acquire this compound from food or gut microflora.23 For example, the rumen protozoan Tetrahymena pyriformis has the ability to synthesize 3.90,91 Accordingly, ruminants can acquire 3 for incorporation into lipids alongside 38 due to the relatively relaxed specificity of the enzymes involved. This can also be performed artificially: the larvae of the common housefly can incorporate 3 into phosphonolipids when fed this compound.93 This occurs through the Kennedy pathway for phospholipid biosynthesis following chemical logic shown in Scheme 2: 3 is activated as the cytidine monophosphate by CTP:phosphocholine cytidyltransferase, followed by transfer to diacylglycerol by choline/ ethanolamine phosphotransferase to form the phosphonolipid.23 Analogous enzyme promiscuity can account for the occurrence of the bile acid 49 in gall bladders of cows, where 3 replaces the normal substrate taurine 50.94 2.4. Bioactive Pn Secondary Metabolites

Beyond incorporation of simple Pn into macromolecules, a large number of complex and bioactive Pn secondary metabolites are biosynthesized by bacteria and fungi. As shown in Figure 1, the majority of these molecules are alkylphosphonic acids (RPO3H2) such as fosfomycin 9. However, Pn secondary metabolites are notable for having explored functionalization of the phosphonic acid group. This

Figure 5. Examples of Pn natural products that inhibit enzymes. The enzyme target of each Pn is given in parentheses, while the substrate of target enzyme is shown below in blue. The peptidase cleavage site on 11 is shown with a dashed line. 5710


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Scheme 3. Overview of Pn Secondary Metabolite Biosynthesisa

Inset: Compounds of “unknown provenance” likely arise from an alternative mechanism for forming the CP bond. Enzyme cofactors are given in parentheses. Abbreviations: PEP mutase, phosphoenolpyruvate mutase; Ppd, phosphonopyruvate decarboxylase; MpnS, methylphosphonate synthase; PLP, pyridoxal phosphate; TPP, thiamin pyrophosphate.

a

oxidase, and DXP synthase.95,96,99 The latter bacterial enzyme catalyzes the first step of the nonmevalonate pathway for isoprenoid biosynthesis, forming deoxyxylulose phosphate 54. Intriguingly, the nonmevalonate pathway is also the target of FR-900098 12a, FR-31564 (fosmidomycin) 12b, and FR-32863 12c. These related compounds inhibit the second enzyme on the pathway, DXP reductoisomerase, as mimics of the substrate 54. Finally, the molecule (Z)-L-2-amino-5-phosphono-3pentenoic acid 14a inhibits the pyridoxal phosphate dependent enzyme threonine synthase as a mimic of homoserine phosphate, where the labile phosphoryl O−P bond has been replaced by a C−P bond.100

reactions in turn lead to building blocks that can be used in divergent ways to create Pn natural products. It must be noted that one other mechanism for synthesizing a C−P bond is currently known, that of P-methyl transferase, a cobalamindependent methyl transferase that transfers a methyl group to a phosphinate nucleophile. This enzyme will be discussed later in the section on the biosynthesis of 10a (section 4.2.11). 3.1. Phosphonopyruvate (1)

The thermodynamic bias for 24 over 1 frustrated early attempts to identify PEP mutase. A breakthrough came when the laboratories of Dunaway-Mariano, Knowles, and Seto realized that 1 was the better substrate to follow PEP mutase activity, as conversion of 1 to 24 is thermodynamically downhill. This allowed the activity guided isolation of PEP mutase from the rumen protozoan Tetrahymena pyriformis,91,101 the source of the first discovered naturally occurring Pn 3.68 PEP mutase was also isolated by Seto from Streptomyces hygroscopicus, the producer of 10b, using a similar activity assay.103 The T. pyriformis enzyme has kinetic parameters of kcat = 5 s−1 and KM = 770 μM for the forward reaction converting 24 to 1.104 In the reverse direction converting 1 to 24 the parameters are kcat = 100 s−1 and KM = 3.5 μM. The enzyme requires Mg2+ for activity (KM = 6 μM) but is also active with Mn2+, Co2+, and Zn2+. Binding of the metal ion precedes 1 in an ordered fashion.105 PEP mutase is a founding member of the PEP mutase/ isocitrate lyase superfamily (cd00377) that is comprised of enzymes that catalyze either the formation or cleavage of C−C

3. BIOSYNTHESIS OF PN BUILDING BLOCKS The primary reaction that forms the C−P bond of Pn natural products is catalyzed by phosphoenolpyruvate mutase (PEP mutase). This enzyme catalyzes the equilibrium between 24 and phosphonopyruvate 1 (Scheme 3). Due to the greater bond dissociation energy of the O−P bond in 24 versus the C− P bond in 1 (17−24 kcal mol−1),41,42 the equilibrium favors 24 by a factor of at least 500.101 In order for the PEP mutase reaction to be biosynthetically useful, a biosynthetic pathway must overcome the unfavorable equilibrium between 24 and 1. This is achieved by coupling the formation of 1 with an irreversible reaction. As outlined in Scheme 3, several strategies have evolved, including decarboxylation, transamination, and aldol reactions. In the case of valinophos 18, the formation of 1 may be coupled to reduction or phosphorylation.102 These 5711


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Scheme 4. Phosphonopyruvate Mutase (PEP Mutase)a

a

(A) Active site scheme of PEP mutase bound to phosphoenolpyruvate 24. Boxed residues belong to the loop that closes over the active site upon substrate binding. (B) Active site scheme of PEP mutase bound to phosphonopyruvate 1. (C) Stepwise mechanism for PEP mutase involving a metaphosphate intermediate. Isotopes are shown to highlight the stereochemistry of this reaction (18O, full circle; 17O, circle with line; 16O, clear circle). (D) Alternative mechanism for phosphoryl transfer involving a dissociative transition state with concerted bond cleavage and formation.

mechanisms were proposed using either a substrate oxygen or an enzyme side chain as a nucleophile to account for retention of stereochemistry.109,112,114 However, it was not possible to trap a covalent phospho-enzyme intermediate,104 and the active site of PEP mutase did not contain a side chain close enough to the phosphoryl group of 1 to act as a nucleophile in a doubledisplacement mechanism.107 D58, H190, and R159 were additionally ruled out as possible nucleophiles as the R159A, D58N, and H190A variants were still weakly active.107 Based on these results Dunaway-Mariano and Herzberg proposed that the reaction proceeds through a highly dissociative mechanism involving a metaphosphate intermediate (Scheme 4C).107 Tight anchoring of metaphosphate by N122, R159, and H190 would prevent movement and force the same face of this trigonal planar species to form and break bonds with C3 and the C2 oxygen, leading to retention of configuration. During the lifetime of this intermediate, it was proposed that rotation about the C1−C2 bond of the substrate would allow the exchange of C2 enolate oxygen and C3 carbanion nucleophiles. Rotation around the C1−C2 bond and a highly dissociative phosphoryl transfer reaction is also supported by computational modeling of the PEP mutase reaction.115 However, while phosphoryl transfer reactions with dianion monophosphate esters like 24 are known to proceed through highly dissociative transition states, little evidence exists for transfer involving a discrete metaphosphate intermediate.116,117 Therefore, a more conservative view of the PEP mutase mechanism would involve a highly dissociative, metaphosphate-like transition state with concerted bond cleavage and formation (Scheme 4D).

or C−P bonds. This family also includes phosphonopyruvate hydrolase (PPH), carboxy PEP (CPEP) mutase, and oxaloacetate hydrolase. X-ray crystal structures of PEP mutase from the blue mussel Mytilus edulis were solved by Hertzberg with the enzyme in free form and bound to sulfopyruvate (KI = 22 μM), a nonreactive analog of 1.106,107 This enabled the creation of models of the binding modes of 24 and 1 in the active site of PEP mutase (summarized schematically in Scheme 4A and B, respectively). The phosphoryl group of the substrate is tightly held by the side chains of R159, H190, and N122 as well as shielded from solvent by L124. A single hexacoordinate Mg2+ ion is bound by three structural water molecules and D85. The substrate binds to the final two ligand sites of Mg2+ in a bidentate mode using oxygen atoms at C1 and C2. Substrate binding is accompanied by closure of a loop over the active site that brings into position K120, N122, and L124. 106 Substitutions made at R159, D58, H190, L124, and N122 produced PEP mutase variants that were either inactive with 1 as substrate or exhibited drastically diminished kcat values and substantially increased KM values.106,107 Earlier work using isotopically labeled 1 and 24 established the stereochemical course of the PEP mutase reaction. Using (S)- and (R)-[16O,17O,18O] labeled 1, Knowles determined that the phosphoryl group was transferred with retention of stereochemical configuration.108,109 The same conclusion was reached by Dunaway-Mariano using (S)- and (R)-[16O,18O]thiophosphonopyruvate.110−112 By labeling 24 at C3 with a deuterium atom, Hammerschmidt demonstrated that the phosphoryl group is transferred from the C2 oxygen to the si face of C3.113 Various double-displacement phosphoryl transfer 5712


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Scheme 5. Phosphonopyruvate Decarboxylase (Ppd)a

a

(A) Model of the active site of Ppd bound to thiamine pyrophosphate and 1. (B) Proposed mechanism for Ppd.

Scheme 6. 2-Aminoethylphosphonic Acid (3): Pyruvate Aminotransferasea

a

(A) Proposed mechanism for the aminotransferase catalyzing the equilibrium between 3 and L-Ala. (B) Active site scheme for the S. typhimurium aminotransferase (PDB ID: 1M32).131

Mg2+-dependent enzymes that include benzoylformate decarboxylase, pyruvate decarboxylase, and acetolactate synthase.121 Two Ppd homologues have been heterologously produced in E. coli for characterization in vitro. Bacteroides f ragilis Ppd is from the biosynthetic pathway that forms the phosphonoglycan 46. This enzyme is a homotrimer in solution and requires TPP and a divalent cation for catalysis, either Mg2+, Mn2+, or Ca2+, with greater specificity exhibited with Mn2+. In an assay containing a mixture of Mg2+ and Mn2+ the B. f ragilis Ppd converts 1 to 2 with the kinetic parameters kcat = 10.2 s−1, KM = 3.2 μM, kcat/

3.2. Phosphonoacetaldehyde (2)

The conversion of 1 to phosphonoacetaldehyde 2 is catalyzed by phosphonopyruvate decarboxylase (Ppd). Ppd was first identified and isolated by Seto from the 10b producer Streptomyces hygroscopicus.118 The isolated enzyme required Mg2+ and thiamine pyrophosphate (TPP) for activity. Shortly after the gene encoding Ppd was identified by Wohlleben in another 10b producer, S. viridochromogenes Tü494119 followed by Seto in S. hygroscopicus.120 By amino acid sequence Ppd comprises one of nine subfamilies within a superfamily of TPP/ 5713


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KM = 3.2 × 106 M−1 s−1.122 Slow turnover is also observed with pyruvate and sulfopyruvate as substrates. By sequence homology to TPP-dependent enzymes with known structures, D260 was predicted to be involved in binding the metal ion, which in turn is used to bind the pyrophosphate group of TPP. The D260A variant had a 1000-fold reduced kcat value, confirming its importance to catalysis. A neighboring residue, D258, is conserved only in the Ppd subfamily. The corresponding D258A variant had a 1000-fold larger KM value for 1, suggesting a role in substrate binding. An X-ray crystal structure for Ppd has not yet been solved. However, an active site model of the S. viridochromogenes Tü494 Ppd was proposed based on sequence homology to pyruvate decarboxylases with known structures (Scheme 5A).123 This allowed the prediction of residues important for metal ion binding (D265, N293, G294), generation of the cofactor ylid (E48), and substrate binding/catalysis (S25, H110, D297). The importance of each of these residues was confirmed through active site substitutions. D263 of S. viridochromogenes Tü494 Ppd is conserved with D258 of the B. f ragilis enzyme. The D263A variant was inactive using Mg2+, but regained partial activity with Mn2+ as the metal ion, suggesting that this residue may in fact be involved in metal ion binding rather than direct substrate binding. The mechanism of Ppd likely follows a classical TPP-dependent decarboxylation where protonation of the enamine-carbanion intermediate leads to formation of the aldehyde 2 (Scheme 5B). Residues mediating general acid−base catalysis and recognition of the phosphonate group remain to be determined. Likewise, it remains to be seen whether Ppd exhibits promiscuity toward electrophilic acceptors other than a proton, such as aldehydes or ketones, which is commonly seen in TPP-dependent decarboxylases.124 Such acyloin-like reactions with 1 would have intriguing synthetic applications.

aeruginosa enzyme, is highly specific for the reverse reaction of 3 and 53 or the forward reaction of 2 and L-Ala.130 The reaction has Keq = 0.5 and kinetic parameters of kcat (forward) = 7 s−1, kcat (reverse) = 9 s−1, KM (3) = 1.11 mM, KM (53) = 0.15 mM, KM (2) = 0.09 mM, KM (L-Ala) = 1.4 mM. Very low activity is observed with L-Asp or D-Ala as alternative amino donors and α-ketoglutarate as an amino acceptor. Substitution of amino acid residues that are conserved in aminotransferases identified D168, K194, R340 as critical for catalysis. The structure of the S. typhimurium aminotransferase bound to PLP and 2 was solved by the Hertzberg lab.131 This allowed creation of a model of the Schiff base intermediate formed between 2 and PLP (Scheme 6B). The phosphonate moiety is recognized by a polar pocket created by S65, G66, and S67. The dimeric strucutre of the aminotransferase appears to play a role in substrate recognition as T243 of one monomer reaches into the active site of the other to provide an additional polar interaction with the phosphonate. K194 is appropriately positioned to form a Schiff base with PLP, as well as a general acid/base catalyst during the transamination reaction. The orientation of 2 as the aldimine orients the C2 pro-S hydrogen for abstraction by K194, while the pro-R hydrogen is protected by a pocket formed by P11 and Y329. Interestingly, the model of corresponding L-Ala-PLP Schiff base indicated that the αcarboxylate of L-Ala would collide with P11 and Y329, suggesting that a conformational change involving these residues would be required to accommodate this substrate. Y329 was also highlighted as a key residue for dictating the specificity for pyruvate over other α-ketoacids in the reverse reaction. 3.4. 2-Hydroxyethylphosphonic Acid (4)

Reduction of 2 by an NAD(P)H-dependent oxidoreductase yields 2-hydroxyethylphosphonic acid 4. This is a key intermediate in the biosynthesis of methylphosphonic acid 8, fosfomycin 9, phosphinothricin 10a (and its peptide derivatives like bialaphos 10b), dehydrophos 11, and phosphonoglycans (Scheme 3). The biosynthesis of 9 in a Streptomyces wedmonrensis fomC mutant could be restored by provision of 4 but not 3, the latter a cellular equivalent to 2 as a result of the nonspecific activity of PLP-dependent transaminases.132 This suggested 2 was converted to 4 by FomC. Likewise, the phpC mutant of Streptomyces viridochromogenes is deficient in the biosynthesis of 10b and instead accumulates 3 by way of the accumulation of 2.133 FomC and PhpC belong to a family of metal ion dependent alcohol dehydrogenases (Pfam # PF00465) that include E. coli 1,2-propanediol oxidoreductase FucO, Zymomonas mobiliz alcohol dehydrogenase II, and S. cerevisiae ADH4.134,135 These dehydrogenases often require an active site Fe2+ ion for activity but members that are dependent on Zn2+ are also known.136 FomC, PhpC, as well as DhpG from the pathway for 11 in Streptomyces luridis have been shown to convert 2 to 4 in vitro.133,137 Interestingly, the enzymes differ in their cofactor specificities: FomC and DhpG require NADPH and Fe2+ for activity, while PhpC is NADH and Zn2+ dependent. Significant differences are observed in kcat/KM values for the conversion of 2 to 4, with FomC showing the greatest efficiency (4.1 × 104 M−1 s−1) followed by PhpC (2.2 × 103 M−1 s−1) and DhpG (1.1 × 103 M−1 s−1). The kcat values are similar (0.4 to 1.3 s−1); therefore, the differences in efficiency are primarily due to differences in KM for 2, with FomC having the lowest value (31 μM), followed by PhpC (185 μM) and DhpG (414 μM). All

3.3. 2-Aminoethylphosphonic Acid (3)

3, also known as ciliatine, is the most commonly found Pn in Nature and can be regarded as the genesis of Pn natural product chemistry with its discovery in 1959 by Horiguchi and Kandatsu in sheep rumen.68 3 is generated from 2 by a pyridoxal phosphate (PLP) dependent aminotransferase (EC 2.6.1.37) that uses pyruvate 53 and L-alanine as the amino acceptor and donor, respectively (Scheme 6A). Because this reaction is reversible, this enzyme is found in Pn biosynthetic and catabolic pathways. The aminotransferase was first detected by Nauze in cell free extracts of Bacillus cereus as the first step in a pathway for degrading 3 into Pi.125 Later, a homologue of this enzyme was successfully purified from Pseudomonas aeruginosa and shown to be exquisitely specific for 3 and 53, converting the former to 2.126 The aminotransferase was also completely inhibited by aminooxyacetate, a nucleophile that reacts with the Schiff-base linkage that is formed between the enzyme active site lysine and PLP. Using (R) and (S)-[2-2H]-3 as substrate, it was shown that the P. aeruginosa aminotransferase removes the pro-S hydrogen from C2 of 3.127 This step is rate limiting as a kinetic isotope effect was also observed with (S)-[2-2H]-3. The gene phnW encoding the 3-specific aminotransferase was identified by Wanner in the pathogenic bacteria Enterobacter aerogenes128 and Salmonella enterica serovar Typhimurium LT2.129 In both bacteria phnW appears in a cluster of genes encoding the catabolism of 3 by the phosphonatase mechanism discussed later in this review.130 Heterologously produced PhnW is a homodimer in solution, and like the P. 5714


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homology with enzymes that perform α-hydroxylation of structurally similar substrates. This includes E. coli TauD (αhydroxylation of taurine 50),140,141 FzmG from the 13a−b pathway in Streptomyces sp. WM6372 (α-hydroxylation of 7),142 and DphA from the 11 pathway in Streptomcyes luridis (αhydroxylation of 4). A PhnY* homologue is also predicted to be encoded in the biosynthetic gene cluster for 19b.102 PhnY* has been shown in vitro to catalyze the stereospecific αhydroxylation of 3 to form (R)-5.143,144 As an α-ketoglutarate/ nonheme iron dioxygenase, PhnY* is proposed to catalyze hydroxylation using a highly reactive ferryl oxygen (Fe(IV) O) (Scheme 7).141 Such enzymes use an active site Fe(II) ion to bind α-ketoglutarate 58 followed by molecular oxygen. The latter is reduced to an iron-bound superoxide species (Fe(III)− O-O•−), which in turn catalyzes the oxidative decarboxylation of α-ketoglutarate 58 to succinate 59, thus driving the formation of Fe(IV)O. The PhnY* Fe(IV)O species is then proposed to abstract the α-hydrogen of 3 to form a carbon-centered radical. Rebound of the Fe(III) bound hydroxide with the radical leads to (R)-5 and regeneration of Fe(II). Formation of a Fe(IV)O intermediate and subsequent C−H bond cleavage has been directly observed in the analogous TauD reaction.145,146 A structure has yet to be determined for PhnY, FzmG, or DphA, which would be highly useful to provide greater insight into the stereo and substrate specificity of these related enzymes.

three enzymes are specific for 4 in the reverse reaction, with no activity observed with hydroxymethylphosphonic acid 35 or 3hydroxypropylphosphonic acid. A homology structural model for each enzyme was created using E. coli FucO and Thermotoga maritima dehydrogenase as templates, allowing the identification of key active site residues (Figure 6).137 Despite differences in metal ion specificity, all

Figure 6. Partial active site scheme for phosphonoacetaldehyde dehydrogenase FomC (GenBank accession ACG70833) based on a structural alignment with E. coli lactaldehyde reductase FucO (PDB ID: 5br4). FomC residues shown in bold and aligned FucO residues given in parentheses.

three enzymes use the same four amino acids for metal ion coordination (D189, Q193, H257, and H271 in FomC). The fifth ligand site is predicted to accept the substrate C2 oxygen, thereby increasing the electrophilicity of the carbonyl of 2. Although iron prefers to be hexacoordinate, a sixth ligand is not predicted for FomC or DhpG, nor is it observed for FucO.138 Although the stereospecificity of reduction of the carbonyl of 2 is not known, the homology model predicts that the pro-R hydride of the nicotinamide ring is delivered during reduction.132 Two residues, Y261 and S157, are conserved in FomC, PhpC, and DhpG. These are predicted to hydrogen bond to the phosphonate oxygens and appear to be unique in homologues of these enzymes that appear in Pn biosynthetic pathways.137 For this reason genes encoding dehydrogenase sequences with these residues can be used to identify Pn biosynthetic gene clusters in microbial genomes.

3.6. Phosphonoalanine (6)

6 is one of the most commonly occurring natural Pn. This compound has been identified in a variety of simple organisms including the rumen protozoan Tetrahymena pyriformis,147,148 sea anemones,147,149 and mussels.149 6 is formed reversibly from 1 through the action of a transaminase. This enzyme has not yet been characterized in vitro, although palB encoding this activity has been identified by Quinn in a gene cluster encoding the degradation of 6 as a source of Pi, carbon, and nitrogen in Variovorax sp.150 PalB is pyridoxal phosphate (PLP) dependent enzyme that is related by sequence to the aspartate aminotransferase superfamily (cl18945) that catalyzes transfer of ammonia from an amino acid donor to an α-ketoacid acceptor.151 This enzyme appears to be specific, as Variovorax sp. can only catabolize the L-enantiomer of 6.152

3.5. 1-Hydroxy-2-Aminoethylphosphonic Acid (5)

3.7. Phosphonoacetate (7)

5 has been observed as a polar headgroup in phosphonolipids such as 41 and is possibly a precursor to hydroxynitralophos 19b (Figure 4 and Scheme 3).65,66,102 An enzyme that hydroxylates the α-carbon of 3 to form 5, PhnY*, was first identified in a pathway for catabolizing 3 into Pi and glycine (discussed later in this review).139 PhnY* belongs to the αketoglutarate/nonheme iron dioxygenase superfamily, thus the asterix is used to distinguish this enzyme from the NAD+ dependent dehydrogenase PhnY.16 PhnY* shares sequence

The aldehyde of 2 can be further oxidized to 7, which is a substrate for catabolism into Pi, or an intermediate in the biosynthesis of fosfazinomycins 13a−b. The conversion of 2 to 7 is most often seen in the catabolic context, as discussed in more detail in section 5.1.3. The phnY gene encoding this transformation was first identified in the genome of Sinorhizobium meliloti.153 The gene appeared in a phnWAY locus encoding the catabolism of 3 to Pi, with phnW and phnA encoding the aminotransferase and phosphonoacetate hydro-

Scheme 7. Proposed Mechanism for the Dioxygenase PhnY*

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Scheme 8. Phosphonoacetaldehyde Dehydrogenase PhnYa

a

(A) Active site scheme for S. meliloti PhnY (PDB ID: 4i3u)154 and proposed mechanism. (B) Alternative substrates for S. meliloti PhnY.

lase activities, respectively. Purified PhnY was shown to oxidize 2 to 7 by 31P NMR spectroscopy with a dependency on NAD+ and Zn2+ for catalysis. Kinetic parameters for this reaction were kcat = 2 s−1, KM (2) = 2.6 μM, KM (NAD+) = 104 μM, and kcat/ KM (2) = 7.5 × 105 M−1 s−1. X-ray crystal structures of S. meliloti PhnY bound to 2, 7, and NAD+ provided remarkably detailed insight into the oxidative mechanism.154 2 is observed to bind covalently to the active site of PhnY as the (R)-thiohemiacetal via the thiol side chain of C291, with the alkoxide stabilized by an oxyanion hole created in part by N158 (Scheme 8A). The importance of C291 serving a role in covalent catalysis was supported by the lack of activity observed with a PhnY C291A variant. The negatively charged phosphonate group is bound by a positively charged pocket created by H159, R108, R290, and R447. The side chain of E254, a putative general base, and the nicotinamide ring of the cofactor compete for the same space in the active site, and are observed in different orientations depending on whether 2/ NAD+, 7/NAD+, or 2 alone are bound. When the nicotinamide ring of NAD+ is in the active site, the re face of C4 is positioned appropriately to accept a hydride from the thioacetal to form a thioester intermediate and NADH. When the nicotinamide ring swings out of the active site (but remains bound to PhnY by virtue of the nucleotide portion of the cofactor), the side chain of E254 is observed to enter the active site in a position appropriate to deprotonate water, thus catalyzing deacylation. A role for general base catalysis is supported by the lack of activity observed for the PhnY E254A variant, as well as accumulation of the acyl-enzyme intermediate as detected by ESI-MS.

PhnY belongs to the nonphosphorylating aldehyde dehydrogenase (ALDH) superfamily. This family includes glyceraldehyde-3-phosphate dehydrogenase (GAPDH) from the glycolysis pathway, and the phosphonoformaldehyde dehydrogenase PhpJ from the phosphinothricin 10a pathway (described later in this review). Remarkable active site structural homology is observed between PhnY and Streptococcus mutans GAPDH (PDB ID: 2ESD). The key active site residues of the two enzymes are identical, with the exception that H159 of PhnY appears as Y155 in S. mutans GAPDH. This structural homology translates to catalytic promiscuity in PhnY. The “longer” substrate 60 (Scheme 8B) was was oxidized to the corresponding acid with a similar kcat value (1.5 s−1) to that for 2, but with a 1000-fold increase in KM (3300 μM). Even 61, the substrate for GAPDH, is slowly oxidized by PhnY (kcat = 0.1 s−1, KM = 97 μM). In contrast, GAPDH is weakly active toward 60, but inactive toward 2. Such crossover of activities strengthens the idea that a number of biosynthetic Pn reactions evolved from their metabolic analogs.9 7 is also a precursor to the fosfazinomycins 13a−b (Scheme 3). However, in this case, 7 appears to be synthesized by the promiscuous action of an α-ketoglutarate/nonheme iron dioxygenase FzmG in the pathway for 13, discussed in section 4.5.155 The primary biosynthetic role for FzmG is stereospecific hydroxylation of the α−carbon of methyl phosphonoacetate 7b to form (S)-62 (Scheme 9A). However, FzmG will also convert 2 to 7 (Scheme 9B). Intriguingly, the desmethyl version of 62, 1-hydroxyphosphonoacetate, is predicted to be encoded by gene clusters in several species of Burkholderia, but in this case as the polar headgroup of a phosphonolipids.72 5716


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Scheme 9. Reactions Catalyzed by the Dioxygenase FzmG in the Biosynthesis of Fosfazinomycins (13a−b)

Scheme 10. Reactions Catalyzed by Methylphosphonate Synthase MpnS with Substrate Analogs

3.8. Methylphosphonic Acid (8)

The simplest Pn is best known as the origin of the “ocean methane paradox”.156 While 8 can be produced through abiotic chemistry (Scheme 1), a biological pathway was proposed by Karl to explain the occurrence of supersaturating concentrations of methane near the surface, or euphotic zone, of the world’s oceans.156 Although anaerobic microbial pathways to methane production are known, these oxygen sensitive pathways could not account for the production of methane in the oxygen rich environment that is found in the euphotic zone. One half of the ocean methane paradox was solved when Karl et al. demonstrated that euphotic marine bacteria could convert 8 to methane and Pi through the carbon−phosphorus lyase pathway (discussed later in this review).156 The answer to the second half of this paradox, the biogenic origin of 8, came in the discovery that the genome of the marine archaeon Nitrosopumilus maritimus contained a ppm gene encoding PEP mutase within a biosynthetic gene cluster encoding the biosynthesis of 4.62 In addition to these genes was mpnS encoding a homologue of hydroxypropylphosphonate peroxidase (HppE) and hydroxyethylphosphonate dioxygenase (HEPD) from the pathways for 9 and 10a, respectively. Both enzymes are dependent on a nonheme Fe(II) cofactor for catalyzing oxidation reactions of their substrates. Using MS and 13 C NMR spectroscopy MpnS was shown to convert 4 into 8 and CO2 (or bicarbonate) in vitro.62 8 was also observed by 31P NMR spectroscopy in cell extracts of N. maritimus, as well as associated with an exopolysaccharide as an ester. The mpnS gene was detected within a similar genomic context in Pelagibacter sp.,62 which is the most commonly found genus of bacterium in marine euphotic zones.157 Remarkably, other species of Pelagibacter will also actively convert 8 to Pi and methane under Pi limiting conditions; thus this genus has the capacity to biosynthesize and catabolize 8.157 MpnS catalyzes an unprecedented C−C bond cleavage reaction. Only C2 of 4 is oxidized during C−C bond cleavage, thus both C2 hydrogens are removed during catalysis. Analysis of MpnS in vitro revealed that the Fe(II) cofactor does not require external electrons for catalysis.158 Because one mole of O2 is consumed per mole of 8 produced, the C−C cleavage step is therefore coupled to a four-electron reduction of O2. The kinetic parameters for the conversion of 4 to 8 were KM (O2) = 11 μM, KM (4) = 4.5 μM, and kcat = 0.2 s−1. Intriguingly, reaction with the substrate analog (R)-63 produced the ketone 64 (Scheme 10A). MpnS was also inactivated during the reaction with (R)-63, while no reaction was observed with (S)63 (Scheme 10B). This latter result implied that the pro-S C2 hydrogen of 4 is abstracted during catalysis. Running the MpnS reaction in D2O with 4 indicated that 8 did not receive a hydrogen from the solvent or a solvent exchangeable residue on

the enzyme. Instead, (R)-[2-2H1]-4 is converted to [1-2H1]-8, indicating that the pro-R hydrogen 4 is transferred to the methyl group group of 8 during catalysis (Scheme 10C). Kinetic isotope effect studies with MpnS suggest that activation of molecular oxygen is rate determining. Reaction with deuterium labeled (R)- and (S)-[2-2H1]-4 did not produce a KIE, suggesting that H-abstraction from C2 of 4 is not rate determining.158 However, a competitive KIE is observed using 18 O2.159 Because a competitive KIE reports on kcat/KM (O2) for the MpnS reaction, the observed 16k/18k value of 1.0158 ± 10 with 4 reflects the first irreversible chemical step for the reduction of O2 by the active site Fe(II). Remarkably, the 16 18 k/ k value increased to 1.0189 ± 13 using dideuterium labeled [2-2H2]-4, suggesting that reduction of O2 by the Fe(II) ion is only partly rate limiting and is more fully expressed with the deuterium label; thus, O2 reduction by MpnS occurs during or before the hydrogen abstraction step. Both 16k/18k values are large compared to other enzymes that activate molecular oxygen, leading to the hypothesis that tunnelling is involved in hydrogen transfer to an Fe(III)−O-O•− species. Greater insight into the mechanism of MpnS came indirectly from analysis of HEPD, an enzyme that converts 4 to hydroxymethylphosphonate 35. MpnS is unusual in missing the Glu or Asp residue of the 2-His-1-carboxylate facial triad that is used to bind the active site Fe(II) ion. Instead, this residue appears to be Gln based on structural homology to HEPD. Remarkably, substitution of the corresponding E176 residue in HEPD for histidine created an MpnS like-variant that converted 4 to a 50/50 mixture of 8 and 35.160 To explain this switch, it was proposed that a methylphosphonate radical is formed as a common intermediate by MpnS and HEPD which can be diverted to different products by changes within the active site. This hypothesis was strengthened by the observation that (R)-[2-2H1]-4 is converted to monodeuterium labeled 8, just like MpnS, but with a dramatic change in product ratio in favor of 35 (1:10). This indicates that the methylphosphonate radical can be diverted to 35 due to the higher activation barrier to transferring deuterium from C2 of 4 to the methyl group of 8 during catalysis. Based on these studies a mechanism for MpnS can be proposed (Scheme 11). 4 is believed to bind to the Fe(II) ion in a bidentate mode, followed by molecular oxygen which is reduced to form Fe(III)−O-O•− (intermediate II). The pro-S hydrogen is abstracted by the superoxide, forming a ketyl-like radical at C2. Steps flanking intermediate II are proposed to be partly rate limiting based on the KIE measurements noted 5717


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Scheme 11. Methylphosphonate Synthase MpnSa

a

Proposed mechanism for MpnS shown along the outer ring. X = unknown active site ligand. Intermediate VII is shared with 2hydroxyethylphosponate dioxygenase HEPD from the phosphinothricin 10a pathway.

enzyme catalyzes the ligation of phosphinopyruvate 97 with acetyl-CoA.164 All three enzymes are sequence homologues of homocitrate synthase (EC 2.3.3.14), which ligates 58 and acetyl-CoA to form homocitrate 65 as part of the αaminoadipate pathway for lysine biosynthesis in certain fungi and archaebacteria (Scheme 12B).165 FrbC and Pfs2 have been heterologously produced in E. coli and shown to convert 1 to 56 in vitro by 13P NMR spectroscopy.162,163 Although these enzymes require a divalent metal ion for activity, their metal ion specificity has yet to be determined. However, PMM synthase is active with Co2+ and Mn2+, with lower activity obtained with Ca2+, Fe3+, and Mg2+,164 while the homocitrate synthase from Schizosaccharomyces pombe (ScHCS) is active with Zn2+.165 A number of key active site residues are conserved in FrbC, Pfs2, and PMM, which can be used to create a model of the active site based on the X-ray crystal structure of ScHCS in complex with 58 (Scheme 12C).165 In the case of FrbC, the metal ion is predicted to be bound by H194, H196, D15, and a water molecule. 1 is bound to the metal ion in a bidentate mode via the carboxylate and ketone groups, thereby increasing the electrophilicity of C2 toward attack by the enolate of acetylCoA. Active site substitutions in ScHCS suggest that the FrbC residues R13 and Q18 interact with the carbonyl of acetyl-CoA to promote formation of the enolate. Likewise, by analogy to the proposed roles of conserved Glu and Arg active site residues in ScHCS,165 the corresponding residues E137 and R13 of FrbC may serve as general acid and base catalysts for enolate formation, aldol condensation, and hydrolysis of the thioester of 56 (Scheme 12D).

above. The ketyl radical is proposed to reduce ferric ion of Fe(III)-O−OH to form an aldehyde and Fe(II)-O−OH (intermediates III to IV). Attack of the bound peroxide on the aldehyde forms a peroxide-hemiacetal (V). Reductive homolytic cleavage of the peroxide bond yields an alkoxyl radical, which in turn initiates β-scission of the C1−C2 bond of the substrate, forming a methylphosphonate radical and formate (VI to VII). The methylphosphonate radical (VII) is proposed to be an intermediate in common to MpnS and HEPD that can be quenched in two different ways. In the case of MpnS, a hydrogen is abstracted from the bound formate ion, yielding a carbon dioxide radical anion and 8 (VII to VIII). The former is a strong reducing agent which can reduce the Fe(III) ion back to Fe(II) (VIII to I). In the case of HEPD, the methylphosphonate radical undergoes a more typical ’rebound’ reaction with the Fe(III) bound hydroxide, yielding 35, formate, and regenerated Fe(II) (VII to IX). It is noteworthy that the deviation from hydroxylation by MpnS in intermediate VII has a strong parallel with α-ketoglutarate/nonheme Fe(II) dependent halogenases, which quench an alkyl radical with an iron bound halide ion.161 3.9. Phosphonomethylmalic Acid (56)

The production of 1 can be coupled irreversibly by phosphonomethylmalic acid synthase in an aldol-like reaction with acetyl-CoA to form phosphonomethylmalic acid 56 (Scheme 12A). Phosphonomethylmalic acid synthase occurs as FrbC in the biosynthetic pathway for 12a in Streptomyces rubellomurinus162 and as Pfs2 in the pathway for 9 in Pseudomonas syringae.163 A homologous enzyme, phosphinomethylmalic acid (PMM, 98) synthase, discussed later in this review, is found in the pathway for 10a, but in this case the 5718


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Scheme 12. Phosphonomethylmalic (PMM) Acid Synthasea

a

(A) Reaction catalyzed by the PMM synthases FrbC and Pfs2. (B) Reaction catalyzed by homocitrate synthase. (C) Active site scheme for FrbC based on homology to the homocitrate synthase ScHCS (PDB ID: 3ivt).165 FrbC active site residues shown in bold, conserved residues in ScHCS shown in parentheses. (D) Proposed mechanism for PMM synthases.

Scheme 13. Rhizocticin Biosynthetic Aldolase RhiGa

3.10. 2-Keto-4-hydroxy-5-phosphonopentanoic Acid (57)

A second aldol-reaction appears in the pathways for rhizocticins 14a−d, plumbemycins 14d−e, and phosacetamycin 14g (Scheme 3). In this case the enolate of pyruvate 53 reacts with the aldehyde of 2, forming 57. An aldolase catalyzing this reaction, RhiG, was first identified as a product of the gene cluster encoding the biosynthesis of 14a−d in Bacillus subtilis ATCC 6633.100 RhiG has highest sequence similarity to 4hydroxy-2-oxovalerate aldolases, including DmpG, an enzyme that appears in a catechol catabolic pathway in Pseudomonas.166 RhiG is also homologous to PamG, which is predicted to catalyze the identical reaction in the biosynthesis of 14g.167 Aldolases can be divided into two classes.168 Class I aldolases are typically found in eukaryotes and use an active site Lys to stabilize formation of the enolate as a Schiff base.169 RhiG belongs to the class II aldolase family, which are of microbial origin and use a divalent metal ion (e.g., Zn2+, Fe2+, Mn2+, Co2+, and Mg2+) to stabilize the enolate, which is typically derived from pyruvate 53.166,170 RhiG was successfully produced in E. coli for in vitro studies.100 The purified enzyme was unreactive with 53 as donor and 2 as acceptor (Scheme 13B) but successfully ligated oxaloacetate 66 and 2 to form 57 (Scheme 13A), suggesting that decarboxylation was used by the enzyme to drive formation of the enolate. Indeed, incubation of RhiG with 66 alone produced the expected decarboxylation product 53 (Scheme 13C). RhiG was unreactive with 58 as the donor (Scheme 13D). The requirement for 66 as an aldol donor

a

(A−D) Reactions catalyzed by RhiG with substrate analogs. (E) Active site model for RhiG and proposed mechanism.

substrate is unusual as other class II aldolases such as HpaI and BphI can deprotonate 53 to form the enolate.171 Although the 5719


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Scheme 14. Mechanism of MurA and Inactivation by Fosfomycin 9

metal ion dependence of RhiG was not examined, Mg2+ was included in all reaction solutions. This is undoubtedly a requirement for catalysis as RhiG shares conserved metal ion binding residues (D12, H192, and H194) with DpmG (Scheme 13E).166 Considering the broad synthetic utility of aldolases,168,172 further exploration of the Pn acceptor and aldol donor specificity of RhiG and its homologues would be warranted.

follows an addition−elimination pathway wherein the 3′−OH of UDP-GlcNAc adds to C2 of 24 with concomitant protonation of C3 to form a tetrahedral intermediate (Scheme 14).184 This is followed by elimination of the proton at C3 and Pi at C2 to form the enoylpyruvyl ether of UDP-MurNAc.185 The use of PEP analogs with fluorine at C3 have revealed that the addition and elimination steps involve considerable oxocarbenium ion character.186−188 Central to the action of 9, but long complicating the MurA mechanism, is an essential active site Cys residue (Cys115 in E. coli). There has been a long and evolving discussion over the role of Cys115 in MurA, either in the context of a nucleophile,189 a general acid/base catalyst,186,190 in electrostatic stabilization of oxacarbenium ion intermediates,186 in product release,185,191 or combinations thereof. In particular is the relevance of the tetrahedral thioketal intermediate that is observed to form by reaction of the Cys115 side chain with PEP prior to transfer to UDP-GlcNAc.192−194 Although the covalent Cys115-thioketal intermediate turns over in the presence of UDP-GlcNAc,193,194 thus exhibiting competence, this intermediate can be bypassed to give the ketal intermediate on UDP-GlcNAc directly,186 and Cys115 can be replaced with Asp to yield a functional enzyme.195 For this reason the Cys115 covalent intermediate is proposed to be an adventitious but reversible branch-point off the normal reaction pathway.186 9 is designed to exploit the nucleophilicity of Cys115 as a mimic of 24. Upon binding of UDP-MurNAc and 9 to MurA, C2 of the epoxide reacts with the Cys115 and covalently inactivates the enzyme (Scheme 14). An X-ray crystal structure of inactivated MurA has been determined (PDB ID: 1uae).196 Interestingly, a close sequence homologue of MurA, enolpyruvyl shikimate 3-phosphate synthase (EPSP synthase or AroA) catalyzes an analogous enoylpyruvyl transfer from 24 to the 5′hydroxyl shikimate 3-phosphate, forming EPSP (Figure 7). EPSP synthase does not have a corresponding Cys residue and is proposed to promote enoylpyruvyl transfer by electrostatically stabilizing the oxacarbenium ion intermediates (or transition states)197 for addition and elimination using a pair of carboxylate side chains, Asp 313 and Glu 341.198 These active site differences lead to contrasting specifies toward phosphonate inhibitors. EPSP synthase is potently inhibited by the phosphonate herbicide glyphosate 23 by virtue of the protonated secondary amine interacting with an ionized Glu

3.11. Natural Product Pn of Unknown Provenance

Several Pn natural products with intriguing structures have unknown biosynthetic origins. Elegant labeling and MS experiments by Bachmann revealed that L-tyrosine is the precursor to 21a in Actinomycete sp. K-26.173 This suggests that the phosphonyltyrosine derivatives 21a and 21b arise from a new C−P bond forming reaction. The single example of a chlorinated Pn, fosfonochlorin 20, is produced by several species of Fusarium.174 Although this compound was discovered over 25 years ago, its biosynthetic origin and mechanism of halogenation remains unknown. SF-2312 22, discovered 30 years ago in cultures of Micromonospora sp.,175 is the only example of a Pn natural product where the phosphonate group is directly attached to a ring. Although the structure of 22 has been confirmed through total synthesis,176 its biosynthetic pathway is also unknown.

4. PATHWAYS FOR SYNTHESIZING COMPLEX PN NATURAL PRODUCTS 4.1. Fosfomycin (9)

9, a product of several species of Streptomyces177,178 and Pseudomonas,179,180 has the distinction of being the first discovered Pn natural product to show antibiotic activity. As a drug, 9 (also known as Monuril, Monurol, and Monural) is active against a wide array of Gram-positive and Gram-negative bacteria and has long been used to treat lower urinary tract infections.181 9 is enjoying a resurgence for treating other types of infections, including treating multidrug resistant bacteria in combination with other antibiotics.182,183 4.1.1. Mechanism of Action. 9 is an active site directed inactivator of UDP-GlcNAc enoylpyruvyl transferase (MurA). This is the first enzyme in the pathway of peptidoglycan biosynthesis that catalyzes the condensation of 24 and UDPGlcNAc to form UDP-MurNAc 67. The mechanism of MurA 5720


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because the minimal biosynthetic pathway in S. wedmorensis and S. f radiae was initially thought to be encoded by fom1, fom2, fom3, and fom4, implying that the aldehyde of 2 was directly methylated to form (S)-63 (Scheme 15).205 Early work by Seto identified a mutant of S. wedmorensis (strain A16) that was defective in the biosynthesis of cobalamin as well as 9.206 Production of 9 could be restored with the addition of hydroxycobalamin to the culture medium of this mutant, and addition of 14CH3-labeled methylcobalamin produced 14CH3labeled 9, suggesting that this cofactor was the donor of the methyl group.206 However, this also implied that methylcobalamin delivered a nucleophilic methyl anion to the aldehyde of 2 to form (S)-63. 8,207 Such a mechanism would be unprecedented, as methylcobalamin typically acts as source of a methyl radical or a methyl electrophile in homolytic or SN2 reactions, respectively.208,209 This unusual methyl transfer reaction was assigned to Fom3 based on its sequence homology to methylcobalamin dependent enzymes.205 A key to resolving this mechanistic puzzle was the early observation that 4 is efficiently converted to 9 by S. f radiae.210,211 This was also the case with the mutant NP-7 strain of S. wemorensis that is defective in the biosynthesis of 2.212 Initially 4 was thought to be a shunt product of 2 through reversible oxidation of the alcohol to the aldehyde by a nonspecific dehydrogenase.206,213 However, this adventitious model was at odds with the observation that conversion of labeled [2-18O]-4 by S. f radiae produces 9 that largely retains the 18O label (50%).207 If 4 was oxidized to 2 prior to conversion to 9, the label would be expected to wash out rapidly through hydration of the aldehyde. However, retention of the 18O label in 9 does make sense if 4 is not a shunt product of the pathway but is a genuine intermediate that follows the reduction of 2. Support for this proposal came with the discovery that the minimal biosynthetic gene cluster for 9 included fomC, encoding the NADH and Fe(II) dependent alcohol dehydrogenase discussed in section 3.4.201 The fomC gene was shown

Figure 7. Comparison of the herbicide glyphosate (23) and the oxacarbenium ion intermediate formed by EPSP synthase.

341 (Figure 7),199,200 but is not inhibited by 9. Conversely, MurA is not inhibited by 23. Interestingly, MurA can be rendered resistant to inactivation by 9 through the Cys115Asp substitution described above.195 4.1.2. Biosynthesis of Fosfomycin. The biosynthesis of 9 in Streptomyces involves five chemical steps. Genetic studies involving S. wedmorensis and S. f radiae revealed fom1, fom2, fomC, fom3, and fom4 to encode the corresponding biosynthetic enzymes.201 Each of these steps, outlined in Scheme 15, have been reconstituted in vitro. The three step conversion of 24 to 2-hydroxyethylphosphonate 4, an intermediate that is common to other Pn biosynthetic pathways,137 is discussed in mechanistic detail within section 3.4. In the context of fosfomycin biosynthesis, Fom1 (PEP mutase) and Fom2 (TPP dependent decarboxylase), draw the 24:1 equilibrium toward the production of 2. The aldehyde of 2 is subsequently reduced by the NADH and Fe(II) dependent dehydrogenase FomC to form 4.132,137,202 This is followed by methylation of 4 by the methylcobalamin dependent enzyme Fom3203 to form (S)-63. Finally the epoxide ring of 9 is formed by by the nonheme iron dependent peroxidase Fom4.204 Fom3 and Fom4 are discussed in more detail below. 4.1.3. Cobalamin-Dependent Methyltransferase Fom3. A longstanding puzzle in the biosynthesis of 9 was the mechanism for installing the methyl group. This was in part

Scheme 15. Comparison of the Proposed Pathways for Fosfomycin 9 Biosynthesis in Streptomyces (Upper Pathway) and Pseudomonas (Lower Pathway)

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Scheme 16. Radical SAM Methyltransferasesa

a

(A) Homolytic cleavage of S-adenosylmethionine (SAM) by the 4[Fe−S] cluster. (B) Methylation of the cobalamin cofactor by SAM.

Scheme 17. Proposed Mechanism for Methyltransferase Fom3

phosphorus atom of phosphinates.214 Like other enzymes in this family, the radical SAM domain of Fom3 contains a CxxxCxxC sequence motif that is used to bind an [4Fe-4S] cluster. Three of the Fe atoms of the cluster are bound by the Cys side chains, while the fourth Fe atom has two free ligand sites that are used to bind the α-amino and α-carboxylate groups of SAM. Upon binding SAM in the radical SAM domain, the reduced form of the cluster, with an [4Fe-4S]+1 oxidation state, can deliver a single electron to reductively cleave the S−C(5′) bond, generating L-methionine and a 5′deoxyadenosyl radical 68 (Scheme 16A and Scheme 17, I to II).215 The radical 68, or one subsequently generated on a side chain in the active site of Fom3, will presumably abstract a C2 hydrogen from 4, forming 5′-deoxyadenosine 72 (Scheme 17, II to III). The resulting C2 radical of 4 can then receive a methyl group from methylcobalamin 70 (III to IV). 70 is formed in the cobalamin binding domain through an SN2 reaction between the highly nucleophilic Co(I) ion of 69208

to be essential for the biosynthesis of 9 in S. wedmorensis, and that 4 would restore the biosynthesis of 9 in a S. wedmorensis fomC mutant.132 As discussed in greater detail in section 3.4, FomC was shown to reversibly convert 2 to 4 using NADPH as the hydride donor (Figure 6).137,202 Therefore 4 is a true intermediate of 9 biosynthesis and the substrate for the methylcobalamin dependent transferase Fom3. By sequence homology Fom3 belongs to a subclade within the radical S-adenosylmethionine (SAM) superfamily. This subclade is comprised of radical SAM methyltransferases (RSMTs), which presently contains over 100 000 members that further divide into four subclades, A-D, according to their cofactor dependence, domain composition, the type of substrate that is methylated, and their proposed mechanisms.214−216 Fom3 falls into the B-class by virtue of a Cterminal radical SAM domain and an N-terminal cobalaminbinding domain. The B-class RSMTs are the largest subfamily, with members that methylate sp3 and sp2 carbons as well as the 5722


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and the methyl group of SAM, producing S-adenosylhomocysteine 71 as the second product (Scheme 16B). Hammerschmidt demonstrated S. f radiae will convert (S)- but not (R)-63 to 9.210 This means that in the absence of inversion of the C2 radical, the pro-S C2 hydrogen of 4 is abstracted by Fom3, and the methyl group is delivered from 70 to form (S)-63.132 Additional evidence for 4 as an intermediate in the biosynthesis of 9 arrived with reconstitution of Fom3 activity in vitro.203 Fom3 from S. wedmorensis was produced heterologously in E. coli as an insoluble inclusion body (commonly observed with radical SAM enzymes) which necessitated purification under denaturing conditions.203 To reconstitute the [4Fe-4S] cluster, Fom3 was refolded aerobically in the presence of SAM, ferrous ammonium sulfate, sodium sulfide, and the reducing agent DTT. Subsequent experiments were conducted anaerobically. EPR spectroscopy of refolded Fom3 indicated the presence of an oxidized [4Fe4S]+3 cluster (EPR signal g-value 2.09; for an excellent comprehensive listing of radical SAM [4Fe-4S] cluster EPR signals, see ref 215). Upon addition of DTT this signal disappeared, suggesting reduction of the cluster to the resting state [4Fe-4S]+2. Finally addition of the stronger reductant dithionite produced a signal (g = 1.91) that was diagnostic of the reduced [4Fe-4S]+1 state of the cluster. Upon addition of SAM, 70, and 4, the signal at g = 1.91 reduced in intensity, suggesting oxidation to the [4Fe-4S]+2 state as one would expect if a single electron transfer occurred to reductively cleave SAM. Analysis of the reaction mixture by 31 P NMR spectroscopy revealed conversion of 4 to 63. Conversion was not complete, likely due to the instability of the enzyme and oxidation of the [4Fe-4S] cluster. Reconstitution of the activity of Fom3 will enable deeper insight into the mechanism of methyl transfer. Only a handful of enzymes from the class B family of RSMTs have been studied in vitro, including Fom3, PhpK (sp3 P-methylation), GenK (sp3 C-methylation),217 ThnK (sp3 C-methylation),218 and TsrM (sp2 C-methylation).219−221 One interesting question is whether methylation of 4 by 70 in the Fom3 reaction occurs in a homolytic or heterolytic fashion. Because a C2 hydroxymethylene radical is generated on 4 (Scheme 17, III), both mechanisms are possible.132 Each have been proposed by Liu and co-workers for GenK, which performs a similar transformation during the biosynthesis of gentamicin.217 Applying the heterolytic mechanism to the Fom3 reaction requires ionization of the C2-hydroxyl of the substrate radical to form a C2 ketyl radical (Scheme 17, III to IV). The carbanion resonance contributor would then attack the methyl group of 70 in an SN2 fashion, producing 69 and the alkoxy radical of (S)-63 (IV to V). The latter would receive an external electron and a proton to form (S)-63 (V to VI). Overall one would expect the Fom3 reaction to require two external electrons, one to reduce the [4Fe-4S] cluster (VI to I), and the second to reduce the substrate radical (V to VI) (or alternatively the cobalamin cofactor from Co(II) to Co(I) in a homolytic methyl transfer mechanism132). Additionally, two equivalents of SAM are expected to be consumed per reaction cycle, one to generate the 5′-deoxyadenosyl radical 68, the other to methylate cobalamin 69, leading to 5′-deoxyadenosine 72 and S-adenosylhomocysteine 71 as products in equimolar quantities. This is indeed observed in the GenK reaction.217 4.1.4. 2-Hydroxypropylphosphonate Epoxidase Fom4. The 2-hydroxypropylphosphonate epoxidase Fom4, also known as HppE, performs an unusual dehydrogenation

reaction of a secondary alcohol to form the epoxide ring of 9. Early work by Seto demonstrated that 63 was the substrate for this reaction,213 while Hammerschmidt revealed that the reaction is stereospecific for (S)-63.210 That this reaction involved dehydrogenation, and not incorporation of oxygen into the substrate, was first inferred from the observation that S. f radiae converted 18O-labeled 4 to 18O-labeled 9, indicating that the C2-hydroxyl of 4 was utilized to form the epoxide (Scheme 18).207 Fom4 was assigned as the likely enzyme catalyzing this Scheme 18. Stereochemistry of Epoxide Formation in 9 Examined by Conversion of Labelled Precursors in S. f radiaea

a18

O is shown as a solid sphere.

reaction based on its weak sequence similarity (21% amino acid identity) to alcohol dehydrogenases.205 Hammerschmidt also fed deuterium labeled precursors to S. f radiae to demonstrate that formation of the epoxide ring by Fom4 is stereospecific. S. f radiae converts (S)-[1-2H]-4 to 9 with retention of the deuterium label at C1, whereas the label is lost upon conversion of the (R)-[1-2H]-4 (Scheme 18).210,211 This means that Fom4 substitutes the C1 pro-R hydrogen of (S)-63 with with the C2 hydroxyl oxygen. Because 9 has R-stereochemistry at C1, formation of the epoxide ring occurs with inversion of stereochemistry at C1. Several years passed before the Liu group successfully reconstituted the activity of S. wedmorensis Fom4 in vitro.222 Initially the activity of Fom4 was so low that it was necessary to use a bioassay against E. coli to detect the conversion of (S)-63 to 9.223 A metal ion screen revealed Fe(II) was required for activity, and that an external electron source, in the form of NADH and a flavin (FAD or FMN) or the NADHdependent flavoprotein reductase E3, significantly increased activity. This allowed for sufficient 9 to be produced in vitro for spectral characterization, as well show in vitro that Fom4 converted 18O-labeled (S)-63 to 18O labeled 9, confirming the origin of the epoxide oxygen.223 Until only recently it was believed Fom4 catalyzed the dehydrogenation reaction as an oxidase due to the apparent requirement for external electrons. The demand for electrons appeared to be stoichiometric, with one equivalent of NADH consumed per turnover of (S)-63 to 9.224 Accordingly, mechanisms were proposed where an active site Fe(II) ion would bind and activate molecular oxygen to form either an iron-superoxo (Fe(III)−O-O•−), iron-hydroperoxo (Fe(III)− O−O-H), or ferryl oxygen (Fe(IV)O) as the reactive species that would abstract the C1-pro-R hydrogen of (S)-63.204,223 The resulting C1 radical would then undergo radical ring closure with the Fe-coordinated C2-oxygen to form the epoxide ring of 9. All three schemes require the input of 2 external electrons, either to form the reactive iron-oxo species or to regenerate Fe(II), and overall 4 electrons are used to reduce molecular oxygen to water. However, the requirement for external electrons was troubling because Fom4 did not bind to flavin cofactors which might mediate electron transfer to the iron cofactor, and even with optimization of the external electron source the reconstituted activity remained relatively low (kobs = 1.3 min−1).224 5723


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Scheme 19. Reaction Mechanisms for Fom4 (HppE) with Substrate (A) and Substrate Analogs (B−D)

motif in mononuclear nonheme iron dependent enzymes (Figure 8).223 (S)-63 binds in a bidentate mode to Fe(II) by

The issue concerning the electron requirement was resolved nearly a decade later when it was determined that Fom4 was in fact not an oxidase, but a peroxidase.204 This transpired through the serendipitous discovery that the addition of the stronger reducing agent dithionite increased Fom4 reaction rates 1000fold (kobs = 10−100 s−1). It was determined that dithionite, like the previously used reducing systems, produced H2O2 in situ by reducing molecular oxygen. The production of 9 was stoichiometric on a 1:1 molar basis with the consumption of H2O2. The kcat/KM value of Fom4 toward H2O2 was estimated to be 5 × 105 M−1 s−1 at 21 °C, which is only 10- to 100-fold less than that of bovine liver catalase, an enzyme that has specifically evolved to react with H2O2.204 This discovery set the mechanism of Fom4 on a new footing: as a nonheme iron dependent peroxidase, Fom4 does not require external electrons to form an Fe(IV)O species that could be used for abstraction of the C1 pro-R hydrogen of (S)-63. Instead, the Fe(IV)O species can be generated by direct oxidation of Fe(II) by H2O2 (Scheme 19A). Likewise external electrons are not required to regenerate the Fe(II) ion as the required 2 electrons are obtained by dehydrogenation of the substrate during the catalytic cycle. X-ray crystal structures of Fom4 bound to (S)-63 added additional detail and intrigue to this enzyme.225,226 By amino acid sequence similarity, Fom4 belongs to the cupin superfamily.227 The structure of Fom4 is tetrameric, with each monomer comprised of the expected cupin fold that contains a single Fe(II) ion. The metal ion is bound by a 2-His, 1carboxylate facial triad (H138, H180, E142) that is a conserved

Figure 8. Active site scheme of the fosfomycin biosynthetic enzyme Fom4 bound to NO.

one of the phosphonate oxygens and the C2 hydroxyl group, the latter likely ionized to an alkoxide. This binding mode was confirmed by EPR studies of Fom4 with 17O labeled substrates and analogs.228 The phosphonate group is additionally bound by hydrogen bonds and electrostatic interactions to K23, R97, Y105, and N135. Insight into activation of molecular oxygen and the stereopecificity of the Fom4 reaction came from a complex with (S)-63 and nitric oxide.226 The sixth and remaining ligand site of the Fe(II) ion was occupied by NO, likely mimicking the binding site of H2O2. A key stereochemical feature is that the C1 pro-R hydrogen of (S)-63 points toward the NO ligand, with only 3.5 Å separating the oxygen of NO and C1. Therefore, the pro-R hydrogen is within reach of where an Fe(IV)O oxygen would be generated during catalysis. Additionally, a large conformational change is observed upon 5724


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binding (S)-63 that almost entirely seals the active site, with the exception of a small opening that presumably allows access of H2O2. Closure of the active site is proposed to protect the reactive Fe(IV)O species that is formed during catalysis. At the time of this structural study NO was thought to mimic molecular oxygen.226 However, in light of Fom4 being a peroxidase,204 this result raises the question as to how Fom4 distinguishes between molecular oxygen and hydrogen peroxide as substrates, given the similarity in size of the two molecules. The specificity for H2O2 is all the more intriguing when it is considered that the closest structural homologue to Fom4 is HEPD, a nonheme iron dependent oxygenase that oxidatively cleaves the C1−C2 bond of 4.204 Like Fom4, HEPD does not require cofactors beyond Fe(II) or the input of external electrons.229 However, HEPD is specific for molecular oxygen as the oxidant, not H2O2. HEPD will be discussed in more detail in the section on the biosynthesis of phosphinothricin 10a. A second mechanistic mystery concerned the mechanism of epoxide ring closure by Fom4. It was difficult to envision how substitution of the C1 pro-R hydrogen with the C2-hydroxyl oxygen of (S)-63 would occur with inversion of stereochemistry to form an (S)-configured C1 in 9 if the mechanism were purely radical based. This is because following abstraction of the C1 pro-R hydrogen, the resulting C1 radical would need to undergo inversion.204 A more satisfying explanation for stereochemical inversion at C1 would involve a carbocation intermediate. In this mechanism the Fe(IV)O species would abstract the pro-R hydrogen at C1, followed by single electron transfer from the C1 radical to Fe(III), forming a planar C1 carbocation and regenerated Fe(II) (Scheme 19A). The Fe(II) bound C2-alkoxide would then attack the neighboring C1 carbocation as a nucleophile and generate the required (S)-C1 configuration in 9. Evidence for a carbocation intermediate benefited from the relaxed substrate specificity of Fom4, which allowed the mechanism to be probed with a suite of substrate analogs.228,230−232 In a key experiment, it was observed that the 1-hydroxy analog (R)-73 was converted by Fom4 to the aldehyde 74 (Scheme 19B).230,231 Such a product could only arise through a 1,2-shift of the phosphonate moiety. Analogous to a 1,2 alkyl shift, this would most likely be promoted by forming a carbocation at C2. Using C2 deuterium labeled (R)73 it was established that the pro-R C2 hydrogen is abstracted by Fom4 en route to forming the carbocation.230 An X-ray crystal structure of Fom4 bound to (R)-73 confirmed that the C2 pro-R hydrogen is nearest to where the Fe(IV)O oxygen is generated, while the C1 hydrogen projects away from this position.230 That a C2 carbocation promoted the phosphonate shift, and not a C2-radical, was inferred from the products of model synthetic reactions. Treatment of the C2 bromide 76 under ionizing conditions with silver triflate led to the same 1,2migration product produced by Fom4 (Scheme 20A), whereas generation of a C2 radical using a combination of the initiator AIBN and tributyltinhydride led to an alkene (Scheme 20B).230 Additional transformations are catalyzed by Fom4 that likewise depend on the regiospecificity of hydrogen abstraction. This is dramatically shown by the reaction of Fom4 with (R)63 and (S)-73. If (R)-63 is used as a substrate, the 2oxopropylphosphonic acid 64 is produced as a product (Scheme 19C).232 Similarly (S)-73 is converted to the 1oxopropylphosphonic acid 75 (Scheme 19D).230,231 In both cases these substrates bind to the Fe(II) ion in a way that

Scheme 20

presents either C2 or C3 hydroxymethylene hydrogens nearest to where the Fe(IV)O oxygen is generated.226,230 However, in these examples, hydrogen abstraction is predicted to generate a ketyl radical that subsequently reduces Fe(III), producing 64 or 75 and regenerated Fe(II). 4.1.5. Biosynthesis of Fosfomycin in Pseudomonas sp. While the biosynthetic pathways for 9 in Streptomyces and pseudomonads share the identical P−C bond forming and epoxidation mechanisms (Scheme 15), strikingly different routes are taken in between these initial and final reaction steps. This divergence was revealed through genome sequencing of Pseudomonas syringae PB-5123.163 A biosynthetic gene cluster for 9 was readily identified from genes encoding homologues of PEP mutase (Psf1) and the epoxidase Fom4 (Psf4). However, genes encoding homologues of the Streptomyces enzymes Fom2 (decarboxylation), FomC (reduction), and Fom3 (methylation) were absent. Heterologous expression of the cluster from a fosmid in Pseudomonas aeruginosa produced 9, confirming that all essential biosynthetic genes had been identified. An alternate pathway was suggested by psf 2 encoding a homologue of FrbC from the FR-900098 12a pathway and PMS from the phosphinothricin 10a pathway. Like these enzymes, Psf2 was shown in vitro to catalyze an aldol-like reaction between acetyl-CoA and 1, followed by hydrolysis of the CoA thioester, to form 2-phosphonomethylmalate 56 (Scheme 12). The mechanism of this enzyme is discussed in greater detail in Section 3.9. This biosynthetic divergence reveals that the C−C bond formation step leading to the C3 methyl group of 9 is fundamentally different: an aldol-like reaction catalyzed by Psf2 versus a radical methyl transfer reaction catalyzed by Fom3. The subsequent steps in P. syringe to form (S)-63 are currently speculative. A chemically reasonable pathway is proposed where C2 of 56 is hydroxylated to form 78, possibly by Psf5, which belongs to the same cupin-like superfamily as the epoxidase Psf4 (37% sequence identity). This would set the stage for elimination of CO2 and water between C1 and C2 to form 79, perhaps assisted by phosphorylation of the C2-OH by the predicted kinase Psf7. Alternatively, Psf7 may play a role in self-resistance through phosphorylation of 9. Tautomerization of the resulting enol to the β-ketoacid 80 would allow decarboxylation to form the ketone 64. Finally, stereospecific reduction of the C2 ketone, perhaps by one of the predicted reductases Psf3 or Psf6, would furnish (S)-63. Although Psf3, Psf5, Psf6, and Psf7 were successfully produced in soluble form in E. coli, their activities could not be reconstituted with various substrates, thus their functions remain to be determined. It is possible that catalysis by these enzymes requires attachment of a biosynthetic intermediates to a nucleotide, which has precedence in the pathway for the biosynthesis of 12a233 and 10a,133 and more compellingly in the appearance of adenylated 9 (called fosfadecin) in cultures of Pseudomonas PK-5.180 5725


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4.1.6. Mechanisms of Resistance to Fosfomycin. Several resistance mechanisms toward 9 have evolved in bacteria. Producing strains of Streptomyces or Pseudomonas inactivate 9 through mono- or bisphosphorylation of the phosphonate group by the ATP-dependent kinases FomA and FomB (Scheme 15).163,234,235 A series of high resolution X-ray crystal structures including FomA in complex with ATP/9, MgADP/9/vanadate, and MgADP/monophosphorylated 9, are consistent with an in-line mechanism for transfer of the γphosphate of ATP to the phosphonate moiety of 9.236,237 The extensive degree of negative charge stabilization of the ATP phosphoryl oxygens by Mg2+ and polar active site residues (K9, K18, K216, and H58) is suggestive of an associative (pentavalent) transition state for phosphoryl transfer. Interestingly, FomA may have evolved from isopentenyl phosphate kinases (IPKs), which catalyze the phosphorylation of isopentenyl phosphate to form isopentenyl diphosphate, a precursor to isoprenoids in archaebacteria. IPKs are the closest structural homologues of FomA within the amino acid kinase superfamily and share the conserved array of polar residues (K9, K18, K216, and H58 in FomA).236 This structural homology inspired a search for promiscuous activity in IPK from Thermoplasma acidophilum, which indeed turned out to be active toward 9 as an alternate phosphoryl acceptor.238 Although the reaction with 9 (kcat/KM = 2.6 M−1 s−1) was 5 orders of magnitude slower than with the native substrate, this activity would likely be sufficient as a starting point to evolve a FomA based resistance following horizontal gene transfer. Microorganisms targeted by 9 have also evolved resistance mechanisms. This topic has been recently reviewed.27,28 Resistance mechanisms include substitutions in the active site of the target enzyme MurA 195 (Scheme 14) or the glycerophosphate transporter GlpT, which provides the main route for importing 9 into the cell. A strain of Rhizobium huakuii was identified by McGrath that not only resists 9 but can use this antibiotic as a sole source of Pi, hinting at a novel mechanism for C−P bond cleavage.239 Resistance is also achieved by expressing enzymes that deactivate the epoxide ring of 9 by reaction with thiols (FosA240and FosB241) or water (FosX242). FosA, B, and X are members of the vicinal oxygen chelate superfamily of metal ion dependent enzymes, which includes glyoxyalase I. All three enzymes use metal ions as Lewis acid catalysts to catalyze SN2 reactions at C1 of the epoxide ring of 9.243 FosA is a Mn2+ and K+ dependent enzyme that utilizes the thiol side chain of glutathione as the nucleophile (Scheme 21).244 FosB has flexible metal ion dependence (Mg2+, Ni2+, Zn2+, and Mn2+) and preferentially catalyzes the addition of the L-cysteinyl moiety of bacillithiol 83 to 9.241 L-Cysteine is also accepted as a substrate by FosB, albeit with lower rates. In contrast, FosX is a Mn2+-dependent enzyme that catalyzes the hydrolysis of the epoxide of 9.242 The occurrence of these enzymes is strain specific, with FosA appearing in Gram-negative bacteria, such as Pseudomonas aeruginosa, FosB occurring in Gram-positive organisms like Bacillus subtilis and Staphylococcus aureus, and FosX appearing in the human pathogens Listeria monocytogenes, Clostridium botulinium, and Brucella melitensis. The occurrence of FosA and FosB likely explains their substrate specificity: unlike Gramnegative bacteria, Gram-positives do not synthesize glutathione and instead use 83 as a low molecular weight thiol for the detoxification of electrophiles. The 3-dimensional structures of FosA, FosB, and FosX are highly similar, reflecting their common evolutionary ances-

Scheme 21. Reactions Catalyzed by the Fosomycin Resistance Enzymes FosA, FosB, and FosXa

a18

O isotope is shown as a filled circle.

try.241,243,245 The enzymes form domain swapped dimers; in the case of FosA the side chains of T9 and H7 of one monomer contribute to the active site of the other (Figure 9A). H7, E110,

Figure 9. Active site schemes for FosA (A) and FosX (B). These enzymes form domain-swapped dimers. The residues highlighted in boxes are donated by one monomer into the active site of the other.

and H64 of FosA are used to bind the Mn2+ ion. Upon binding 9, a fourth Mn2+ ligand site is taken by the substrate phosphonate oxygen while the epoxide oxygen is poised to form a fifth ligand interaction.245 The phosphonate moiety of 9 is further recognized by a polar pocket that includes R118. It was proposed that in the transition state for epoxide ring opening the metal ion will stabilize negative charge development on the epoxide oxygen and assume a more ideal five coordinate, trigonal bipyrimidal geometry.245 T9 is positioned to provide further stabilization of the epoxide oxygen as a general acid or hydrogen bond donor,245 while an ionized Y39 is proposed to provide general base assistance to the incoming thiol nucleophile.246 The proposed five coordinate metal ion transition state for FosA was was bolstered by a high-resolution crystal structure of FosA complexed with transition state analog phosphonoformate 30 (KI = 410 nM).54 FosX appears to be the evolutionary progenitor of FosA and FosB. FosX has a very similar active site (Figure 9B), with the exception that E44 appears at a position corresponding to a glycine residue in FosA. Mutagenesis studies suggest E44 acts as a general base for the hydrolysis reaction.242,243 Interestingly, a gene mlr3345 encoding a FosX homologue occurs in the genome of Mesorhizobium loti, associated with a phn operon 5726


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Scheme 22. Glutamine Synthetase Mechanism of (A) Catalysis and (B) Inhibition by 10a and (C) 86

encoding carbon−phosphorus lyase. The M. loti FosX is a poor enzyme for the hydrolysis of 9,242 and thus likely serves an alternative role in Pn catabolism. Nevertheless, DNA shuffling of mlr3345 with the gene encoding the P. aeruginosa FosA afforded a hybrid gene that was 90% similar to mlr3345, but encoded a FosX variant with FosA activity that was sufficient to confer resistance to 9 in E. coli.240 Most of the mutations encoded changes around the active site, including the acquisition of conserved residues used by FosA homologues to bind glutathione.

media, its lack of therapeutic value as an antimicrobial may be attributed to the reversal of its inhibitory effects with hostsupplied glutamine.249,251 However, plant death resulting from inhibition of plant GS has made 10a a successful herbicide that is typically sold as the ammonium salt (glufosinate ammonium) under trade names such as Basta and Liberty. More recently, 10a and other GS inhibitors have been studied as potential treatments for tuberculosis252 and neurological diseases.253 4.2.1. Mechanism of Action. 10a is a potent ATPdependent inactivator of GS, a critical and highly regulated enzyme catalyzing the incorporation of free ammonia into glutamate to form glutamine. GS can be classified into at least three families that share little sequence identity, termed GSI, GSII, and GSIII.254,255 The GSI enzymes are the best characterized family, with structures from Salmonella typhimurium,256 Mycobacterium tuberculosis,257−259 and Escherichia coli260 all revealing dodecameric architectures. GSII enzymes are decamers found only in eukaryotes and a few soil-dwelling bacteria, with structures available for mammalian and maize enzymes.261,262 GSIII enzymes occur in some anaerobic bacteria and cyanobacteria, and structures from Bacteroides f ragilis263 and Bacillus subtilis264 reveal a dodecameric structure different from that of GSI. Despite low sequence similarity and differing homo-oligomeric structures, all GS enzymes possess intersubunit active sites with remarkable structural conservation, suggesting similar mechanisms.256,261 Each active site is defined by an hourglass, or “bifunnel”, shape open at both ends

4.2. Phosphinothricin (10a)

Phosphinothricin (PT) 10a (commonly referred to as glufosinate) is a nonproteinogenic amino acid that was discovered as a component of the PT-Ala-Ala tripeptide 10b (called PTT, SF-1293, or bialaphos) in the early 1970s from S. viridochromogenes DSM 40736 and S. hygroscopicus ATCC 21705.247,248 It was subsequently found in the PT-Ala-Leu tripeptide 10c (phosalacine) from Kitasatosporia phosalacinea DSM 43860249 and the PT-Ala-Ala-Ala tetrapeptide 10d (trialaphos) from S. hygroscopicus KSB-1285.250 As a structural analog of a glutamine synthetase (GS) catalytic intermediate (Figure 5), 10a potently inhibits this enzyme but has poor activity as the free amino acid, probably due to poor uptake. However, “Trojan horse” peptides 10b−d are readily imported into cells where the active inhibitor 10a is released by peptidases. Although 10a inhibits bacterial GS on minimal 5727


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Scheme 23. Proposed Pathway for Phosphinithricin Peptide Biosynthesis

the enzyme. Although 10a binds slowly to the enzyme, subsequent phosphorylation by ATP yields 86 as a tightly bound analog of the transition state and/or the tetrahedral intermediate (Scheme 22B).262,270−272 The corresponding reactivity of 85 in the GS active site produces the phosphorylated inhibitor 87 (Scheme 22C).273 Slow binding is rationalized by the tetrahedral geometry of 10a, which mimics the GS transition state but not the ground state substrate geometry of glutamate.270 Notably, the same major shift of the Glu237 loop that closes the active site on the glutamate substrate is also observed to trap 10a and may contribute to the observed slow and tight binding of the inhibitor. 4.2.2. Biosynthesis of Phosphinothricin and Derivatives (10a−d). Relative to phosphonates the amino acid phosphinothricin 10a represents the only known naturally produced phosphinate, which, together with its commercial value, has motivated biosynthetic investigations for more than three decades.9,274,275 DNA sequencing has revealed nearly identical gene clusters among 10b-producing strains of S. viridochromogenes (the php gene cluster)276,277 and S. hygroscopicus (the bcp gene cluster),278 as well as the 10c producer Kitasatospora phosalacinea.278 This extensive work has collectively established a proposed biosynthetic pathway comprising at least 18 steps, but many key transformations remain unknown (Scheme 23). To simplify the following discussion, the php nomenclature of the best-characterized gene cluster will be used. Overall, the pathway is initiated by the PEP mutase encoded by ppm to afford 1, from which 2 is generated by the ppd-encoded decarboxylase prior to reduction by the gene product of phpC to yield 4 as detailed in section 3.

to respectively accommodate glutamate and ATP, with two or three metal ions (Mg2+ or Mn2+) observed in the middle of the hourglass. The proposed two-step mechanism involves initial phosphorylation of glutamate by ATP to afford a γ-glutamyl phosphate intermediate 84 (Scheme 22A). Indirect evidence for the existence of this intermediate comes from detection of its cyclization product pyrrolidine-5-carboxylate and various partitioning products of GS reactions in the absence of ammonium.265−267 Additional evidence for the intermediate comes from burst kinetics268 and positional isotope exchange (PIX) experiments with [18O]ATP showing glutamate-dependent and reversible cleavage of ATP to ADP in the active site.269 In the second step, ammonia displaces phosphate from the γglutamyl phosphate intermediate to generate glutamine. Significant structural changes occur during the catalytic cycle, most notably a major glutamate-induced shift of a loop to position Glu237 (S. typhimurium numbering) adjacent to the site of tetrahedral intermediate formation. This shift essentially closes the active site to prevent hydrolysis of the reactive γglutamyl phosphate intermediate. ATP binding and coordination of the γ-phosphate to the metal n2 accompanies a relocation of Asp50′ from another subunit. The ammonium ion binds to a carboxylate-rich region and is deprotonated by Asp50′ for nucleophilic addition to the γ-glutamyl phosphate to afford the tetrahedral intermediate. The aminium (NH3+) ion of the tetrahedral intermediate is deprotonated by Glu237 before or during phosphate elimination.256 The inhibition of GS by 10a is noncovalent but is considered practically irreversible. Like the other well-studied inhibitor of GS, methionine sulfoximine (85), 10a is a structural analog of glutamate and so competes with glutamate for its binding site in 5728


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Scheme 24. Isotope Labeling Experiments with HEPDa

a

Starting from the initial C2-centered radical I (labeled III in Scheme 11) derived from the substrate (S)-[1-2H1]-4, mechanistic proposals included: (A) hydroperoxylation and Criegee rearrangement (B) hydroxylation to yield the methyl anion resonance-stabilized as an enolate; and (C) oxidation to generate a bridged alkylperoxide and methylphosphonate radical that could freely rotate to produce the observed racemic mixture of [2H1]-35. (D) Stereochemical fidelity of the [2H1]-35 oxidation reactions ruled out V-mediated racemization in mechanism A.

4.2.3. Hydroxyethylphosphonate Dioxygenase (HEPD) Encoded by PhpD. The transformation of 4 is catalyzed by the phpD-encoded enzyme hydroxylethylphosphonate dioxygenase (HEPD or PhpD) to afford hydroxymethylphosphonate 35 and formate in a striking C(sp3)−C(sp3) bond cleavage reaction (Scheme 23). Initial characterization and structural elucidation of HEPD in 2009 revealed that, despite lacking sequence homology to known proteins, it was clearly a new member of a large family of enzymes possessing a His2-

Interestingly, the next 7 steps in the pathway completely jettison all PEP-derived carbon atoms in service of furnishing the first phosphinate intermediate, phosphinopyruvate (97). Subsequent P-methylation and nonribosomal peptide synthetase (NRPS)-catalyzed peptide assembly produce N-acetylated hydrogen-phosphinate substrates. N-Acetylation confers selfresistance to the producing bacterium, a strategy that has also been exploited to develop glufosinate resistant crops. 5729


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Scheme 25. Reaction Mechanism Identified from QM/MM Studies that Accounts for Acetylphosphate Formation from 1-HEP

acid facial triad that coordinates a Fe(II) ion.279 As with some other nonheme iron dioxygenases in this family, HEPD requires only O2 and Fe(II) for activity280 and is similar to 2hydroxypropylphosphonate epoxidase (HppE or Fom4) involved in fosfomycin 9 biosynthesis (section 4.1.4) and methylphosphonate synthase (MpnS). The mechanism of the latter enzyme is almost identical to that of HEPD (Scheme 11) and is discussed in detail in section 3.8. The discovery of HEPD was precipitated by the realization that 4 is a bona fide biosynthetic intermediate en route to 10b.133 Previous proposals invoked direct transformation of phosphonoacetaldehyde 2 to 35 via an unknown mechanism, and any 4 accumulating in the culture broth was considered to arise from a side reaction.275,281 However, Blodgett et al. observed that 4 accumulated in a ΔphpD mutant of S. viridochromogenes, and that overexpression of PhpD in E. coli was necessary and sufficient for direct conversion of 4 to 35.133 Purified phpD-encoded HEPD was subsequently shown to be inactive without Fe(II) and O2, demonstrating that 4 is the source of all four electrons required for reducing molecular oxygen.279 The X-ray crystal structure of HEPD was reported in 2009 and followed by a series of mechanistic studies. The crystal structure possesses a fold characteristic of the cupin superfamily282 with bidentate binding of 4 to Cd2+ in the active site.279 4 is thought to bind before O2 because steady-state kinetic analysis of HEPD variants revealed saturation with the former but not the latter,283 and bidentate coordination of substrate prior to oxygen binding has been proposed for several nonheme iron enzymes.280 Although the overall structure and active site binding mode were reminiscent of the substrate 63 in the active site of Co(II)-HppE (see section 4.1.4),225 63 was converted to 2-oxopropylphosphonate 64 in a process that

inactivated HEPD.279 Isotopic labeling experiments with 18O2 demonstrated that HEPD is a dioxygenase based on the incorporation of one 18O atom into 35 and one into formate. However, mechanistic interpretation was confounded by the observation that the primary alcohol of 35 had incorporated some 16O from the solvent. Because this exchange did not occur when 35 was incubated under the reaction conditions, the results were initially rationalized by two possible mechanistic trajectories diverging from a carbon centered radical intermediate (intermediate I in Scheme 24, intermediate III in Scheme 11): (i) hydroperoxylation followed by Criegee rearrangement (Scheme 24A) or (ii) hydroxylation to afford an exchangeable Fe(IV)O ferryl intermediate (Scheme 24B).279 The former hypothesis was initially favored after Criegee rearrangement products were directly observed upon incubating substrate analogs with HEPD; for example, 1-hydroxyethylphosphonate 36 yielded acetylphosphate 88.229 However, this proposal necessitated an unusual formyl ester hydrolysis (III, Scheme 24A) to generate solvent exchangeable oxygen for incorporation into the primary alcohol of 35, and no enzymecatalyzed hydrolysis was observed when the proposed catalytic intermediate O-formyl-35 was supplied to the enzyme.229 Furthermore, QM/MM studies invoked a proton coupled electron transfer mechanism to account for 88 formation from 36 (Scheme 25)284 rather than the previously proposed Criegee rearrangement, which would occur similarly to the reaction depicted in Scheme 24D. Results of isotopic labeling experiments on the HEPD reaction ruled out the Criegee rearrangement mechanism and ultimately led to the currently accepted mechanism presented in Scheme 11.285 Consistent with active site geometry and the bidentate binding of 4 observed in the Cd(II)-HEPD crystal structure, stereospecific abstraction of the pro-S hydrogen was 5730


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Scheme 26. Alternative Mechanism for HEPD Catalysis Involving a Ferryl Intermediate V as Proposed Based on QM/MM Calculationsa

a

The mechanism also satisfies the experimental evidence for a methylphosphonate radical and solvent exchangeable ferric hydroxide in intermediate III. W represents a water molecule.

confirmed by incubating (R)- and (S)-[2-2H1]-4 with HEPD and observing deuterated formate only from incubation with the (R)-enantiomer. To distinguish the hydroperoxylationCriegee rearrangement mechanism of Scheme 24A from the hydroxylation mechanism of Scheme 24B, both enantiomers of [1-2H1]-4 were supplied to HEPD. Surprisingly, all of the stereochemical information at C1 of either (R)- or (S)-[1-2H1]4 was lost in the racemic [2H1]-35 products, which ruled out the expected inversion resulting from the hydroperoxylationCriegee rearrangement mechanism (Scheme 24A). Loss of stereochemical information was also unexpected for the hydroxylation mechanism (Scheme 24B) because the anticipated predominance of one of the two possible resonance stabilized enolate intermediates VII would impart stereospecificity to afford either (R)- or (S)-[2H1]-35. The observation of racemic product inspired an additional mechanistic proposal involving electron transfer to iron from the initial C2 centered radical to generate the aldehyde VIII (Scheme 24C). Nucleophilic attack by the peroxide would then afford a bridged alkylperoxide intermediate IX that could cleave homolytically to produce a methylphosphonate radical X.285,286 The presumed lower barrier to rotation of the methylphosphonate radical X (Scheme 24C) relative to the enolate-like carbanion VII of Scheme 24B could explain the observed racemization at the deuterium labeled carbon. A series of experiments further supported the methylphosphonate radical and ruled out the intermediacy of either phosphite or 8. The racemization of label could have been explained by Criegee rearrangement if the resulting product 35 underwent transient and reversible cleavage to phosphite and formaldehyde (V in Scheme 24). Rotation about the CO bond of formaldehyde prior to nucleophilic attack by phosphite would scramble the stereochemistry of the isotopic label to yield a racemic product. This possibility was tested by cleverly exploiting the ability of HEPD to also catalyze the oxidation of 35 to Pi and formic acid.229 (R)-[2H1]-35 was incubated anaerobically with HEPD to allow equilibration between intermediates V and VI before oxygen was added to facilitate the slow conversion of 35 to Pi and formic acid (Scheme 24D).283 The Criegee mechanism with racemization via the phosphite intermediate V (Scheme 24A) was ruled out because no deuterium was detected in the formate product. This result was consistent with the previously observed deuterium incorporation into formate only when using the (S)-[2H1]-35 substrate, and an observed primary KIE of 7.6 ± 0.4 due to slower turnover of (R)-[2H1]-35.285 This experiment therefore ruled out the Criegee mechanism with phosphite-mediated racemization of 35. Similarly, the intermediacy of 8 was tested by looking for deuterium incorporation into 35 when the HEPD reaction was carried out in D2O. If methylphosphonate

8 occurred on the reaction coordinate, captured either by intermediate VI or enolate VII in Scheme 24B, then deuterium would end up in the product 35 if a label-free substrate was used. The inability to detect deuterium in the product was evidence against the intermediacy of 8, and further strengthened the methylphosphonate radical mechanism outlined in Scheme 24C. Key evidence supporting a methylphosphonate radical was obtained from the Y98F variant of HEPD. Consistent with the role of Y98 in binding the substrate, this variant exhibited an elevated KM for 4. Critically, the Y98F variant was slowly inactivated by catalysis and produced small amounts of 8 that possessed no deuterium when the reaction was conducted in D2O. These results suggested that active site changes due to the Y98F substitution generated a methylphosphonate radical that was unable to efficiently access the ferric hydroxide and instead irreversibly inactivating the enzyme by abstracting hydrogen from the protein. Finally, as discussed in Section 3.8, a common methylphosphonate radical intermediate was proposed based on its preferential “rebound” trajectory toward the ferric hydroxide en route to 35 versus abstracting deuterium from formate en route to 8 in the MpnS-catalyzed reaction (see intermediate VII in Scheme 11).160 The unusual reaction catalyzed by HEPD has motivated several computational studies that have provided mechanistic insights. Indeed, the currently accepted mechanism of Scheme 24C was first proposed by Hirao and Morokuma in 2010.286 Density functional theory (DFT) calculations using an active site model of HEPD demonstrated that the bridged alkylperoxide intermediate and homolytic cleavage mechanism of Scheme 24C were energetically favored over the hydroxylation pathway of Scheme 24B, indicating that a high valent Fe(IV)O intermediate was not needed to facilitate HEPD chemistry. The calculations reported a rate-limiting initial H-abstraction (i.e., from II to III in Scheme 11) although later work failed to detect substrate deuterium KIEs on kcat/KM for the substrate 4.160 However, competitive KIEs with 18O2 increased 16k/18k from 1.0148 ± 9 with 4 to 1.0231 ± 26 for 2[2H2]-4 KIE, consistent with slower turnover of deuterated substrate “unmasking” an 18O isotope effect on an O2 reduction step occurring before or with C−H cleavage.159 In addition, the DFT studies suggested the H-abstraction step occurs on 4 that retains its hydroxyl proton,286 while subsequent QM/MM studies suggested this proton is absent during H-abstraction (i.e., II in Scheme 11).287 Finally, a QM/MM investigation invoked an Fe(IV)O intermediate that also produces the methylphosphonate radical (Scheme 26),288 but distinguishing between mechanisms involving a high valence ferryl species (II, Scheme 26) versus a bridged alkylperoxide (IX, Scheme 24C) awaits experimental verification. 5731


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Scheme 27

Scheme 28. Proposed Mechanisms of (A) Phosphoglycerate Mutase from Sulfolobus sofataricus and (B) PhpG

4.2.4. PhpE, PhpJ, and PhpF Catalyze Phosphonoformate (30) Synthesis and Activation. In vivo gene inactivation studies demonstrated the conversion of 35 to phosphonoformate 30 by the sequential actions of PhpE and PhpJ. This is followed by PhpF-catalyzed activation to afford the CMP-phosphonoformate 93 (Scheme 23).133 The phpE gene possesses sequence similarity to 2-hydroxyacid alcohol dehydrogenases and the ΔphpE mutant of S. viridochromogenes accumulated 35 in the culture broth, suggesting PhpE catalyzes the oxidation of 35 to phosphonoformaldehyde 91. Although PhpE activity could not be detected after overexpression in either E. coli or S. lividans, further evidence for PhpE-catalyzed oxidation was obtained from the accumulation of aminomethylphosphonate 92 in the ΔphpJ mutant. Despite not directly observing the accumulation of the proposed product 91, the accumulation of 92 in the culture broth was reminiscent of 3 accumulation in the ΔphpC mutant. These results imply that the aldehydes 2 and 91 are rapidly aminated either nonenzymatically or by promiscuous aminotransferases to accumulate 3 and 30 in cultures of ΔphpC and ΔphpJ, respectively. The nucleotidyl transferase homologue encoded by phpF was similarly inactivated, and the resulting ΔphpF mutant was found to accumulate 30 and its spontaneous decarboxylation product phosphite 34, as well as small amounts of 35. Additional in vivo evidence for the PhpF-catalyzed transformation of 30 to 93 was obtained when 10b production was restored by feeding 30 to cultures of upstream gene mutants ΔphpD and ΔphpE, but production was not restored by feeding 30 to the downstream mutant ΔphpF. In vitro characterization of PhpF confirmed that this enzyme could transform CTP and 30 to 93 and pyrophosphate, and that the reaction could be driven to near completion by including inorganic pyrophosphatase to hydrolyze pyrophosphate to Pi (Scheme 27).133 These experiments collectively provide a model where phpE and phpJ respectively encode HMP dehydrogenase and phosphonoformaldehyde dehydrogenase

to furnish phosphonoformate 30, which is then activated as the CMP-conjugate 93 by the phpF-encoded CTP:phosphonoformate cytidylyltransferase. 4.2.5. Glycolytic Enzyme Homologues PhpG and PhpH Generate CPEP (95). Nonproducing mutants of S. hygroscopicus provided the first evidence for the biosynthetic intermediacy of phosphonoformate 30,281 which led to the initial proposal that carboxyphosphonoenolpyruvate (CPEP, 95) originated from a condensation between 30 and 24.103 In vivo gene disruption studies implicated a so-called “CPEP synthase” encoded by bcpE in S. hygroscopicus, which is homologous to phpH in S. viridochromogenes.289 However, homology to the enolase family290 suggested that a direct displacement transesterification reaction was unlikely to be catalyzed by phpH, and apparent cotranscription with the phosphoglycerate mutase homologue phpG implied parallels with glycolytic reactions leading to 24 (boxed reactions, Scheme 23).277 Interestingly, PhpG is closely related to a 2,3bisphosphoglycerate-independent phosphoglycerate mutase (iPGM) from the archaeon Sulfolobus solfataricus. This enzyme transfers the γ-phosphate from ATP or GTP to the 2-hydroxyl group of 3-phosphoglycerate 89 via a phosphoserine intermediate to afford 2-phosphoglycerate 90 (Scheme 28A).291 PhpG has been proposed to catalyze the analogous transfer of phosphonoformate from 93 to a three-carbon unit such as 89 to afford 94 (Scheme 28B). Although neither ΔphpG nor ΔphpH mutants accumulated Pn intermediates, both enzymes must act after 30 formation because feeding this intermediate to the mutants was unable to restore phosphinothricin production.133 While there is not yet any direct biochemical evidence for the PhpG and PhpH-catalyzed reactions, the known PhpF-catalyzed production of 93 together with the in vivo data is most consistent with the three-step formation of CPEP 95 from 30 as outlined in Scheme 23. The proposed PhpG and PhpH transformations thus appear to represent 5732


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Scheme 29. Proposed Similarities in Pathway Logic between 10a Biosynthesis (Bottom) and TCA Cycle and Glutamate Synthesis (Top)a

a

TCA cycle enzymes isocitrate dehydrogenase and glutamate dehydrogenase may also catalyze the similar steps during 10a biosynthesis.

Scheme 30. Reactions Catalyzed by (A) (S)-Citrate Synthase and Aconitase of the TCA Cycle, (B) (R)-Citrate Synthase and Aconitase of the TCA Cycle in Some Anaerobic Bacteria, (C) Pms and Pmi during Phosphinothricin Biosynthesisa

a

The boxed reactions catalyzed by Pmi have not been demonstrated in vitro, nor has 99 been isolated.

CPEP mutase may catalyze a rate-limiting step in 10b biosynthesis.294 Although CPEP mutase and PEP mutase catalyze similar reactions, they exhibit strict substrate specificities. Additionally, CPEP mutase is weakly inhibited by PnPy 1 while PEP mutase is weakly inhibited by 95 and 97.292 Knowles and co-workers subsequently synthesized 95 and demonstrated that it is a substrate for CPEP mutase in vitro.295,296 A continuous coupled assay was developed using malate dehydrogenase and NADH to reduce the product 97, which yielded a kcat of 0.020 s−1 and KM of 0.27 mM. The low turnover number was proposed to result from the endergonic conversion of 95 to the intermediate carboxyphosphinopyruvate 96 prior to exergonic decarboxylation, which is similar to the unfavorable conversion of 24 to 1 catalyzed by PEP mutase. The solution stability of 96 ruled out spontaneous decarbox-

additional examples of chemical logic in phosphonate biosynthesis that is borrowed from primary metabolism.9 4.2.6. Carboxyphosphonoenolpyruvate (CPEP) Mutase. CPEP mutase catalyzes the transformation of CPEP 95 to phosphinopyruvate 97 (Scheme 23) via a rearrangement similar to that catalyzed by PEP mutase (see section 3.1) and was discovered in nonproducing mutants of S. hygroscopicus.103,292−294 The mutant NP213 accumulated CPEP 95 and its cell extract restored 10b production to mutant NP71. This enabled purification of CPEP mutase from NP71 using a bioassay that monitored 10b production from NP213 mycelia that had been supplemented with enzyme fractions. The purified enzyme converted 95 to phosphinopyruvate 97 with a requirement for Mn2+ or Mg2+ metal ions, and its overproduction in a high-producer strain HP5−29 suggested that 5733


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dehydrogenase converting 99 to 100, and glutamate dehydrogenase or other aminotransferases catalyzing transamination to 101. First, there are no obvious gene candidates or homologues of these enzymes encoded by the gene clusters sequenced to date,278 and the extensive work by Seto and co-workers failed to generate mutants blocked at these steps.275 Second, 98 was converted to 101 by the glutamic acid producer Brevibacterium lactofermentum, which does not produce 10 and therefore would rely on TCA cycle enzymes to accomplish this transformation. Moreover, 100 was a substrate, albeit a relatively poor one, for transamination to 101 by the three common primary metabolic enzymes aspartate aminotransferase, alanine aminotransferase, and glutamate dehydrogenase.275,300,309 Notably, this is consistent with the apparent promiscuity of aminotransferases during phosphinothricin biosynthesis as evidenced by the accumulation of 3 and 92 instead of aldehydes 2 and 91, respectively (Scheme 23). Overall, the collective evidence suggests that the TCA cycle enzyme isocitrate dehydrogenase catalyzes decarboxylation of 99 to 100, followed by transamination by glutamate dehydrogenase or another common aminotransferase. It is curious that only the first two of the four enzymes catalyzing the 97 to 101 transformation are apparently dedicated to this pathway. The presumed stereochemistry of 99 (Scheme 30) and its efficiency as a substrate for isocitrate dehydrogenase remains unknown. 4.2.8. Phosphinothricin Acetyltransferase (PAT): From Self-Protection to Crop Protection. Analysis of S. hygroscopicus mutants with blocked phosphinothricin biosynthetic pathways revealed the unexpected accumulation of Nacetylated intermediates, with 102 as a major intermediate common to all mutants.313,314 N-Acetyltransferases were then found to be encoded by the bar gene in S. hygroscopicus315 and the pat gene in S. viridochromogenes.316,317 Subsequent sitespecific mutagenesis indicated that inactivating the bar gene in S. hygroscopicus abolished phosphinothricin production and accumulated 101,318 which further implied that the substrate for peptide bond formation is N-acetylated. Phosphinothricin acetyltransferase (PAT) encoded by the bar gene displayed similar kcat values (although the values were not reported), but the KM varied widely among substrates, with 10a and 101 possessing the lowest values of 0.06 and 2 mM, respectively.315 Other substrates tested and found to be acetylated by PAT were 10b, glutamate, 85, and ∂-hydroxylysine, while desmethyl-10b was not acetylated. Although the apparent specificity of PAT for 10a over 101 might at first appear confusing with respect to the timing of acetylation in the biosynthetic pathway, it can be readily understood in the context of acetylation as a mechanism of self-protection against the toxic end product 10a (discussed below). PAT-like enzymes have been widely identified in bacterial genomes and in vitro studies have uncovered varying specificities to several substrates (Figure 10). The pat gene product from S. viridochromogenes shares 85% sequence identity with that of bar and its in vivo expression is phenotypically identical despite not being extensively characterized in vitro.316,317 A PAT homologue (ScPAT) from the prolific natural product producer S. coelicolor A3(2) shares ∼30% sequence identity with the bar gene product, but the genomic context of bar has no apparent relation to phosphinothricin and the enzyme exhibits a high KM (∼1 mM) toward 10a. Moreover, it confers resistance only when expressed from a high copy number plasmid and glutamate is not a substrate,

ylation and its turnover by CPEP mutase demonstrated chemical competence. However, the slow turnover of 96 (kcat = 7.6 × 10−4 s−1, KM = 2.2 μM) revealed that the free intermediate was not kinetically competent, which could be explained if the enzyme does not release 96 during catalysis.296 4.2.7. Similarities to the Citric Acid Cycle en Route to Demethylphosphinothricin (101). The four transformations from 97 to 101 bear striking resemblance to those catalyzed by citrate synthase, aconitase, isocitrate dehydrogenase, and glutamate dehydrogenase of the tricarboxylic acid (TCA) cycle (Scheme 29). This relationship was recognized by Seto and co-workers upon isolating the oxaloacetate analog 97297 and the glutamate analog demethylphosphinothricin 101;298 the intermediacy of the citrate analog 98 was subsequently demonstrated after it accumulated in the presence of the aconitase inhibitor monofluoroacetic acid.299 Insight into the citrate synthase homologue catalyzing the transformation of 97 to 98 was provided by the intriguing discovery that 98 possessed the R-configuration arising from a Claisen condensation with acetyl-CoA occurring at the re-face of 97 (Scheme 30C).300 Apart from a few (R)-citrate synthases (also called Re-citrate synthases) in anaerobic bacteria that carboxymethylate into the pro-R position of citrate (Scheme 30B),301−303 this contrasts with the well established sistereofacial specificity for oxaloacetate in citrate synthase (Scheme 30A).304−307 Indeed, purification of the enzyme phosphinomethylmalate (PMM) synthase from S. hygroscopicus confirmed the production of (R)-98 from 97,164 while the same substrate presented to canonical (S)-citrate synthases (also called Si-citrate synthases) yielded (S)-98.300,308 PMM synthase is similar to (R)-citrate synthase in its metal ion requirements (preference for Co2+ or Mn2+) and inhibitor sensitivities, although only (R)-citrate synthases (present in anaerobes) are inactivated by oxygen.164 An evolutionary relationship between PMM synthase and (R)-citrate synthase is also suggested based on cross reactivity of their respective substrates164,309 and their coclustering in a subgroup of Claisen condensation-like enzymes within the DRE-TIM metallolyase superfamily.310 Apart from phosphinomethylmalate isomerase (Pmi) encoded by the pmi gene, the evidence for the remaining transformations en route to 101 is sparse. Pmi from S. viridochromogenes shares 52% sequence identity with the TCA cycle aconitase AcnA of the same organism and presumably originates from gene duplication of an ancestral aconitase.311,312 Aconitase possesses a [4Fe-4S] center that is unusual because: (i) it is required in a nonredox role as a Lewis acid to catalyze a simple elimination-addition reaction307 and (ii) it responds to oxidative stress by disassembling to an inactive [3Fe-4S] form.312 Pmi can be produced heterologously in S. lividans but not E. coli, which has been attributed to specific accessory proteins that are required to assemble the [4Fe-4S] cluster. However, no aconitase activity could be detected from purified Pmi using citrate as a substrate.311 Although 98 was not available to the researchers to test as a substrate, the results of in vivo experiments were consistent with Pmi specifically catalyzing the conversion of 98 to 99; introducing the pmi gene did not restore the wild-type phenotype to a S. viridochromogenes ΔacnA mutant, whereas pmi, but not acnA, could restore 10b production to the Δpmi mutant. Overall, these experiments suggest that Pmi and not the orthologous TCA cycle aconitase AcnA catalyzes the transformation of 98 to 99. Several lines of evidence suggest that TCA cycle enzymes catalyze the final transformations to 101, with isocitrate 5734


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4.2.9. Breaking the NRPS Specificity Code: PhsA, PhsB, and PhsC. The three proteins PhsA, PhsB, and PhsB are stand-alone nonribosomal peptide synthetases (NRPSs) that catalyze the attachment of amino acids to 10a. NRPSs are modular enzymes comprising an adenylation (A) domain that uses ATP to activate and load a specific amino acid onto the phosphopantetheinyl (Ppant) thiol of a peptidyl carrier protein (PCP); extension modules also include a condensation (C) domain catalyzing peptide bond formation between PCP-linked thioesters.335 Characterization of the phosphinothricin pathway NRPS enzymes was initiated by inactivating the phsA gene in S. viridochromogenes, which abolished 10b production and led to the accumulation of 102 (Scheme 23).336−338 In vitro characterization of PhsA revealed a strict specificity for Nacetylated substrates 102 and N-acetyl-10a, and minor activity toward N-acetyl-glutamate.339,340 This preference for Nacetylated substrates is unusual because most NRPS systems incorporate α-amino acids. Insight into this deviation came from a recent analysis that uncovered a novel ‘N-Ac(DM)PT’ (N-acetyl(dimethyl)phosphinothricin; i.e. either 102 or Nacetyl-10a) specificity code that is shared among PhsA enzymes in the three sequenced 10a-producing strains (S. viridochromogenes, S. hygroscopicus, and K. phosalacinea).278 Interestingly, the A-domain of PhsA possesses Val in place of a conserved Asp residue that has only been substituted in rare cases where the A-domain selects a non α-amine substrate.278,341 These studies demonstrated that PhsA is a rare N-acetylamino acid selecting initiation module, but it remains unclear whether it selects and installs 102 or N-acetyl-10a onto its PCP domain. Although 102 was the implied substrate based on its accumulation in a ΔphsA mutant of S. viridochromogenes,336,337 another mutant producer strain of S. hygroscopicus accumulated N-acetyl-10a and 104a,313 suggesting that PhsA can accept Pmethylated substrates. In addition, ATP-PPi exchange assays revealed a 6-fold greater kcat/KM value for racemic N-acetyl-10a over 102. This result was further supported by the preference for loading the N-acetyl-10a onto the Ppant arm of PhsA.340 However, the accumulation of N-acetyl-10a may result from the action of peptidases that are ultimately responsible for its activation.342 The identity of the “true” PhsA substrate may ultimately be revealed by understanding the pathway timing of the P-methyltransferase (Section 4.2.11), which may possess sufficient substrate promiscuity to supply both substrates to PhsA. In S. viridochromogenes, both PhsB and PhsC incorporate alanine as predicted from the product structure and their Adomain specificity codes.98,339,341 The recent sequence of the 10c biosynthetic gene cluster from K. phosalacinea finally provided insight into the longstanding puzzle regarding the ordering of thiotemplate assembly. Comparing the A-domain specificity codes for all three modules showed that only PhsC from K. phosalacinea was predicted to activate Leu, which suggests that PhsC acts last. Overall, the data suggest a model in which the initiation module PhsA generates a product for PhsB followed by the attachment of the final amino acid by PhsC (Scheme 23).278 Interestingly, phsB and phsC are apparently cotranscribed in a region about 20 kb from phsA.98 This genetic organization is consistent with a modular biosynthetic logic employing a common 10a-specific PhsA initiation module that can be combined with different extension modules to fashion diverse 10a-linked peptides. 4.2.10. Putative Transacylase PhpL and Putative Thioesterase PhpM. Initial characterization of the genetic

Figure 10. Substrates for N-acetyltransferases with structural homologues glutamate and methionine.

suggesting an alternative substrate and physiological role for some PAT-like enzymes.319 One possible role was finally suggested in 2007 when Brown and colleagues reported the structure and activity of the so-called pita acetyltransferase from Pseudomonas aeruginosa.320 Like the PAT enzymes from 10b biosynthetic pathways, pita belongs to the GNAT (GCN5related N-acetyltransferase) family of enzymes that are widely distributed among Gram-negative and Gram-positive bacteria and possesses ∼37% sequence identity to the bar gene product of S. hygroscopicus. However, pita possessed no activity toward 10a but rapidly acetylated methionine sulfoximine 85 and methionine sulfone (112) with KM values of 1.3 mM for both substrates and respective kcat values of 505 and 610 s−1. A similar structure and substrate profile were found for an enzyme from Acinetobacter baylyi ADP1.321 Likewise, a similar substrate preference for 85 and 112 was found for the methionine derivative detoxifier A (MddA) enzyme from Salmonella enterica, with no activity detected toward 10a, methionine sulfoxide 113, buthionine sulfoximine, methionine, or glutamine.322 In contrast, a so-called MAT (methionine sulfone Nacetyltransferase) enzyme from Nocardia sp. acetylated both 10a and 112 with respective KM values of 2.2 and 14.3 mM, although full kinetic parameters were not reported.323 Full kinetic constants toward both 10a (kcat = 131 min−1, KM = 76 μM) and 85 (kcat = 29.8 min−1, KM = 156 μM) were reported for the RePAT enzyme isolated from 10a-resistant Rhodococcus sp. from a marine environment.324 As with PAT specificity, the possible physiological roles of PAT-like enzymes are varied. Like 10a, 85 occurs naturally as a toxic component of plants in the genus Connaraceae325 and therefore may require detoxification when encountered by soil bacteria. It was also identified as a toxic byproduct of flour bleaching that has been proposed to contribute to neurological disorders.326,327 Interestingly, the soil bacterium P. putida KT2440 possesses both 10a- and 85-specific acetyltransferase enzymes PhoN1 and PhoN2, respectively, suggesting that this species has evolved in the presence of both of these natural compounds.328 Methionine oxidation in proteins can generate 113,329 suggesting that these acetyltransferases might be mobilized in response to oxidative stress. It is not clear why acetylation is required prior to peptide bond formation during phosphinothricin biosynthesis (Section 4.2.9) but may reflect the toxicity of 101 and/or incompletely extended peptides released as shunt metabolites. GNAT-catalyzed acetylation is a common mechanism of protection against a myriad of toxins, and N-acetylated biosynthetic intermediates have been observed or proposed prior to peptide bond formation for several natural products.330−332 Indeed, the protective effect of N-acetylation has been exploited by the crop protection industry in the development of 10a-resistant plants expressing the bar or pat gene.333,334 5735


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Scheme 31. Proposed Roles for GXCXG-Containing Type II TEs PhpL, SyrC, and BtaH in Transferring Amino Acids from a Single PCP Domain Module to a Module Possessing Two PCP Domains

domain of PhsB, which is an unusual module because it possesses two PCP domains.278 Interestingly, PhpL homologues SyrB1 and BtaH are also proposed to transfer amino acids to dual PCP-containing modules in bactobolin and syringomycin biosynthesis, respectively (Scheme 31). Overall, this suggests that PhpL catalyzes intermodular acyl transfer while PhpM is the TE that catalyzes peptide release (Scheme 23). 4.2.11. Cobalamin-Dependent P-Methyltransferase PhpK. In contrast to the relative ubiquity of N-, C-, and Omethyltransferases, P-methylation is another fascinating example of unprecedented enzymology giving rise to the only C−P− C linkage known in Nature.10,216 A role for cobalamin (vitamin B 12 ) in P-methylation was initially proposed due to accumulation of 10b in Co2+-containing cultures of S. hygroscopicus, while 101 and desmethyl-10b accumulated either without Co2+ or in vitamin B12 auxotrophs.275,298 Enzyme activity was observed by heterologous expression of a complementing DNA fragment in S. lividans, which also demonstrated the requirement for N-acetylated substrates 102 or 104a and the vitamin B12 derivative methylcobalamin (MeCbl) as the methyl donor by incorporation of [14CH3]MeCbl.351 The apparent ability of PhpK to accept either 102 or 103a is consistent with the observed ability of PhsA to accept either N-acetyl-10a or 104a,313 suggesting that PhpK may act either before or after peptide bond formation. Sequencing revealed that putative methyltransferases encoded by bcpD from S. hygroscopicus and phpK from S. viridochromogenes and K. phosalacinea276−278,352 are related to the Fom3 methyltransferase involved in fosfomycin biosynthesis that was discussed in section 4.1.3. Like Fom3, PhpK is a member of the class B radical SAM methyltransferase (RSMT)

organization of the 10b biosynthetic pathways revealed clustering of the acetyltransferase (section 4.2.8) and deacetylase (section 4.2.12) genes with two genes encoding putative thioesterases (TEs).343,344 Recent sequencing demonstrated conserved organization among S. viridochromogenes, S. hygroscopicus, and K. phosalacinea and the TE genes were named phpL and phpM.278 Apparent translational coupling of the deacetylase and the two TEs implies stoichiometric production consistent with these three enzymes catalyzing sequential steps in the biosynthetic pathway.344 Gene disruption and complementation experiments showed that phpL and phpM were not essential but were required for efficient production of 10b, suggesting that PhpL and PhpM function as editing TEs to optimize biosynthetic flux by removing aberrant intermediates from the NRPS complex.343 Comparative bioinformatic analysis of PhpL and PhpM from the three bacterial species above implies respective transacylase and thioesterase functions.278 In contrast to the integrated Type I TEs that typically hydrolyze mature peptides from the final module of an NRPS assembly line, the 10a biosynthetic gene clusters possess PhpL and PhpM as stand-alone, or Type II, TEs.345 Significantly, although PhpM possesses the α/βhydrolase fold and GXSXG motif that is typical of many TE enzymes, PhpL has a variant GXCXG motif that is associated with increased transacylase activity.346 Indeed, similar GXCXGcontaining Type II TEs in several biosynthetic gene clusters are proposed to function as transacylases to shuttle PCP-tethered intermediates between modules,347,348 including biochemically verified PCP-to-PCP shuttles operating during coronamic acid349 and syringomycin350 biosynthesis. Based on these similarities Blodgett et al. proposed that PhpL catalyzes transfer of 102 from the PCP domain of PhsA to the N-terminal PCP 5736


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Scheme 32. Proposed Mechanisms for PhpK Involving Either (A) Homolytic or (B) Heterolytic Catalysis

Recently the Wang group characterized a new PhpK-like Pmethyltransferase from the marine bacterium Shewanella denitrif icans OS217.356 In contrast to the apparently strict requirement of PhpK for N-acetylated substrates, this methyltransferase, called SD_1168, catalyzed P-methylation of both 101 and 102. Although using titanium(III) citrate instead of sodium dithionite as a reductant elicited a second turnover of the enzyme that was not observed in the previous analysis of PhpK from K. phosalacinea,354 significant technical hurdles remain to be overcome before this fascinating enzyme can be fully understood and exploited. 4.2.12. Deacetylase. Among the mutants of S. hygroscopicus SF-1293 characterized by Seto and co-workers, the NP8 mutant accumulated the late-stage intermediate 104a together with N-acetyl-10a.313 This suggested that NP-8 was unable to produce the deacetylase required to remove the Nacetyl group and furnish 10a in the final biosynthetic step (Scheme 23). The isolation of N-acetyl-10a could arise from Pmethyltransferase-catalyzed methylation of 102, which was observed for PhpK354 (section 4.2.11) and is consistent with the apparent preference of PhsA for N-acetyl-10a in the subsequent peptide bond formation step (section 4.2.9).340 However, when considered together with the observed accumulation of 102 in a ΔphsA mutant of S. viridochromogenes,336,337 deacetylation occurring as the final step as shown in Scheme 23 is most probable. Although characterization of the deacetylase gene product of either bah from S. hygroscopicus or dea from S. viridochromogenes could clarify the issue of pathway timing, in vitro experimental data for this enzyme has not been obtained. Thompson and coworkers showed that a 392 bp fragment of the bah gene restored 10b production to the NP-8 mutant,357 and bioinformatic analysis revealed significant sequence similarity to lipases,344 including the characteristic GXSXG motif that defines the nucleophilic elbow of the α/β-hydrolase enzyme family.358,359 Similarly, abolished production upon disruption of the dea gene in S. viridochromogenes could be restored by cosynthesis with a mutant possessing a functional copy of the dea gene.337 In addition, the absence of membrane-spanning sequence features in the Dea protein suggests that 10b is generated inside the cell prior to export.277 Interestingly, deacetylation of N-acetyl-10a has been explored as a male sterility system for use in plant breeding. In contrast to the proposed deacetylation of 104a by Dea or Bah, the male sterility system employs tapetum-specific expression of

subgroup of the broader radical SAM enzyme superfamily.214−216,353 In contrast to the methylation of nucleophilic acceptors catalyzed by typical SAM-dependent methyltransferases, RSMTs have expanded Nature’s methylation capacity to include a variety of electrophilic targets. Although class B RSMTs are a large subfamily, it was only recently that PhpK from K. phosalacinea became the first member to be reconstituted in vitro.354 The enzyme was heterologously expressed in E. coli and the insoluble protein was anaerobically refolded and purified, and subsequent anaerobic reconstitution of the [4Fe-4S] cluster was confirmed by EPR spectroscopy. Under anaerobic conditions in the presence of SAM, MeCbl(III), and sodium dithionite, a single turnover of PhpK apparently transformed 102 to N-acetyl-10a based on the appearance of a new 1H−31P cross peak observed in a two-dimensional gradient heteronuclear single-quantum correlation NMR experiment. The inability to catalyze multiple turnovers was presumed to result from a side reaction of dithionite to reduce MeCbl(III) to a catalytically incompetent MeCbl(II), while dithionate would also be required to reduce the inactive oxidized +2 state of the [4Fe-4S] cluster thought to arise during the RSMT catalytic cycle. Indeed, no turnover was observed in the absence of dithionite, which is consistent with a radical-dependent methylation mechanism. Evidence for the role of MeCbl as the methyl donor was obtained by the appearance of new cross-peaks corresponding to N-acetyl-10a when [13CH3]-MeCbl was used in the reactions. A subsequent study with [13CH3]-MeCbl as the methyl donor directly demonstrated PhpK-catalyzed formation of the 13CH3−31P bond by multiple quantum 1H−13C−31P (HCP) NMR.355 Like other class B RSMTs such as Fom3 and GenK,217 both heterolytic and homolytic mechanisms can be proposed for PhpK-catalyzed methylation (Scheme 32). Although homolytic mechanisms employing MeCbl as methyl radical donors are unusual (Scheme 32A), it is consistent with the requirement of the [4Fe-4S] cluster and external reductant. This contrasts with the relatively straightforward SN2 displacement mechanism of Scheme 32B lacking an obvious role for the [4Fe-4S] cluster. Curiously, either mechanism is energetically expensive because it would require two equivalents of SAM to generate one molecule of product. This stoichiometry is supported by the observation of nearly equivalent quantities of methylated product, 5′-deoxyadenosine 72, and S-adenosylhomocysteine 71 in the GenK-catalyzed reaction.217 5737


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the argE gene from E. coli, which encodes an N-acetyl-Lornithine deacetylase possessing a dinuclear Zn(II) active site.360,361 In order to selectively kill the anthers and prevent pollen production, the system takes advantage of the accumulation of N-acetyl-10a in the upper parts of the plant and the tapetum-specific deactylase activity.362−364 Overall, the use of ArgE to catalyze a deacetylation of N-acetyl-10a instead of Dea or Bah is consistent with the latter enzymes’ role in deacetylating the tripeptide in the final step. 4.2.13. Resistance Mechanisms. Because 10a inhibits GS (section 4.2.1), it slows cell growth by interfering with normal nitrogen metabolism. This inhibition can be countered by an exogenous supply of glutamine or by two resistance mechanisms that are known to occur in 10a-producing organisms. The first mechanism involves detoxifying the antibiotic by N-acetylation, which has been exploited by the crop protection industry as described in section 4.2.8. The second mechanism of resistance arises from overproduction or modification of the GS target enzyme. Many soil bacteria, including 10a producers S. hygroscopicus365 and S. viridochromogenes,366 possess GSII enzymes common to eukaryotes in addition to the typical bacterial GSI enzyme, and overexpression of this additional GSII enzyme in S. viridochromogenes conferred resistance to 10a.366 Mutations in plant GS enzymes have been identified and shown to confer resistance in maize, yeast, and Arabidopsis.367,368

then hydrolyzed inside the cell by peptidases to liberate the phosphonate warhead.342 4.3.1. Mechanism of Action. The mechanism of action of 11 is probably comparable to that of the structurally similar peptide alaphosphin 117 (Scheme 33). As described above, the Trojan horse peptide 117 is imported and cleaved by peptide permeases and peptidases, respectively, to release the alanine racemase inhibitor 153 inside the cell (Scheme 33A).372,373 Similarly, 11-resistant deletion mutants of Salmonella enterica were used to reveal that specific peptide transporters and peptidases are also required for growth inhibition by 11.99 Incubation of an S. enterica peptidase with 11 led to accumulation of methyl acetylphosphonate (52), a pyruvate (53) analog known to inhibit pyruvate dehydrogenase, pyruvate oxidase,96 and bacterial 1-deoxy-D-xylulose 5-phosphate (DXP) synthase (Scheme 33B).374−376 This experiment also revealed a transient phosphonate intermediate with a 31P NMR chemical shift of 6.2 ppm probably corresponding to the direct cleavage product 1-aminovinylphosphonate O-monomethyl ester 118 or its tautomer 1-iminoethylphosphonate O-monomethyl ester 119. This suggests that, in the case of 11, the peptide has a dual role of facilitating uptake and preventing enamine-imine tautomerization of the vinyl amine.99 The cellular targets of 11 remain unknown, but the demonstrated importance of the O-methyl and vinyl moieties revealed by structure−activity relationship analysis377 suggests that 52 is the primary active compound, although the toxicity of electrophiles 118 and/or 119 cannot be ruled out.99 As a structural analog of pyruvate 53, 52 is expected to disrupt pyruvate-processing enzymes, with the reduced negative charge of the methylated phosphonate monoanion378 possessing a charge distribution more representative of a carboxylic acid substrate.377 Indeed, 52 inhibits pyruvate dehydrogenase (PDH) with a Ki of 5 × 10−8 M, which is 125-fold lower than unmethylated acetylphosphonate.96 Extensive characterization of the thiamine pyrophosphate (TPP, 120)-dependent reaction catalyzed by the E1 subunit of the E. coli pyruvate dehydrogenase (PDH) complex has unveiled the mechanism of inhibition by 52 (Scheme 34). As a typical TPP-dependent decarboxylase, the E1 subunit initiates the catalytic cycle by forming the ylid of 120, which attacks the carbonyl carbon of pyruvate 53. The resulting tetrahedral substrate-TPP adduct 2(2-lactyl)TPP 121 is readily decarboxylated to generate 2-(1hydroxyethyl)TPP 122, which is then transferred to the lipoyl moiety of the E2 subunit.307,378 Chemical synthesis of 123 and its addition to PDH enzyme lacking TPP 120 restored activity to the enzyme,379 demonstrating reversibility of the previously observed inhibition by 52.380 Significantly, crystal structures have been obtained for 123 alone381 and in complex with the E1 subunit of E. coli PDH382 and pyruvate oxidase from Lactobacillus plantarum.383 All structures reveal that the C−P bond remains intact and is perpendicular to the thiazolium ring, representing a conformation expected to promote decarboxylation of 121 by maximizing orbital overlap between carboxylate-derived electrons and the π-system of the thiazolium cation.378 Recently, 52 has also been identified as a potent inhibitor of 1-deoxy-D-xylulose 5-phosphate (DXP) synthase from the nonmevalonate, or MEP, isoprenoid biosynthetic pathway in E. coli and Plasmodium species.374−376 4.3.2. Biosynthesis of 11. As described above, the Omethyl group of 11 attenuates phosphonate negative charge to more closely resemble the charge distribution of a carboxylic acid substrate or an enzymatic tetrahedral intermediate thereof.

4.3. Dehydrophos (11)

Dehydrophos 11 was first isolated by Eli Lilly from the culture supernatant of Streptomyces luridus as a broad-spectrum antibiotic with efficacy against Salmonella in a chicken model.369 Originally called A53868, its initially proposed chemical structure 114 was revised first to 115370 and finally to 11 in 2007 to reveal a novel O-methylated 1-aminovinylphosphonate moiety that, based on its structural similarity to dehydroalanine (116), resulted in 11 being renamed dehydrophos (Figure 11).371 Interestingly, synthesis of 115 as

Figure 11. Initial (114), reassigned (115), and final structural reassignment of dehyrophos (11) highlighting in blue the structural similarity to dehydroalanine (116).

part of the final structural reassignment revealed that this compound was active against E. coli.371 Due to the amide linkage of the aminovinylphosphonate to a Gly-Leu dipeptide, 11 joins an expanding group of peptidyl phosphonates such as phosphinothricin peptides 10b−d, APPA-containing peptides 14b−h (e.g., rhizocticins, plumbemycins, and phosacetamycin), argolaphos 15, fosfazinomycins 13, and the synthetic dipeptide alaphosphin 117.372 The appended peptides are believed to enhance cellular uptake via peptide permeases by disguising the phosphonate antibiotic in a “Trojan horse” peptide, which is 5738


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Scheme 33. Hydrolytic Processing of Trojan Horse Peptides (A) Alaphosphin 117 to Release Alanine Analog 153 and (B) Dehydrophos 11 to Liberate Pyruvate Analog 52

Scheme 34. Mechanism of Thiamine Pyrophosphate-Mediated Decarboxylation of Pyruvate (Route a) and Inhibition by 52 (Route b) by the E. coli Pyruvate Dehydrogenase Complex

Scheme 35. Proposed Biosynthetic Pathway for Dehydrophos 11a

Boxed areas represent two possible pathways: route a involves 127 as a discrete intermediate formed by an unknown enzyme (denoted by “?”); route b represents reactions occurring in the DhpH-N active site as detailed in Scheme 37.

a

gene cluster consists of 17 genes required to heterologously produce 11, beginning with the usual suspects encoding the PEP mutase (DhpE), decarboxylase (DhpF), and alcohol

This carboxylate mimicry is reminiscent of the C−P−C moiety of 10a, but a much simpler biosynthetic strategy is employed during the biosynthesis of 11 (Scheme 35).384 The biosynthetic 5739


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Scheme 36. Comparison of (A) Initially Proposed Roles of DhpB, DhpC, DhpD, and DhpHa

a

Dotted lines and gray shading indicated reactions and/or products not observed. (B) Serine biosynthetic pathway enzymes that correspond to Dhp enzymes directly above. (C) DhpD reactions that have been characterized in vitro.

results. First, 124 was generated upon treatment of this intermediate with alkaline phosphatase, demonstrating that the intermediate is a phosphate ester of 124. Second, the absence of P−P splitting in the NMR spectrum indicates that phosphorylation occurs at the more distal 2-hydroxyl group. In contrast to DhpB and DhpC, the functional assignment of the PLP-dependent aminotransferase DhpD was less straightforward. Although ΔdphD deletion mutants failed to produce 11, they surprisingly did not accumulate the expected intermediates 124−126, suggesting a possibly alternative function for DhpD.384 Important insight was obtained from in vitro characterization of the enzymatic reaction using purified His6-DhpD.95 Surprisingly, no reaction occurred when 127 was incubated with a variety of amino acceptors, implying that DhpD does not physiologically catalyze the reversible formation of 127 from 126. However, 153 and the phosphonate analog of serine (Ser(P), 139) were respectively transformed to 129 and 1-oxo-2-hydroxyethylphosphonate (OH-EP, 138) with pyruvate as the amino acceptor (Scheme 36C). Steady-state kinetic parameters were obtained for amination of 129 by coupling pyruvate formation with NADH oxidation using lactate dehydrogenase, and the same assay was used to show 100-fold lower catalytic efficiency toward 52 (Scheme 36C). Overall, these results support DhpD catalyzing 153 formation as shown in Scheme 35 instead of 127 as initially proposed (Scheme 36A). 4.3.5. DhpH: a tRNA-Dependent Peptide Ligase also Catalyzing PLP-Dependent Elimination. DhpH is a twodomain protein consisting of an N-terminal PLP domain (DhpH-N) with similarity to aminotransferases and a Cterminal GCN5-related N-acetyltransferase (GNAT) domain (DhpH-C) found in acetyl- and peptidyltransferases.95,384 This dual activity was initially interpreted to support the pathway shown in Scheme 36A because it elegantly prevented loss of the unstable eneamine 128 resulting from tautomerization and hydrolysis to 129 (Scheme 35). Specifically, the unstable intermediate 128, generated by β-elimination activity of the DhpH-N PLP domain,385 could be protected from hydrolysis via immediate capture of the reactive amine by the adjacent DhpH-C peptidyltransferase domain. Indeed, both His6-DhpH and His6-DhpH-N generated 129 as the expected product of βelimination, but 136 was not observed when rac-127 was

dehydrogenase (DhpG) that generate HEP 4. In contrast to initial proposals based on biochemical precedent,384 subsequent work on the remainder of the pathway revealed fascinating enzymes contributing to an unexpected biosynthetic logic.95 The second phase of biosynthesis consists of many unique reactions and generates the intermediate 153, which was previously recognized only as a manmade compound used as the bioactive warhead of alaphosphin 117. The final four steps involve unusual enzymes catalyzing nonribosomal tRNAdependent formation of two peptide bonds during the installation of leucine and glycine, O-methylation of the phosphonic acid, and carbon−carbon double bond formation to furnish the vinyl moiety. 4.3.3. Dioxygenase DhpA. The biosynthetic role of DhpA was assigned based on a combination of in vivo and in vitro experiments.384 The accumulation of 4 in the supernatant of a ΔdphA mutant suggested that DhpA catalyzes the hydroxylation of 4, and this protein possesses homology to known αketoglutarate-dependent dioxygenases such as PhnY* discussed in section 3.5. Incubation of purified His6-DhpA with 4 in the presence of α-ketoglutarate and ferrous ammonium sulfate generated the hydroxylated product 124 (Scheme 35). DhpA could also catalyze hydroxylation of the O-phosphonomethylester of 4 to afford the corresponding O-phosphonomethylester of 124. 4.3.4. Kinase DhpB, Dehydrogenase DhpC, and Aminotransferase DhpD. Because reaction sequences analogous to those of primary metabolism feature prominently in several phosphonate biosynthetic pathways,9 the homology of DhpB, DphC, and DphD to serine biosynthetic enzymes guided initial proposals for the biosynthetic pathway of 11 (Scheme 36).384 Homology of DhpB to glycerate kinase suggested that this enzyme could phosphorylate 124 to afford 125, and the accumulation of the former in a ΔdphB mutant was consistent with this hypothesis. Unfortunately, direct enzymatic characterization of DhpB has been thwarted by its inability to express in E. coli.95 DhpC possesses homology to malate dehydrogenase, which catalyzes an oxidation similar to the proposed oxidation of 125 to 126. Analyzing the supernatant of the ΔdphC deletion mutant by 31P NMR showed accumulation of 124 and another phosphonate intermediate identified as 125 based on two experimental 5740


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Scheme 37. Proposed Catalytic Mechanism of DhpH-N for Generating 129 from 126

provided to His6-DhpH in the presence of Leu-tRNALeu (Scheme 36A).95 However, 130 was produced when DhpD was added to the His6-DhpH reaction, suggesting that 153 is the physiological substrate for DhpH-C as shown in Scheme 35. It is not known how the producing organism protects itself from the alanine racemase inhibitor 153,373 but the timing of the DhpD-catalyzed step between those catalyzed by DphH-N and DphH-C suggests a proximity of the three active sites that may limit free diffusion of this toxic metabolite. Although it has been clearly demonstrated that DhpH and DhpD can together transform 127 to 130, it remains unclear whether 127 exists as a discrete biosynthetic intermediate. Although 127 was detected in a ΔdphH mutant,384 this would require the action of an aminotransferase from outside the dehydrophos cluster to catalyze transamination from 126 (route a in Scheme 35). Nonspecific transaminations are often observed in phosphonate biosynthesis (e.g., 2 to 3 in the 10 and 11 pathways), and the similarity of 126 to 134 of the serine biosynthetic pathway intriguingly implicates phosphoserine aminotransferase (Scheme 36). Alternatively, an additional aminotransferase would not be required if DhpH-N could directly generate 129 from 126 (route b in Scheme 35). Based on observed aminotransferase activity, Bougioukou et al. proposed that DhpH-N could generate a pyridoxamine phosphate (PMP) capable of catalyzing elimination in the presence of an amine donor such as alanine (Scheme 37).95 Although this fascinating issue could be resolved by testing the activity of DhpH toward 126, this has been hampered by the inability to express and purify DhpB.95 4.3.6. Phosphonate O-Methyltransferase DhpI. Although DhpI catalyzes standard SAM-dependent O-methylation, it is unusual in that it acts on a phosphonate substrate.

Indeed, 11 harbors the only known example of a natural phosphonate monomethyl ester. In vitro characterization of DhpI revealed it has broad substrate tolerance, but strongly prefers tripeptide substrates typical of late stage biosynthetic intermediates (Figure 12).386 For example, DhpI preferentially methylated Gly-L-Leu-L-Ala(P) (140) over hydroxylated ethylphosphonates (4, 36, and 124) by 2−3 orders of magnitude (Figure 12A) and exhibited little to no activity toward aminoethylphosphonates 3 and 153 or tripeptide Gly-L-LeuSer(P) (145) (Figure 12B). These three poor substrates suggest that DhpI does not accept substrates with significant positive charge or bulky substituents close to the reacting phosphonate functional group, and this was further supported by methylation of 144, which possesses an amino group that has been neutralized by N-acetylation (Figure 12C). The promiscuity of DhpI holds promise for structural diversification and discovery of phosphonate natural products. DhpI was active toward known phosphonate natural products 9 and 12b, indicating that phosphonate methyl esters with improved pharmacokinetic properties may be readily generated for a variety of natural phosphonates.386 As noted previously, additional advantages may arise from attenuating phosphonate negative charge to more closely resemble carboxylic acid substrates or their enzymatic tetrahedral intermediates. DhpI promiscuity was also exploited to identify two previously unknown phosphonates 142 and 143 (carboxylate methyl esters of 7 and 62, respectively) that led to the discovery of new 13 producer strains as described in section 4.5 (Figure 12C and Scheme 41).142 The biological origins and extent of this unusual enzymatic transformation remains unknown, but intriguing clues lie in the observed activity of DhpI toward phosphoserine (135) and its 5741


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phosphonate biosynthesis based on the absence of a gene encoding PEP mutase, suggesting that these DhpI homologues may catalyze methylation of phosphate groups.386 4.3.7. Desaturase DhpJ Installs the Vinyl Moiety. The formation of 130 by the tandem action of DhpH and DhpD suggested that vinyl installation occurs on a peptidyl substrate at some point after DhpH-C. DhpJ shares significant sequence similarity with known α-ketoglutarate-dependent aspartyl/ asparaginyl β-hydroxylases,393 but was found to have evolved desaturase activity.95 The 1R-diastereomer of 130 (L-Leu-LAla(P)) was initially considered as a likely physiological substrate for DhpJ because it was the only substrate accepted by DhpJ (Scheme 38B) among a series of unreactive substrates (Scheme 38A). However, generated together with the expected product 136 was a dead-end side product proposed to be the αhydroxylation product 146 (Scheme 38B). The inability of DhpJ to convert 146 to 136 implied that the former is not a catalytically competent intermediate and that furthermore 130 is not the physiological substrate. To test for the possibility that DhpI methylates 130 to afford 131 as a substrate for DhpJ (Scheme 35), the latter compound was enzymatically prepared from 52 via the coupled DhpD/DhpH-C reaction and was shown to cleanly generate the desaturated product 132 (Scheme 38C). The observed desaturase activity of DhpJ toward the phosphonate methyl ester 131 clearly implies that DhpI functions before DhpJ in the biosynthetic pathway as shown in Scheme 35. 4.3.8. Peptide Ligase DhpK. In vitro enzyme assays demonstrated that DhpK catalyzes tRNA-dependent peptide bond formation in a manner similar to DhpH-C (Section 4.3.5). The ability of DhpK to ligate Gly to the N-terminus of 130, 132, and 136, but not to 15395 was consistent with DhpK catalyzing the final step in 11 biosynthesis (Scheme 35). Although DhpK and DhpH-C share only 16% sequence identity, both are most closely related structurally to the tRNA-dependent FemX peptidyl transferase from Weissella viridescens.394,395 DhpK and DhpH-C represent an emerging class of tRNA-dependent peptide ligases that expropriate aminoacyl-tRNAs from protein biosynthesis in order to elaborate natural product scaffolds. Additional examples include aminoacyl-tRNA involvement in the biosyntheses of valanimycin,396 pacidamycin,397 and streptothricins,398 as well as tRNAdependent cyclopeptide synthetases.399,400

Figure 12. Compounds tested as substrates for DhpI. (A) Substrates for which steady-state kinetic parameters were obtained. (B) “Poor substrates” were reported to have little to no activity. (C) Although kinetic parameters were not determined, reactivity was clearly observed toward the “Active” substrates.

nonhydrolyzable phosphonate analog 187, also known as LAP4 (Figure 12C).386 Although the phosphate moiety of 135 was methylated less efficiently than that of 187, this cross reactivity implies a possible evolutionary relationship. Just as 11 possesses the only known natural phosphonate methyl ester, Omethylation of phosphates is also very rare.387−390 However, from the limited examples of characterized phosphate methyltransferases, the only one with sequence information shows no homology with DhpI.391,392 Interestingly, proteins with highest homology to DhpI were not associated with Scheme 38. Substrate Tolerance of DhpJa

a

(A) Seven peptides that were not transformed by DhpJ. (B) The 1R-diastereomer of 130, L-Leu-L-Ala(P), transformed to 146 and proposed side product 147. (C) Enzymatically prepared 131 was cleanly converted to 132. 5742


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Scheme 39. Proposed Biosynthesis of FR-900098 (12a)a

a

Box: N-acetylation by FrbF competes with hydroxylation by FrbG.

4.4. FR-900098 (12a), Fosmidomycin (12b), and FR-32863 (12c)

FrbA, a homologue of aconitase, followed by either FrbB or FrbE, both homologues of isocitrate dehydrogenases, are proposed to catalyze these steps. As an analog of αketoglutarate 58, 186 could incorporate a 3-amino group via transamination to form the glutamic acid analog 187 as a biosynthetic intermediate. Interestingly, a parallel TCA cycle enzyme logic is used during phosphinothricin biosynthesis to convert the phosphinate oxaloacetate analog 97 to the glutamic acid analog 101 (Scheme 29). To test this hypothesis, the f rbABCDEFGHIJ-dxrB gene cluster was refactored for expression in E. coli.233 The genes f rbA-J along with the selfresistance gene dxrB were individually expressed from three plasmids, each under the control of a T7 promoter. Production of 12a was successful, and all the more notable because this represented the first Pn natural product to be heterologously produced in E. coli. The inclusion of dxrB was essential for high titers of 12a, whereas f rbI and f rbJ were confirmed to be dispensable. Remarkably, titers of 6.3 mg/L culture of E. coli were achieved by expression of f rbA-H-dxrB. In comparison 22.5 mg/L culture (purified yield) is observed in the producing strain S. rubellomurinus.401 Surprisingly, upon feeding 187 to E. coli expressing f rbH, the cytidine-5′-monophosphate ester 189 was observed by LC-MS, whereas expression of f rbHFG led to the formation of 191, 193, and 12a. 186 was also observed in culture extracts, indicating that an endogenous transaminase reversibly formed 187. These experiments confirmed the intermediacy of 187 as a biosynthetic intermediate but more surprisingly revealed the use of nucleotide conjugates as biosynthetic handles. A similar strategy is also seen in the Pn degradation pathway for CP-lyase, described in section 5.2. FrbH was the most likely source of the nucleotide conjugates. The amino acid sequence for FrbH was novel in being comprised of a pyridoxalphosphate (PLP)-dependent

The antimalarial and antibacterial compound FR-900098 12a is produced by a strain of Streptomyces rubellomurinus,401 while the related compounds FR-31564 12b, also known as fosmidomycin, and the dehydro analog FR-32863 12c are produced by Streptomyces lavendulae.402,403 Interestingly, 12b and 12c are considerably more potent against a wider variety of bacterial strains than 12a.403 The biosynthetic gene cluster encoding 12a in S. rubellomurinus was identified by Metcalf by screening a fosmid library for a gene encoding PEP mutase.162 The gene cluster was verified by heterologous production of 12a in S. lividans transformed with the ppm-containing fosmid. The cluster consists of 11 genes, f rbABCDEFGHIJ-dxrB. Resistance to 12a may be afforded by the gene dxrB, which encodes a homologue of the target enzyme, DXP reductoisomerase. Mutagenesis determined that f rbA-H was the minimal set of genes that encoded the biosynthesis of 12a. PEP mutase was encoded by frbD, while f rbA, f rbB, f rbC, and f rbE appeared to encode homologues of enzymes from the TCA cycle. This was a significant discovery, as it suggested that phosphonopyruvate 1, an analog of the TCA intermediate oxaloacetate, would be irreversibly transformed by a series of reactions that parallel the TCA cycle. This was confirmed by demonstrating that purified FrbC, a homologue of citrate and homocitrate synthases, condensed 1 and acetyl-CoA to form phosphonomethylmalic acid 56 (Scheme 12A). The phosphonomethylmalic acid synthase FrbC is discussed in more detail in section 3.9. The chemical logic of the first half of the TCA cycle would also allow a one-carbon chain extension to form the 3aminopropylphosphonic acid backbone of 12a. On this basis, 56 was predicted to be converted to 186 through a sequence of isomerization and oxidative decarboxylation (Scheme 39). 5743


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Scheme 40. Proposed Mechanism for N-Acetylation by FrbF

with 190 or a Pn ligand was not obtained, a large, solvent exposed and positively charged cleft is observed to funnel toward the thioester of acetyl-CoA. Presumably this is the binding site of the nucleotide substrate 190. FrbF is highly specific for acetyl-CoA (kcat = 1.1 min−1, KM = 20 μM), but low activity is also observed toward propionyl-CoA, n-butyryl-CoA, malonyl-CoA, and acetoacetyl-CoA.404,405 Despite the 15-fold smaller kcat/KM value toward propionyl-CoA (kcat = 0.09 min−1, KM = 24 μM), this promiscuous activity was a sufficient starting point to biosynthesize the N-propionyl analog of 12a in E. coli. 405 This new compound, called FR900098P, is a significantly stronger inhibitor of the target enzyme DXP reductoisomerase (KI = 0.92 nM) than the parent compound 12a (KI = 3.7 nM) and thus points the way to developing more potent antimalarial drugs. The final step in the biosythesis of 12a is hydrolysis of the phosphoanhydride bond of 191 (Scheme 39). This bond is hydrolytically sensitive, thus slow, spontaneous hydrolysis of 191 to form 12a is observed in vitro. However, the addition of FrbI, a homologue of nucleotide hydrolases, was shown to increase the hydrolysis rate substantially above the background rate.233 Surprisingly, FrbI also catalyzed the hydrolysis of 188, 189, and 190. Depending on the relative rate constants, this nonspecific activity would appear to be unhelpful in the context of the overall pathway due to the requirement of the nucleotide handle. It might be possible that the nucleotide transferase domain of FrbH can accept other Pn acceptors to allow the reformation of these key intermediates. However, Pn acceptors other than 187 have not been reported. In the context of heterologous production of 12a in E. coli, FrbH is not essential due to the presence of an nonspecific nucleotide hydrolase in this bacterium. FrbJ is a member of the α-ketoglutarate/nonheme Fe(II) dependent dioxygenase family. This family includes PhnY*, which performs the hydroxylation 3 to form (R)-5 (section 3.5 and Scheme 7). The f rbG gene is not essential for the biosynthesis of 12a in E. coli, implying that this enzyme serves a tailoring role. This was confirmed by showing that purified FrbG hydroxylates C2 of 12a to form 192,233,406 also known as FR-33289, which had been observed previously in cultures of Streptomyces rubellomurinus.402 FrbJ is unreactive toward 191 in vitro but converts 12a to (R)-192 with kinetic parameters of kcat = 147 min−1, KM = 289 μM, and kcat/KM = 8.48 × 103 M−1 s−1.406 This demonstrates that hydroxylation by FrbJ follows hydrolysis of the nucleotide conjugate 191 by FrbI. FrbJ is also active toward the N-formyl analog 12b, with the kinetic parameters kcat = 57 min−1, KM = 700 μM, and kcat/KM = 1.4 × 103 M−1 s−1. The hydroxylation of 12b by FrbJ may explain the biosynthesis of 12c by Streptomyces lavendulae through a sequence of hydroxylation and dehydration. However, identification of biosynthetic gene cluster for 12c would be necessary

decarboxylase domain and a nucleotide transferase domain. Reaction of purified FrbH with 187 produced no product, but inclusion of CTP resulted in the formation of 188 and 189. Thus, the two domains of FrbH were proposed to catalyze a sequence of nucleotide transfer to 187 to form 188 followed by PLP dependent decarboxylation to form 189. This was confirmed by the formation of 188, but not 189, by a FrbH K466A variant, which lacks the conserved lysine residue used by PLP-dependent enzymes to form the Schiff base with the cofactor. The nucleotide transferase domain likewise contains a conserved lysine, K38, that is shared with IspD, an enzyme that catalyzes the formation of a nucleotide intermediate in the nonmevalonate pathway for isoprenoid biosynthesis. The FrbH K38A variant was essentially inactive, confirming that formation of a nucleotide handle is required for the subsequent decarboxylation reaction. In vitro studies also established a sequence of N-acetylation and N-hydroxylation to form 190 and 191. Initially it was thought N-acetylation of 189 occurred first due to the observation of 193 in E. coli cultures expressing f rbHFG. However, FrbG, a FAD and NADPH dependent monooxygenase, was unreactive toward 193 (Scheme 39) and instead was specific for N-hydroxylation of 189 (kcat = 6 min−1, KM = 15 μM, kcat/KM = 6.7 × 103 M−1 s−1). In contrast, the acetylCoA dependent acyltransferase FrbF was active toward 189 (kcat = 3.5 min−1, KM = 35 μM, kcat/KM = 1.7 × 103 M−1 s−1) and 190 (kcat = 2 min−1, KM = 50 μM, kcat/KM = 6.7 × 102 M−1 s−1), but reaction with the former produces a dead-end intermediate 193 that accumulates in cell culture. An interesting biosynthetic competition for 189 thus occurs between FrbG and FrbF, but FrbG ultimately drives the biosynthesis in a productive direction due to its higher kcat/KM value. FrbF is notable for belonging to an uncharacterized family of N-acetyltransferases. An X-ray crystal structure of FrbF revealed that the closest structural homologues were putative aminoglycoside N-acetyltransferases from Bacillus anthracis and Bacillus subtilis.404 A costructure with acetyl-CoA identified H45, T190 and H193 as potential catalytic residues for acyltransfer, which was confirmed by the loss of activity in FrbF H45A (1.5% of wild-type FrbF), T190A (inactive), T190C (inactive), and H193A ( Fe2+ > Zn2+.470 Such metal ion promiscuity is a common feature of members of the alkaline phosphatase superfamily. PhnA from P. f luorescens sp. 23F471 and S. meliloti sp. 1021153,470 have been heterologously produced in E. coli for kinetic and structural analysis. The S. meliloti PhnA had kinetic parameters of kcat = 0.9 s−1, KM = 22 μM, and kcat/KM = 4.1 × 104 s−1 M−1 for the hydrolysis of 7 at 30 °C.153 The P. f luorescens PhnA has virtually identical kinetic parameters with 7, and was additionally shown to have low hydrolytic activity toward ATP and p-nitrophenylphosphate.471 Using [γ32P]ATP as a substrate, it was shown that T64 of P. f luorescens PhnA was phosphorylated during hydrolysis, but not in an inactive T64A variant, or in the presence of the competitive inhibitor phosphonoformate 30 (KI = 50 μM). T64 corresponds to a conserved nucleophile residue in the alkaline phosphatase superfamily (and T68 in S. meliloti PhnA), thus these experiments indicated that PhnA likewise formed a covalent phosphoryl intermediate during the hydrolysis of 7. The X-ray crystal structures of PhnA from P. f luorescens and S. meliloti reveal dinuclear metal ion active sites that have high structural similarity with other members if the alkaline phosphatase superfamily.470,471 A high-resolution structure of the S. meliloti enzyme in complex with the transition state analog orthovanadate (VO42−) enabled a direct comparison with complexes with X. axonopodis nucleotide phosphodiesterase (PDB ID: 2GSO) and E. coli alkaline phosphatase (PDB ID: 1B8J). The orthovanadate is bound in a distorted pentacoordinate trigonal bipyramidal configuration (Figure 13). Strict identity is observed in the His and Asp residues

stabilized in the transition state. In enzymes that catalyze hydrolysis of P−O bonds the leaving group oxygen is apical relative to the nucleophile in the transition state and negative charge development on the departing atom is stabilized directly by metal ion M1. In the case of PhnA, a much poorer carbanion leaving group (pKaLG ≈ 30)472 is stabilized as the acetate enolate where negative charge develops further away on the carboxylate oxygens (Scheme 53). A complex of 7 with the inactive S. meliloti PhnA T68A variant suggested that one carboxyl oxygen of the enolate would be stabilize by M1.470 Intriguingly, the distance between M1 and M2 in S. meliloti or P. f luorescens PhnA is on average significantly greater (4.5 to 4.6 Å) than that observed in other enzymes of the alkaline phosphatase superfamily (4.3 Å),470,471 suggesting that the active site has been tuned to accommodate the development of negative charge further away from where bond cleavage occurs. Additional stabilization of the second acetate enolate oxygen may derive from a hydrogen bond to a water molecule held in position by the amide carbonyl of I287 in the S. meliloti PhnA.470 In contrast, K126 and K128 in P. f luorescens PhnA are proposed to stabilize the enolate oxygens electrostatically.471 These Lys residues are conserved in the P. f luorescens and S. meliloti enzymes and their substitution creates inactive PhnA variants, implying a critical catalytic or structural role.470,471 Subsequent hydrolysis of the phosphoryl enzyme intermediate to release Pi is proposed to proceed through a similar transition state. In this case a nucleophilic water, bound and activated by M1, performs in-line attack on the Thr-phosphate ester intermediate where the departing Thr alkoxide is stabilized by M2. 5.2. Radical C−P Bond Cleavage by Carbon−Phosphorus Lyase

Carbon−phosphorus (CP) lyase is distinguished by its ability to cleave unactivated C−P bonds through a radical mechanism (Scheme 54). For example, the reaction of methylphosphonic acid 8 with CP-lyase affords methane and Pi. Equally impressive is the activity of CP-lyase toward a wide array of structurally diverse Pn, which may account for its wide distribution in bacterial species. The activity of CP-lyase was first detected in E. coli in 1963,473 but it is only recently that the mechanism of CP bond cleavage has come to light.32 Over the intervening years many laboratories failed to observe CP-lyase activity in cell free extracts, thus preventing purification of active enzyme. This technical hurdle stemmed from several factors, including the complexity of the CP-lyase pathway, the necessity of maintaining intricate protein−protein interactions for catalysis, and the oxidative sensitivity of a [4Fe-4S] cluster in the CPbond cleaving enzyme PhnJ. 5.2.1. Substrate Specificity and Reaction Products. Unlike every other known pathway for catabolizing Pn, CPlyase is astonishing for its lack of substrate specificity. Figure 14 is a summary of the range of Pn that can sustain various bacterial species, such as E. coli, Agrobacterium radiobacter, Kluyvera sp., Klebsiella sp., Pseudomonas sp., and Streptomyces sp., as a sole source of Pi through the activity of CPlyase.425,473−478 Viable Pn substrates include simple, branched, and substituted alkylphosphonic acids. As discussed earlier, the catabolism of 8 by marine bacteria is a major source of oceanic methane.62 CP-lyase is also active with 3 and 7, the substrates that are processed by the phosphonatase PhnX and phonoacetate hydrolase PhnA pathways, revealing that Nature has evolved multiple solutions for catabolizing the same

Figure 13. Active site scheme of PhnY bound to orthovanadate. Key bond lengths given in angstroms.

used to bind metal ions M1 and M2. Likewise, the nucleophile T68 is observed bound to M2, which would favor ionization to form a better nucleophile. This structure is consistent with an associative transition state for phosphoryl transfer during C−P bond cleavage of 7. In this mechanism the hydroxyl of T68, likely as the alkoxide, attacks the phosphorus center of 7 to form a pentavalent, trigonal bipyramidal transition state, where the nucleophile and carbon leaving group are apical, and the three phosphoryl oxygens are equatorial. One of the equatorial oxygens is stabilized by forming a bridging interaction between the two metal ions, a feature that is conserved in the alkaline phosphatase superfamily. However, variation is observed in stabilizing the development of negative charge on the other two equatorial phosphoryl oxygens: in PhnA a pair of water molecules and the amide N−H bond of the nucleophile T68 serves this purpose, whereas in X. axonopodis nucleotide phosphodiesterase and E. coli alkaline phosphatase this is performed by Asn and Arg side chains, respectively. PhnA also deviates from other members of the alkaline phosphatase superfamily in how the leaving group atom is 5754


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Scheme 53. Proposed Mechanism for Phosphonoacetate Hydrolase PhnAa

a

Residue numbering for S. meliloti PhnA shown, with P. f luorescens PhnA numbering in parentheses.

synthetic L-alanine derivative, alaphospin 117,372 but has more recently been observed as an intermediate in the biosynthesis of dehydrophos 11.95 Alaphosphin has antibacterial properties due to the release of 153 in the cell, which as a mimic of D-Ala inhibits peptidoglycan biosynthesis. It is therefore notable that the action of CP-lyase on 153 not only deactivates the compound but also allows its use as a sole source of Pi by E. coli.425 This is not the only example of CP-lyase appearing to afford a resistance and catabolic role, as shown by the use of fosfomycin 9 as a Pi source by A. radiobacter.475 The producers of 9, Streptomyces wedmorensis and Streptomyces fradiae, can also

Scheme 54. General Reaction Scheme for CP-Lyase

compounds. In addition to 3, CP-lyase can cleave the CP-bonds of a wide range of aminoalkylphosphonic acids, including glyphosate 23 and (S)-1-aminoethylphosphonic acid 153. The catabolism of 23 by CP-lyase is one of two routes for degrading this herbicide by soil bacteria (reviewed in ref 14). 153 was originally known as the hydrolytic cleavage product of the

Figure 14. Examples of substrates for CP-lyases from various bacterial species. 5755


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slowly grow on 9 as a Pi source.477 In addition to cleaving C−P bonds with sp3 carbons, CP-lyase can also cleave C−P bonds with sp2 and sp carbons, such as 157, 158, and 160. This is notable considering the radical mechanism of catalysis by CPlyase, as these C−P bonds become progressively stronger as measured by bond dissociation energies (see Figure 3). Alkylphosphinic acids, which require the cleavage of two C− P bonds, are also Pi sources for A. radiobacter.475 Finally, CPlyase of E. coli can cleave the H−P bond of 34 to produce Pi, revealing a relative indifference to the presence of an alkyl group for productive binding energy during catalysis.478 Equally notable are Pn that are not substrates for CP-lyase. This can be strain specific. For example, A. radiobacter will vigorously grow on 23 and 161 as Pi sources, whereas Kluyvera, Klebsiella, and Pseudomonas will not grow at all on these compounds.475 This may reflect differences in the ability of the strains to transport the Pn into the cell, or differences in Pn specificity of the corresponding CP-lyases. The latter is observed in E. coli at the stereochemical level where (S)-153 can serve as a Pi source, but not the corresponding (R)enantiomer.425 In this case the difference can be traced to the stereochemical preference of the N-acetyltransferase PhnO in the CP-lyase pathway. Substrate specificity may also explain why 165, 166, and 167 are not substrates for E. coli CP-lyase, despite the fact that the resulting alkyl radicals following C−P bond cleavage would benefit from stabilization by resonance or hyperconjugation.474 Finally, alkylphosphonate esters such as 148 are not a substrates for CP-lyase due to the requirement of a free phosphonic acid for processing, as discussed in detail below. Early feeding experiments with E. coli and other bacteria quickly pointed to a radical mechanism for C−P bond cleavage by CP-lyase. Frost observed that catabolism of alkylphosphonic acids by E. coli afforded low yields of alkene products (0.5 to 1%) in addition to the major alkane product (Scheme 55A).474 Likewise, conversion of cyclopropylmethylphosphonic acid 152 by E. coli, Klebsiella oxytoca, and Kluyvera ascorbata affords cyclopropylmethane as the major product as well as 0.3−2% of the rearrangement product 1-butene (Scheme 55B).474,479 It was also shown that K. oxytoca would convert cis-1,2-dideuterio219 to primarily to the cis-alkene along with 1% of the trans isomer (Scheme 55C).479 Such minor products cannot be readily explained as arising from heterolytic or oxidative CPbond cleavage, but instead are consistent with homolytic C−P bond cleavage and formation of a carbon-centered radical. This proposal was strengthened by the observation that similar mixtures of alkanes and alkenes can be produced through the radical based dephosphorylation of alkylphosphonic acids with lead acetate.474 Quenching of the carbon radical by CP-lyase appears to be highly efficient. Because cyclopropylcarbinyl and propenyl radicals rearrange or isomerize, respectively, at known rates of 108 to 109 s−1, the yields of 1-butene and trans-alkene indicate that the carbon radical formed upon C−P bond cleavage receives a hydrogen at a rate of 109 to 1011 s−1.474,479 In the case of A. radiobacter, the CP-lyase appears to be 100% efficient in capturing the carbon radical, as rearranged or isomerized products are not observed using 152 or 219 as substrates. 479 More conclusive evidence for a radical intermediate came from the observation of substantial racemization of stereochemistry at C1 following conversion of (R) or (S)-[1-2H,1-3H]-147 by E. coli CP-lyase (Scheme 55D). Although 67% retained stereochemistry was observed at C1 of the ethane product (keeping in mind Cahn-Ingold-Prelog

Scheme 55. Use of Substrate Analogs to Probe the Formation of Radical Intermediates by CP-Lyase

rules of priority), 33% inversion was observed. Significant inversion of configuration would occur if a neutral ethyl radical with sp2-like character at C1 was formed and proceed to rapidly rotate about the C1−C2 bond prior to receiving a specific hydrogen from the enzyme active site. In contrast, this result cannot be easily explained by carbocation or carbanion intermediates, which would be expected to be electrostatically restrained in the active site.480 5.2.2. Structural Diversity of the phn Operon. The name notwithstanding, CP-lyase is actually comprised of several enzyme activities that form a catabolic pathway. E. coli has a number of phosphate starvation inducible (psi) genomic loci that are under control of the phosphate (pho) regulon.481 Wanner and Walsh identifed the psiD locus as encoding catabolism of phosphonates and phosphite by CP-lyase,482 and subsequentely renamed this locus as phn.483 Sequencing and mutagenesis of this locus revealed a 14-gene cistron phnCDEFGHIJKLMNOP encoding the CP-lyase pathway.483−488 It was subsequently shown that phnCDEFGHIJKLMNOP was required by E. coli to grow on minimal media with Pn or phosphite 34 as the only Pi source. However, expression phnGHIJKLM was sufficient to encode C−P bond cleavage.489 These pioneering studies thus established that phnCDE encoded a Pn transport system, phnF a likely regulatory protein, phnGHIJKLM the CP-lyase activity, and phnNOP accessory catabolic enzymes. A comparison of phn genes from various bacterial species indicates that this operon has undergone extensive structural changes.14,15 Such changes are the evolutionary outcome of lateral transfer of phn genes between bacterial species either as plasmids or by transposition.15 Unlike the tight organization of the phn operon in E. coli, phn genes are often more loosely 5756


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arranged in other species. In all cases phnGHIJKLM is strictly conserved, but these genes often appear in different orders, are transcribed as separate cistrons, or are present in multiple copies. For example Mesorhizobium loti has phnCDEFGHIJKLMN distributed across the chromosome and a plasmid. Several of these genes present in multiple copies while the genes phnO and phnP are missing. Pseudomonas stutzeri WM88 has two CP-lyase pathways encoded on separate phn and htx cistrons, with phnO missing in both cases.14,490,491 The marine cyanobacterium Trichodesmium IMS101 has a phn operon comprised of phnCDEEGHIJKLM.492 Fragments of the phn operon have also been co-opted by bacteria for alternative or unknown functions. For example, a locus containing the genes phnCDEF is found in Mycobacterium smegmatis even though this bacterium cannot use Pn as a Pi source. In this case the phnCDEF genes appear to have been adopted to encode the transport of Pi, rather than Pn, into the cell when Pi is limiting.493,494 Likewise phnO is found in Salmonella enterica adjacent to genes encoding PhnX and PhnA pathways for the catabolism of 3 and 7, respectively.495 As discussed below, phnO encodes an N-acetyl transferase, a chemistry that does not lend an obvious purpose to the PhnX and PhnA pathways. Equally intriguing is the supplementation of phn operons with genes encoding novel or seemingly redundant chemistries. For example, Enterobacter aerogenes IFO 12010 has operons encoding CP-lyase and PhnX pathways for the conversion of 3 to Pi.128 Pseudomonas stutzeri WM88 has a gene within the htx operon, htxA, encoding a nonheme Fe2+/α-ketoglutarate dependent dioxygenase that performs the oxidation of hypophosphite 168 to phosphite 34 (Scheme 56).490,496

for example, a gene encoding a metal ion dependent hydrolase from the polymerase and histidinol phosphatase family (cog0613) is observed in place of phnP in Clostridium dif f icile, Egerthella lenta, and Oceanobacillus iheyensis.515 The enzyme encoded by this gene, Elen0235, was shown to perform the same role as PhnP, hydrolysis of the cyclic-phosphodiester product formed by the C−P bond cleaving enzyme PhnJ, but with different regioselectivity and greater promiscuity.515 Similarly, a gene of unknown function, duf1045, appears in association with phnN in several species, including, M. loti, Agrobacterium tumefaciens, Burkholderia pseudomallei, and Ralstonia eutropha,15 but the function of the encoded enzyme has yet to be determined. 5.2.3. Regulation of the phn Operon. As noted above the phn operon is a member of the phosphate (Pho) regulon. Control of the Pho regulon in bacteria has been reviewed by Wanner26,516 and others17,18 and will only be briefly described here. In E. coli the Pho regulon consists of at least 30 genes arranged in 9 transcriptional units that are expressed under conditions of Pi starvation.26 Expression of the Pho regulon is controlled by a two-component system consisting of PhoR, a transmembrane sensory histidine kinase, and PhoB, the transcriptional response regulator. PhoB and PhoR work in conjunction with the phosphate specific transport system, PtsSCAP through the mediation of PhoU, a chaperone-like inhibitory protein of PhoR. Under Pi-rich conditions ([Pi] > 4 μM), PtsSCAP is in a phosphorylated state that inhibits PhoR/ PhoB via PhoU. When Pi levels fall ([Pi] < 4 μM), PtsSCAP is dephosphorylated, leading to release of PhoU from PhoR. PhoR is now active and phosphorylates PhoB, causing a conformational change517 that allows PhoB to bind to Pho regulon promoters. A deactivated state also exists for PhoR/ PhoB when Pi levels increase, which leads to reseting the PstSCAB/PhoU/PhoR/PhoB to its default inhibited state. Mutations within pstSCAP-phoU lead to constitutive expression of the Pho regulon. This has proven very useful for studying the CP-lyase pathway in vivo as the E. coli pst mutant will catabolize Pn even in Pi-rich media.425,478,518 In marine environments bacteria are in a chronic state of Pi starvation18 and accordingly operons encoding the utilization of Pn and phosphite 34 are actively expressed.492,519,520 The phn operon is frequently observed to contain phnF encoding a transcriptional regulator. The precise role of phnF is not known in the context of a complete phn operon encoding CP-lyase. PhnF encoded by a partial phnFDCE operon in Mycobacterium smegmatis has been shown to repress the transcription of phnDCE when Pi levels are high.494 Therefore, PhnF may provide a second, more specific, level of control over the expression of a phn operon in addition to PhoR/PhoB. M. smegmatis PhnF has been shown in vitro to bind at sites within the phnD-phnF intergenic region.521 However, because M. smegmatis appears to use PhnDCE to import Pi rather than Pn, it is not clear what ligand binds to PhnF to cause release of the protein from its DNA binding sites. The X-ray crystal structure of PhnF consists of a homodimer with a fold that is conserved with GntR/HutC-family transcriptional regulators.521 A sulfate ion was observed in a pocket within the C-terminal domain of PhnF, which may represent the binding site of the natural phosphorylated molecule that triggers derepression of phnDCE. 5.2.4. CP-Lyase Pathway. The overall pathway encoded by phnCDEFGHIJKLMNOP is shown in Scheme 57 using 3 as an example. The key feature of this catabolic pathway is incorporation of the Pn substrate into D-ribose (170), as well

Scheme 56. Oxidation of Hypophosphite (168) and Phosphite (34) by the Oxidative Enzymes HtxD, PtxD, and E. coli Alkaline Phosphatase

Although 34 is known to be a substrate for CP-lyase, P. stutzeri has an additional ptx operon that encodes the import and oxidation of 34 to Pi.490,491 The oxidation reaction is performed by an NAD+ dependent dehydrogenase encoded by ptxD (Scheme 56).497,498 PtxD has been extensively characterized499−504 and is the subject of a review.505 The interest in PtxD is driven by its biotechnological potential for NADH regeneration506−509 and as a selection marker in transgenic plants and microorganisms using 34 as a Pi source.510−512 Desulfotignum phosphitoxidans also has ptxD and phn gene clusters, but in this case the former is used not to exploit 34 as a Pi source, but as a source of electrons for metabolic energy.513 E. coli also has catalytic redundancy toward 34, but in this case relies on the promiscuity of a well-known enzyme; in addition to CP-lyase, 34 is also a substrate for alkaline phosphatase, which has been shown to perform the hydrolysis of 34 to Pi and H2.514 Individual phn genes are also subject to substitution; 5757


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Scheme 57. Overview of the CP Lyase Pathway

As PhnD can be considered a “gatekeeper” protein, its specificity toward Pn can dictate the spectrum of substrates that are catabolized by the rest of the CP-lyase pathway. The unique sequence of PhnD has proven to be a useful marker to detect the potential for Pn catabolism by bacteria,492,519,520 although the expression of phnD loci does not appear to be responsive to Pi limitation in some strains of marine bacteria.522 The affinity of E. coli PhnD for various ligands has been assayed in vitro based on isothermal titration calorimetry or changes in tryptophan fluorescence, with dissociation constants measured for 3 (KD = 20 nM), ethylphosphonic acid 147 (KD = 200 nM), methylphosphonic acid 8 (KD = 1300 nM), glyphosate 23 (KD = 650 μM), Pi (KD = 50 μM) as well as several other Pn.523,524 The high affinity for 3 suggests that exposure to this Pn has been a driving force for evolving substrate specificity. However,

as N-acetylation in the case of aminoalkylphosphonic acids, prior to C−P bond cleavage. In most CP-lyase pathways the phosphonate phosphorus ends up not as free Pi in the cell, but as the anomeric phosphate of 5′-phospho-α-D-ribosyl-bisphosphate (PRPP, 174). This is a general glycosyl donor used in the biosynthesis of nucleotides, histidine, and tryptophan. The broad specificity of CP-lyase may partly be explained by incorporating the Pn into a common ribosyl handle, which can provide a fixed amount of transition state stabilization energy for CP-bond cleavage, irrespective of the structure of the Pn substrate. 5.2.4.1. PhnCDE. PhnCDE comprise an ABC-type Pn transporter with PhnD as the periplasmic binding protein, PhnE the transmembrane domain, and PhnC the ATPase component that drives the influx of 3 across the cell membrane. 5758


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of substrates acted as competent amino acceptors, including 92, (S)-153, 3, 155 (Figure 14) and taurine 50 (Figure 4). Surprisingly, the highest specific activity was observed with (S)153 (kcat = 13 s−1, KM = 0.17 mM, kcat/KM = 7.8 × 104 M−1 s−1). This was followed by 3 and 92, which had essentially identical kinetic parameters (kcat = 9 s−1, KM = 1.8 mM, kcat/KM = 5 × 103 M−1 s−1 for 3). Based on kcat/KM PhnO is selective for (S)-153 over 3 by a factor of 15. This preference is even stereoselective as (R)-153 not a substrate for S. enterica PhnO, and a very poor one for E. coli PhnO. The efficiency of PhnO with 1-aminoalkylphosphonates 92 and (S)-153 was a puzzling discovery as this suggested an evolutionary pressure that arose by exposure of the host bacterium to these or similar compounds. (S)-153 is an analog of D-Ala and thus is bacteriocidal due to interference with peptidoglycan biosynthesis. Therefore, it seemed possible that PhnO served a detoxification role rather than a catabolic one; this would explain why phnO in S. enterica appears not with a phn operon, but in association with genes encoding PhnX and PhnA pathways.495 This hypothesis was confirmed by experiments with E. coli PhnO.425 E. coli does not require phnO to grow on 3 or other aminoalkylphosphonates as a Pi source, with the exception of 92 and (S)-153. E. coli phnO mutants could could be rescued from the bactericidal action of (S)-153 by the addition of D-Ala and Pi to the culture medium. However, the requirement of phnO to catabolize 92, which is not toxic to E. coli, suggests N-acetylation is also required by the CP-lyase pathway to process 1-aminoalkylphosphonates. This may include the homolytic C−P bond cleaving step. As the 1amino group of 92 and (S)-153 will likely have a positive charge at pH 7, a destabilizing inductive effect, estimated to be 4−5 kcal mol−1, will be experienced by the resulting carboncentered radical.525,526 However, N-acetylation would neutralize the 1-amino group, as well as provide resonance stabilization of the resulting radical (Scheme 58). In contrast, the 2-amino

ligand binding specificities are observed to vary considerably in PhnD homologues. For example, Prochlorococcus MIT9301 has two copies of PhnD encoded on separate ptxABCD-phnY*Z and phnCDE operons.522 The ptx locus encodes genes for importing and utilizing phosphite 34 as a Pi source with ptxB encoding one copy of PhnD, while the phnCDE locus encodes a second copy as part of a Pn transporter. Using ITC it was shown that the two PhnD homologues did not bind to 3; instead, PtxB was specific for 34 (KD = 120 nM) while PhnD was specific for methylphosphonic acid 8 (KD = 800 nM). The high affinity of Prochlorococcus PtxB for 34 is consistent with the ability of this organism to use 34 as a Pi source.522 In general, the low KD values that are observed for PhnD variants with their best ligands correspond well with with the 70−140 nM concentrations of dissolved organic phosphorus typically found in the ocean, of which 25% is composed of Pn.18,492 The X-ray crystal structures of E. coli PhnD in apo form and bound to 3 have also been solved, revealing the origins of ligand specificity.524 Like other periplasmic binding proteins, the PhnD has a bilobal structure with the ligand binding cleft formed between the two lobes. These lobes rotate approximately 70° upon binding 3. The negatively charged phosphonate moiety is tightly bound at the end of the cleft by a polar pocket comprised of Y47, S29, H157, S127, Y93, T128, and a water molecule (Figure 15). The 2-amino group of

Figure 15. Active site scheme for E. coli PhnD bound to 3.

Scheme 58. Proposed Role for N-Acetylation by PhnO for Enabling CP-Bond Cleavage of 1-Aminoalkylphosphonates by CP-Lyase

3, which is likely to be positively charged at pH 7, interacts with the two negatively charged side chains of D205 and E177. Five water molecules extend away from the 2-amino group along a channel to the surface of the protein, suggesting that longer Pn can be accommodated by the binding pocket. Interestingly, D205 and E177 are not conserved in PhnD homologues, suggesting different Pn specificity. Likewise, Agrobacterium radiobacter PhnD has proline residues in place of H157 and Y47, suggesting the creation of a larger, less polar pocket for the phosphonate moiety. A. radiobacter is the only bacterium known to catabolize phosphinic acids (161 through 164, Figure 14),475 thus this substitution may partly explain how the additional steric bulk of this type of substrate is accommodated. Following Nature’s lead, the structure of PhnD has served as a starting point for the development of fluorescent biosensors for the detection of Pn.523,524 5.2.4.2. PhnO. PhnO performs N-acetylation of aminoalkylphosphonic acids using acetyl-CoA as the acetyl donor. This step has been shown to occur before the Pn is incorporated into D-ribose.425 The enzyme from E. coli425 and Samonella enterica495 has been characterized in vitro. The S. enterica PhnO requires a metal ion for activity, with Mn2+, Ni2+, and Co2+ affording the highest specific activities. S. enterica PhnO was shown to accept acetyl-, propionyl-, and malonylCoA as acyl donors, with acetyl and propionyl-CoA having essentially identical kinetic parameters (kcat = 13 s−1, KM = 60 μM, kcat/KM = 2.2 × 105 M−1 s−1). A surprisingly wide variety

group of 3 would have a weaker inductive effect and thus would not require N-acetylation for C−P bond cleavage. The specialized role of PhnO in dealing with 1-aminoalkylphosphonic acids as well as the frequency of occurrence of phnO in microbial genomes suggests that these compounds may be more prominent in the environment than is currently appreciated. Currently (S)-153 is known to occur as an intermediate in the biosynthesis of dehydrophos 11,95 while 92 appears as a microbial catabolite of glyphosate 23,14 and as a component of argolaphos 15.102 5.2.4.3. PhnI. PhnI catalyzes the ribosylation Pn, such as 169, to form an α-D-ribosyl phosphonate, 170 (Scheme 57). Versions of 170 have been observed in E. coli cultures growing 5759


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Scheme 59. Phosphonate Glycosyltransferase PhnIa

a

(A) Role of PhnGHL in dictating the glycosyl acceptor specificity of PhnI. (B) A comparison of the hypothetical PhnI transition state and the transition state analogue inhibitor 181.

on Pn as a Pi source,425,478,527 but until recently the origin and purpose of α-D-ribosyl phosphonates in the CP-lyase pathway was unknown. This changed when Raushel recognized that PhnI has weak sequence homology to nucleoside hydrolases.528 As described in section 5.2.5, PhnI is also a component of the PhnG2H2I2J2K complex.529 For this reason PhnI is insoluble when synthesized alone in E. coli. However, soluble PhnI can be obtained when fused to glutathione-S-transferase (GST). Raushel demonstrated that PhnI catalyzes the hydrolysis of ATP (kcat/KM = 1.3 × 104 M−1 s−1) and GTP (kcat/KM = 8.3 × 104 M−1 s−1) to form D-ribose-5-triphosphate 179 and the corresponding base, even if Pi or a Pn is present in the reaction mixture (Scheme 59A).528 With remarkable insight, Raushel then demonstrated that formation of a PhnGHIL complex can change the nucleophile selectivity of PhnI from water to Pn. Specifically, when PhnG, H, L, and I are combined in a reaction mixture, followed by a protease to cleave away the GST tags, a PhnGHIL complex can be formed in situ which then catalyzes the ribosylation of methylphosphonic acid 8 to form 180. Essentially identical kinetic parameters were observed with ATP or GTP as glycosyl donors and 8 as acceptor, with kcat/KM = 3.5 × 105 M−1 s−1 measured with ATP. The glycosyltransfer reaction occurs with inversion of stereochemistry with water or Pn as the nucleophile. Highlighting the glycosyltransfer mechanism of PhnI is the potent inhibition (KI = 20 nM) of the enzyme by the oxacarbenium ion transition state analog 181 (Scheme 59B).530 5.2.4.4. PhnM. PhnM catalyzes the hydrolysis of the α−β phosphoryl bond in 170 to form the 5-phospho-α-D-ribosyl alkylphosphonate 171. PhnM belongs to the amidohydrolase superfamily, which includes hydrolases of phosphate esters. The enzyme can be produced in soluble form without a solubility tag. Raushel demonstrated that PhnM catalyzed the hydrolysis of the methylphosphonate analog 180 with a kcat/KM value of 1.1 × 105 M−1 s−1.528 D-Ribose-5-diphosphate and D-ribose-5triphosphate were also substrates but hydrolyzed at slower rates. Running the PhnM reaction in 18O-labeled water indicated that hydrolytic attack occurs at the α-phosphate of 171. The addition of other phn-encoded enzymes did not change the kinetic parameters of PhnM, thus this enzyme does not appear to be influenced by the CP-lyase complex like PhnI.

5.2.4.5. PhnJ. PhnJ catalyzes the intriguing C−P bond cleaving step, converting 171 to the α-D-ribosyl 1,2-cyclic phosphate 172. PhnJ is also a component of the PhnG2H2I2J2K complex described below in Section 5.2.5.529 The substrate and product for PhnJ were first identified by 31 P NMR spectroscopic analysis of cultures of E. coli mutants. E. coli ΔphnP is observed to accumulate 171 and 172 in the culture medium, while E. coli ΔphnJ accumulates 171.425,478,518 Raushel recognized that a sequence of four Cys residues in PhnJ, CX2CX21CX5C, had a resemblance to the CX3CX2C motif that is used by radical SAM enzymes to form a [4Fe-4S] cluster, and thus was a prime candidate for cleaving a C−P bond in the homolytic fashion.528 PhnJ was produced in soluble form as a GST fusion and copurified with 2.2 eq of iron per monomer. Incubation of PhnJ under anaerobic conditions in the presence of excess FeSO4 and Na2S produced what appeared to be an intact [4Fe-4S]2+ cluster with an absorbance maximum at 400 nm. Subsequent addition of dithionite caused this absorbance to disappear, consistent with reduction of the cluster to the [4Fe-4S]+ state and the form necessary to initiate radical chemistry. Finally, in an experiment that had evaded researchers for decades, incubation of GST-PhnJ in the [4Fe-4S]+ state with the α-D-ribosyl-1-methylphosphonate substrate 182, along with SAM, dithionite, and protease (to remove the GST tag in situ) produced 172 and methane. SAM is converted to 5′deoxyadenosine 71 and L-methionine. PhnJ, once released from the GST tag, slowly precipitated and was estimated to perform only four turnovers of substrate. This instability reflects the normal context of PhnI as part of the PhnG2H2I2J2K complex. Detailed labeling and mutagenesis studies by Raushel identified the key catalytic residues involved in C−P bond cleavage.531 Substitution of any of the four Cys residues for Ala produced inactive PhnJ. C241, C244, and C266 were shown to be essential to form the [4Fe-4S] cluster. However, C272 appeared to have a different catalytic role as PhnJ C272A could still form the cluster according to EPR and absorbance spectroscopy, but would not perform C−P bond cleavage. As discussed previously, radical SAM enzymes initiate radical chemistry by generating a 5′-deoxyadenosyl radical 68 by donation of an electron from the [4Fe-4S]1+ cluster to SAM (Scheme 16A). By running the PhnJ reaction in D2O it was 5760


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Scheme 60. Proposed Mechanism for CP Bond Cleavage by PhnJ of the CP-Lyase Pathway

studies described in section 5.2.1. A key aspect of this mechanism is formation of a covalent phosphothioester intermediate IV during catalysis. Turnover of this intermediate is achieved by anchimeric assistance of the 2-hydroxyl of ribose, forming the α-D-ribosyl-1,2-cyclic phosphate 172. An attempt was made to trap the covalent intermediate by reacting PhnJ with the 2-deoxy analog 183. Methane was produced from 183, but attempts to detect the resulting covalent intermediate by mass spectrometry were unsuccessful. However, 31P NMR spectroscopic analysis of tryptic digest of PhnJ treated with 183 revealed a downfield resonance consistent with formation of phosphothioester. In several respects the C−P bond cleaving mechanism proposed for PhnJ resembles the C−C bond cleaving mechanism of pyruvate formate lyase, including the use of a glycyl radical to generate a thiyl radical, homolytic bond cleavage performed by the thiyl radical, and formation of a covalent thioester intermediate.532 However, the stereospecificity of hydrogen abstraction by 68 is opposite in the case of PhnJ, and the abstraction of both hydrogens of Gly32 is without precedent.531 5.2.4.6. PhnP and PhnPP. The cyclic α-D-ribosyl phosphate 172 that is produced by the PhnJ reaction is a dead-end catabolic intermediate. PhnP, a member of the metallo-βlactamase superfamily, performs the regiospecific hydrolysis of the cyclic phosphate of 172 to afford α-D-ribosyl-1,5diphosphate 173, a metabolite that can be used by the cell. (Scheme 57). Although PhnP is not directly involved in C−P bond cleavage, mutation of phnP results in the accumulation of 172 and 171 in E. coli cultures and thus inadvertently led to the initial identification of these key intermediates in the CP-lyase pathway.425,478,518 PhnP can be overproduced in soluble form in E. coli and copurifies with Mn2+ and Zn2+.533 A metal ion

determined that 68 was not converted to 72 by abstraction of solvent exchangeable hydrogen in the enzyme active site. However, by replacing every Gly residue in PhnG with [2,2-2H2]-Gly (designated PhnJ-Gly-2H2) 68 was converted to 72 containing a single deuterium atom. This indicated that a glycyl radical was formed during catalysis, and that one of the Gly hydrogens was abstracted by 68. Labeling studies also led to the identification of the hydrogen that is abstracted by the methyl radical upon C−P bond cleavage of the methylphosphonate substrate 182. When PhnJ turns over multiple times with 182 in D2O, CH3D is formed, but under single turnover conditions CH4 is formed. If PhnJ-Gly-2H2 is allowed to turn over a single time with 182, CH3D is formed. Thus, the source of the hydrogen in methane was assigned to one of the Gly residues. Single turnover reactions with PhnJ labeled with either (R)- or (S)-[2-2H]-Gly demonstrated that the pro-R hydrogen of the Gly residue was used to form 72, while the pro-S hydrogen was used to form methane during the first reaction cycle. Analysis of deuterium incorporation into PhnJ peptides by ESI-MS/MS along with site directed mutagenesis led to the identification Gly32 as the catalytic residue. The working model for the mechanism of PhnJ based on these experiments is summarized in Scheme 60.32 The 5′adenosyl radical 68 abstracts the pro-R hydrogen of Gly32, forming an active site glycyl radical (I), which subsequently abstracts the thiol hydrogen of Cys272 to form a thiyl radical (II). This sets up the C−P bond cleaving step where the thiyl radical is proposed to displace a methyl radical from the phosphorus center of the α-D-ribosyl methylphosphonate 182 (II to IV). Subsequently the methyl radical abstracts the pro-S hydrogen of Gly32 to form methane; this step is very fast, proceeding at a rate of 109 to 1011 s−1 based on the radical clock 5761


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the first evidence that CP-lyase comprised a pathway and also foreshadowed the conversion of 171 to 174 based on Frost’s earlier isolation of the ethylphosphonic acid analog 171 from E. coli.527 5.2.4.8. PhnGHLK. The functions of these four proteins remain unknown. However, PhnGHK appear as part of a greater CP-lyase complex described below. PhnG and PhnH have unique amino acid sequences, whereas PhnL and PhnK have homology to the nucleotide binding domains of ATPbinding casette transporters. Every chemical step shown in Scheme 57 for the transformation of Pn into Pi has been reconstituted in vitro. Only in the step involving PhnI do three of these proteins, PhnGHL, directly influence chemistry by changing the glycosyl acceptor specificity of PhnI from water to Pn. How this is achieved is unknown. 5.2.5. PhnG2H2I2J2K Complex. Throughout phn operons phnGHIJKLM is strictly conserved. In part this is explained by the key chemical steps performed by PhnM, PhnI, and PhnJ in the CP-lyase pathway. However, a subset of this conserved set of genes also encodes a multiprotein complex. Hove-Jensen was the first to isolate a PhnGHIJK complex from E. coli that had overexpressed phnGHIJKLM from a plasmid. 529 The PhnGHIJK complex survived a sequence of cell lysis, centrifugation, ion-exchange chromatography, ammonium sulfate precipitation, and size exclusion chromatography to afford pure proteins as judged by SDS-PAGE. The PhnGHIJK complex was also purified by affinity chromatography by encoding a His6 tag at the C-terminus of PhnK. A molecular mass of 640 kDa was estimated by size exclusion chromatography, leading to proposed stoichiometry of PhnG4H2I2J2K. Smaller complexes of PhnGHIJ and PhnGI could also be produced. Isolation of the former suggested that PhnK was loosely associated and not essential for structural stability of the overall complex. PhnH was shown to be protected from crosslinking with glutaraldehyde as part of the PhnGHIJK complex but not in isolation. This suggested that reactive amino groups of PhnH were protected by tight association with the complex. Later, Raushel and Russell proposed a model for assembly of the PhnGHIJK complex based on detailed cross-linking, ultracentrifugation, and hydrogen-deuterium exchange studies. In this model a PhnG2I2 core receives two copies of PhnH and PhnJ to form PhnG2H2I2J2, followed by a copy of PhnK to produce a final complex of PhnG2H2I2J2K.536 The X-ray crystal structure of the PhnG2H2I2J2 complex was determined by Brodersen in 2015.537 This milestone paper generates as many questions as answers as to how CP-lyase performs catalysis. Overall the PhnG2H2I2J2 complex has 2-fold symmetry (Figure 16). Only the PhnI monomers make direct contact and form the waist of the complex. The PhnG monomers form extensive interactions with PhnI, and extend an α-helix followed by a strand toward PhnJ. PhnJ is observed to form the top and bottom of the complex, binding to PhnI, the latter extending N- and C-terminal α-helices well into PhnJ. PhnH is observed at the periphery of the complex, making a distinct α-helix/α-helix interaction with PhnJ. This is a surprising result as PhnH forms a very similar α-helix mediated homodimer when produced and purified as a single polypeptide.538 This observation questions how homodimerization of PhnH is prevented from competing with assembly of the complex. Overall the structure of PhnG2H2I2J2 corresponds well with the assembly model proposed by Raushel and Russell,536 but the exposure of PhnH to solvent is surprising in light of the resistance of this component to cross-linking.529

screen revealed that PhnP is highly active with Ni2+ and Mn2+, while poor activity was observed with Zn2+. PhnP has wide substrate specificity, hydrolyzing a variety of diaryl- and arylmethylphosphate diesters. 534 With bis(p-nitrophenyl)phosphate and Mn2+, PhnP has kinetic parameters of kcat = 1.2 s−1, KM = 2.3 mM, kcat /KM = 410 M−1 s−1. Much greater specific activity is observed with 2′,3′-cyclic nucleotides, such as 2′,3′-cAMP (kcat = 1.7 s−1, KM = 0.11 mM, kcat /KM = 1.6 × 104 M−1 s−1). This is almost entirely due to a drop in KM. The reaction of PhnP with 2′,3′-cAMP is regiospecific, forming 3′AMP. Initially the higher specificity toward cyclic-nucleotides was puzzling until it was determined that 172 was the true substrate for PhnP.478,518 The structure of PhnP bound to malate and orthovanadate has been determined by X-ray crystallography, revealing a dinuclear metal ion active site that is used to activate water and the phosphate diester in the hydrolysis transition state.533,534 A model for binding 172 in the active site has been proposed.534 Raushel noted that in several bacterial phn operons phnP had been replaced by a gene encoding a histidinol phosphatase from the amidohydrolase superfamily. 515 The enzyme from Eggerthella lenta, Elen0235, was shown to catalyze the hydrolytic ring opening of 172 to form 175, and thus with regiospecificity opposite to PhnP. Surprisingly, the enzyme also functioned as a monoesterase, catalyzing the specific hydrolysis of the 2-phosphate of 175 to form 176. Running the reaction in H218O demonstrated that hydrolysis occurred by attack at the phosphorus atom in both steps. Equally surprising was that the kinetic parameters for both steps were very similar (kcat/KM = 1.4 × 105 M−1 s−1 and 3.3 × 105 M−1 s−1). Therefore, this enzyme has the paradoxical distinction of being promiscuous, acting as a diesterase and monoesterase with nearly equal efficiency, as well as highly specific: 175 and 176 are substrates, whereas 173 is not. To denote this unique activity, Elen0235 and its homologues were named cyclic phosphate dihydrolases or PhnPP. In the context of an overall CP-lyase pathway, the Pn phosphorus in this case is released as free Pi in the cell. 176 presumably can be isomerized to 177 by phosphoribomutase DeoB, followed by phosphorylation by ribose-1-phosphokinase RPK to form 173.535 5.2.4.7. PhnN. PhnN catalyzes the regiospecific phosphorylation of 173 to form 5-phospho-α-D-ribosyl-1-diphosphate, 174, also known as PRPP. This the end of the CP-lyase pathway in E. coli, and thus the Pn phosphorus ends up within the pyrophosphate leaving group of 174, which is used as a glycosyl donor to synthesize NAD+, guanosine, uridine, histidine, and tryptophan. The use of the CP-lyase pathway to directly support primary metabolism is intriguing as PRPP synthase, encoded by prs, is the primary enzyme responsible for generating 174 in the cell. This metabolic redundancy led to the surreptitious discovery of the function encoded by phnN. Hove-Jensen observed that an E. coli prs mutant continued to grow on media that was supplemented with guanosine, uridine, histidine, and tryptophan, but lacking NAD+. It was determined that the E. coli prs mutant had acquired a second mutation in the pst-phoU operon, which led to constitutive expression of the phn operon and thus phnN. It was subsequently shown that E. coli prs-phn or E. coli prs-phnN mutants required supplementation with guanosine, uridine, histidine, tryptophan, as well as NAD+ and therefore were entirely deficient in the synthesis of 174. Purified PhnN was subsequently shown to catalyze the conversion of 173 to 174 in vitro using ATP as the phosphoryl donor. This study provided 5762


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to 72 and is predicted to abstract a hydrogen from C272, the PhnG2H2I2J2 complex must be highly dynamic in order to bring G32 and C272 into proximity. A second potential active site was identified by a Zn2+ ion bound at the interface of PhnI and PhnJ. The Zn2+ ion was held by H328 and H333 of PhnI, while a third residue, H108, of PhnJ was observed 4.5 Å from the metal center. These three His residues appear to be critical for the function of CP-lyase; not only are they conserved but their substitution with Ala created CP-lyase variants that were unable to process Pn in vivo. The location of this second active site at the midpoint of a dead-end, solvent accessible tunnel is also intriguing. A sulfate ion is observed at the end of the tunnel, which may be a proxy for the phosphonate moiety of a Pn substrate. The role of PhnK and PhnL in the complex is an additional mystery. PhnK has the conserved sequence motifs of a nucleotide-binding domain that one finds in the ABC transporter family, including the α-helical and RecA-like domains. Nucleotide binding domain proteins, such as PhnC (Scheme 57), bind to the transmembrane component of the transporter and drive the transport of solutes across the membrane by hydrolysis of ATP or other nucleotides. This has compelling potential because nucleotides and phosphorylated ribose derivatives appear to enter and exit the PhnG2H2I2J2 complex during catalysis (Scheme 57). Specifically, PhnI and PhnJ of the complex must receive ATP and 171, respectively, while 170 must leave the complex to be hydrolyzed by PhnM. Although crystals of a PhnG2H2I2J2K complex could not be grown for X-ray crystallography, this complex was successfully observed by electron microscopy, revealing PhnK to be associated with PhnJ.537 Deletion of the domain of PhnJ that interacted with PhnK created a PhnG2H2I2J2 complex that no longer copurified with PhnK. Zhang and Raushel followed up with a higher resolution structure of PhnG2H2I2J2K using single-particle cryo-electron microscopy. 539 A positively charged surface on PhnK, comprised of a helix-turn-helix motif, is observed to bind to a negatively charged surface on PhnJ formed by an α-helix and a loop. Despite the presence of two identical binding sites, only one copy of PhnK can bind to the complex at one time as the RecA-like domain sterically hinders access to the second copy of PhnJ. The asymmetry introduced by PhnK suggests that the active sites of PhnI and PhnJ on one-half of the complex will have different reactivity than the other, depending on the location of PhnK. Intriguingly, binding of PhnK causes a conformational change in PhnJ that exposes G32 to solvent. This change does not happen in the unbound copy of PhnJ. Although exposure of G32 would be necessary to form a glycyl radical during catalysis, this is at odds with the observation that PhnK is not an essential component of any reaction of the CP-lyase pathway, at least in vitro. How PhnL interacts with the complex

Figure 16. Structure of the PhnG2H2I2J2 complex. The overall complex is shown in the center. The individual components are shown in isolation along the periphery at the same scale and relative orientation. The scale of PhnJ 2 is increased for clarity. PhnJ G22 is shown as red spheres, Cys residues as sticks. Zn ions in PhnI and PhnJ are shown as yellow spheres.

Two potential active sites were identified on PhnI and PhnJ. The two active sites (four overall in the complex) are consistent with the glycosylation and C−P bond cleaving reactions that are predicted to be catalyzed by the complex. Four Zn2+ ions are observed in the structure, two bound by the PhnJ monomers, and two others bound at the PhnI-PhnJ interface. Because the complex was purified and crystallized under aerobic conditions, an intact [4Fe-4S] cluster is not observed in the structure. Instead, the key Cys residues C241, C244, C266, and C272 of PhnJ are observed to bind a Zn2+ ion. As the thiol side chain of C272 is predicted to be a hydrogen donor during catalysis, its interaction with the Zn2+ ion as a thiolate is not likely an accurate representation of its catalytic orientation. Nevertheless this site is the likely location of the [4Fe-4S] cluster and where the 5′-deoxyadenosyl radical 72 is generated by reductive cleavage of SAM. Surprisingly, G32 is located 30 Å away from the active site in the vicinity of PhnH. Moreover, this residue is buried. As G32 was shown to donate a hydrogen

Scheme 61. Catabolism of 3 by the Oxidative Enzymes PhnY* and PhnZ

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Figure 17. Active site schemes for PhnZ in two conformations involving Y24 and E27.

reacquired from the substrate by the end of the catalytic cycle. PhnZ is specific for the (R)-enantiomer of 5, which is stereospecifically formed by PhnY*.139,143,144 The kinetic parameters for the conversion of (R)-5 are kcat = 11 min−1 and KM = 0.17 mM.143 PhnZ is equally active toward (R,R)-2amino-1-hydroxy-propylphosphonic acid 184 (kcat = 7 min−1, KM = 0.16 mM) but is inactive toward 1-hydroxyethylphosphonic acid 36. These results reveal a dependence on the 2amino group of the substrate for catalysis, and in part explains why the PhnY*/PhnZ pathway is specific for 3.540 The structure of PhnZ revealed that two metal ions are used to oxidatively cleave the C−P bond of (R)-5. X-ray crystal structures have been solved independently by two groups with PhnZ bound to the buffer additives L-tartrate and citrate as well as the substrate (R)-5.143,144 Two Fe ions (denoted Fe1 and Fe2) are observed in the active site, with (R)-5 bound in a bidentate fashion to Fe2 through the α-hydroxyl and phosphonyl oxygens (Figure 17A).143 Both Fe ions are hexacoordinate, and likely oxidized to the Fe(III) state during crystallization, which would account for the lack of turnover of (R)-5. In addition to the substrate, the metal ions are bound by a conserved set of four histidines, two aspartates, and a bridging water molecule or hydroxide ion. D59 also bridges the Fe ions, which along with H58 comprise the ’HD’ motif that defines this superfamily. In addition to binding the metal ions, the phosphonate group of (R)-5 is recognized by a polar pocket, while the α-hydroxyl interacts with H62. This fifth active site histidine is conserved in PhnZ homologues, but not in the other members of the HD phosphohydrolase superfamily. The PhnZ H62A variant (kcat = 2 min−1, KM = 0.4 mM) is a significantly poorer catalyst than the wild type PhnZ, indicating that this residue is important for catalysis.143 Elegant EPR and Mössbauer spectral studies by Wörsdörfer, Bollinger, and Pandelia revealed that the diiron atoms are electronically coupled and, more surprising, that the mixed oxidation state, Fe(II)/Fe(III), is the catalytically competent form of the cofactor.144 This is unusual, as other diiron oxygenases, such as methane monooxygenase, use a fully reduced, Fe(II)/Fe(II) cofactor to reduce molecular oxygen, whereas the corresponding Fe(II)/Fe(III) state is not stable. Therefore, PhnZ has the unique ability to stabilize this mixed oxidation state. The structure of PhnZ also suggests that a unique induced-fit mechanism is used to coordinate binding of substrate with dioxygen. Two loops that flank the active site fold inward upon binding (R)-5. One loop, containing two conserved residues, Y24 and E27, is observed to adopt two orientations in response to the 2-amino group of the substrate. In one conformation the phenolic oxygen of Y24 is observed bound to the apical position of Fe1, completing the hexacoordination state of this metal ion, while E27 is oriented outside of the active site

remains unresolved. PhnL does not copurify with the complex, but as described above, a weak interaction must occur as PhnL is required to dictate the glycosyl acceptor specificity of PhnI. 5.3. Oxidative C−P Bond Cleavage by PhnY*/PhnZ

The third known mechanism for Pn catabolism utilizes molecular oxygen and ferrous ion to oxidatively cleave the C−P bond (Scheme 61). The phnY* and phnZ genes encoding this pathway were first identified through a functional screen of a marine metagenomic DNA. Fosmids comprising a libary of this DNA were screened for their ability to complement an E. coli phn mutant for growth on 3 as a Pi source.540 Based on amino acid sequence PhnY* was predicted to be a nonheme iron/α-ketoglutarate dependent dioxygenase. This enzyme, discussed earlier in Section 3.5, catalyzes the hydroxylation of the α-carbon of 3 to form (R)-5 (Scheme 7). More puzzling was the sequence similarity of PhnZ to the HD superfamily of metal ion dependent phosphohydrolases, which implied a hydrolytic mechanism for C−P bond cleavage. The phnY* and phnZ genes appear most commonly in marine bacteria such as Plantomyces maris DSM8797, Plesiocystic pacif ica SIR-1, and Prochlorococcus sp. MIT9303 and MIT930116,540 but can also be found in the fungus Aspergillus niger. There is evidence that the substrate specificity encoded by phnY-phnZ varies across species. For example, these genes appear in association with a phosphite utilization operon, ptxABCD-phnY*Z, in Prochlorococcus sp.522 However, Prochlorococcus sp. phnY*Z does not encode the catabolism of 3 or a variety of other Pi sources such as hypophosphite 168, phosphite 34, phosphonoalanine 6, phosphonoacetate 7, and methylphosphonic acid 8.522 Also, phnZ frequently appears in association with phnW and phnX encoding the phosphonatase pathway for degrading 3,144,540 suggesting a different catabolic role. Likewise, a phylogenetic analysis of PhnZ revealed a number of distinct clades that are associated with Pn catabolism and likely represent different substrate specificities.144 An examination of the metal ion dependence of PhnZ led to the discovery that this enzyme was an oxygenase rather than a hydrolase.139 Only Fe(II) enabled PhnZ to convert 5 to Pi in vitro. ICP-MS analysis also indicated that PhnZ copurified with 1.2 mol of iron per mol of enzyme. While Fe(II) dependent hydrolases are known, a more common role for Fe(II) is activation of dioxygen. This indeed proved to be the case with PhnZ, which was shown to produce glycine along with Pi upon reaction with 5 (Scheme 61). Formally PhnZ is classified as a monooxygenase as only one atom of dioxygen is incorporated into glycine.144 Because PhnZ performs a four electron oxidation of a secondary alcohol to a carboxylic acid without the need for an external source of electrons, the two electrons donated by the Fe(II) ion to reduce dioxygen must be 5764


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Scheme 62. Proposed Mechanism for CP Bond Cleavage by PhnZ

(Figure 17A).143 The interaction of Y24 with Fe1, which is likely in a Fe(III) oxidation state, creates a charge transfer complex that affords crystals and concentrated solutions of PhnZ with a pink hue (λmax = 510 nm). This particular conformation of PhnZ appears to represent a precatalytic state as the 2-amino group of (R)-5 is not engaged by an active site side chain, while dioxygen is prevented from binding to either Fe ion due to the hexacoordinate state of both metal centers. However, a second conformation was observed in a crystal of PhnZ where E27 is observed to flip into the active site, forming an electrostatic interaction with the 2-amino group of (R)-5 (Figure 17B). This movement levers Y24 out of the active site (and disrupts the charge transfer complex), thereby freeing the apical site on Fe1 for a ligand. In the crystal structure a water molecule is observed at this position, but presumably during catalysis this would be the dioxygen binding site. The α−C-H bond of (R)-5 projects toward the water molecule, with only 3.1 Å separating the α-carbon and the water oxygen atom. Accordingly, this conformation likely represents a catalytic state poised to bind dioxygen at Fe1, which would be ideally positioned to stereospecifically oxidize the α-carbon of (R)-5. The precise role of Y24 in the PhnZ reaction is uncertain and all the more intriguing in the light of mutagenesis studies. While PhnZ E27A is significantly impacted as a catalyst (kcat = 3 min−1, KM = 1.1 mM), the PhnZ Y24F variant has essentially wild type activity.143 Although this can be interpreted as arising from a direct interaction between (R)-5 and E27, but not Y24, during catalysis, it is surprising that both residues are conserved in PhnZ homologues as Y24 does not appear to be important. However, E24 and Y24 appear to be specific to Pn catabolism, as a distant homologue of PhnZ, the myo-inositol oxygenase (MIOX) discussed below, does not have these residues. Nevertheless, the induced-fit mechanism by which PhnZ triggers the creation of a vacant dioxygen binding site on Fe1 is analogous to how other nonheme iron dependent oxygenases, such as TauD, bind their substrates prior to binding

and reducing dioxygen.141,541 More striking is the parallel observed in cytochrome cd1 nitrite reductase, where the active site residue Y25 controls access of dioxygen or nitrite to the heme iron.542,543 When the heme is in an oxidized, resting state, a strong electrostatic interaction is observed between the Fe(III) ion and the tyrosyl oxygen. Upon reduction to the active Fe(II) state, the interaction is weakened and Y25 is released, allowing dioxygen or nitrite access to the metal ion. In these examples, and perhaps with PhnZ, control of dioxygen binding presumably prevents spurious oxidation of the enzyme or the host cell in the absence of a substrate. PhnZ is also an illustrative example of how evolution can use a common scaffold to develop different chemistries. The closest structural homologue to PhnZ is MIOX, a mammalian enzyme that oxidatively cleaves the C1−C6 bond of myo-inositol to form D-glucuronic acid. MIOX is also a member of the HD phosphohydrolase superfamily but is only distantly related to PhnZ on the basis of amino acid sequence (15% sequence identity). The enzyme has been extensively studied (reviewed in544) and structures of the human545 and mouse546 enzymes have been determined. Like PhnZ, MIOX uses a mixed-valence Fe(II)/Fe(III) cofactor for catalysis, with dioxygen binding to Fe1 and the substrate to Fe2. The active site residues used to bind the metal ions are conserved in MIOX and PhnZ, with the exception that MIOX does not have a tyrosine side chain that protects the apical position of Fe1. Instead, a water molecule is only observed at this position. MIOX also engages metal ion bound hydroxyl of its substrate with a Lys side chain, analogous to H62 of PhnZ. While the evolution of oxidative C−P and C− C bond cleaving reactions can be rationalized as a change in substrate specificity, the jump between oxidative C−P and hydrolytic O−P cleavage is more interesting. In this regard an evolutionary link between hydrolytic and oxidative chemistry in the HD phosphohydrolase family might be represented by the Fe(II) dependent phosphodiesterase MptB from Methanocaldococcus jannaschii.547 MptB shares the conserved Fe(II) 5765


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of peptide bonds in dehydrophos 11. Many other “simple” chemical puzzles remain to be deciphered, such as the biosynthesis of phosphonochlorin 20 or nitrilaphos 19a.102 Likewise, while the most common way of synthesizing a C−P bond is by PEP mutase, the existence of K-26 21a and IB52 21b indicates that there is another mechanism waiting to be discovered. In addition to exploring new reactions, Pn biochemistry also borrows heavily from the familiar reactions of primary metabolism. It is tempting to view this biosynthetic plagiarism as evidence of Pn as ancient molecules that were available to early forms of microbial life as chemical building blocks. A second, more humbling lesson is that this field is still in its infancy. Unlike other natural product families, such as nonribosomal peptides, polyketides, terpenoids, and alkaloids, comparatively few Pn natural products have been characterized. Since the discovery of 3 in 1959, only 21 or so distinct structural classes of Pn have been identified (Figure 1), and far fewer have been linked to a defined biosynthetic pathway. The slow rate of discovery can be primarily attributed to the polar character of Pn, as well as the lack of chromophores, which presents a serious challenge to extracting, concentrating, and detecting these compounds, especially when culture titers are low. While 31P NMR spectroscopy can take advantage of the high chemical shift values that distinguish Pn natural products from phosphorylated metabolites,20 the sensitivity of this technique limits its use when dealing with dilute samples. Therefore, there is an acute need for methods that selectively enrich and detect Pn in order to advance this field. The discovery of phosacetamycin 14g through the use of immobilized metal ion affinity chromatography and precursor ion scanning mass spectrometry,167 and the fosfazinomycins 13a−b by chemoselective methylation,142 represent important steps toward addressing this problem.21 Genomics, in contrast, has advanced much more quickly and has emphasized that we have explored only a small part of Pn chemical space, and by extension, the associated enzymology. As has been observed in other natural product families such as indolocarbazoles555 or enediynes,556 enzymes encoded by gene clusters tend to coevolve, thus the phylogenetic relationship of a key biosynthetic enzyme will often mirror the chemical structural diversity of a natural product class. Because PEP mutase is the most common path to synthesizing a C−P bond, the corresponding gene can be used as a marker to identify Pn biosynthetic gene clusters, as well as gauge Pn chemical space.21,72 Gratifyingly, this chemical space is quite large. From a data set of microbial genomes, metagenomic samples, and soil clone libraries Metcalf has estimated that there are 656 distinct groups of Pn structures.72 Even within one genus of bacterium there is much to be uncovered. In a study that sequenced 10 000 actinomycete genomes,102 Metcalf identified 278 strains (2.8%) with the genetic capacity to synthesize Pn, which are predicted to form 64 chemically distinct classes of compounds. The majority of the biosynthetic gene clusters encode new compounds. Extrapolating from this data set, it was estimated that actinomycetes have a total genetic potential to synthesize 125 distinct classes of Pn natural products. Therefore, in contrast to other natural product families, the rediscovery of known compounds will be less of a problem in the Pn field for some time to come! There is also compelling evidence that additional pathways for degrading Pn exist in the microbial world. The human pathogens Campylobacter jejuni and Helicobacter pylori can use

binding residues and even has a Lys residue that corresponds to H62 of PhnZ. The Fe(II) specificity of MptB likely reflects the reducing and anaerobic environment that is inhabited by M. jannaschii. As well, Fe(II) rather than Fe(III) may achieve a better balance between Lewis acidity and electrostatic attraction when forming a hydroxide ion as a nucleophile. A switch from Lewis acid to oxidative chemistry could thus be envisioned by creating a vacancy on one of the Fe(II) ions to allow activation of dioxygen.143 The proposed C−P bond cleaving mechanism for PhnZ is summarized in Scheme 62.139,143 Initially Fe1 and Fe2 are proposed to be in Fe(III) and Fe(II) oxidation states (I), respectively, to enable a strong interaction to form between Y24 and Fe1. Upon binding (R)-5 an electron transfer is proposed to occur between the metal centers (I to II), reversing their oxidation states. The weaker interaction between Fe1 in the ferrous form and Y24 would favor the release of this side chain from the dioxygen binding site, which would be reinforced by a favorable interaction between E27 and the 2amino group of (R)-5. Additionally, the greater Lewis acidity of Fe2 in a ferric oxidation state would be better suited for binding (R)-5. With H62 acting as a general base, Fe2 may also promote the ionization of the α-hydroxyl group. Such changes in ligand environment and oxidation states of the Fe(II)/ Fe(III) cluster would account for the distinct changes in the EPR spectra that are observed with PhnZ upon binding (R)5.144 The free ligand site on Fe1 would allow the reduction of dioxygen to form a ferric-superoxide (Fe(III)−O-O•−) species (III). Such an intermediate is observed to form on MIOX, which subsequently cleaves the C−H bond of the substrate.548 In the case of PhnZ, the radical-like transition state for α−C-H bond cleavage would be stabilized by an ionized α-hydroxyl group, and thus would resemble a ketyl anion. In the corresponding MIOX step, DFT calculations suggest that a substrate radical intermediate is not formed, but instead the single electron is donated to the ferric Fe2 ion.549 In the context of PhnZ this leads to the acylphosphonate intermediate (IV). Next, attack by the Fe1 bound hydroperoxide on the acylphosphonate would form the Criegee-like intermediate V. Calculations on MIOX549 suggest that instead of undergoing a Baeyer−Villiger rearrangement to form an acylphosphate, the lower activation barrier corresponds to homolytic cleavage of the peroxide bond, producing a Fe(IV)O species at Fe1 (VI). This step is also predicted to be rate determining. Abstraction of the hemiketal hydroxyl hydrogen by the ferryl oxygen would lead to an alkoxy radical that can initiate β-scission of the C−P bond (VI to VII). Carbon or alkoxy radicals preferentially decompose through beta-scission reactions,550,551 and the corresponding radical beta-scission of C−P bonds is also well-known,552,553 thus this step is chemically reasonable. The phosphonyl radical produced by this step is predicted to reduce Fe2 back to the initial ferrous state, analogous to how Cu(II) can be reduced to Cu(I) by phosphonyl radicals.554 Glycine and metaphosphate are produced by C−P bond cleavage (VIII), with the latter captured by solvent to produce Pi.

6. SUMMARY AND OUTLOOK If there is one overriding lesson in Pn biochemistry, it is that deceptively simple structures and transformations are often the result of unusual enzyme chemistries. Examples include the installation of the methyl and epoxide groups of fosfomycin 9, oxidative cleavage of the C−C bond in 4, the synthesis and degradation of methylphosphonic acid 8, or even the formation 5766


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acknowledges support from NSERC, the Canadian Foundation for Innovation, the Canadian Glycomics Network, and Wilfrid Laurier University.

Pn as a Pi source, yet genes encoding known Pn catabolic pathways are not found in the genomes of these organisms.557−559 In the case of H. pylori, this catabolic capacity affords resistance toward N-phosphonoacetyl-L-aspartate,560 a potent transition state analog inhibitor of aspartate carbamoyltransferase.561 Such pathways are of particular interest for discovering new mechanisms for cleaving C−P bonds. However, in contrast to Pn biosynthesis, one cannot rely on marker genes to discover new catabolic pathways. Instead, functional genomic strategies that screen microbial and metagenomic DNA for genes encoding the liberation of Pi from Pn will likely provide the most efficient means of discovering these new pathways.540

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AUTHOR INFORMATION Corresponding Authors

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

The authors declare no competing financial interest. Biographies David L. Zechel studied chemistry at the University of Toronto before moving to the University of British Columbia for his Ph.D. (2001) with Professor Stephen G. Withers where he investigated glycosidase mechanisms. This was followed by postdoctoral studies with Professor Gideon J. Davies (York University, U.K.), where he explored transition-state analogue inhibitors by X-ray crystallography, and with Professor Andreas Plückthun (University of Zürich), where he studied directed evolution of protein function. In 2004 he began his independent career at Queen’s University in the Department of Chemistry. His programs seek to identify and characterize new enzymes in catabolic and biosynthetic pathways, with a particular emphasis on phosphonate related enzymes. Geoff P. Horsman studied agricultural chemistry and biochemistry at the University of Saskatchewan before obtaining his M.Sc. in Chemistry from McGill University studying directed evolution of enantioselective enzymes with Professor Romas J. Kazlauskas. After his Ph.D. (2008) with Professor Lindsay D. Eltis at the University of British Columbia investigating the mechanistic enzymology of aromatic compound degradation, he completed a postdoctoral fellowship in natural products biosynthesis with Professor Ben Shen at the University of WisconsinMadison. In 2011 he moved to Wilfrid Laurier University where his research program focuses on the enzymology of natural product biosynthesis with an interest in phosphonate biochemistry.

ACKNOWLEDGMENTS We would like to thank our students and collaborators for their essential and inspired contributions to our research on phosphonate related enzymes. It has been a privilege to work scientists from a number of international laboratories, including Zongchao Jia, Queen’s University; Alexander Yakunin, University of Toronto; Friedrich Hammerschmidt and Katharina Pallitsch, University of Vienna; Edward F. DeLong and Asuncion Martinez, MIT; Peter B. Wyatt, Queen Mary University of London; and Bjarne Hove-Jensen, Metropolitan University College, Copenhagen. D.L.Z. is additionally grateful to NSERC for a Discovery Grant and Directed Accelerator Supplement to support his research into phosphonate catabolism and natural product biosynthesis. G.P.H. gratefully 5767


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