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Published in final edited form as: J Nat Prod. 2025 Dec 30;89(1):198–207. doi: 10.1021/acs.jnatprod.5c01272

CURE-ating the Substrate Scope and Functional Residues of Nonheme Iron(II) α-Ketoglutarate-Dependent Hydroxylase BesE

Austin R Hopiavuori 1,†, Beau S Andre 1,†, Emely D Avalos 1,†, Surina L Beal 1,†, George R Beck 1,†, Charles J Choi 1,†, Elizabeth M Dolzhansky 1,†, Michelle X Du 1,†, Isabella S Gallardo 1,†, Cyrus J Ghorbani 1,†, Kyle M Hinaga 1,†, Analynn T Nguyen 1,†, Edward H T Pham 1,†, Shaun M K McKinnie 1
PMCID: PMC13207679  NIHMSID: NIHMS2164910  PMID: 41467595

Abstract

Nonheme iron(II) α-ketoglutarate-dependent dioxygenases (Fe/αKGs) play important roles in functionalizing biological substrates from individual amino acids to macromolecules. BesE, a unique homologue from the actinobacterial β-ethynylserine biosynthetic pathway, catalyzes a highly selective hydroxylation on dipeptide substrate γ-l-glutamyl-l-propargylglycine (1). Inspired by this transformation, our year-long Course-based Undergraduate Research Experience (CURE) laboratory interrogated BesE catalysis using an interdisciplinary approach. After establishing a modular chemical synthesis of 1, we rationally designed 14 non-native analogues with key alterations to specific substrate moieties putatively involved in enzymatic recognition. Following in vitro enzymology and the application of chemical derivatization techniques compatible with all substrate analogues and putative products, we determined that the terminal alkyne moiety is not essential while reinforcing the significance of the γ-glutamyl moiety for hydroxylation activity. Thirteen rationally designed BesE mutants established the importance of polar active site residues for substrate recognition and catalysis. This work establishes a baseline of BesE recognition from both a chemical and biochemical perspective and contributes to the growing understanding of Fe/αKG recognition on biomedically relevant targets. Moreover, this contributes to the growing examples of using natural products and their biosynthetic enzymology as a vibrant platform for the interdisciplinary training of early career biomedical researchers.

Graphical Abstract

graphic file with name nihms-2164910-f0001.jpg

Nonheme iron(II) α-ketoglutarate-dependent dioxygenases (Fe/αKG) are a diverse family of enzymes that perform important roles in both primary and secondary metabolism.1 Within primary metabolism, these enzymes are involved in the modification of small molecules up to macromolecules and play important physiological regulatory roles, including oxygen-sensing, stabilizing structural proteins like collagen, and modifying nucleic acid bases.2 Fe/αKG enzymes exhibit expanded chemical reactivity within secondary (or specialized) metabolism and contribute broadly to the diversity observed within natural product biosynthesis.3,4 Through a conserved catalytic cycle involving cofactor iron(II) and cosubstrates α-ketoglutarate and molecular oxygen, Fe/αKGs generate a high valency Fe(IV) oxo species capable of radical hydrogen abstraction, typically from an unactivated, aliphatic C–H bond.5 Following substrate radical formation, native or engineered Fe/αKGs can perform fascinating and synthetically useful reactions, including halogenation, epimerization, desaturation, endoperoxidation, carbon–nitrogen, and carbon–carbon bond formations.3,6,7 However, the most common reaction pathway involves radical rebound from the Fe(III)–OH intermediate to form hydroxylated products with high regio- and stereoselectivity. We have been interested in understanding how this family of oxidative enzymes modulates substrate hydroxylation and other chemistries within natural product biosynthetic and focused biocatalytic directions.8

We were particularly inspired by the biosynthetic pathway of known antimetabolite β-ethynylserine (βes) from Streptomyces cattleya, discovered and fully elucidated by the Chang group in 2019 (Figures 1A and S1).9 Through the action of three biosynthetic enzymes (BesBCD), l-lysine is transformed into terminal alkyne-containing l-propargylglycine (l-Pra). Subsequent ligation of this noncanonical amino acid (ncAA) to the γ-carboxylate of l-glutamic acid by ATP-dependent ligase BesA formed dipeptide γ-l-glutamyl-l-propargylglycine (γ-l-Glu-l-Pra, 1). BesE performs the final stereoselective β-hydroxylation to generate γ-l-glutamyl-l-β-ethynylserine (γ-l-Glu-l-βes, 2). Two Fe/αKGs with diverging chemistries are highlighted within 2 biosynthesis: BesD initiates the biosynthetic pathway by performing a stereoselective γ-chlorination on l-lysine, whereas BesE catalyzes a conventional hydroxylation reaction to create the threonine-mimicking antimetabolite pharmacophore.10 Considerable mechanistic, structural, and protein engineering efforts have provided insight into BesD halogenation catalysis and how it diverges from conventional l-lysine hydroxylation.11,12 We were instead interested in the unusual features associated with the more canonical hydroxylase BesE. As noted in its original characterization, this Fe/αKG did not appreciably hydroxylate l-Pra in vitro, suggesting an important role for the γ-l-glutamyl moiety.9 Moreover, the ability of BesE to catalyze radical hydrogen abstraction without rearrangement of the adjacent alkyne moiety was unusual from a chemical reactivity perspective (Figure S2). BesE itself was characterized via a coupled assay during its initial discovery9 and subsequent cell-free lysate assays,13 thus providing an opportunity to specifically interrogate it individually. In addition, given the structural similarity of βes mimicking l-threonine, we were curious if native or engineered BesE could produce other non-native antimetabolites in appreciable yields.

Figure 1.

Figure 1.

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(A) BesE catalyzed β-hydroxylation of γ-l-glutamyl-l-propargylglycine (γ-l-Glu-l-Pra, 1) to form γ-l-glutamyl-l-β-ethynylserine (γ-l-Glu-l-βes, 2) in the context of the β-ethynylserine biosynthetic pathway. The structure of l-threonine is shown for comparison to the related antimetabolite β-ethynylserine. (B) General project overview for our Course-based Undergraduate Research Experience to interrogate an improved chemical, biochemical, and analytical understanding of BesE catalysis.

Through this study, our year-long Course-based Undergraduate Research Experience (CURE) lab of 12 undergraduate researchers investigated the key features for BesE catalysis using both a chemical and biochemical approach (Figure 1B). After developing a modular four-step chemical synthesis of 1, undergraduate student researchers followed comparable organic chemistry and analytical techniques to individually design and synthesize 14 non-native analogues and assess the extent of substrate engineering. Student pairs followed established protocols to heterologously express and purify native BesE and then identified amino acid residues of interest within the putative active site pocket following structural prediction. Individual researchers designed mutagenesis primers, completed site-directed mutagenesis, and purified 13 mutants to probe the biochemical features of hydroxylation catalysis. Moreover, we collaboratively generated a robust analytical assay and chemical derivatization approach to detect and quantify the extent of conversion of multiple, polar non-native substrates via in vitro BesE assays. These cumulative efforts provide an improved understanding of this unique and significant Fe/αKG hydroxylase homologue and this broader enzyme family while serving as a template to strategically integrate interdisciplinary natural product research into upper division undergraduate research environments.

RESULTS AND DISCUSSION

The chemical synthesis of native substrate 1 was initiated by an acid-catalyzed methyl ester formation of l-propargylglycine in 2,2-dimethoxypropane (Figure 2A). The resulting l-propargylglycine methyl ester hydrochloride salt (3) was subsequently coupled to the γ-carboxylic acid side chain of Boc-l-Glu-OtBu using hexafluorophosphate azabenzotriazole tetramethyl uronium (HATU) in the presence of triethylamine, isolating protected amide product 4 following silica flash column chromatography. Sequential methyl ester hydrolysis using aqueous lithium hydroxide followed by trifluoroacetic acid (TFA) deprotection of Boc- and tert-butyl ester groups afforded 1 as a TFA salt in an appreciable yield (approximately 10% yield after four synthetic steps).

Figure 2.

Figure 2.

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(A) Modular four-step synthesis of native BesE substrate 1. (B) Structures of non-native BesE substrate analogues (5–18) grouped by region of modification at either the Pra side chain (blue), Pra α-carboxylic acid (purple), or γ-glutamyl moiety (green).

Following the successful synthesis of 1, a series of non-native analogues were designed with modifications to either the glutamic acid or propargylglycine moieties. These were selected by individual undergraduate researchers and used to assess the importance of key substrate features for BesE catalysis and selectivity (Figure 2B). Alteration of the propargylglycine amino acid side chain generated five novel side chain modified analogues nearest the site of hydroxylation. Adapting the previous synthetic scheme to use either l-allylglycine (l-Alg) or l-norvaline (l-Nva) allowed the generation of substrate analogues with the same carbon chain length but either a terminal alkene (γ-l-Glu-l-Alg, 5) or a linear saturated alkane (γ-l-Glu-l-Nva, 6), respectively. Other amino acid starting materials were selected, beginning with l-homopropargylglycine (l-Hpg) to retain the terminal alkyne feature but extend it by one methylene unit (γ-l-Glu-l-Hpg, 7), and a simplified l-alanine containing analogue (γ-l-Glu-l-Ala, 8) was created to assess hydroxylation activity lacking this unique chemical feature. The importance of the Pra l-stereocenter was investigated by adapting the same synthetic workflow with d-propargylglycine to generate an epimerized analogue (γ-l-Glu-d-Pra, 9). Efforts to replace the terminal alkyne with a nitrile were stymied due to chemical incompatibility during the methyl esterification reaction on β-cyano-l-alanine (Figure S3), and this analogue was not pursued further.

Another series of four analogues were developed to interrogate the significance of the l-Pra α-carboxylic acid for BesE recognition. Intermediate 4 was directly subjected to TFA deprotection, generating methyl esterified analogue γ-l-Glu-l-Pra-OMe (10). Coupling Boc-l-Glu-OtBu with chiral amine (R)-pent-4-yn-2-amine (Pya), purification of the amide intermediate, and subsequent TFA deprotection enabled the synthesis of an analogue 11 (γ-l-Glu-Pya, 11) that replaced this moiety with an α-methyl group while retaining the desired propargyl group stereochemistry. Adapting a similar workflow beginning with but-3-yn-1-amine (Bya) generated analogue 12 (γ-l-Glu-Bya) that lacked this C-terminal functionality altogether. Lastly, C-terminal amide analogue γ-l-Glu-l-Pra-NH2 (13) was synthesized by Fmoc-solid phase peptide synthesis on Rink Amide resin by the following sequence: loading Fmoc-l-Pra-OH on the resin by HATU coupling; Fmoc-deprotection using piperidine; HATU coupling of Boc-l-Glu-OtBu; and global TFA deprotection and concomitant resin cleavage.

The last series of analogues were designed to modify chemical features on the γ-l-glutamic acid side of the substrate. Four putative substrates were synthesized by HATU coupling key intermediates to 3 and then following necessary deprotection steps, resulting in the removal of the following: the α-carboxylic acid (GABA-l-Pra, 14); the α-amine (glutarate-l-Pra, 15); both α-moieties (But-l-Pra, 16); or a side chain methylene (β-l-Asp-l-Pra, 17). Lastly the same synthetic workflow to 1 was altered to use Boc-d-Glu-OtBu, generating an epimerized analogue (γ-d-Glu-l-Pra, 18) to investigate the importance of the Glu stereocenter. With established substrate 1 and 14 novel analogues in place, we next focused on the biochemical and analytical aspects of BesE interrogation.

To generate our recombinant BesE enzyme, we synthesized the gene as an Escherichia coli optimized construct and cloned it into a pET28a vector with an N-terminal hexahistidine tag. Recombinant BesE was expressed inE. coli BL21(DE3) cells following conventional IPTG-induction and purified to near homogeneity using Ni-NTA affinity chromatography, producing typical yields of 40 mg/L. Any copurified iron(II) was removed by treatment with EDTA prior to size exclusion chromatography to minimize any potential aerobic BesE oxidation and putative inactivation, a strategy that we routinely use for other Fe/αKG homologues.14 In vitro BesE assays were established following adapted literature conditions from either the one pot BesAE9 or cell free protein synthesis13 publications and analyzed by ultrahigh performance liquid chromatography-mass spectrometry (UPLC-MS). Initial assays with 1 (1.0 mM) were set up in 50 mM K2HPO4 pH 8.0 buffer with 5 mol % BesE (50 μM) and all necessary cofactors and cosubstrates (FeSO4, αKG, l-ascorbate, and O2). Subtle optimization efforts revealed that increasing the concentrations of both l-ascorbate and FeSO4 improved the overall in vitro turnover of BesE based on relative extracted ion chromatogram (EIC) intensities (Figures S4 and S5). Despite this optimization, complete in vitro consumption of 1 was not achieved, which was consistent with previous literature efforts.9,13 To improve the reproducibility of peak shapes, intensities, and retention times for polar analytes 1 and 2 following in vitro assays, we adopted two chemical derivatization methods (Figure 3A). Primary amines were derivatized with 2,4-dinitro-5-fluorophenyl-l-alanine amide (l-FDAA, Marfey’s reagent) following established conditions (Figure S6),15,16 and terminal alkyne-containing substrates were reacted via copper(I)-catalyzed azide alkyne cycloaddition (CuAAC) with coumarin azide (Figure S7).9 Importantly, at least one derivatization approach would be compatible with each of the synthetic non-native substrates and their putative hydroxylated products to improve the chromatographic retention properties and analytical reproducibility.

Figure 3.

Figure 3.

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(A) BesE in vitro enzyme assay with native substrate 1 and subsequent derivatization with either 7-azido-4-methylcoumarin (left) or Marfey’s reagent (right). Extracted ion chromatograms of derivatized β-hydroxylated product 2 for assays incubated in the presence (red) and absence (black) of purified BesE (EICs: 460.13; 511.13 ± 0.3 m/z [M + H]+ for 7-azido-4-methylcoumarin and Marfey derivatized products, respectively). (B) Extracted ion chromatograms of Marfey derivatized hydroxylated substrate analogues 5, 6, and 18, for assays incubated in the presence (red) and absence (black) of purified BesE (EICs: 513.15; 515.16; 511.13 ± 0.3 m/z [M + H]+ respectively). (C) Structures of substrate analogues which were not appreciably hydroxylated by BesE in vitro.

With the in vitro assay and derivatization conditions in hand, we next looked to identify the scope of BesE hydroxylation biochemistry on non-native substrates. Each non-native substrate (5–18; 500 μM) was incubated with 4 mol % wild-type BesE (20 μM) for 3 h under optimized cofactor and cosubstrate concentrations and an aerobic environment (Figures S8-S21). BesE assays in the presence of 1, and in the absence of enzyme, corresponded to positive and negative controls, respectively. Following chemically appropriate derivatization and UPLC–MS analysis, we identified that three analogues (5, 6, and 18) showed a significant [M + 16] m/z feature that was absent in the no enzyme control (Figures 3B, S8, S9, and S21). Substrates 5 and 6 had modifications directly near the native site of hydroxylation, with allyl- and n-propyl-amino acid side chains, respectively. The epimerized γ-d-Glu-l-Pra derivative 18 also showed putative activity supportive of hydroxylation. Analogues that failed to be appreciably hydroxylated in vitro included the following: methylene extended 7; simplified l-Ala derivative 8; epimerized d-Pra-containing 9; any alteration to the Pra carboxylate (10–13); and all other modifications to the γ-l-Glu moiety (14–17) (Figures 3C and S10-S20). While modest from a substrate scope perspective, these experiments highlighted the strong preference of BesE for its native substrate 1 and underscored the importance of the γ-Glu moiety for enzymatic recognition and direct hydroxylation chemistry.

After assessing the extent of hydroxylation chemistry on non-native substrates, we next examined the biochemical features that are significant for BesE catalysis. In the absence of a crystal structure, we used an AlphaFold 3 model17 of BesE, generated a structural model of 1 using Avogadro,18 and docked the substrate into the putative active site using AutoDock.19 Using the 25 lowest energy docked structures and filtering for orientations that positioned the l-Pra β-hydrogen in a productive position (12 of 25 structures), we hypothesized key BesE amino acid residues that could be contributing to substrate recognition and catalysis. Sequence and structural alignment identified the conserved catalytic active site triad (H153, D155, and H238) involved in binding iron(II) for oxidative chemistry. While it is difficult to precisely predict where the organic substrate binds, especially given the dynamics associated with Fe/αKG catalysis, this enabled us to generate testable hypotheses to probe BesE substrate recognition. A total of 13 residues were selected by individual students for substitution based on their proximity (within 6 Å) to docked 1 (Figures 4A and S22) and sequence conservation in other putative actinobacterial besE homologues (Figure S23). The two facial triad histidines (H153, H238) were left untouched; however, D155 was selected given that its substitution contributes to halide coordination in Fe/αKG halogenase homologues.20 Other selected residues were broadly categorized into putative hydrophobic pocket shaping (M70, V146, A175, F232, V240, and F255) or hydrogen bonding residues (R144, T150, Q173, N177, Y207, and R249). Most selected residues were mutated to alanine, with the exception of A175W, which was designed to assess if a bulkier side chain impeded 1 hydroxylation activity.

Figure 4.

Figure 4.

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(A) Depiction of the BesE AlphaFold 3 model, with mutant residues assayed shown as spheres. Mutation sites of hydrophobic residues are shown in blue; mutation sites of polar residues are shown in red. Mutations N177A and F232A, which were produced in lower titers and were not able to be assayed, are shown in gray. The catalytic triad residues (black) and docked substrate 1 (gray) are shown as sticks. (B) Bar graph displaying in vitro mutant activity in comparison to wild-type BesE. Mutants are grouped and colored by hydrophobic (blue) and polar side chains (red) in the same manner as above. Variants that did not show any conversion to product are indicated with an “X”. Assays were run in quadruplicate (n = 4) for each BesE mutant.

Following successful site-directed mutagenesis (Table S1) and sequence confirmation, BesE mutants were transformed into E. coli BL21(DE3) cells and successfully heterologously expressed and purified following analogous protocols. All 13 purified BesE variants were isolated in >90% purity by SDS-PAGE (Figure S24). However, mutants N177A and F232A were isolated in lower concentrations, showed poor solubility, precipitated during efforts to further concentrate them, and were excluded from being functionally assessed in vitro. The remaining 11 BesE variants were incubated with 1 in quadruplicate following previously established in vitro assay conditions, l-FDAA derivatization, and analysis by UPLC–MS. The relative production of derivatized 2 was standardized to an internal chloramphenicol control to correct for deviations in ionization efficiency and compared to wild-type BesE activity (Figure 4B). While all mutants showed reduced in vitro 1 hydroxylation activity (Figures S25-S35), an interesting trend was uncovered. Alanine substitution of hydrophobic pocket shaping residues retained some 2 production, with F255A showing the highest relative activity (56%) and a continued decrease across the V146A, M70A, and V240A BesE variants. These indicate that these residues are important for efficient substrate recognition but not essential for catalysis. The A175W mutant had detectable hydroxylation activity but at ~1% wild type conversion. This suggested that this normally small residue is helping to shape a critical portion of the substrate bonding pocket and putatively occludes 1 from constructive binding. In contrast, interrogating alanine mutations of residues with polar side chains nearly abolished the 1 hydroxylation activity. T150A was the only variant from this second group of mutants that retained any in vitro activity (23% wild type), potentially due to its more distant position from the predicted active site. All other polar side chain BesE mutants (R144A, D155A, Q173A, Y207A, and R249A) showed a complete loss of 2 production, indicating their importance in 1 recognition via hydrogen bonding or ion pairing interactions. We assessed D155A for its halogenation potential on 1 by extracting for a putative chlorinated product m/z value; however, no appreciable signal was observed (Figure S36). This corroborates multiple experimental and computational reports that additional engineering efforts are needed in on top of mutating the facial triad aspartic acid to enable efficient Fe/αKG halogenase activity.11,12,20-22 While none of our single point mutations improved in vitro BesE activity, they provided valuable functional insight into the significance of key hydrogen bonding and pocket shaping residues for efficient 1 hydroxylation catalysis. Moreover, they highlight the increasing utility of using structural prediction models to form experimentally testable hypotheses.

Overall, our study identified the extent of in vitro BesE hydroxylation catalysis on a non-native substrate library and suggested putative structural features that are important for 2 production. These focused substrates helped identify that BesE has limited tolerance for alteration at key regions of native substrate 1. None of our analogues with substitutions to the propargylglycine α-carboxylic acid (10–13) were appreciably hydroxylated by BesE in vitro. While epimeric γ-d-Glu analogue 18 showed some hydroxylation, negligible activity was observed on other γ-glutamyl modified analogues 14–17. This highlighted that the presence of both α-amine and α-carboxylate moieties on the glutamyl moiety is essential for BesE recognition. Moreover, the absence of a putative hydroxylated product on the methylene-truncated β-l-Asp-l-Pra 17 indicates that the chain length of the glutamyl moiety is important. These results complement the BesE mutant assays, where the alanine substitution of polar residues in the putative substrate binding pocket abolishes efficient 1 hydroxylation. While prediction in the absence of structural data is hypothetical, these cumulative data suggest there are important recognition elements, potentially hydrogen bonding or ionic interactions, that must be satisfied on both the substrate and enzyme end to afford this level of specificity. This further corroborates the initial report that l-Pra itself does not serve as a BesE substrate, underscoring the importance of this conserved γ-glutamylation step by BesA prior to stereoselective hydroxylation. The addition of chemical handles to standalone amino acid substrates prior to Fe/αKG reactions has been previously observed for the SadA amino acid β-hydroxylase,23 which showed promiscuity across a variety of N-succinylated hydrophobic amino acids and has served as a vibrant platform for further biocatalytic diversification.24,25 The presence of a dedicated hydrolase to liberate the hydroxylated product26 does not seem to be observed within the bes BGC but could speak to additional recognition, self-protection, or assisting with export of the precursor to the βes antimetabolite.

Intriguingly, substrate modifications occurring near the site of hydroxylation were tolerated provided the side chain carbon length was conserved. Both alkene (5) and alkane (6) derivatives showed reproducible hydroxylation albeit with a relative decrease in conversion compared to the native substrate. Although slightly counterintuitive, this indicates that the terminal alkyne functionality itself is not essential for BesE recognition and catalysis. However, the absence of hydroxylated products for methylene extended analogue 7, alanine-derived analogue 8, and epimerized analogue 9 show the limitations of additional side chain modification. While mutation of hydrophobic BesE residues retained some level of in vitro 2 production, both valine mutations V146A and V240A displayed substantially reduced activity despite being two of the more conservative mutations screened in this study. Molecular docking suggested that these residues flank a hydrophobic pocket near the catalytic site that could be significant for properly orienting the propargyl side chain. Additional targeted mutagenesis fueled by structural insight would provide further credibility for this hypothesis. While low in vitro turnover prevented the isolation of hydroxylated 5 and 6 products, efforts to structurally characterize them and assess their ability to serve as antimetabolites in other biological systems are underway.

Our interdisciplinary investigation into BesE catalysis provides insight into necessary features on both the substrate and macromolecular levels. Terminal alkyne containing scaffolds are relatively scarce in Nature and, when present, are typically observed at the end of fatty acid chains.27,28 The ability of this unique Fe/αKG homologue to selectively modify alkyne-containing amino acids generates additional tools for their application in natural product and biorthogonal labeling contexts.29 While this work provides an initial baseline into the biocatalytic capacity of BesE, building upon these results with experimentally validated structures and expanded mutant and substrate libraries would comprehensively identify its potential applications for generating hydroxylated terminal alkyne-containing amino acids and peptides. Additional efforts via directed evolution30,31 and high-throughput screening methods32 would rapidly diversify the testable BesE variants in relation to this goal. Nonheme iron enzymes have proven to have broad utility in both biocatalysis and chemoenzymatic synthesis;33-36 moreover, a remarkable array of inspiring transformations have also been observed from alteration of Fe/αKGs on complex meroterpenoid37 and tropolone38 substrates. An improved understanding of their fundamental biochemical and chemical scope features empowers these impactful applications in diverse non-native substrates. The application of multiple interdisciplinary technical skills while encouraging individual goal setting and completion on a focused research project creates a vibrant platform for the training of early career biomedical researchers (Tables S2 and S3). Moreover, this study adds to the growing examples of leveraging natural products and their associated metabolomics, genomics, molecular biology, protein biochemistry, and biosynthetic enzymology in CURE or comparable upper division laboratory settings.39-42

EXPERIMENTAL SECTION

General Experimental Procedures

All chemicals, including solvents and media components, were used as received from commercial suppliers (Millipore Sigma, Thermo Fisher Scientific, and Enamine). Reagents were used without further purification unless otherwise noted.

Reactions were monitored using thin-layer chromatography (TLC, Merck, 60 F254). TLC plates were visualized with a UV lamp at 254 nm and stained with either potassium permanganate (1.5 g of KMnO4, 10 g of K2CO3, 1.25 mL of 10% NaOH in 200 mL of water) or ninhydrin (1.5 g of ninhydrin, 3 mL of AcOH in 100 mL of n-butanol) staining solutions. TLC Rf values were rounded to the nearest 0.05. Silica ((Alfa Aesar, 60 (215–400 mesh)) was used when flash column chromatography was employed to purify compounds. Rotary evaporation was used to concentrate samples under reduced pressure. Additionally, polar final products were concentrated and dried via lyophilization (Labconco FreeZone 4.5 Liter). Nuclear magnetic resonance (NMR) spectra were obtained by using an Avance III HD spectrometer (Bruker) equipped with a BBFO SmartProbe at 500 MHz (1H NMR) or 125 MHz (13C NMR) using CDCl3 or D2O as solvents. Chemical shifts (δ) are reported in parts per million and referenced to methanol (δ = 3.31 ppm for 1H, δ = 49.0 ppm for 13C NMR) as an internal standard for samples in D2O or the CDCl3 solvent signal (δ = 7.26 ppm for 1H, δ = 77.2 ppm for 13C NMR) for samples in CDCl3. NMR data are reported as follows: s = singlet, d = doublet, t = triplet, q = quartet, m = multiplet, and J = coupling constant in Hz.

General LCMS measurements were measured on a Bruker Elute UHPLC system coupled with a Bruker amazon SL ESI-Ion Trap mass spectrometer in positive mode. Compounds were separated via reversed-phase chromatography on a Bruker Intensity Solo C18(2), 2 μm-2 × 100 mm column with the eluents water + 0.1% formic acid (solvent A) and acetonitrile + 0.1% formic acid (solvent B). The LC method uses a flow rate of 0.3 mL/min and the following gradient: 5% B (3 min), 5–15% B (3 min), 15–100% B (3 min), 100% B (2 min), 100–55% B (1 min), 5% B (2 min).

General Chemical Synthetic Methods for Native BesE Substrate 1 and Analogues

General Reaction A. Methyl Esterification of Amino Acids.

Approximately 0.1 g of l-propargylglycine or analogue was suspended in 10 mL of 2,2-dimethoxypropane. To the suspension was added 1 mL of concentrated (12 N) HCl. The mixture was stirred at room temperature (RT) overnight (16–18 h). The mixture was concentrated via rotary evaporation at 50 °C. The dark brown residue was resuspended in 10 mL of water and washed with EtOAc (3 × 10 mL). The aqueous layer was frozen in a dry ice/acetone bath and lyophilized to afford the product without further purification.

General Reaction B. Coupling Reaction.

To approximately 0.05 g of O-methyl-l-propargylglycine (3) or analogue stirring in 10 mL CH2Cl2 was added Boc-l-glutamic acid-α-tert-butyl-ester or alternative coupling compound (1 equiv) and HATU (1.05 equiv). Triethylamine (6 equiv) was added dropwise, and the reaction was left stirring at room temperature overnight (16–18 h). The reaction mixture was transferred to a separatory funnel and washed with saturated NH4Cl (2 × 10 mL), water (10 mL), and brine (10 mL). The organic layer was dried over MgSO4, filtered, concentrated in vacuo, and purified via silica flash chromatography (2:1 hexanes/EtOAc).

General Reaction C. Methyl Ester Deprotection.

To approximately 0.035 g of Boc-l-Glu-OtBu-l-Pra-OMe (4) or similarly protected analogue in 2.5 mL of tetrahydrofuran (THF) at 0 °C was added an aqueous solution of 1 N LiOH at once. The reaction was brought to room temperature after 15 min and stirred for an additional 2 h until the starting material was consumed. The reaction was concentrated in vacuo to remove THF. The LiOH was neutralized by the addition of aqueous 1 N HCl to a pH of 4. Organic compounds were extracted with EtOAc (3 × 10 mL). The organic layer was dried over MgSO4, filtered, concentrated in vacuo, and immediately used in the next reaction (general reaction D) without further purification.

General Reaction D. Boc- and/or O-tert-Butyl Ester Deprotections.

The product of reaction C was dissolved in 5 mL of CH2Cl2, and 1 mL of trifluoroacetic acid (TFA) was added at once. The reaction was stirred for 2 h at room temperature. The mixture was concentrated in vacuo, redissolved in minimal water, and lyophilized to afford the product without further purification. Compound characterization details for each isolated intermediate and final product are outlined in the Supporting Information.

Molecular Biology and Biochemical Methods

A pET28a(+) plasmid containing an E. coli codon-optimized N-terminal hexahistidine (His6) tagged besE gene from S. cattleya (Twist Biosciences) was transformed into both E. coli DH10β and BL21(DE3) chemically competent cell lines for plasmid production and protein expression, respectively. Transformation was performed via heat shock according to the following protocol: 1 μL of plasmid was added to 100 μL chemically competent cells, and the mixture was incubated on ice for 30 min. The cells were then heated to 42 °C for 55 s and placed on ice again for 5 min; 700 μL of LB medium was added in the tube, and the cells were incubated for 45 min at 37 °C and 200 rpm of agitation. After this step, 300 μL of the recovered cells were plated on LB agar plates supplemented with kanamycin (50 μg/mL). The plates were incubated at 37 °C overnight. For plasmid purification, a single colony was used to inoculate 5 mL of LB supplemented with kanamycin (50 μg/mL) and grown overnight at 37 °C and 200 rpm of agitation. Plasmids were purified following the protocol of Plasmid DNA Purification QIAprep Spin Miniprep Kit (QIAGEN). Plasmid concentrations were measured by NanoDrop UV–vis spectrophotometry and stored at −20 °C.

A single colony of BesE transformed BL21(DE3) cells was used to inoculate 20 mL of LB medium supplemented with kanamycin (50 μg/mL). The culture was grown overnight at 37 °C and 200 rpm. 10 mL of the overnight culture was then added to 1 L of Terrific Broth supplemented with kanamycin (50 μg/mL). The 1 L culture was grown in a shaking incubator at 37 °C and 200 rpm until the OD600 reached ~0.6. The flasks were then cooled to 18 °C for 1 h, at which point 0.1 mM IPTG was added to induce protein expression. The cultures were left to grow at 18 °C and 200 rpm overnight (16–18 h) and pelleted in a centrifuge at 3000g for 30 min at 4 °C. The media supernatant was discarded, and the cell pellet was resuspended in 40 mL of cold buffer A (1 M NaCl, 20 mM Tris–HCl pH 8.0) and immediately purified or stored at −80 °C.

The BesE-resuspended cell pellet was thawed and lysed on ice via sonication at 40% amplitude for 10 cycles of 15 s on and 45 s off (1 min total time per cycle). The lysate was pelleted via centrifugation at 4 °C and 16,000g for 30 min. The cleared lysate was loaded onto a 5 mL HisTrap FF column (GE Healthcare Life Sciences) that was pre-equilibrated with 5 CV of buffer A. After loading, the column was washed with buffer A until the UV absorbance reached <50 mAU. The column was then washed with 10% buffer B (1 M NaCl, 20 mM Tris–HCl pH 8.0, 250 mM imidazole) for 5 CV to remove any nonspecifically bound proteins. BesE was then eluted using a linear gradient of 100% buffer A to 100% buffer B over 60 mL, and 5 mL fractions were collected. The flow rate through the column was 2 mL/min. Fractions were assessed for the presence of protein and purity using SDS-PAGE. Fractions containing BesE had 2 mM EDTA added and were pooled and concentrated by Amicon ultra centrifugal filters (10 kDa molecular weight cutoff) to a volume less than 2.5 mL. The concentrated protein was then further purified via a PD-10 desalting column (Cytiva) pre-equilibrated with GF buffer (50 mM HEPES-KOH pH 8.0, 300 mM KCl) following the default gravity filtration protocol. Enzyme concentration was determined via Bradford assay, and pure enzyme was either used immediately for assays or aliquoted and stored at −70 °C.

Site-directed mutagenesis PCR of the besE plasmid was performed using the Q5 Site-Directed Mutagenesis Kit (New England Biolabs) following the manufacturer’s protocol and using the primers listed in Table S1. The concentration of the besE plasmid used as the template was 1 ng/μL. The general PCR protocol for each primer set was as follows: initial denaturation 98 °C for 30 s, then 30 cycles of the following: denaturation at 98 °C for 10 s, annealing at the 3–5 °C less than the Ta for 10 s (where Ta is the annealing temperature determined for each primer pair), and extension at 72 °C for 2 min, lastly, a final extension at 72 °C for 2 min followed by a hold at 4 °C. Upon completion of the PCR, a KLD reaction was performed to linearize the resulting plasmid containing the mutant gene and digest residual wild-type besE plasmid. The 10 μL reaction was prepared using KLD Enzyme Mix and KLD Reaction Buffer (New England Biolabs) and followed the manufacturer’s KLD Enzyme Mix reaction protocol. The reaction was left to incubate at room temperature for 5 min. The entirety of the reaction mixture was then added to a 100 μL aliquot of E. coli DH10β chemically competent cells and transformed as previously described. A single colony was used to generate a 5 mL inoculum from which plasmid was purified as described previously. Successfully generated mutants were confirmed via Sanger sequencing (Azenta Life Sciences) and transformed into E. coli BL21(DE3) chemically competent cells as previously described.

BesE mutant expression was carried out in a way identical to that of wild-type BesE without modification. Resuspended cell pellets in buffer A were immediately purified or stored at −80 °C. Each individual resuspended BesE mutant pellet was lysed via sonication and centrifuged as previously described for the wild-type BesE. The clarified supernatant was then loaded onto an Econo-Column Chromatography column (2.5 × 10 cm; Bio-Rad Laboratories) containing 5 mL of nickel-NTA batch resin that was pre-equilibrated with 3–5 CV of buffer A (20 mM Tris 1 M NaCl pH 8.0). After loading, the resin was washed with an additional 5 CV buffer A. The protein was then eluted with 5 CV 20% buffer B followed by 5 CV 50% buffer B, followed by 5 CV 100% buffer B (20 mM Tris 1 M NaCl 250 mM imidazole pH 8.0). All fractions were assessed for purity using a 10% SDS-PAGE gel (Figure S3). The 100% B fraction containing BesE had 2 mM EDTA added to remove any bound iron and was concentrated and further purified via the PD-10 desalting column as previously described for wild-type BesE. Enzyme concentration was determined via Bradford assay, and pure mutant enzyme was aliquoted and stored at −80 °C.

Analytical BesE Assay Methods

Analytical BesE enzyme assays were conducted in 50 mM potassium phosphate (KPi) buffer (pH 8.0) with 10 mM l-ascorbate, 5 mM α-ketoglutarate, and 1 mM FeSO4 with 1 mM of substrate 1 and 50 μM of purified BesE. Enzyme followed by FeSO4 were the last additions to the assay, and the reaction was allowed to run for 3 h. The total volume for the reaction was 100 μL. The reaction was then either quenched with 0.1 mM chloramphenicol in methanol (1 equiv) or further derivatized via Marfey’s reagent or Cu-catalyzed azide–alkyne cycloaddition (CuAAC) with 7-azido-4-methylcoumarin. Quenched and/or derivatized reactions were centrifuged at 13,000g for 5 min, and the supernatant was then subjected to analysis by UPLC–MS. For analytical assays involving substrate analogues and BesE mutants, the same procedure was followed, except 0.5 mM of substrate and 20 μM of purified enzyme were used.

Marfey’s derivatization procedure: to 50 μL of enzyme assay reaction mixture were added 20 μL of saturated sodium bicarbonate and 100 μL of 1% w/v Marfey’s reagent in acetone. The reaction was then incubated at 37 °C for 90 min followed by quenching with 30 μL of 1 N HCl. The sample was then centrifuged at 13,000g for 10 min, and the supernatant was subjected to analysis by UPLC–MS.

CuAAC derivatization procedure: to 25 μL of the enzyme assay reaction mixture were added the following: 1.5 mM 7-azido-4-methylcoumarin, 0.2 mM CuSO4, and 0.6 mM sodium ascorbate. The total reaction volume was then brought to 50 μL with Milli-Q water and the mixture was left for incubation at room temperature for 1 h. The sample was then centrifuged for 5 min at 13,000g, and the supernatant was subjected to analysis by UPLC–MS.

Supplementary Material

2026-JNP-BesE-McKinnie-SI
2026-JNP-Raw NMR data files

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jnatprod.5c01272.

General materials and methods, supplementary figures, chemical synthesis, tables, and NMR spectra of characterized compounds (PDF)

Raw NMR data of compounds 1, 3–18 and intermediates SI-1-SI–16 (ZIP)

ACKNOWLEDGMENTS

This work was supported by the University of California, Santa Cruz Physical and Biological Sciences division (CURE lab start up funding) and the National Institutes of Health (R21-GM-148870, R35-GM-147235) for general McKinnie lab work in the field of nonheme iron enzymology and other biosynthetic metalloenzymes, respectively. We gratefully acknowledge S. Rubin, F. Pavlovici, P. Ngoi, and M. Membreño for additional biochemical support and supervision, H.-W. Lee for maintenance of nuclear magnetic resonance spectroscopy facilities, and R. Dunkin for support conceptualizing CURE lab assessments and pedagogy (all University of California, Santa Cruz). The authors also acknowledge the National Science Foundation Division of Undergraduate Education (DUE 2150444) for supporting general CURE lab research at the University of California Santa Cruz.

Footnotes

The authors declare no competing financial interest.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

2026-JNP-BesE-McKinnie-SI
NIHMS2164910-supplement-2026-JNP-BesE-McKinnie-SI.pdf (10.7MB, pdf)
2026-JNP-Raw NMR data files
NIHMS2164910-supplement-2026-JNP-Raw_NMR_data_files.zip (13.2MB, zip)

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