Riboflavin Biosynthesis: Mapping the Tagged Lines of a Complex Metabolic Design

Abstract

Nearly 150 years ago, riboflavin was first isolated as the water-soluble, yellow-green, fluorescent “lactochrome,” a discovery that added the earliest bright stroke to what would become a complex metabolic mural. The ribityl-isoalloxazine structure was fully defined nearly a century ago, and from the 1970s through the 1990s, a series of elegant biochemical studies began tracing the lines of biosynthesis with increasing definition, including isotopic labeling experiments (¹³C, ¹⁵N) that effectively tagged substrate atoms to follow their movement through the pathway. In more recent years, presteady state analyses and high-resolution crystal structures have brought sharper focus to the proposed enzymatic mechanisms, adding depth and shading to the emerging picture. The purpose of this article is to define the current state of our understanding of the enzymatic mechanisms that compose the riboflavin biosynthetic pathway, bringing the latest evidence together as an updated rendering of this intricate biochemical (master)­piece.

1. Introduction

Riboflavin, or vitamin B2, is the universal precursor to cofactors FAD (flavin adenine dinucleotide) and FMN (flavin mononucleotide) which are essential for critical cellular processes such as DNA repair, metabolism, cellular respiration, redox homeostasis, and bioluminescence. If riboflavin intake is insufficient in humans, ariboflavinosis develops. Ariboflavinosis causes neuropathy, vision distortions, and sores of the lips, mouth, and throat. , As such, ingestion of riboflavin rich foods such as leafy greens, eggs, lean meats, milk or supplements containing riboflavin is necessary. Just as riboflavin is required for human health, agriculture relies on the production of riboflavin to be used as a dietary supplement for animal feeds. − Industrial production of riboflavin is accomplished using microorganisms such as Ashbya gossypii, Bacillus subtilis, and Candida spp. to biosynthetically generate riboflavin using specialized riboflavin biosynthetic enzymes. ,

Riboflavin biosynthetic enzymes are found exclusively in micro-organisms, fungi, and plants. Riboflavin biosynthesis requires one molecule of GTP, one molecule of NADPH, and two molecules of ribulose 5-phosphate to produce one molecule of riboflavin (Figure ). The riboflavin biosynthetic pathway is composed of two branches that converge at the penultimate step of riboflavin production. There is significant disparity in naming conventions of the enzymes across different organisms, so we will use the Escherichia coli naming conventions throughout for clarity. Using the gene names from E. coli, RibA is GTP cyclohydrolase II and the first enzyme of the left arm of the convergent pathway. RibA catalyzes a deformylation that excises the bridging methyne (C8) of the imidazole heterocycle of the GTP guanosine while also first hydrolyzing pyrophosphate and then converting the pyrophosphate to two molecules of inorganic phosphate. The product of the RibA reaction is DARP or 2,5-diamino-6-β-pyrimidinone 5′-phosphate. RibD transforms DARP by an aminohydrolase reaction on the pyrimidine 2-amino and reduction of the ribose to form a ribityl, generating ARAPDP or 5-amino-6-ribitylamino-pyrimidinedione 5′-phosphate. ARAPDP is dephosporylated to form ARAPD (5-amino-6-ribitylamino-pyrimidinedione). Interestingly, some bacteria use ARAPD to generate deazaflavin, a precursor for the coenzyme F420 that is important for methanogenesis and antibiotic biosynthesis. RibB is the initiating enzyme of the right arm and catalyzes a deformylation of ribulose-5-phosphate that eliminates the (internal) 4-carbon as formate to form 3,4-dihydroxy-2-butanone 4-phosphate (DHBP). The two arms converge at the enzymatic step catalyzed by RibE or lumazine synthase, in which ARAPD (product of RibA, RibD and the phosphatase) and DHBP (product of RibB) condense and cyclize to form a pyrazine that will be the center ring of the isoalloxazine ring system of riboflavin. The final step is the formation of the xylene ring, catalyzed by riboflavin synthase (RibC). RibC performs a disproportionation reaction of two lumazine molecules to generate one riboflavin and regenerate one ARAPD that can be recycled for the lumazine synthase reaction.

1.

The biosynthesis of riboflavin from GTP and ribulose 5-phosphate (Ru5P) and subsequent production of flavin mononucleotide and flavin adenine dinucleotide from riboflavin. The enzymes are named according to the E. coli naming convention, RibA-E. DARP = 2,5-diamino-6-ribosylamino-pyrimidinone 5′-phosphate; ARAPDP = 5-amino-6-ribitylamino-pyrimidinedione 5′-phosphate; ARAPD = 5-amino-6-ribitylamino-pyrimidinedione; DHBP = 3,4-dihydroxy-2-butanone 4-phosphate; LUM = 6,7-dimethyl-8-ribityllumazine.

The enzymes of the riboflavin biosynthetic pathway are notoriously slow, with turnover numbers more appropriately reported in per minute, as opposed to hundreds to thousands per second for enzymes of primary metabolism. Indeed, the steady state kinetics of RibA, RibB, RibC, and RibE are determined to have k _cat values of 0.06, 0.037, 0.056, 0.024 s^–1, respectively. The kinetic parameters for the bifunctional RibD reactions, both reduction and deamination are reported per hour. That means, with one copy of each enzyme, the fastest a molecule of riboflavin is generated is about 1 every 2 min, ignoring caveats such as diffusion between steps, which lengthen the time. The copy number estimates for these proteins in E. coli range from ∼275 (RibD) to ∼2700 (RibE). If we multiply the turnover numbers by the copy numbers to adjust our estimate, then we would calculate that an E. coli generates 10 riboflavin molecules per second, or 12,000 riboflavin per doubling time (20 min). This is consistent with earlier estimates. However, E. coli copy number estimates for important flavin containing enzymes, such as lipoamide dehydrogenase of the pyruvate dehydrogenase complex (∼9500) and succinate dehydrogenase of the citric acid cycle and the electron transport chain (∼1600) would indicate that the amounts of riboflavin that the biosynthetic enzymes can produce is insufficient for cellular metabolism. This is compounded by data indicating that flavins are produced by E. coli in excess but not stored, with more excreted than retained. So, the purified enzymes of riboflavin biosynthesis have kinetic parameters that are significantly slower than the same enzymes in vivo.

Whereas much is documented about the individual enzymes, how they come together to efficiently generate riboflavin to support life has not yet been determined. This has broader implications for the generation of riboflavin as a food supplement. In addition, riboflavin has long been considered an ideal antimicrobial target as many bacterial pathogens rely on endogenously synthesized riboflavin because they lack efficient riboflavin uptake machinery , However, despite dedicated effort to generate inhibitors to the riboflavin biosynthetic enzymes, , no antibiotics have been developed that target this pathway. Considering the large gap in fundamental knowledge as to efficient riboflavin formation and how the translation of such knowledge would benefit human health, there is still a lot to be learned. We provide here a summary of the current field and highlight exciting avenues of future research.

2. Genomic Organization and Regulation

The majority of bacteria synthesize riboflavin de novo, with a few notable exceptions including spirochetes, which have riboflavin uptake machinery. The five committed enzymes of riboflavin biosynthesis are organized in bacterial genomes in a variety of ways. All five can be clustered in operons, as is the case with B. subtilis. , In contrast, E. coli and Aquifex aeolicus provide examples of organisms whose riboflavin biosynthetic genes are scattered throughout their genomes. , Interestingly, an additional enzyme, and ARAPDP phosphatase, is also required to produce riboflavin, − but the identity and specificity of this phosphatase varies between organisms.

Some organisms possess only one copy of each of the riboflavin biosynthetic genes, while others may possess multiple copies. Two examples of redundant riboflavin biosynthetic genes are RibA of Streptomyces coelicolor and lumazine synthase of Burkholderia mayonis. In the case of S. coelicolor, three copies of genes with high sequence similarity to RibA were identified with two of the three demonstrating GTP cyclohydrolase II activity and the third exhibited a completely new function that is unrelated to flavin biosynthesis. Meanwhile, the redundant copy of lumazine synthase identified in B. mayonis was found clustered with genes encoding enzymes to synthesize an antimicrobial phosphonate. Interestingly, increased expression of lumazine synthase, regardless of organism of origin, conferred resistance to this phosphonate, suggesting that the extra lumazine synthase gene was expressed as a means of resistance against the toxic phosphonate.

All the required riboflavin biosynthetic enzymes of B. subtilis are located in a single operon (Figure ) with the exception of the ARAPDP phosphatase. While some organisms possess two separate polypeptides to catalyze the first committed steps of each of the two independent branches, some organisms like B. subtilis and A. aeolicus contain a single ribBA gene that encodes for a bifunctional enzyme with GTP cyclohydrolase II activity and DHBP synthase activity. , Notably, there is an additional gene, annotated ribT, which encodes for an N-acetyltransferase. RibT homologues have been observed in multiple organisms including E. coli. The role of RibT in riboflavin biosynthesis is a new avenue for exploration. In B. subtilis, RibT, which acetylates lysine 29 of lumazine synthase, is proposed to play a novel regulatory role in modulating enzyme activity.

2.

B. subtilis riboflavin biosynthetic operon. Genes are labeled using the E. coli naming convention. Created in BioRender. Hoffpauir, Z. (2026) https://BioRender.com/0t9i7at.

Riboflavin biosynthesis is regulated at the transcriptional and translational levels, but the enzymes of riboflavin biosynthesis are not directly regulated by increased cellular riboflavin concentrations. ,, At the 5′ end of the B. subtilis riboflavin operon, located between the promoter and the coding region, there is a noncoding region dubbed ribO (Figure ). After transcription to mRNA, this region of the mRNA strand adopts a fold that binds to FMN and FAD, called a riboswitch, which attenuates the expression of the riboflavin biosynthetic enzymes. Indeed, the antimicrobial pigment roseoflavin targets the riboswitch to inhibit riboflavin biosynthesis. The riboswitch regulatory control mechanism explains observations that increased riboflavin production is achieved by inactivating riboflavin kinase, which generates FMN, and the bifunctional riboflavin kinase/FAD synthetase, or by introducing mutations to this ribO regulatory region. In E. coli, which lacks a riboflavin biosynthetic operon, an FMN riboswitch has been identified upstream of the RibB gene that regulates expression of RibB.

3. RibA (GTP Cyclohydrolase II) – GTP → DARP

The left of the two branches of the riboflavin biosynthetic pathway begins with the conversion of GTP to 2,5-diamino-6-ribosylamino-4­(3H)-pyrimidinedione phosphate (DARP), releasing formate and two inorganic phosphate ions as byproducts of the reaction (Figure ) by GTP cyclohydrolase II (GCHII), often referred to by the gene name RibA.

Early studies recognized the mechanistic similarity between RibA and GTP cyclohydrolase I (GCHI). GCHI catalyzes the first committed step of tetrahydrofolate biosynthesis, , an enzyme for which ample structural analyses had been completed. − Figure shows a side-by-side comparison of the GCHI and GCHII reactions and the dissimilar protein architecture. , The homodecameric GCHI has D ₅ symmetry that builds a β-barrel of 20 antiparallel strands making a tunnel, which is surrounded α-helical segments. The last α-helix from each monomer lines the tunnel (Figure A). The active site of GCHI contains one zinc per subunit, chelated by three cysteine residues, to facilitate the guanine ring opening. , Despite a ring opening step, the ultimate result of the reaction is a ring expansion upon closure of the ring (Figure D). The 21.8 kDa structure of RibA is composed of an α/β fold with six α-helices surrounding the central seven β-strands that form a mixed parallel and antiparallel β-sheet (Figure B). Two catalytically required divalent metal cations reside in the active site, a zinc and a magnesium. A structure with the nonhydrolyzable GTP analogue GMPPNP shows the alignment of the substrate such that the hydrolytic cleavage of the base releasing formate occurs at the zinc site, whereas pyrophosphate cleavage occurs at the magnesium. An early mechanism proposed that the enzyme formed a covalent bond between an arginine residue and the α-phosphorus of the substrate because of the proximity of the arginine side chain (Figure A), ,− but recent transient state investigations found no evidence to support the formation of the adduct and an alternate mechanism is proposed (Figure B).

3.

Comparison of GTP cyclohydroase I and II. (A) GTP cyclohydrolase I decamer. (B) GTP cyclohydroase II (RibA) dimer. (C) Overlay of GCHI and GCHII monomers. (D) Deformylation reactions catalyzed by GCHI and GCHII.

4.

Proposed mechanisms for RibA. (A) Early mechanism dependent on an arginine residue that covalently displaces pyrophosphate, generating a covalent intermediate (adapted from ref ). (B) Current mechanism in which no covalent intermediate is observed, and the initial pyrophosphate is hydrolyzed to two phosphate ions during a single turnover (adapted from ref ).

Initially, the coproduct of RibA was reported to be pyrophosphate. , However, recent work using ³¹P NMR and transient state kinetics provides evidence that rather than producing pyrophosphate, RibA has catalytically relevant pyrophosphatase activity that yields inorganic phosphate in the time of a single turnover. Smith et al. showed that RibA adopts an inactive, relaxed conformation in solution and that binding of substrate stimulates a conformational change to the active conformation. After the cleavage of the α-β phosphoanhydride bond (the rate limiting step of the reaction , ), approximately 90% of the GMP intermediate undergoes the deformylation at the Zn site with concomitant cleavage of the pyrophosphate to inorganic phosphate at the Mg site. However, 7–10% of the GMP and pyrophosphate dissociate from the RibA active site. , The formation of GMP as an alternate product is consistent with earlier reports. ,, The small amount of pyrophosphate that dissociates is proposed to rebind and hydrolyze in a separate, slow step and may aid in stabilizing the active conformation of RibA in vivo along with the millimolar levels of magnesium within the cell. ,

RibA is an incredibly slow enzyme, with a reported turnover number of 0.06 s^–1 , or around 1 turnover every 17 s. Cell-based studies report that riboflavin production is limited by GTP , suggesting that riboflavin biosynthesis is dictated by flux through RibA, and a mathematical kinetic model supports that RibA is a rate-limiting step of riboflavin biosynthesis.

4. RibB – Ru5P → DHBP

The right branch of the riboflavin biosynthetic pathway (Figure ) is initiated by RibB, which catalyzes the deformylation of d-ribulose 5-phosphate (Ru5P) to afford 3,4-dihydroxy-2-butantone 4-phosphate (DHBP). RibB is a homodimeric enzyme consisting of two ∼23.5 kDa monomers. The protein topology of RibB is quite unique although the foundation is a standard α + β structure in which α-helices pack against a central β-sheet. Two highly flexible loops interact with bound substrate and catalytically required metal (Figure A). RibB is considered to be one of the rate-limiting steps of riboflavin biosynthesis with reported turnover rates reported on the order of 6 min^–1 (0.1 s^–1) to 1 turnover every 5 min (0.003 s^–1) depending on source organism and conditions. ,

5.

RibB or 3,4-dihydroxy-2-butantone 4-phosphate (DHBP) synthase. (A) Structure of the V. cholerae RibB dimer (PDB: 7UF1). Purple spheres are manganese atoms bound to the active sites. (B) RibB chemical mechanism (adapted from ref .).

Studies using ¹³C labeled compounds demonstrated RibB accomplishes the deformylation reaction by eliminating carbon number 4 of the 5 carbon Ru5P molecule. − The chemical mechanism is shown in Figure B. In brief, the reaction begins with tautomerization to form a 2,3-endiol, dehydration of the 1-carbon to yield the 2,3-diketone, followed by a skeletal rearrangement to the branched intermediate and the ultimate elimination of formate to yield the DHBP product. This mechanism, proposed in 1991, was later supported by acid quenched single turnover reactions monitored by ¹³C NMR in 2022.

Early crystal structures indicated that two metals were bound to the active site and facilitated catalysis, , so the proposed mechanisms reflected the presence of two metals bound and interacting with the substrate. , Many of the crystal structures reported in the PDB did not represent active states of RibB, because they contain noncatalytic metals such as Zn²⁺, and Ca²⁺ in place of the catalytic Mg²⁺ or Mn²⁺. − The more recent work robustly demonstrated that only one metal is required per active site using a combination of activity assays, fluorescence titration, electron paramagnetic resonance (EPR), and time-resolved X-ray crystallography that captured RibB midturnover bound to intermediates of the deformylation reaction.

5. RibD – DARP → ARAPDP

Most bacteria possess a bifunctional RibD that catalyzes the consecutive deamination and reduction of DARP to 5-amino-6-ribitylamino-pyrimidinedione 5′-phosphate (ARAPDP) (Figure ). However, some archaea and fungi use two separate polypeptides to catalyze these steps independently. Interestingly, the order of the two reactions varies among organisms with reduction being reported to precede deamination in yeast, whereas deamination is the initial step in bacteria. The structure of the bifunctional B. subtilis RibD reveals the N-terminal deaminase domain and a C-terminal reductase domain that can be individually expressed and purified to yield active enzymes. The quaternary structure of RibD from B. subtilis is composed of 40.3 kDa monomers that form a tetramer, whereas the E. coli RibD is dimeric (Figure ).

6.

Structural comparison for RibD. (A) B. subtilis RibD monomer shown in mint green (PDB: 4G3M). (B) B. subtilis RibD (mint green) forms a tetramer with the tetramer interface on the deaminase domains, whereasE. coli RibD (PDB: 2G6V), shown in gray, forms a dimer.

The RibD deaminase domain of E. coli has a high degree of similarity to members of the cytidine deaminase (CDA) superfamily, which hydrolytically deaminate nucleotides with a zinc-activated water molecule. , The proposed mechanism of the E. coli RibD deamination reaction is based upon the CDA superfamily’s mechanism in which the zinc-activated water molecule attacks the C-2 carbon to produce a tetrahedral intermediate that collapses to release ammonia and produce the pyrimidinone product (Figure A). However, providing evidence to support this proposed mechanism has been challenging, because substrate analogues derived from inosine triphosphate without the exocyclic amine group that is deaminated show no binding.

7.

Proposed deamination and reduction mechanisms for RibD. (A) The proposed RibD deamination mechanism in which a Zn activated water attacks C2 to produce a tetrahedral intermediate that collapses to release ammonium. (B) Several mechanisms of RibD reduction have been proposed. The initially proposed mechanism involved an Amadori rearrangement, but recent studies indicate that RibD likely catalyzes the reduction via a Schiff base or by direct transfer of the hydride from the NADPH coenzyme (adapted from ref ).

In 2008, there were 3 proposed mechanisms for the reduction reaction: Amadori pathway, Schiff base pathway, and direct transfer pathway (Figure B). The Amadori pathway was initially proposed due to in vivo studies using a riboflavin-requiring mutant of B. subtilis in which the accumulation of an Amadori derivative was observed. Interestingly, these results were not reproduced using the enzyme from the filamentous fungiA. gossypii. While the reactions catalyzed by RibD occur on structural distinct areas of the molecule, it is important to note that the order of reduction and deamination could impact mechanism, and whether the eubacterial and fungal RibD enzymes are mechanistically similar is yet to be established. Regardless, the work in A. gossypii supports the formation of a Schiff base intermediate, suggesting the furanose ring opening is due to the removal the proton from the nitrogen atom adjacent to the C-1′ of the sugar followed by reduction of the resulting Schiff base (Figure B). Finally and most recently, a direct transfer mechanism was proposed; however, discrimination between the Schiff base and direct transfer pathways has been difficult, and, as of this review, there are still many exciting knowledge gaps surrounding the mechanism of RibD, for both the deamination and reductase reactions.

6. RibZ – ARAPDP Dephosphorylation

After the synthesis of ARAPDP, the phosphate group on the ribityl tail must be removed (Figure ). Lumazine synthase can use ARAPDP as a substrate to condense with DHBP to form lumazine 5-phosphate, albeit with much lower efficiency than the ARAPD reaction. However, lumazine 5-phosphate is not a substrate for riboflavin synthase to catalyze the formation of riboflavin or riboflavin 5-phosphate (FMN). Therefore, an ARAPDP phosphatase is an essential component of riboflavin biosynthesis.

Despite the other steps of the riboflavin biosynthetic pathway being catalyzed by a dedicated riboflavin biosynthetic enzyme, there is limited evidence that there is a dedicated phosphatase to dephosphorylate ARAPDP. Indeed, ARAPDP phosphatases have been identified in E. coli, B. subtilis, Bacteroides thetaiotaomicron, and Arabidopsis thaliana. In each case, haloacid dehalogenase (HAD) family phosphatases were identified to perform this essential chemistry, but the HAD superfamily of phosphatases is notoriously nonspecific (Figure ). Indeed, deletion of these phosphatases from the genomes of these organisms does not prevent the organism from synthesizing riboflavin, suggesting there are redundant phosphatases available to dephosphorylate ARAPDP. Further, 30% of HAD phosphatases can dephosphorylate FMN to yield riboflavin, demonstrating how numerous members of this family could nonspecifically dephosphorylate compounds like FMN and intermediates of the riboflavin biosynthetic pathway.

8.

Enzymes that perform the required dephosphorylation step. Phosphatases identified to perform the dephosphorylation of ARAPDP comprise 3 distinct families of phosphatases. (A) The histidine family phosphatase from A. aeolicus (PDB: 9MMH). (B) The A. aeolicus myo-inositol monophosphatase (PDB: 9MMI). (C) A representative haloacid dehalogenase from Pseudomonas syringae (PDB: 3VAY) that has high sequence similarity to the ARAPDP phosphatase of E. coli, YigB.

Only in A. aeolicus have phosphatases that are not part of the HAD superfamily been identified to dephosphorylate ARAPDP: a histidine family phosphatase (AaHFP) and an inositol-monophosphatase (AaIMP) (Figure ). Although both AaHFP and AaIMP dephosphorylate ARAPDP, only AaHFP did not indiscriminately dephosphorylate the substrates of the pathway, Ru5P and NADPH. Indeed, only AaHFP could facilitate riboflavin formation when combined with the substrates and other enzymes of the pathway.

7. Lumazine Synthase – ARAPD + DHBP → LUM

RibE, or lumazine synthase, catalyzes the condensation of 5-amino-6-ribitylamino-pyrimidinedione (ARAPD) with 3,4-dihydroxy-2-butanone 4-phosphate (DHBP) to form 6,7-dimethyl-8-ribityllumazine (lumazine). In the proposed chemical mechanism (Figure ), the carbonyl group of DHBP forms a Schiff base with the 5-amino group of ARAPD. The resulting Schiff base intermediate is dephosphorylated and tautomerizes to allow for an intramolecular condensation ring closure that yields the lumazine product. RibE catalyzes the condensation of the 5′-phosphorylated 5-amino-6-ribitylamino-pyrimidinedione (ARAPDP) to DHBP to form 6,7-dimethyl-8-ribityllumazine 5′-phosphate at 1/100th the rate of the physiological ARAPD-DHBP condensation; however, riboflavin synthase (RibC) does not recognize 6,7-dimethyl-8-ribityllumazine 5-phosphate as a substrate to form FMN (riboflavin 5-phosphate).

9.

Proposed chemical mechanism of RibE. Substrates are proposed to form a Schiff base prior to elimination of the orthophosphate group. Keto–enol tautomerization precedes the intramolecular condensation reaction to close the ring to generate LUM (adapted from ref ).

One of the most fascinating characteristics of RibE is the quaternary structure. The approximately 17 kDa monomers assemble into pentamers with the active site located between adjacent subunits of the pentamer. Interestingly, RibE pentamers from some organisms spontaneously assembles into T = 1 icosahedral capsids with an external diameter of 154 Å and an inner diameter of 80 Å. The stability and dimensions of these protein capsids are dependent on buffer conditions. Larger capsids are observed by cryogenic electron microscopy under certain conditions, such as changes in buffer or variant forms of the enzyme. Crystal structures of a T = 1 icosahedral capsid from A. aeolicus, B. subtilus, and spinach have been reported, ,, and EM has additionally been used to determine the structure of A. aeolicus RibE.

Much of the current knowledge of capsids comes from the study of viral capsids and mathematical modeling. Icosahedral capsids are comprised of 12 pentamers separated by a variable number of hexamers, and the number of hexamers increases with the triangulation number, T. The previously reported RibE capsid structures are T = 1 icosahedral capsids, with 12 pentamers. The next larger capsid composed exclusively of pentamers, no hexamers, as would be expected for the larger RibE capsids, is the T = 7d capsid conformation. Examples of biological T = 7d capsids include papillomaviruses and polyomaviruses, − which are made exclusively of pentamers with gaps between some of the adjacent pentamers to allow for formation of the triangulation facet (Figure ). These large gaps in the putative T = 7d RibE capsid may provide the answer to the long-standing debate about substrate entry and product egress from the capsid. The pores of the T = 1 RibE capsid have not been considered large enough. Therefore, a catalytically efficient RibE may form T = 7d capsids. The much larger capsids were observed when the charges of RibE were altered or internal cargo molecules were used as a scaffold for higher order oligomerization. , The Hilvert group has previously reviewed efforts to utilize the RibE enzyme as a platform for nanotechnology development exploiting RibE’s malleable scaffold.

10.

Relative size of RibE oligomers. The largest structure of RibE is from a variant of A. aeolicus (PDB: 5MQ7), which forms a T = 7d capsid (left). A. aeolicus typically forms a T = 1 capsid (PDB: 7X7M, middle). Finally, some forms of RibE purify as pentamers, for example the Mycobacterium tuberculosis pentamer (PDB: 2VI5, right).

Some RibE homologues purify as pentamers, including the clinically relevant lumazine synthase of Mycobacterium tuberculosis. , The observation of some RibE existing as stable pentamers rather than capsids could be an artifact of purification or storage conditions, but sequence analysis has been employed to illuminate differences between the pentameric and icosahedral RibE homologues. Indeed, the stability of the capsids may play a crucial role in facilitating enzymatic activity, and post-translational modification may result in altered capsid formation.

The genomes of many microorganisms, including B. subtilis, encode a riboflavin biosynthetic operon that includes an additional gene, annotated as ribT, which encodes an N-acetyl transferase (RibT) that is absent in organisms such as A. aeolicus. RibT homologues have been identified in organisms that lack a riboflavin biosynthetic operon, such as E. coli. While deletion of ribT does not render an organism unable to produce riboflavin, the absence of ribT leads to decreased riboflavin production. , RibT is a structural homologue of N-acetyl transferases that catalyze the transfer of an acetyl group from acetyl coenzyme A to a variety of substrates (proteins or small molecules). While many bacterial enzymes are acetylated nonenzymatically due to high cellular concentrations of acetyl coenzyme A or acetyl phosphates, enzymatic protein acetylation is a posttranslational modification that can provide functional heterogeneity by modulating the activity. One target of B. subtilis RibT is lumazine synthase on the lysine at position 29. K29 is located at the interface of three lumazine synthase pentamers as they assemble into a T = 1 icosahedral capsid (Figure ). Rotter et al. hypothesize that acetylation at this position would destabilize the capsid and promote pentamer formation in vivo and that pentamers would have a higher turnover rate because the pentameric structure would favor product release rather than lumazine being trapped inside the capsid. An alternate hypothesis is that acetylation at K29 leads to the transition from T = 1 capsids to T = 7d capsids. The larger capsid with the open triangulation facet may provide an alternative pathway for generating expedited product release. The impact of RibT acetylation of lumazine synthase should be the subject of future investigations.

11.

Acetylation of lumazine synthase. (A) RibT is an N-acetyl transferase found across multiple species of bacteria (PDB: 5XXS) that is reported to acetylate RibE. (B) The site of acetylation on B. subtilis RibE capsid (PDB: 1RVV) is K29 (red), which is hypothesized to destabilize the capsid and promote the formation of pentamers.

8. Riboflavin Synthase – 2 LUM → Riboflavin + ARAPD

The final enzyme of the riboflavin biosynthetic pathway, riboflavin synthase, is called RibC. RibC catalyzes a dismutation reaction that results in the removal of the 4 carbons of one lumazine molecule and covalently attaches these carbons to a second lumazine molecule to form one molecule of riboflavin and regenerate one molecule of ARAPD that can be recycled by lumazine synthase to condense with a new molecule of DHBP to produce lumazine (Figure ). The 4 carbon atoms removed from the initial lumazine molecule were those provided by DHBP, meaning that all the carbons that form the third (xylene) ring of riboflavin were originally from DHBP, and are joined with a head-to-tail orientation. ,

Despite having defined how the carbons of the two lumazine molecules are arranged in the products riboflavin and ARAPD, the mechanism by which carbons are linked in the RibC dismutation reaction is currently an ongoing area of investigation. Interestingly, the RibC reaction will proceed with good yields in the absence of the enzyme under diverse conditions including when lumazine is refluxed under nitrogen in phosphate buffer at pH 7.3. , The proposed mechanism of RibC (Figure ) is that two lumazine molecules, an acceptor and donor of the 4 carbons to be transferred, bind to the active site. A nucleophilic cysteine residue forms a covalent bond to the donor lumazine at the C7 carbon. This allows for the 7-exomethylene carbon on the acceptor to attack the donor at the C6 position, forming a pentacyclic intermediate. Finally, the pentacyclic intermediate is cleaved such that the 4 carbons on the donor molecule are attached to the acceptor and the covalent bond between the donor and the enzyme is broken, forming one molecule of riboflavin and one molecule of ARAPD. Support for this mechanism comes from quenching the reaction catalyzed by a RibC variant with a significantly decreased catalytic rate. The pentacyclic intermediate was isolated and the chemical identity was determined using NMR.

12.

The proposed mechanism of riboflavin synthase. Two LUM molecules bind to the RibC active site. The donor LUM, shown in black, will form a covalent bond to the enzyme. The acceptor LUM, shown in blue, will attack the donor LUM to form a covalent bond between the two lumazine molecules leading to the formation of the pentacyclic intermediate, which resolves to form riboflavin and ARAPD (adpated from ref ).

There are two structurally distinct riboflavin synthases. The most extensively studied are trimeric RibC isoforms from E. coli, B. subtilis, A. thaliana, and Brucella abortus. − The general structure is shown in Figure A, an asymmetric trimer of 23 kDa monomers, with the active site located between two adjacent subunits. Each subunit forms N- and C-terminal β-barrels. Interestingly, these two barrels have pseudo 2-fold symmetry which can be predicted based on the high sequence similarity of the N- and C- terminal domains. The final 20 amino acids of each chain form a 32 Å-long triple stranded coiled coil. Less common are the pentameric riboflavin synthases (17 kDa monomers) from Methanocaldococcus jannaschii and Methanothermobacter thermoautrophicus, which have a high degree of sequence similarity to lumazine synthases and a very similar structure (Figure B). Despite the very different structures of the trimeric and pentameric riboflavin synthases, they catalyze the same reaction and likely use the same chemical mechanism. However, the trimeric RibC produces a pentacyclic intermediate with (S,R) stereochemistry, whereas the pentameric RibC generates a pentacyclic intermediate with (R,S) stereochemistry (Figure ).

13.

Structural comparison for RibC. (A) RibC trimer from E. coli (PDB 1I8D). (B) RibC pentamer from Methanocaldococcus jannaschii (PDB: 2B98). RibC trimers generate pentacyclic reaction intermediates with inverted stereochemistry of the RibC pentamers (intermediates shown below their corresponding enzyme).

9. Conclusions

Once riboflavin is made, riboflavin kinase performs a phosphorylation reaction to convert riboflavin to flavin mononucleotide (FMN). FAD synthetase converts FMN to flavin adenine dinucleotide (FAD) with an adenylation reaction or a dual function riboflavin kinase/FAD synthetase sequentially phosphorylates and adenylates riboflavin to generate flavin FAD. These kinase and synthetase enzymes are ubiquitous in all forms of life (Figure ). Slow enzymes have their own kind of charm: like deliberate strokes in a careful mural, their unhurried pace allows enzymologists to observe chemical details that might otherwise blur past. Yet a lingering question remains: how do the riboflavin biosynthetic enzymes, sluggish in vitro, operate with such surprising speed in the cellular environment? As we define the chemistry, ranging from enzymatic mechanisms to macromolecular architecture and even hints of higher-order organization, we move closer to resolving the puzzle. Exciting avenues for exploration include elucidating the role of quaternary structure on the kinetic mechanism, determining the role of post-translational modifications on regulation of the pathway, and determining how these proteins interact to facilitate the expedited riboflavin synthesis in vivo. By answering these questions, we also open paths to improved industrial vitamin production through systems biology or enzyme engineering approaches and to designing new antimicrobial strategies against bacterial and fungal infections.

Z.A.H.: Conceptualization, writing of the manuscript (original draft, reviewing, and editing), and funding acquisition; A.L.L.: Conceptualization and writing of the manuscript (original draft, reviewing, and editing), supervision, project administration, and funding acquisition.

Research reported in this publication was supported by the National Center for Advancing Translational Sciences of the National Institutes of Health under Award Number T32TR004544 to Z.A.H. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. This publication was made possible by funds from the National Science Foundation Award Number CHE-2204080 to A.L.L. The content is solely the responsibility of the authors and does not necessarily represent the views of the National Science Foundation.

The authors declare no competing financial interest.