Skip to main content
NCBI home page
UKPMC Funders Author Manuscripts logoLink to UKPMC Funders Author Manuscripts
. Author manuscript; available in PMC: 2020 Mar 16.
Published in final edited form as: Nat Chem. 2019 Sep 16;11(11):1049–1057. doi: 10.1038/s41557-019-0324-8

Enzymatic control of cycloadduct conformation ensures reversible 1,3 dipolar cycloaddition in a prFMN dependent decarboxylase

Samuel S Bailey 1, Karl A P Payne 1, Annica Saaret 1, Stephen A Marshall 1, Irina Gostimskaya 1, Iaroslav Kosov 1, Karl Fisher 1, Sam Hay 1,*, David Leys 1,*
PMCID: PMC6817360  EMSID: EMS83924  PMID: 31527849

Abstract

The UbiD enzyme plays an important role in bacterial ubiquinone (coenzyme Q) biosynthesis. It belongs to a family of reversible decarboxylases that interconvert propenoic or aromatic acids with the corresponding alkenes or aromatic compounds using a prenylated flavin (prFMN) cofactor. This cofactor is suggested to support (de)carboxylation through a reversible 1,3-dipolar cycloaddition process. Here we report an atomic-level description of the reaction of the UbiD related ferulic acid decarboxylase with substituted propenoic and propiolic acids (data ranging from 1.01 to 1.39 Å). The enzyme is only able to couple (de)carboxylation of cinnamic acid-type compounds to reversible 1,3-dipolar cycloaddition, while formation of dead-end prFMN cycloadducts occurs with distinct propenoic and propiolic acids. The active site imposes considerable strain on covalent intermediates formed with cinnamic and phenylpropiolic acids. Strain reduction through mutagenesis negatively affects catalytic rates with cinnamic acid, indicating a direct link between enzyme-induced strain and catalysis that is supported by computational studies.

Many enzymes make use of covalent catalysis to achieve substantial rate enhancements, often by recruiting cofactors such as PLP1, TPP2 and flavins3. To ensure high turnover, these enzymes are inherently required to ensure both rapid and reversible cofactor-ligand adduct formation. In the case of the UbiD enzyme family, reversible decarboxylation has been suggested to occur via a 1,3-dipolar cycloaddition process between the substrate and the UbiD-cofactor, prenylated FMN (prFMN)4, enabled by the azomethine ylide character of the latter. While 1,3-dipolar cycloaddition is commonly used in organic synthesis5, here we describe this particular process in enzymatic catalysis. Diels-Alder reactions have also recently been found to be catalysed by dedicated enzymes6, but these are not reported to be reversible. Similarly, 1,3-dipolar cycloaddition reactions such as biocompatible ‘click’-chemistry are not considered reversible7, although the possibility of force-induced cycloreversion is a recent and controversial topic8–11. In the case of the fungal ferulic acid decarboxylase (Fdc) UbiD-enzyme12, 1,3-dipolar cycloaddition between prFMNiminium and the cinnamic acid substrate (Fig 1) is proposed to lead to an initial pyrrolidine cycloadduct (Int1) that supports decarboxylation concomitant with ring opening, resulting in formation of a distinct prFMN-alkene adduct (Int2). A conserved Glu residue is proposed to donate a proton to the alkene moiety, leading to a second pyrrolidine cycloadduct (Int3). Finally, the reaction cycle concludes with cycloelimination of Int3, leading to alkene product formation and release. The reaction is readily reversible at elevated [CO2] 12. There is no direct experimental evidence that any of the proposed intermediates exist, although indirect evidence supporting this mechanism has been reported13. Alternative mechanisms proposed to occur with distinct forms of the cofactor have been discounted14. However, it remains unclear how the enzyme is able to ensure reversibility of the 1,3-dipolar cycloaddition steps. We aimed to determine whether detailed structural insights could be gained into catalysis by A. niger Fdc1, an enzyme used as a model prFMN decarboxylase by virtue of the fact it yields atomic resolution crystal structures12.

Fig 1. A. niger Fdc proposed enzyme mechanism.

Fig 1

Open in a new tab

The decarboxylation of cinnamic acid (in red) or the alkyne substrate analogue phenylpropiolic acid (in blue) by covalent catalysis using the prFMNiminium cofactor (in black) is shown. Following the binding step, a 1,3-dipolar cycloaddition between the dipolarophile ligand and the azomethine ylide cofactor leads to the first cycloadduct species Int1. The additional unsaturated bond shown in blue highlights the reaction with the alkyne substrate analogue. Decarboxylation occurs concomitant with ring opening and formation of Int2. The bound CO2 leaves the active site and is replaced by the E282 side chain (shown in green). Protonation via E282 leads to formation of a second cycloadduct species Int3. A cycloelimination process leads to formation of styrene and the entire reaction cycle is freely reversible. The labels for species for which crystal structures are determined here are underlined.

Results

Structure of the AnFdc1 substrate complex

Catalytically competent AnFdc1 crystals were made by avoiding illumination of the protein during sample preparation and crystallisation, preventing photo-induced cofactor isomerisation and inactivation14. Incubation of crystals with cinnamic acid for brief periods of time (typically 10-15 seconds) led to rapid sample deterioration, likely due to styrene accumulation. Electron density obtained with diffraction data from cinnamic acid soaked crystals (prior to crystal dissolution) unambiguously revealed the location of the substrate benzyl moiety above the prFMNiminium isoalloxazine ring system, but revealed density corresponding to multiple conformations of the substrate acrylic acid moiety (vide infra). In contrast, rapid soaking and flash-cooling of crystals with alpha-fluorocinnamic acid revealed clear density for the substrate positioned directly above the prFMNiminium azomethine ylide functionality (i.e. Sub complex, to 1.1 Å resolution; Fig 2a). We could confirm that cinnamic acid binds to the protein in the same conformation, by using inactive AnFdc1 crystals that contained FMN rather than prFMNiminium (to 1.26 Å resolution; Supplementary Fig 1a). In the case of the alpha-fluorocinnamic acid prFMN complex, the substrate Cα and Cβ carbons are located directly above the prFMNiminium C1’ and C4a (Fig 1), at distances of 3.0 Å and 3.4 Å respectively, in a reactive conformation compatible with the proposed cycloaddition.

Fig. 2. Crystal structures of A. niger Fdc with propionic acid substrates.

Fig. 2

Open in a new tab

a. Active site structure of Fdc in complex with alpha-fluorocinnamic acid. b. Omit electron density contoured at 4.5 sigma and corresponding model of the cinnamic acid Int2 in the E282Q mutant. c. Overlay between the Int2 E282Q covalent adduct formed and the adduct obtained with phenylpyruvate (PDB 4ZA9) d. Omit electron density contoured at 3 sigma and corresponding model of the cinnamic acid Int3. (populated at 30%) trapped by rapid flash-cooling of WT crystals following exposure to cinnamic acid (Supplementary Fig. 1 shows the product complex for comparison). e. Omit electron density contoured at 4 sigma corresponding to Int3crotonic. f. Overlay between the Int3 and Int3crotonic acid covalent adducts formed.

The E282Q AnFdc1 variant allows structure determination of the Int2 covalent adduct

In order to determine whether covalent adducts (either Int1 or Int2) might accumulate in the absence of the Glu282 mediated protonation step, we sought to determine the crystal structure of the inactive AnFdc1 E282Q variant14 in complex with cinnamic acid and cinnamic acid derivatives. Indeed, UV-Vis spectroscopy of AnFdc1 E282Q incubated with cinnamic acid leads to formation of a distinct spectral species, indicative of adduct formation (Supplementary Fig 2a). The E282Q crystal structure obtained by co-crystallisation with cinnamic acid reveals a covalent prFMN-substrate complex, corresponding to the proposed Int2 species following decarboxylation (to 1.18 Å resolution; Fig 2b). The substrate alkene bond retains the trans configuration but substantially deviates from ideal planar geometry, with a torsional angle of τ = ~112° (as opposed to 180° for a planar trans configuration) and an elongated Cα=Cβ bond length of 1.37 Å. Exchange of the bound cinnamic acid for alpha-fluorocinnamic acid or pentafluorocinnamic acid readily occurs in crystallo, demonstrating a Sub ⇌ Int2 + CO2 equilibrium exists in E282Q (Supplementary Fig 1b,c). The alpha-fluorocinnamic acid Int2 structure (to 1.13 Å resolution) directly demonstrates syn-pyramidalisation15 of the substrate alkene Cα (pyramidalisation angle of ϕ ~20°), which is likely to be a result of the torsional strain imposed. The observed alkene distortion is a likely consequence of the firm hold the enzyme active site exerts on the substrate phenyl moiety, held in place by I187, I327 and F437 at a position directly above and stacked with the prFMN plane. For all E282Q Int2 structures, the prFMN C1’ is located further out of the prFMN plane by ~0.4 Å when compared to other prFMN single bond adduct species (i.e. PDB 4ZA9; Fig 2c). In the case of cinnamic acid E282Q Int2, this places the Cβ directly above the prFMN C4a at a distance of 3.04 Å, while the Cα atom is in close proximity (~3.2 Å) of the location occupied by the Glu282 carboxylate moiety in the wild-type enzyme structure.

Structure of cycloadducts Int1’ and Int3’ formed with alkyne compounds

The substrate and product alkyne analogues phenylpropiolic acid and phenylacetylene were used to determine whether cycloadducts Int1 and/or Int3, respectively can be observed in the wild-type enzyme. These compounds can also act as dipolarophiles, but cycloaddition will lead to the double bond-containing 3-pyrroline Int 1’ rather than the pyrrolidine cycloadducts (Fig 1). Upon addition of either alkyne compound, a clear shift is observed in the enzyme UV-Vis spectrum indicating modification of the prFMNiminium cofactor, with the formation of a species distinct from the E282Q Int2 adduct (Supplementary Fig 2b). Stopped-flow experiments reveal the formation of this alkyne intermediate occurs with kobs = 101.5 ± 0.4 s-1 (Supplementary Fig 3), but following prolonged incubation no product formation could be observed (Supplementary Fig 2c). To investigate the phenylpropiolic acid complex prior to reaction (i.e. Inhib complex), inactive crystals of AnFdc1 (with prFMN in a semiquinone radical form) were used, revealing the inhibitor (to 1.29 Å resolution) binds in a similar manner to the alpha-fluorocinnamic acid:prFMN or cinnamic acid substrate:FMN complexes obtained. This positions the alkyne Cα and Cβ directly above the prFMNradical C1’ and C4a at distances of 3.3 Å and 3.7 Å (Supplementary Fig 1d). In contrast, crystals of active AnFdc1 co-crystallised with phenylpropiolic acid (to 1.01 Å resolution) or soaked with phenylacetylene (to 1.10 Å resolution) revealed clear density for a 3-pyrroline cycloadduct (Int3’, analogous to the proposed Int3) formed between the alkyne moiety and the prFMNiminium (Fig 3a). Hence, surprisingly, the phenylpropiolic acid derived adduct lacked density for the carboxylate moiety, and was highly similar to that obtained with phenylacetylene (Supplementary Fig 1e). This suggests decarboxylation has occurred over the prolonged time frame required for co-crystallisation (typically exceeding 24 hours). Indeed, when using a rapid soaking procedure as an alternative approach, a distinct cycloadduct (Int1’; to 1.21 Å resolution) is observed that retains the carboxyl moiety attached to the prFMN C1’ group (and thus resembles the proposed Int1; Fig 3b). We used a mass spectrometry-based detection method to demonstrate slow decarboxylation occurs within a 24 hour time period in solution, confirming the conversion of Int1’ to Int3’ mimicking the natural activity of the enzyme with cinnamic acid occurs (Supplementary Fig 4a). We then took advantage of the distinct spectral change associated with cycloadduct formation to determine if cycloaddition with natural substrates can also be monitored in solution. Indeed, rapid mixing of the enzyme with cinnamic or sorbic acid substrates revealed spectral changes in the UV-Vis spectrum of prFMN similar to those observed with the alkyne inhibitors (Supplementary Fig 3b,c). This suggest that, for these substrates, cycloadduct(s) accumulate under multiple turnover conditions.

Fig 3. Crystal structures of A. niger Fdc with propiolic acid substrate analogues.

Fig 3

Open in a new tab

a. Omit electron density contoured at 5 sigma and corresponding model of the phenylpropiolic acid derived Int3’ obtained through co-crystallisation (Calpha-C1’ 1.52 Å and Cbeta-C4a 1.66 Å bond distances). b. Omit electron density contoured at 4 sigma and corresponding model of the phenylpropiolic acid derived Int1’ obtained through soaking and rapid flash-cooling (Calpha-C1’ 1.59 Å and Cbeta-C4a 1.70 Å bond distances). c. Overlay between the phenylpropiolic acid derived Int1’ and Int3’ covalent adducts formed. d. Omit electron density contoured at 4 sigma and corresponding model of the phenylpropiolic acid derived Int1’ for the L439G variant (Calpha-C1’ 1.48 Å and Cbeta-C4a 1.86 Å bond distances). e. Omit electron density contoured at 4 sigma and corresponding model of the phenylpropiolic acid derived Int1’ for the F437L variant (Calpha-C1’ 1.51 Å and Cbeta-C4a 1.79 Å bond distances). f. Overlay between the Int1’ adduct obtained in the WT and the F437L variant. g. Omit electron density contoured at 4 sigma and corresponding model of the 2-butynoic acid derived Int1’butynoic (Calpha-C1’ 1.52 Å and Cbeta-C4a 1.53 Å bond distances). h. Overlay between the Int1’ and Int1’butynoic adduct.

The AnFdc1 active site constrains cycloadduct conformation

While both Int1’ and Int3’ structures confirm 1,3-dipolar cycloaddition can occur with the prFMNiminium cofactor, several features indicate a significant degree of strain in these cycloadduct(s). The phenylpropiolic acid Int1’ structure deviates from ideal bond parameters both in the position of the inhibitor phenyl group (Cβ out-of-phenyl plane angle is ~24° as opposed to ideal 0°), the deviation of the 3-pyrroline ring (C1’-Cα-Cβ bond angle of ~99°, as compared to ideal 108°), prFMN tetrahedral C4a bond angles (C2-C4a-Cβ bond angle of ~103° versus ideal 109.5°of sp3 C4α) as well as the elongated C4a-Cβ bond length of 1.7 Å (Fig 4a). Following decarboxylation, the position of the Int3’ 3-pyrroline ring shifts (Fig 3c) to alleviate some of the phenyl group strain (Cβ out-of-plane angle reduced to ~16°) as well as the deviation of the 3-pyrroline ring (C1’-Cα-Cβ bond angle of ~111°; Cβ-C4a bond length of 1.65 Å). The protein does not alter in conformation when comparing the Inhib/Int1’ and Int3’ structures, with the notable exception of Glu282, which reorients to occupy the substrate/inhibitor carboxylate binding pocket in Int3’ (a motion similar to the CO2 for Glu282 exchange proposed to occur during catalysis, Fig 1). In the latter, Glu282 forms an interaction with the 3-pyrroline substrate-derived Cα (distance ~3.2 Å to either E282 oxygen; Fig 3a). Glu282 mediated proton donation or abstraction would interconvert Int3’ with the putative prFMN C1’ – alkyne adduct Int2’ (the complex analogous to Int2; Fig 1). However, an ‘unstrained’ linear C1’-alkyne adduct –with a C1’-Cα≡Cβ angle of 180°– is sterically incompatible with the AnFdc1 active site, suggesting a highly strained Int2’ (similar to strained cyloalkyne species15) connects the Int1’ and Int3’ species, providing a likely explanation for the extremely slow rate of Int1’ decarboxylation.

Fig. 4. The role of strain in the Fdc reaction.

Fig. 4

Open in a new tab

Schematic overview of the structural insights gained into the Fdc reaction cycle. The prFMNiminium cofactor and the substrate are shown for the various crystal structures of the Sub/Int1’ (as a model for Int1)/Int2 and Int3 states. The active site restricts the position of the substrate phenyl ring throughout the reaction (as indicated by black arrow labelled F437), leading to formation of strained intermediates Int1/Int2 and Int3. Substantial deviations from ideal bond lengths and angles are highlighted in black. For the Int1’/Int2 and Int3 species, DFT models of the corresponding prFMN adducts free in solution are overlaid (coloured in grey). The differences observed for Int1’ and Int3 with respective DFT models of the corresponding unconstrained prFMN adducts resembles those observed when comparing Int1’ and Int3 with Int1butynoic and Int3crotonic (Figs 3h and 2f).

Mutagenesis alleviates cycloadduct strain and impacts catalysis

As observed for the Int2 E282Q structures, the strain observed in the Int1’ and Int3’ structures appears to be a direct consequence of the inherent restrictions imposed by the rigid AnFdc1 active site. The position of the substrate aromatic group appears constrained to remain π-stacked with the prFMN isoalloxazine ring, by virtue of steric interactions with the Phe437 sidechain located adjacent to the conserved Leu439. This in turn imposes restrictions on the extent by which the prFMN isoalloxazine ring system can deform upon formation of a C4a-Cβ bond with substrate, as the flavin uracil ring is sandwiched between Ala172 on the si face and the substrate aromatic group at the re face. We constructed the F437L and L439G variants and determined the crystal structures in complex with phenylpropiolic acid, as well as the activity with cinnamic acid. We found both mutations severely compromised enzyme activity as judged from steady state turnover. In the case of L439G, a substantial increase of K M was observed, while the F437L variant was only affected in kcat confirming a key role for Phe437 in catalysis (Supplementary Fig 5b,d). In the corresponding phenylpropiolic acid adduct crystal structures for both variants, Int1’ decarboxylation in crystallo was not observed, even following co-crystallisation and prolonged incubation (up to several weeks) prior to X-ray exposure. This lack of decarboxylation activity with Int1’ was confirmed in solution (Supplementary Fig 4b). In the case of the L439G Int1’ crystal structure (to 1.12 Å resolution), a modest reorientation of the substrate carboxyl group along with a minor reduction of the Cβ out-of-plane angle is observed, associated with a corresponding reorientation of the F437 phenyl ring (Fig 3d). The void created by the L439G mutation is filled with a water molecule located within hydrogen bonding distance of the Int1’ carboxylate, reducing the hydrophobicity of the carboxylate binding pocket. In addition, the L439G void was partially filled by the phenyl group of a second phenylpropiolic acid molecule, bound at the entrance of the active site. This observation offers a possible explanation for the drastic increase of the observed K M for cinnamic acid in the L439G (as opposed to F437L) mutant, as similar binding of a second cinnamic acid molecule could be required for decarboxylation by L439G. The hydrophobic nature of the carboxylic acid binding pocket has been implicated in the mechanism of other decarboxylases16,17. In marked contrast, the F437L mutation does not affect the carboxylate binding or conformation, but subtly alters the position of the substrate phenyl moiety (structure obtained to 1.08 Å resolution; Fig 3e,f). This reorientation, while retaining a strained conformation of the Cβ out-of-plane angle of ~25°, is associated with elongation of the Cβ-C4a bond to 1.8 Å. It thus appears the Phe437 side chain counteracts the forces exerted on the prFMN 3-pyrroline ring through enzyme interactions with the Int1’ carboxylate.

Smaller dipolarophile compounds form dead-end cycloadducts

Unfortunately, other mutations at the Phe437 position that could lead to further reduction of strain imposed by the 437 side-chain led to variants that did not bind the prFMN cofactor. We therefore co-crystallised the WT enzyme with 2-butynoic acid, avoiding the enzyme imposed adduct strain by interactions with the missing substrate phenyl moiety. In this case, an Int1’butynoic structure was obtained to 1.02 Å resolution, revealing no decarboxylation had occurred (Fig 3g). Furthermore, the Int1’butynoic is markedly different from the strained phenylpropiolic acid derived Int1’ structure (Fig 3h). Little to no features associated with a strained Int1’ configuration can be observed in the case of Int1’butynoic, a consequence of repositioning of the prFMNiminium C4a and associated uracil plane. This repositioning leads to a near ideal conformation of the Int1’butynoic 3-pyrroline ring, with a reduced Cβ-C4a bond length of 1.56 Å and a C1’-Cα-Cβ bond angle of ~107°, but is incompatible with the presence of a substrate phenyl group held within the context of either the WT or F437L variant structures. As a consequence of the uracil plane repositioning, the prFMN O1 has moved out of the oxyanion binding-pocket formed by Q191 and two water molecules.

Decarboxylation of Int1’, and by similarity Int1, is likely assisted by the relatively hydrophobic nature of the carboxylate binding pocket, and ring opening of the strained cycloadduct to Int2’/Int2. It is however unclear what drives the cycloelimination step from Int3 to product release, a process not observed for Int3’. Careful analysis of electron density associated with rapidly flash-cooled WT crystals incubated with cinnamic acid reveals this can be modeled as a mixture of 30% Int3 and a 40% product complex (the remainder corresponding to the inactive hydroxylated prFMN species), suggesting cycloelimination is the rate limiting step under these conditions, as has previously been suggested13 (to 1.24 Å resolution; Fig 2d). As observed with the Int3’ structure, the Int3 Cβ has an out-of-plane angle of ~13°, with additional strain observed for the Cα-Cβ-Cγ angle of ~122°, both associated with the tight hold the enzyme exerts on the substrate phenyl moiety and resulting in an C1’ endo envelope conformation of the pyrrolidine ring (as opposed to the N endo conformation of the Int3’ enforced by the Cα-Cβ double bond (Fig 4a). Hence, to determine how Int3 strain might assist cycloelimination, we determined the structure of an Int3crotonic species (to 1.39 Å resolution), obtained by co-crystallisation with crotonic acid (2-butenoic acid; Fig 2e). A comparison of Int3crotonic with the cinnamic acid Int3 (Fig 2f) reveals the Int3crotonic pyrrolidine ring adopts a distinct envelope conformation with the N in endo position, that allows the prFMNiminium C1’ to adopt a more in-plane position, resembling the Int3’ geometry. The Int3crotonic conformation cannot be adopted by the cinnamic acid Int3, as severe clashes between the substrate phenyl ring and the active site occur. We suggest the lack of strain in the Int3crotonic species is coupled to the fact no cycloelimination can be observed for this species, while a strained conformation such as that observed for cinnamic acid Int3 is required for progression along the catalytic cycle.

Combining the various crystal structures of Sub/Inhib, Int1’/Int2 and Int3’ complexes with our solution data leads to a comprehensive overview of the AnFdc1 catalytic cycle (Fig 4). The rigid AnFdc1 active site brings the substrate into close proximity of the prFMNiminium cofactor, aligning the Cα-Cβ double bond (or triple bond in case of alkyne inhibitors) with the C1’-N5-C4a azomethine ylide to poise the complex for cycloaddition. Formation of a pyrrolidine adduct between a similar putative encounter complex of cinnamic acid and free prFMNiminium in solution would be accompanied by repositioning of both the Cα-Cβ substituents (i.e. carboxylate/benzyl group), as well as the prFMN uracil moiety. This is a direct consequence of the ring formation and the accompanying sp2 to sp3 change for substrate Cα/Cβ and prFMN C1’/C4a carbon atoms. However, within the confines of the enzyme active site, the extent to which reorientation can occur is limited. In fact, the enzyme active site appears evolved to have maximum complementarity with the substrate and product complexes, and hence constrains both the substrate carboxylate and phenyl moieties to adopt a position resembling the substrate complex throughout catalysis. In the case of the Int1’butynoic and Int3crotonic adducts, the lack of a phenyl moiety allows for a more relaxed conformation of the substrate/prFMNiminium adduct within the confines of the active site. Crucially, neither Int1’butynoic decarboxylation nor Int3crotonic cycloelimination are observed suggesting that strain contributes to catalysis for both these steps.

Computational studies provide further insight into the AnFdc1 catalytic cycle

An active ‘cluster’ site model comprising ~ 230 atoms was built from the E282Q Int2 X-ray crystal structure. The positions of peripheral atoms were fixed to maintain a crystal structure-like geometry, in addition to constraining the position of the F437 and L439 side chains in most calculations to allow their contribution to strain in the adducts to be investigated. Stable models of each species as identified in Fig 1 were found for cinnamic acid, phenylpropiolic acid and crotonic acid. In the case of cinnamic acid, cycloaddition (Sub → Int1) was found to occur as a 2-step reaction, via a ring-open covalent adduct (Int1open), which possesses an unusual Cβ–C4a interaction (2.60 Å) in addition to a Cα–C1’ bond (1.58 Å). The barrier to formation of the ring-closed Int1 intermediate is small (Fig 5), so the reaction may effectively occur as one asynchronous step. In contrast, no stable Int1open-type intermediates were observed for phenylpropiolic or crotonic acid cycloaddition, which also leads to significantly more stable Int1 cycloadducts.

Fig. 5. Potential energy diagram of the Fdc reaction.

Fig. 5

Open in a new tab

Potential energy diagram determined using DFT cluster models (Supplementary Figs 6,7), with potential energy normalised to the Sub/Inhib and Int3 energies. The relaxed cinnamic acid energies were determined for models where restraints on the side chains of Phe437 and Leu439 were removed, which alleviates some of the strain on the Cβ–Ph bond (Supplementary Fig. 8). The two phenylpropiolic acid Int1 species interconvert via rotation of the Cα–CO2 bond and the conformation with the carboxyl group in the phenyl plane is the higher energy species and is not stable in the DFT model used here. The crotonic acid Int3 → Prod transition state actually represents a minimum energy crossing point (Supplementary Figs. 9, 10).

Cinnamic acid decarboxylation occurs from the ring-open Int1open species and the Int1open → Int2CO2 potential barrier is relatively low at ~ 16 kJ mol-1 (Fig 5). The barriers for phenylpropiolic and crotonic acid decarboxylation were not determined, but may be much higher than for cinnamic acid and/or occur via a different mechanism, as these reactions cannot proceed via an Int1open intermediate and the decarboxylation of these is then significantly endothermic. In all cases, the Int2CO2, which contain noncovalently bound CO2, remain ring-open with broken (~3 Å) Cβ-C4a bonds. The crotonic acid derived Int2CO2 is most stable as it can adopt an unstrained ring-open conformation with near-ideal Cα–Cβ geometry. In contrast, both cinnamic acid and phenylpropiolic Int2CO2 intermediates adopt strained conformations with non-linear Cα–Cβ–Phenyl angles. Relaxation of the side chain constraints of Phe437 and Leu439, as a proxy of the F437L and L439G variants, relieves ~14 kJ mol-1 of strain from the cinnamic acid derived Int2CO2 adduct consistent with the role of these residues in constraining the geometry of the phenyl moiety of the adducts.

Once the CO2 has vacated the active site, Glu282 mediated protonation of Cβ leads to the formation of a ring-closed Int3 species for all compounds. The release of CO2 and Cβ protonation were not studied in detail, as these reactions require substantial rearrangement of Glu282, which would require much larger models. Cycloelimination (Int3 → Prod) to form non-covalently bound product complexes is exothermic and likely to be rate limiting in all three cases (Fig 5). Formation of styrene is the least exothermic (38 kJ mol-1), while cycloelimination of phenylacetylene is highly unfavorable, being 81 kJ mol-1 uphill and likely proceeding via a barrier > 100 kJ mol-1 (not determined). Similarly, the barrier to propylene cycloelimination is estimated at ~100 kJ mol-1. This is in agreement with our observation that turnover does not occur with either crotonic or phenylpropiolic acid, with the reaction stalling at the Int3 stage. Crucially, in the case of cinnamic acid, the cycloelimination barrier was determined to be feasible, at 64 kJ mol-1 and 78.5 kJ mol-1 in the constrained and F437/L439-relaxed models respectively. The 14.4 kJ mol-1 difference is at least qualitatively consistent with the ~30 fold decrease in kcat observed in the F439L variant (which retains some element of strain).

Discussion

In the case of the Fdc branch of the UbiD family, our data unambiguously establishes that the enzyme relies on covalent catalysis by the unusual prFMN cofactor through a reversible 1,3-dipolar cycloaddition process. The fact the active site is highly complementary in shape to the substrate/prFMNiminium complex prior to cycloaddition has the serendipitous consequence it does not yield to allow pyrrolidine ring substituents to adopt the ideal conformation. This results in the formation of strained intermediates, ensuring rapid progression through the catalytic cycle. While substrate distortion/ground state destabilization has long been suggested to contribute to enzyme catalysis18,19, it has only been conclusively demonstrated in a limited number of cases20,21, and rarely includes atomic resolution insights22–25. Our study reveals in AnFdc1 distortion occurs specifically for covalent substrate-cofactor adducts only, and the strain is directly linked to progression along the reaction coordinate. Hence, by targeted destabilization of intermediate species only, AnFdc1 harnesses 1,3-dipolar cycloaddition as a readily reversible reaction. This aligns with the proposals from Albery and Knowles26 for evolution of an efficient enzyme by control of the internal thermodynamics of the bound states. Interestingly, many members of the UbiD family act directly on aromatic substrates, and can act in the carboxylative direction as suggested for the anaerobic degradation of benzene27,28. (De)carboxylation of aromatic substrates presents a large inherent barrier to cycloaddition due to transient substrate dearomatisation, as well as scope for formation of a highly stable Int2 adduct by re-aromatisation29. How these problems are overcome, particularly in view of whether the prFMN-dependent catalysis in these enzymes is similar to the Fdc branch, awaits a more detailed analysis of additional members of the UbiD family. Finally, our data suggests mechanochemical reversion of 1,3-dipolar cycloaddition reactions is feasible, providing further support to the possibility of developing mechanoresponsive materials through reversible click-chemistry8–11. Our data show Nature is able to harness and control 1,3-dipolar cycloaddition chemistry by combination of a unique cofactor with strict mechanochemical control.

Methods

See Supplementary Information for methods.

Supplementary Material

Supplementary information

Acknowledgements

This work was supported by the grants BBSRC BB/K017802 and ERC pre-FAB 695013. The authors acknowledge the assistance given by the use of the Manchester Protein Structure Facility. We thank Diamond Light Source for access (proposal numbers MX12788 and MX17773) that contributed to the results presented here. D.L. is a Royal Society Wolfson Merit Award holder.

Footnotes

Author contributions

SSB cloned, expressed and purified AnFdc1 and various variants. SSB determined all crystal structures with guidance from DL. KAPP generated various AnFdc1 variants, and collected solution data with cinnamic acid. AS assisted with crystallisation of the Int3crotonic state. SAM assisted with AnFdc1 reconstitution. IG assisted with crystallisation of FMN substitued AnFdc1 and the Int3butynoic state (with help from IK). KF assisted with protein purification and solution data collection and analysis. SH performed all computational studies. All authors discussed the results and participated in writing of the manuscript. DL initiated and directed this research.

Competing interests

The authors declare no competing interests.

Data availability

The data generated and analysed in this study, including the computational modeling data associated with all figures, are available from the corresponding authors upon reasonable request. The diffraction data and corresponding atomic models have been deposited in the PDB under accession codes 6R3G, 6R3F, 6R3I, 6R2Z, 6R30, 6R3O, 6R2T, 6R2R, 6R3N, 6R3L, 6R3J, 6R2P, 6R34, 6R33 and 6R32.

References

  • 1.Toney MD. Controlling reaction specificity in pyridoxal phosphate enzymes. Biochim Biophys Acta - Proteins Proteomics. 2011;1814:1407–1418. doi: 10.1016/j.bbapap.2011.05.019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Kluger R, Tittmann K. Thiamin diphosphate catalysis: Enzymic and nonenzymic covalent intermediates. Chem Rev. 2008;108:1797–1833. doi: 10.1021/cr068444m. [DOI] [PubMed] [Google Scholar]
  • 3.Piano V, Palfey BA, Mattevi A. Flavins as Covalent Catalysts: New Mechanisms Emerge. Trends Biochem Sci. 2017;42:457–469. doi: 10.1016/j.tibs.2017.02.005. [DOI] [PubMed] [Google Scholar]
  • 4.Marshall SA, Payne KAP, Leys D. The UbiX-UbiD system: The biosynthesis and use of prenylated flavin (prFMN) Arch Biochem Biophys. 2017;632:209–221. doi: 10.1016/j.abb.2017.07.014. [DOI] [PubMed] [Google Scholar]
  • 5.Pellissier H. Asymmetric 1,3-dipolar cycloadditions. Tetrahedron. 2007;63:3235–3285. [Google Scholar]
  • 6.Jeon B-S, et al. Investigation of the mechanism of the SpnF-catalyzed [4+2]-cycloaddition reaction in the biosynthesis of spinosyn A. Proc Natl Acad Sci U S A. 2017;114:10408–10413. doi: 10.1073/pnas.1710496114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Meldal M, Tornøe CW. Cu-Catalyzed Azide–Alkyne Cycloaddition. Chem Rev. 2008;108:2952–3015. doi: 10.1021/cr0783479. [DOI] [PubMed] [Google Scholar]
  • 8.Jacobs MJ, Schneider G, Blank KG. Mechanical Reversibility of Strain-Promoted Azide-Alkyne Cycloaddition Reactions. Angew Chemie Int Ed. 2016;55:2899–2902. doi: 10.1002/anie.201510299. [DOI] [PubMed] [Google Scholar]
  • 9.Khanal A, Long F, Cao B, Shahbazian-Yassar R, Fang S. Evidence of Splitting 1,2,3-Triazole into an Alkyne and Azide by Low Mechanical Force in the Presence of Other Covalent Bonds. Chem - A Eur J. 2016;22:9760–9767. doi: 10.1002/chem.201600982. [DOI] [PubMed] [Google Scholar]
  • 10.Krupička M, Dopieralski P, Marx D. Unclicking the Click: Metal-Assisted Mechanochemical Cycloreversion of Triazoles Is Possible. Angew Chemie Int Ed. 2017;56:7745–7749. doi: 10.1002/anie.201612507. [DOI] [PubMed] [Google Scholar]
  • 11.Stauch T, Dreuw A. Force-induced retro-click reaction of triazoles competes with adjacent single-bond rupture. Chem Sci. 2017;8:5567–5575. doi: 10.1039/c7sc01562c. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Payne KaP, et al. New cofactor supports α,β-unsaturated acid decarboxylation via 1,3-dipolar cycloaddition. Nature. 2015;522:497–501. doi: 10.1038/nature14560. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Ferguson KL, Eschweiler JD, Ruotolo BT, Marsh ENG. Evidence for a 1,3-Dipolar Cyclo-addition Mechanism in the Decarboxylation of Phenylacrylic Acids Catalyzed by Ferulic Acid Decarboxylase. J Am Chem Soc. 2017;139:10972–10975. doi: 10.1021/jacs.7b05060. [DOI] [PubMed] [Google Scholar]
  • 14.Bailey SS, et al. The role of conserved residues in UbiD/Fdc decarboxylase in oxidative maturation, isomerisation and catalysis of prenylated flavin mononucleotide. J Biol Chem. 2018;293:2272–2287. doi: 10.1074/jbc.RA117.000881. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Borden WT. Pyramidalized Alkenes. Chem Rev. 1989;89:1095–1109. [Google Scholar]
  • 16.Okrasa K, et al. Structure and Mechanism of an Unusual Malonate Decarboxylase and Related Racemases. Chem - A Eur J. 2008;14:6609–6613. doi: 10.1002/chem.200800918. [DOI] [PubMed] [Google Scholar]
  • 17.Jez JM, Ferrer JL, Bowman ME, Dixon RA, Noel JP. Dissection of malonyl-coenzyme A decarboxylation from polyketide formation in the reaction mechanism of a plant polyketide synthase. Biochemistry. 2000;39:890–902. doi: 10.1021/bi991489f. [DOI] [PubMed] [Google Scholar]
  • 18.Blake CC, et al. Crystallographic studies of the activity of hen egg-white lysozyme. Proc R Soc London Ser B, Biol Sci. 1967;167:378–88. doi: 10.1098/rspb.1967.0035. [DOI] [PubMed] [Google Scholar]
  • 19.Vocadlo DJ, Davies GJ, Laine R, Withers SG. Catalysis by hen egg-white lysozyme proceeds via a covalent intermediate. Nature. 2001;412:835–838. doi: 10.1038/35090602. [DOI] [PubMed] [Google Scholar]
  • 20.Anderson VE. Quantifying energetic contributions to ground state destabilization. Arch Biochem Biophys. 2005;433:27–33. doi: 10.1016/j.abb.2004.09.026. [DOI] [PubMed] [Google Scholar]
  • 21.Zhang Y, Schramm VL. Ground-State Destabilization in Orotate Phosphoribosyltransferases by Binding Isotope Effects. Biochemistry. 2011;50:4813–4818. doi: 10.1021/bi200638x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Lehwess-Litzmann A, et al. Twisted Schiff base intermediates and substrate locale revise transaldolase mechanism. Nat Chem Biol. 2011;7:678–684. doi: 10.1038/nchembio.633. [DOI] [PubMed] [Google Scholar]
  • 23.Milić D, et al. Crystallographic snapshots of tyrosine phenol-lyase show that substrate strain plays a role in C-C bond cleavage. J Am Chem Soc. 2011;133:16468–16476. doi: 10.1021/ja203361g. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Vladimirova A, et al. Substrate Distortion and the Catalytic Reaction Mechanism of 5-Carboxyvanillate Decarboxylase. J Am Chem Soc. 2016;138:826–836. doi: 10.1021/jacs.5b08251. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Lüdtke S, et al. Sub-ångström-resolution crystallography reveals physical distortions that enhance reactivity of a covalent enzymatic intermediate. Nat Chem. 2013;5:762–767. doi: 10.1038/nchem.1728. [DOI] [PubMed] [Google Scholar]
  • 26.Albery WJ, Knowles JR. Efficiency and Evolution of Enzyme Catalysis. Angew Chemie Int Ed English. 1977;16:285–293. doi: 10.1002/anie.197702851. [DOI] [PubMed] [Google Scholar]
  • 27.Abu Laban N, Selesi D, Rattei T, Tischler P, Meckenstock RU. Identification of enzymes involved in anaerobic benzene degradation by a strictly anaerobic iron-reducing enrichment culture. Environ Microbiol. 2010;12:2783–2796. doi: 10.1111/j.1462-2920.2010.02248.x. [DOI] [PubMed] [Google Scholar]
  • 28.Luo F, et al. Metatranscriptome of an anaerobic benzene-degrading, nitrate-reducing enrichment culture reveals involvement of carboxylation in benzene ring activation. Appl Environ Microbiol. 2014;80:4095–4107. doi: 10.1128/AEM.00717-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Baunach M, Hertweck C. Natural 1,3-dipolar cycloadditions. Angew Chemie - Int Ed. 2015;54:12550–12552. doi: 10.1002/anie.201507120. [DOI] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Supplementary information
EMS83924-supplement-1.pdf (7.5MB, pdf)

Data Availability Statement

The data generated and analysed in this study, including the computational modeling data associated with all figures, are available from the corresponding authors upon reasonable request. The diffraction data and corresponding atomic models have been deposited in the PDB under accession codes 6R3G, 6R3F, 6R3I, 6R2Z, 6R30, 6R3O, 6R2T, 6R2R, 6R3N, 6R3L, 6R3J, 6R2P, 6R34, 6R33 and 6R32.

RESOURCES