Arylmalonate decarboxylase (AMDase, EC 4.1.1.76), originally isolated from the soil bacterium Bordetella bronchiseptica, catalyzes the decarboxylation of α-aromatic- or α-alkenylmalonic acids to yield the corresponding optically pure mono-acid without the aid of a cofactor. While the wildtype enzyme exhibits strict (R)-selectivity, enzyme engineering afforded efficient (S)-selective or racemizing enzyme variants (Figure 1A). The native function or the natural substrate of AMDase, however, both remain unknown to date. The unique reactivity and broad substrate tolerance allows for the production of diverse aryl- or alkenylaliphatic carboxylic acids in outstanding optical purity, amongst them several α-arylpropionates with non-steroidal anti-inflammatory activity, the so-called profens (Miyamoto and Kourist, 2016).

Miyamoto et al. discovered arylmalonate decarboxylase (AMDase) in the 1990’s in a screening to identify enzymes for the generation of chiral molecules from prochiral malonates by enzymatic decarboxylation (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992a) As the decarboxylation of malonyl-ACP is a key step in metabolism, it appeared likely that the bacterial catabolism might possess malonate decarboxylases. The screening followed the assumption that decarboxylation of α-phenylmalonic acid 1a could represent the initial step in the metabolism, followed by oxidation to yield 2-oxo-2-phenylacetic acid. Accordingly, it was proposed that analogous decarboxylation of disubstituted malonic acids would lead to α-chiral acids, as further oxidation would not be possible (Figure 1B).

An assay for microorganisms with the ability to grow on phenylmalonic acid 1a as the sole carbon source led to the identification of the soil bacterium Alcaligenes bronchisepticus KU 1201 (now: Bordetella bronchiseptica). It could be shown, that whole cells of this bacterium also converted α-methyl-α-phenylmalonic acid 2a and analogous substrates with various aromatic residues (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992b). The decarboxylase was purified from the microorganism and named arylmalonate decarboxylase (AMDase), due to its preference towards α-arylmalonates. Characterization of the purified enzyme revealed that it acts without a cofactor and is biotin-independent. Yet, the native role and substrate of AMDase remained unclear, as aryl malonates are not naturally abundant (Miyamoto and Ohta, 1992b).

Some enzymes with about 30% homology to AMDase were identified via PSI-BLAST (position-specific iterative basic local alignment search tool), all belonging to the class of isomerases (EC 5). Glutamate racemase (Lactobacillus fermenti), aspartate racemase (Streptococcus thermophilus), hydantoine racemase (Pseudomonas sp. strain NS671) and maleate isomerase (Alcaligenes faecalis) all share the conserved cysteine at position 188 with AMDase, while enzymes from the isomerase class have an additional cysteine at around residue 74 (Figure 2A) (Ijima et al., 2005). As the reaction mechanism and crystal structure of glutamate racemase were already well-studied, two cysteine residues on both sides of the substrate were proposed to be essential for the racemizing activity, by either abstracting or donating protons from opposite sites in a so-called two-base mechanism (Glavas and Tanner, 1999).

Inspired by these findings, Ijima et al. decided to introduce a cysteine at position 74 instead of the glycine residue present in AMDase (Ijima et al., 2005). Additionally, previous studies have shown that the mutation G188S led to a drastic decrease of native AMDase activity (Miyazaki et al., 1997). While the proton-donating ability of serine is already low compared to cysteine, the authors proposed that an amino acid without any acidic proton would be beneficial for the enantiomeric excess of the obtained product. Surprisingly, the mutant G188A proved to be completely inactive. Hence, the double-variant G74C/C188S was prepared, and was found to produce the opposite enantiomers in 94–96 %ee as compared to wildtype AMDase. Yet, activity of the G74C/C188S variant even fell below the tremendously reduced activity of C188S (Ijima et al., 2005). Interestingly, also for the (S)-selective variant S36N/G74C/C188S, preference for cleavage of the pro-(R) carboxylate was observed by isotope labeling studies (Terao et al., 2006a), thus confirming that enantioselective reprotonation of the enolate intermediate is decisive for the final configuration of the product (Miyamoto et al., 2007a). As already observed by Ijima et al., the single amino acid exchange G74C rendered arylmalonate decarboxylase a racemase (Terao et al., 2006b). Notably, the decarboxylase activity of AMDase was preserved in the G74C variant, yielding racemic arylpropionates from arylmalonates. Kinetic analysis disclosed that, in terms of catalytic efficiency (k _cat/K _M), decarboxylation (0.96 s⁻¹mM⁻¹) exceeded racemization (0.56 s⁻¹mM⁻¹), which was also reflected by the racemic product already detected at an early stage of the reaction (Terao et al., 2006b).

Despite the crystal structure of AMDase remained unsolved until 2008, considerable insights were already acquired by that time, including suggestions on the enzyme mechanism (Miyamoto et al., 1992b; Miyamoto et al., 2007a; Matoishi et al., 2004), identification of key residues (Matoishi et al., 2004; Terao et al., 2007), switch in enantioselectivity (Ijima et al., 2005; Terao et al., 2006a) and introduction of a racemase activity (Terao et al., 2006b). Yet, the exact mechanism for stabilization of the highly unstable enediolate intermediate remained elusive. While other enzymes with related enolate intermediates use Mg²⁺ for stabilization (Gerlt et al., 2005), activity of AMDase is not dependent on metal ions or other cofactors (Miyamoto and Ohta, 1992b). Further, delocalization of electron density into the aromatic ring system might partly account for stabilization of the enediolate but would be insufficient to explain the observed highly efficient enzymatic decarboxylation (Okrasa et al., 2008).

By solving the first crystal structure of wildtype AMDase (PDB: 3DG9), Okrasa et al. were able to unravel key features of enzyme activity and selectivity in more detail. Interestingly, in the core structure consisting of two four-stranded parallel β-sheets surrounded by several α-helices, a tightly bound phosphate ion was found near the active site cysteine 188. A total of six hydrogen bonds were established by side-chain and backbone interactions to Thr75, Ser76, Tyr126, Cys188 and Gly189, belonging to two adjacent oxyanion holes (Okrasa et al., 2008). The authors suggested that the phosphate might resemble the position of the enediolate intermediate. In retrospective, this structural motif termed “dioxyanion hole” was also found in related enzymes of the isomerase family and mutational studies on Thr75 and Ser76 or Tyr126 of AMDase have already demonstrated the essential role of this region for enzyme activity (Terao et al., 2007; Okrasa et al., 2008).

After reconfirming loss of the pro-(R) carboxylate by a ¹⁸O-labelling strategy, the enediolate intermediate resulting from decarboxylation of α-methyl-α-phenylmalonic acid 2a was placed in the active site. It became obvious, that the phenyl moiety occupies a large solvent accessible pocket, whereas the small methyl substituent is left at an orientation, where only little space is available. Considering these restrictions and the fixed position of the pro-(S) carboxylate entrapped in the dioxyanion hole, it was suggested that the pro-(R) carboxylate must point towards a small hydrophobic pocket (Leu40, Val43, Val156, Tyr48), which could act as a driving force for decarboxylation by hydrophobic destabilization (Figure 2B). Thus, the key for cofactor-free decarboxylation by AMDase was proposed to be the extensive stabilization of one carboxylate, while the other one is destabilized by unfavorable electrostatic interactions (Okrasa et al., 2008).

This proposed binding mode was confirmed when AMDase was co-crystallized with the mechanism-based inhibitor benzylphosphonate (K _i = 5.2 mM) (PDB: 3IP8) (Okrasa et al., 2009). Similarly, the phosphonate dianion was tightly engaged by six hydrogen bonds to the residues of the dioxyanion hole and the phenyl residue was positioned in the large pocket stacked within the Gly189-Gly190 amide bond and Pro14 through van der Waals interactions.

Obata et al. solved the crystal structure of the G74C/C188S variant containing the ligand 2-phenyl acetate (PDB: 3IXL, Figure 3A) and of the G74C variant (PDB: 3IXM, Figure 3C) and WT enzyme (PDB: 3DTV, Figure 3D) in a sulfate associated form (Obata and Nakasako, 2010). While the overall structure of AMDase G74C/C188S was mostly preserved when compared to wildtype structures either containing BnzPO (PDB: 3IP8, Figure 3A) or PO₄ ³⁻ (PDB: 3DG9) in the active site (RMSD of C_α atoms approx. 0.25 Å), considerable conformational differences became obvious by comparing to the SO₄ ²⁻-associated structures of the wildtype and G74C variant (PDB: 3DTV and 3IXM). It was suggested that a positional shift of Cys188-Gly189-Gly190 towards Gly74-Thr75 was induced upon ligand binding, thereby triggering the formation of a hydrophobic network covering the active site (Val43, Thr154, Val156). In the unliganded (SO₄ ²⁻-associated) crystal structures, such cluster was not observed, as the corresponding loop regions are not contacting each other. Additionally, in this conformation, the key residue 188 is rotated away from the ligand towards the hydrophobic pocket in a rather “non-productive” orientation (Figures 3C,D). These observations indicated the presence of either an “open” or “closed” conformation, regulated by ligand binding (Figure 3) (Obata and Nakasako, 2010). A similar behavior between structures of the empty and the ligand-bound enzyme was found for the related glutamate racemase (Puig et al., 2009).

Most important, however, was the observation that the position of Cys74 of the G74C/C188S variant was indeed in mirror symmetry to C188 of AMDase WT with regards to the C_α atom of the enediolate intermediate, which gets protonated during the reaction, thereby confirming the early proposed rationale for inversion of enantioselectivity (Ijima et al., 2005; Terao et al., 2007). In this binding mode, configuration of the final product is only dependent on the enantioface-selective protonation, which occurs from si-face in case of Cys188 or from re-face in case of Cys74, resulting in (R)- or (S)-2-phenylpropionate, respectively in accordance to experimental data (Miyauchi et al., 2011).

The reaction mechanism of AMDase was also subject of computational studies. Lind and Himo studied the mechanism by means of density functional theory (DFT) calculations (Lind and Himo, 2014). By employing the quantum chemical cluster approach, the mechanism, stationary points thereof and enantioselectivity were investigated with the aid of two different truncated active site models. The smaller model only partly contained the residues of the dioxyanion hole and the catalytic Cys188 (81 atoms), whereas the second model additionally included residues from the small and large binding pockets (223 atoms). Results obtained from using the small model indicated that a two-step mechanism proceeding via a planar enediolate intermediate is plausible, with the decarboxylation step being rate-limiting. However, binding modes leading either to cleavage of the pro-(R) or pro-(S) carboxylate were very similar in energy (1.9 kcal mol⁻¹ for initial binding pose; 1.5 kcal mol⁻¹ for transition state) probably due to the absence of the binding pockets determining the orientation of the remaining substituents, therefore being unable to explain the observed enantioselectivity with this small model (>99 %ee corresponds to at least 3 kcal mol⁻¹). By using the larger model, the calculated energies for binding and decarboxylation were 14.1 and 18.3 kcal mol⁻¹ higher for the formation of the (S)-enantiomer than for the (R)-product, mostly caused by unfavorable steric clashes of the large aryl substituent and the residues of the small binding pocket. When the much smaller α-methyl-α-vinylmalonic acid was studied, ambiguous results were obtained concerning absolute transition state energies, probably being a result of the too small and rigid models of the binding pockets. Both binding modes and transition states, however, were much closer in energy as compared to α-methyl-α-phenylmalonic acid 2a. This would indicate that stereoselectivity is exclusively determined by substrate binding in case of bulky aromatic compounds, whereas in case of smaller substrates, the decarboxylation transition state could also contribute to enantiodiscrimination (Lind and Himo, 2014). Just recently, Dasgupta et al. emphasized that geometric constraints such as introduced in quantum-chemical studies can lead to artifacts like imaginary vibrational frequencies and impaired efficiency of the overall optimization process (Dasgupta and Herbert, 2020). The authors thus introduced soft harmonic confining potentials to the terminal atoms of the model, thereby avoiding the artificial strain and rigidity of fixed-atom truncated active site models. By employing this system, they were able to reproduce the results from previous studies (Lind and Himo, 2014) and further claimed that this methodology is easy to implement and can dramatically reduce optimization efforts (Dasgupta and Herbert, 2020).

In the previously described modeling approaches, results were substantially dependent on the size of the truncated enzyme models used (Lind and Himo, 2014; Dasgupta and Herbert, 2020). In order to obtain a more comprehensive overview of the molecular level origin of AMDase selectivity, a full enzyme model would be highly preferable. Particularly, counterintuitive effects observed after iterative saturation mutagenesis within the AMDase active site cavity (Okrasa et al., 2009; Miyauchi et al., 2011; Yoshida et al., 2015) require consideration of the entire first coordination sphere. Thus, the empirical valence bond (EVB) approach was the methodology of our choice, which was further complemented by metadynamics simulations (Warshel and Weiss, 1980; Barducci et al., 2008; Biler et al., 2020). EVB simulations were able to reproduce experimentally observed activation energy barriers (within 3 kcal mol⁻¹) and the preferential cleavage of the pro-(R) carboxylate group which was already determined during substrate positioning in the Michaelis complex. Yet curiously, the (S)-selective CLGIPL variant showed preferential cleavage of the pro-(S) group which still led to the expected formation of (S)-enantiomers due to reacting via a different substrate binding pose. For compounds, which were not or only poorly converted by AMDase and variants thereof, an increased flexibility and motility within the active site was observed during simulations. The resulting inadequate substrate positioning furthermore allowed water to enter the active center, which is likely to destroy the vulnerable interaction network vital for catalysis (Biler et al., 2020). By consulting the grid inhomogeneous solvation theory (GIST) (Nguyen et al., 2012; Nguyen et al., 2014) to analyze the local hydrophobicity within the AMDase active site, we found clear evidence for a mechanism driven by ground-state destabilization by entrapment of the carboxylate to-be-cleaved in a hydrophobic environment. These analyses also revealed the emergence of an additional hydrophobic cavity in the CLGIPL variant, which allows this variant to react via an alternative binding pose and unexpected cleavage of the pro-(S) carboxylate. While the alteration of hydrophobic pockets in the CLGIPL variant was not by design, the targeted engineering of the active site hydrophobicity presents a seminal approach for future engineering of AMDase. WT-MetaD simulations further confirmed that the reactive binding pose (according to EVB simulations) also coincides with the most populated binding pose observed at the Michaelis complex in most of the studied cases. Thus, AMDase selectivity is partly already determined at the stage of substrate binding, but selective destabilization of a distinct carboxylate group (leading to ground-state destabilization) seems to be the true determinant of the eventual reactive transition state (Biler et al., 2020).

Busch et al. used semiempirical QM/MM calculations to study the mechanism of the AMDase racemase variant G74C (Busch et al., 2016). MD simulations revealed that both catalytic cysteines need to be in their deprotonated state for successful catalysis. In contrast to glutamate racemase, where co-catalytic residues activate the cysteines (Glavas and Tanner, 2001), water is suggested to deprotonate the corresponding residues of AMDase G74C prior to substrate binding, which are then further stabilized by thiolate holes. This might also explain the pronounced pH-dependency of the reaction. Further semiempirical calculations indicated a stepwise mechanism similar to decarboxylation, where a delocalized π-electron system within the substrate is necessary to stabilize the enediolate intermediate (Busch et al., 2016). Likewise to previous studies (Lind and Himo, 2014), there was no indication for a concerted mechanism as described for glutamate racemase (Puig et al., 2009).

Another study conducted by Karmakar et al. aimed at the simulation of CO₂ and product release from the active site (Karmakar and Balasubramanian, 2016). By performing (steered) MD simulations it was demonstrated that release of the decarboxylated product is energetically unfavorable and might even surpass the decarboxylation step, thus potentially rendering product release the true rate-limiting step (WT: ∼14 kcal mol⁻¹ for decarboxylation vs. ∼23 kcal mol⁻¹ for product release). The authors claimed that this might also partly account for the reduced activity of the G188S/G74C variant (20,000-fold) (Miyauchi et al., 2011), as the calculated energy barrier for product release was even higher in this case (∼20 kcal mol⁻¹ for decarboxylation vs. ∼37 kcal mol⁻¹ for product release) (Karmakar and Balasubramanian, 2016).

Already after the discovery of AMDase, Miyamoto and Ohta thoroughly characterized the enzyme and its substrate scope (Miyamoto and Ohta, 1992b). These fundamental findings, obtained in absence of any structural and mechanistic knowledge on the enzyme, provided the basis for studies up to the present by demonstrating the essential limitations of AMDase substrates. 1) Steric effects play a crucial role for the small substituent, as mono-substituted malonates were converted faster than the corresponding methylated substrates. Further, the ethyl analogue was not converted. 2) The aromatic substituent is essential for activity, as substrates with a benzyl-, phenoxy- or phenylthio- instead of an phenyl-group were inert (Miyamoto Ohta and, Hiromichi, 1990). This was also true for malonic acid (R^L = H) or methylmalonic acid (R^L = CH₃). 3) Only free malonic acids can act as a substrate, as the corresponding mono- and diesters were not accepted. 4) Electron-withdrawing substituents on the aromatic moiety enhance the reaction rate by better stabilizing the proposed carbanion intermediate. Table 1 summarizes substrates studied in AMDase catalyzed decarboxylation and racemization.

The large substituent R^L offers major variability within the substrate scaffold. Phenyl- (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992b; Miyamoto et al., 1992a; Miyamoto et al., 1994; Miyazaki et al., 1997; Fukuyama et al., 1999; Matoishi et al., 2000; Terao et al., 2006b; Terao et al., 2007; Okrasa et al., 2008; Okrasa et al., 2009; Tamura et al., 2008; Kourist et al., 2011b; Miyauchi et al., 2011; Yoshida et al., 2015; Lewin et al., 2015), 2-naphthyl- (Miyamoto and Ohta, 1992b; Ijima et al., 2005; Terao et al., 2006a; Terao et al., 2006b; Terao et al., 2007; Miyauchi et al., 2011) and thienyl- (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992b; Ijima et al., 2005; Terao et al., 2006a; Terao et al., 2006b; Terao et al., 2007; Lewin et al., 2015) malonic acids are typical substrates of AMDase and omnipresent in literature. Over time, the substrate scope was extended to a vast number of variously substituted phenyl derivatives (o-Me (Miyamoto Ohta and, Hiromichi, 1990), p-Me (Miyamoto and Ohta, 1992b; Kourist et al., 2011b), p-iso-butyl (Yoshida et al., 2015), p-NO₂ (Kourist et al., 2011b), OMe (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992b; Kourist et al., 2011b), F (Miyamoto et al., 1992a; Miyamoto and Ohta, 1992b), Cl (Miyamoto Ohta and, Hiromichi, 1990; Miyamoto and Ohta, 1992b; Miyamoto et al., 1994), CF₃ (Miyamoto et al., 1992a), p-Ph-m-F (Kourist et al., 2011b; Gaßmeyer et al., 2016), m-COPh (Kourist et al., 2011b)), naphthyl derivatives (1-naphthyl (Miyamoto and Ohta, 1992b), 6-methoxynaphthalen-2-yl (Miyamoto Ohta and, Hiromichi, 1990; Kourist et al., 2011b; Miyauchi et al., 2011; Yoshida et al., 2015; Gaßmeyer et al., 2016)) heterocyclic (pyridinyl (Kourist et al., 2011b; Lewin et al., 2015), furanyl and bicyclic systems (Lewin et al., 2015)) and also alkenyl (Okrasa et al., 2009; Kourist et al., 2011b) substrates.

With regards to the small substituent R^S, it was soon recognized that, besides H and CH₃, other small groups like D (Matoishi et al., 2000), F (Miyamoto et al., 1992a), OH or NH₂ (Tamura et al., 2008) were generally tolerated, while larger alkyl groups (ethyl (Miyamoto and Ohta, 1992b) or propyl (Terao et al., 2006b; Kourist et al., 2011b)) were not or only converted at a very low rate (Terao et al., 2006b; Kourist et al., 2011b). This was later on explained by structural analysis, as only limited space is available at the position left for this group (Okrasa et al., 2008; Obata and Nakasako, 2010).

As already exposed by Miyamoto et al. in 1992, presence of the free di-acid is indispensable for decarboxylation to proceed (Miyamoto and Ohta, 1992b). This was also proven valid for the racemization activity of AMDase, as carboxylate derivatives such as alcohols, amides, nitriles (Terao et al., 2006b), esters (Terao et al., 2006b; Kourist et al., 2011b), thioesters, ketones and aldehydes (Kourist et al., 2011b) were completely inactive. Thus, stabilization of the free carboxylate via the complex network within the dioxyanion hole seems crucial, regardless of decarboxylation or racemization. The only exceptional case known so far is (nitromethyl)benzene, which was accepted as a substrate by AMDase racemase G74C and G74C/V43A (Kourist et al., 2011b). It was generally concluded that the substrate structure seems more decisive for successful conversion, than α-proton acidity of the α-aryl propionates (Kourist et al., 2011b).

In terms of spatial arrangement, Miyamoto et al. proposed that a co-planar alignment of the phenyl group and α-substituent would allow optimal overlap of the π-orbitals in order to stabilize the formed charge on the enolate intermediate (Miyamoto et al., 1994). In case of ortho-substituted substrates like α-(o-chlorophenyl)malonic acid, two such conformations come into consideration, namely a syn- and anti-periplanar arrangement (Figures 4A,B).

Kinetic studies revealed that, while the activity of AMDase towards α-(o-chlorophenyl)malonic acid even surpassed α-phenylmalonic acid 1a, the corresponding α-methyl-substituted compound was not accepted at all. This led to the assumption, that a syn-periplanar arrangement is necessary for AMDase-mediated decarboxylation, which might in turn disfavor substrates with an additional α-substituent, due to steric clashes with the ortho-substituent (Miyamoto et al., 1994). Ab initio calculations were thus performed to assess the energy barriers encountered during rotation along the C_α-C_Ar bond. Results obtained for α-(o-chlorophenyl)malonic acid indicated two stable conformations corresponding to a syn- or anti-periplanar orientation (approx. 0° or 360° and 180° respectively). The small energy difference between both structures (<1 kcal mol⁻¹) implies, that steric repulsion between o-Cl and α-H is negligible. Repulsive contributions were mainly caused by interaction of the chlorine residue with the carboxylates (Figure 4B). In case of α-(o-chlorophenyl)-α-methylmalonic acid, an anti-periplanar or perpendicular arrangement of substituents was energetically favored, whereas syn-periplanar-like conformations were found to be unstable (Figure 4B).

It was further proposed that by using the conformationally restricted indane-1,1-dicarboxylic acid (IDA), a syn-periplanar conformation would be mimicked, thereby lowering the activation entropy ΔS^‡ (Figure 4A). Indeed, the model substrate IDA was accepted by AMDase in contrast to the corresponding α-methyl-α-(o-tolyl)malonic acid, thus underlining the inability of the latter to overcome rotational energy barriers towards a suitable syn-periplanar arrangement for conversion (Miyamoto et al., 1994). The remarkably low K _M value observed for IDA in comparison to other studied substrates further affirmed the suitability of such spatial arrangement for AMDase-catalyzed decarboxylation (Miyamoto et al., 1994; Kawasaki et al., 1996). In this context it was also proposed that a hydrophobic pocket within the active site might be responsible for correct positioning of unrestricted substrates via CH-π interactions (Nishio et al., 1995), thereby reducing the activation entropy in a similar manner (Kawasaki et al., 1996).

In view of the dramatically reduced efficiency of the initially created variants with inversed (Ijima et al., 2005) or racemizing activity (Terao et al., 2006b) in comparison to the native enzyme, considerable efforts to recover or even to increase the activity by enzyme engineering have been made until today.

In search of organisms with similar reactivities and characteristics as AMDase from Alcaligenes bronchisepticus (now: Bordetella bronchiseptica) KU1201, Ohta and co-workers were able to identify two further strains from soil samples by their ability to degrade α-phenylmalonic acid 1a (Miyamoto et al., 2007b; Yatake et al., 2008). The strains KU1311 (Miyamoto et al., 2007b) and KU1313 (Yatake et al., 2008) were the most active ones and identified as Achromobacter sp. and Enterobacter cloacae, respectively. Respective genes were amplified from the genomic DNA and heterologously produced in E. coli for characterization. Both genes encoded a protein of 240 amino acids length, consistent with the originally identified AMDase from strain KU1201 (Miyamoto and Ohta, 1992a) and both shared a high sequence homology of 94% (KU1311) and 85% (KU1313). Accordingly, the observed substrate selectivity and activity were comparable to the original AMDase and all shared the same strict enantiopreference (Miyamoto and Ohta, 1992a, 1992b; Miyamoto et al., 2007b; Yatake et al., 2008).

While those three AMDases show considerable similarity, the shifted pH optimum (pH 5.5) towards acidic conditions of AMDase from strain KU1313 was intriguing. It was the only microorganism, which showed AMDase activity when soil samples were screened at acidic pH, others were rather active at neutral to slight basic pH (pH 7–8.5) (Table 7).

Okrasa et al. tried to identify novel AMDase enzymes from protein sequences with a similarity of 30-52% and a requisite cysteine residue at the same relative position as in the enzyme from B. bronchiseptica. The putative AMDases were characterized, however only the enzyme from Mesorhizobium sp. (now: Chelativorans sp.) BNC1, exhibiting the highest similarity, showed satisfactory decarboxylase activity, while the other candidates rather acted as racemases (Okrasa et al., 2008).

The difficulty of identifying novel AMDases accompanied by the issue of inappropriate gene annotation was addressed by the work of Maimanakos et al., who developed a sequence-based search algorithm for a more reliable prediction of enzymes possessing AMDase activity (Maimanakos et al., 2016). Sequence information of confirmed AMDases was used to specify 12 conserved sequence patterns (e.g. residues of the hydrophobic pocket, dioxyanion hole, aryl binding pocket and residues C188 and G74 including their surroundings), which, in turn, were used for concise database screening. In this way, 58 additional ORFs encoding putative AMDase-like enzymes were found and used for generating a Hidden Markov Model (HMM), which furnished six specific motifs (Table 8) and was implemented in the applied search algorithm. To prove the reliability and applicability of these search criteria, the enzyme from Polymorphum gilvum SL003B-26A1 identified via data base search was characterized together with two enzymes from Variovorax sp. HH01 and HH02 identified via screening of soil samples. All three enzymes showed the desired AMDase activity within common mesophilic temperature and around neutral to slightly acidic pH (Table 7) (Maimanakos et al., 2016).

Interestingly, all putative AMDase-encoding ORFs identified by Maimanakos et al. belonged to the class of α- β- and γ-proteobacteria. Interestingly, when AMDase flanking regions were analyzed, genes encoding mandelate racemase/muconate lactonizing enzyme or several transporters (e.g. tripartite tricarboxylate transporter TTT, TRAP or ABC transporters) were commonly found. According to these flanking regions and sequence similarities, AMDases were classified into eight enzyme clusters. While the natural role of this enzyme still remained elusive, the fact, that close relatives of AMDase-producing bacteria often do not encode this enzyme, strongly indicated, that this gene was frequently lost in evolution and might be therefore only temporarily advantageous for organisms under certain physiologic conditions (Maimanakos et al., 2016).

30 years after its discovery, AMDase has arrived at the stage of industrial applications. The history of research on this unique decarboxylase is exemplary for the technological progress in biocatalysis within the last decades. After AMDase discovery, based on a purely functional screen, the first important milestone achieved by the Ohta group was the identification of the corresponding gene, which allowed recombinant production of the enzyme. Enzymological characterization and studies with isotope-labelled pseudochiral malonic acids allowed initial mechanistic insights: During the reaction, only one carboxylate group is cleaved, and this step is not directly determining the stereooutcome of the reaction. Based on homology models, the enantioselectivity was completely inverted, and a promiscuous, unique profen racemase was generated. Yet, elucidation of the crystal structure of AMDase in its liganded form in the early 2010s was groundbreaking for the formulation of a hypothetical decarboxylation mechanism, which was confirmed by several computational studies in recent years. Despite the availability of several crystal structures, it remains exceedingly difficult to accurately predict the influence of specific amino acid substitutions in the hydrophobic pocket on enzyme activity. In lieu of predictability, site-saturation mutagenesis and simultaneous saturation mutagenesis proved to be efficient methods to increase the activity of enzyme variants in the synthesis of both enantiomers of α-aryl propionic acids, and to identify optimal variants for different substrates.

The tendency of α-aryl α-methylmalonic acids to undergo spontaneous decarboxylation proved to be an obstacle for industrial application, particularly regarding the decarboxylation of substrates with large electron-poor substituents. By developing a carefully balanced deprotection strategy for the corresponding malonic acid esters, conditions leading to spontaneous decarboxylation could be avoided. A telescoped deprotection/decarboxylation approach resulted in a robust, practicable reaction that allows to obtain α-aryl propionates in either their (S)- or (R)-form in outstanding optical purity. While the activity of the enzyme is generally high, immobilization greatly increased the stability, resulting in an excellent productivity.

Starting with a completely unknown enzyme with unknown mechanism that produced the “wrong” enantiomer of profens, a combination of biocatalysis, enzymology, structure elucidation, molecular modeling, protein engineering, organic chemistry and process intensification succeeded to create a powerful toolbox of AMDase variants that provide access to a large diversity of α-substituted propionates in their enantiopure form.