l-2-hydroxyglutarate (l-2HG) can be obtained by hydroxylation of the C₅-dicarboxylic acid glutarate at the α-carbon position. Glutarate is known to be a demanded building block for the production of biopolymers, like aliphatic polyamide 6,5 which is mainly used in the construction industry (Navarro et al., 1997). Carbon chain length and functional groups of the monomers are important for the performance of the biopolymer with respect to physical and chemical properties. In the case of l-2HG, the additional hydroxyl group provides greater versatility for further chemical modification with the potential to alter the properties of the resulting polymer. Furthermore, incorporation of hydroxylated monomers enhances the biodegradability of the polyamide (Varela and Orgueira, 2000). Production of l-2HG by microbial fermentation would provide the chemical industry with an environmentally friendly building block for polyamides and polyesters. The addition of functionalities on polyester backbones (hydroxyl, carboxyl, allyl, azido, and acetylene groups) facilitates covalent post-polymerization modification. In 2017, it was shown that the enantiomer d-2-hydroxyglutarate (d-2HG) can be used as a functionalized building block for a polyester after cyclization to an allyl containing lactide (Nan and Feher, 2017). Also, it was demonstrated that electrochromic polyamides with functional hydroxyl groups for homogeneous hybrid films can be produced. The repeating units of hydroxysuccinate in the polymer backbone provided reaction sites for organic-inorganic bonding resulting in homogeneous and transparent hybrid films (Pan et al., 2018).

In medicine, l-2HG is used as a diagnostic biomarker for the characterization of various cancer types (Shim et al., 2014). Under oxygen limitation, l-2HG accumulates as metabolic intermediate in healthy as well as in malignant animal cells (Shim et al., 2014; Oldham et al., 2015; Shelar et al., 2018) facilitating a physiological adaption to hypoxic stress. The accumulation of l-2HG is triggered by increasing concentrations of 2-oxoglutarate (2-OG), which is caused by tricarboxylic acid cycle dysfunction and increased mitochondrial reducing potential. As consequence, the increased cellular l-2HG concentrations lead to inhibition of the electron transport and glycolysis compensating the effects of mitochondrial reductive stress induced by hypoxia (Oldham et al., 2015). The stereospecific reduction of 2-oxoglutarate to l-2HG is catalyzed by lactate dehydrogenase (LDH) and malate dehydrogenase (MDH) under hypoxic stress conditions (Intlekofer et al., 2017). In contrast, the formation of its enantiomer d-2-hydroxyglutarate (dr-2HG) is catalyzed by a mutated version of isocitrate dehydrogenase 1 or 2 (IDH1/2) contributing to the pathogenesis of cancer, whereas l-2HG biosynthesis does neither involve IDH1 nor IDH2. Albeit both enantiomers of 2-hydroxyglutarate display an inhibitory effect on 2-oxoglutarate-dependent enzymes involved in diverse biologic processes (Chowdhury et al., 2011; Xu et al., 2011), their stereospecific biosynthesis differs. Thus, bio-based routes enable stereospecific synthesis of either d-2HG or l-2HG, which is preferred over chemically synthesized racemic d,l-2HG. Beyond the occurrence as a product of cellular redox stress, l-2HG also plays a role in plants and eukaryotic cells as part of the mitochondrial metabolic repair mechanism (Hüdig et al., 2015). A side reaction of MDH yields small concentrations of l-2HG by reduction of 2-oxoglutarate. The mitochondrial FAD-containing oxidase l-2-hydroxyglutarate dehydrogenase (L2HGDH, EC 1.1.5.13) oxidizes l-2HG and the electrons produced in the reaction are transferred to the mitochondrial electron transport chain through the electron transfer protein (ETF) (Wanders et al., 1997; Rzem et al., 2004). An enzyme homologous to L2HGDH was also described in the model organism Escherichia coli (Knorr et al., 2018).

In bacteria, l-2HG mainly accumulates under carbon starvation conditions (Marschall et al., 1998; Metzner et al., 2004; Knorr et al., 2018). It is also an intermediate of the l-lysine degradation pathway in Pseudomonadaceae (Zhang et al., 2018; Thompson et al., 2019). Glutarate is the direct precursor of l-2HG. Hydroxylation of glutarate to l-2HG is catalyzed by a highly regio- and stereospecific Fe(II)/2-oxoglutarate-dependent dioxygenase CsiD (EC 1.14.11.64, also named glutarate hydroxylase) (Hibi and Ogawa, 2014). The co-product succinate is metabolized via the TCA-cycle. Since l-2-hydroxyglutarate oxidase (LghO) regenerates 2-oxoglutarate by oxidation of l-2HG (Knorr et al., 2018; Herr et al., 2019), the combined activities of CsiD and LghO convert the C5-dicarboxylic acid glutarate to the C4-dicarboxylic acid succinate. Thus, the absence of LghO is required for overproduction of l-2HG from glutarate.

Glutarate can be derived from l-lysine by four pathways that converge to 5AVA, which is converted to glutarate by GABA/5AVA aminotransferase (GabT) and the succinate/glutarate semialdehyde dehydrogenase (GabD). The first pathway from l-lysine to 5AVA employs l-lysine-α-oxidase (RaiP) from Scomber japonicus that catalyzes oxidative desamination of l-lysine using molecular oxygen followed by spontaneous decarboxylation (Cheng et al., 2020). The second pathway to 5AVA combines oxidative decarboxylation by l-lysine monooxygenase using molecular oxygen followed by desamidation by γ-aminovaleramidase from P. putida (Adkins et al., 2013). The third pathway is based on l-lysine decarboxylase from E. coli, putrescine oxidase PuO from Rhodococcus qingshengii and the γ-aminobutyraldehyde dehydrogenase from E. coli that catalyze decarboxylation, oxidative deamination using molecular oxygen and NAD-dependent oxidation (Haupka et al., 2020). The fourth pathway does not require molecular oxygen as it cascades l-lysine decarboxylase, 2-oxoglurate-dependent putrescine/cadaverine transaminase PatA, and NAD-dependent γ- aminobutyraldehyde dehydrogenase PatD from E. coli (Jorge et al., 2017). The pathway combinations for LdcC-PuO-PatD-GabT-GabD and LdcC-PatA-PatD-GabT-GabD couple conversion of lysine to glutarate in one or two transaminase reactions, respectively, that generate glutamate from 2-oxoglutarate. This coupling enabled flux enforcement, i.e., growth requires production of glutarate, which was achieved by deletion of gdh, the gene for the major ammonium assimilating enzyme l-glutamic acid dehydrogenase (Pérez-García et al., 2018; Haupka et al., 2020).

Since biosynthesis of l-2HG from glutarate requires molecular oxygen, glutarate biosynthesis from l-lysine by a pathway independent of molecular oxygen was extended by glutarate hydroxylase. The pathways RaiP-GabT-GabD, DavA-DavB-GabT-GabD, and LdcC-PuO-PatD-GabT-GabD involve oxygenases (RaiP, DavA, and PuO, respectively), whereas the pathway LdcC-PatA-PatD-GabT-GabD does not contain an enzyme using molecular oxygen as substrate (Jorge et al., 2017). To construct the six-step cascade LdcC-PatA-PatD-GabT-GabD-CsiD (Figure 1), CsiD enzymes from E. coli MG1655, P. putida KT2440 and Halobacillus sp. BA-2008 were tested since inspection of the genome sequence of C. glutamicum did not indicate that a CsiD homolog is encoded. The absence of LhgO from the C. glutamicum genome was considered beneficial since it indicated that l-2HG may not be degraded. This and the fact that we previously established efficient glutarate production employing the LdcC-PatA-PatD-GabT-GabD pathway provided an ideal starting point to establish fermentative production of l-2HG from renewable resources. This concept may be transferred to fermentative production of the stereoisomer d-2HG by extending the glutarate pathway with enzymes for conversion of glutarate to d-2HG.

In this study, production of the sought after compound l-2HG by the industrial workhorse C. glutamicum was demonstrated after extension of lysine biosynthesis in a six-step cascade employing the metalloenzyme CsiD from P. putida as final step. The glucose-based process was stable in 2 L bioreactor cultivations and a l-2HG titer of 3.5 g L⁻¹ was obtained in fed-batch fermentation. Moreover, l-2HG production based on the renewable feedstock wheat sidestream concentrate was demonstrated.

l-2HG production was achieved by glutarate hydroxylase extending lysine biosynthesis in a six-step cascade, in which only the last committed step catalyzed by oxygen-dependent glutarate hydroxylase requires oxygen. Glutarate hydroxylases belong to the large family of non-heme Fe(II)- and 2-oxoglutarate-dependendent oxygenases, which are essential for diverse biological functions. These enzymes form an Fe(IV)-oxo intermediate to initiate oxidative transformations and can be assigned to four major types of reactions: hydroxylation, halogenation, ring formation and desaturation (Hausinger, 2004). The glutarate hydroxylases which have been studied in this research belong to the enzymes catalyzing a hydroxylation reaction at an unactivated carbon center by incorporation of molecular oxygen (Guengerich, 2015; Martinez and Hausinger, 2015). The CsiD enzymes from E. coli MG1655 and P. putida KT2440 have been recently characterized in different studies displaying their high specificities toward the native substrate glutarate with K_M-values around 0.65 mM in E. coli (Knorr et al., 2018) and around 0.15 mM for P. putida (Zhang et al., 2018). The K_M-values of around 0.1 mM for the co-substrate 2-oxoglutarate are comparable for both enzymes. The higher affinity of glutarate hydroxylase from P. putida for glutarate is reflected by the better conversion of glutarate to l-2HG observed in this study. Even though no K_M value for the glutarate hydroxylase from Halobacillus sp. BA-2008 has been determined yet, it was demonstrated that it converts 5 mM glutarate to l-2HG with comparable efficiency to the other tested hydroxylases with 2-oxoglutarate as co-substrate (Thompson et al., 2019). Although the capability of CsiD from Halobacillus sp. BA-2008 to produce l-2HG from glutarate was demonstrated in C. glutamicum, the efficiency of the codon harmonized version of CsiD was inferior compared to the enzymes derived from other organisms.

It could be demonstrated that l-2HG inhibits the combined GabTD activity with an inhibitory constant of about 25 mM (Figure 5A). Due to its structural similarity to 2-OG, l-2HG potentially inhibits transamination reactions (McBrayer et al., 2018) by competitive inhibition (Intlekofer et al., 2015). Here, possibly competitive inhibition of 2-OG dependent transaminase GabT by l-2HG may have limited l-2HG product titers. Potentially, this may be overcome by enzyme engineering of the 2-OG binding pocket of GabT for better differentiation of this substrate from l-2HG. We have chosen enzymes GabT and GabD from P. stutzeri as they performed better than those from C. glutamicum, P. putida and P. syringae regarding glutarate production (Pérez-García et al., 2018). Since we did not compare GabT and GabD enzymes from various sources for inhibition of their activities by l-2HG, identifying feedback resistant enzymes/variants of the GabT and GabD enzymes in future work may help to increase l-2HG production. On the other hand, glutarate hydroxylase CsiD is subject to weak product inhibition (Knorr et al., 2018). Therefore, CsiD from other sources or variants may be selected that exhibit no or reduced product inhibition in order to improve l-2HG production.

Moreover, a weak substrate inhibition on l-2HG production by glutarate could also be identified (Figure 5B). CsiD from E. coli was recently characterized and the reaction mechanism was described (Knorr et al., 2018; Herr et al., 2019). Since the substrate analogon N-oxalylglycine (NOG), a 2-OG mimic, inhibited CsiD (Knorr et al., 2018), this feedback inhibition may limit l-2HG production. However, since we did not assay feedback inhibition of various glutarate hydroxylases by l-2HG, future work to improve l-2HG production should involve the identification of feedback resistant CsiD enzymes/variants.

Surprisingly, extracellular addition of glutarate boosted glutarate production. This is unlikely due to effects on enzyme activities. C. glutamicum possesses chromosomal copies of gabT and gabD (Pérez-García et al., 2018; Haupka et al., 2020). The PucR-like regulator GabR that requires GABA as coactivator activates transcription of the gabTDP operon (Zhu et al., 2020). Possibly, glutarate mimics GABA as coactivator of GabR activating the endogenous gabTDP operon and increasing GabT and GabD enzyme levels in addition to plasmid-borne expression of gabT and gabD from P. stutzeri.

Cultivation schemes with higher oxygen supply reduced l-2HG production, but improved growth to high biomass concentrations. Glutarate hydroxylases are 2-OG-dependent enzymes and these metalloenzymes specifically require Fe²⁺ as cofactor for their function (Mitchell et al., 2017; Dunham et al., 2018). Previously, it has been shown that increased iron concentrations improve enzyme activity (Fukumori and Hausinger, 1993). Increased Fe²⁺ concentrations improved l-2HG product titers, but slowed growth significantly. This might be due to iron effects such as toxicity mainly due to the formation of hydroxyl radicals as oxidative stress response (Braun, 1997; Touati, 2000; Eid et al., 2017) and/or due to inhibition of 2-OG-dependent dioxygenases and methylases, which play an essential role in DNA/RNA repair (van den Born et al., 2009), by l-2HG (Low et al., 2001; Ledesma-García et al., 2016). Provision of oxygen and iron to the C. glutamicum cell are interlinked. Molecular oxygen is required for 2-OG-dependent dioxygenases such as CsiD to form the Fe(IV)-oxo intermediate, but with too much oxygen decoupling occurs via “non-productive reactions” resulting in inactive Fe(III)-containing enzymes (Hausinger, 2004) as observed with AlkB (Henshaw et al., 2004) and TfdA (Liu et al., 2001). Secondly, an “uncoupled turnover” of the co-substrate 2-OG may occur by decomposition of 2-OG to carbon dioxide and succinate. These “uncoupled reactions” might contribute to the lower titers of l-2HG obtained in 2 L scale fermentative production with the highest oxygen supply. The addition of ascorbic acid is a promising option since ascorbate improved hydroxylase activity (Fukumori and Hausinger, 1993). Alternatively, production of l-2HG may be improved by concomitant overproduction of either glutathione (Liu et al., 2019) or ascorbate in C. glutamicum.

Our results suggested that export of l-2HG involves SucE (Figure 3). Surprisingly, glutarate export does not depend solely on SucE. Overexpression of sucE increased succinate production (Zhu et al., 2014) similar to increased production of glutarate and l-2HG observed here. Deletion of sucE revealed that SucE is the main export system for l-2HG, but not for glutarate. Production of l-2HG may benefit from re-uptake of glutarate secreted to the culture medium as by-product. However, the uptake system for glutarate is unknown. MctC is involved in the uptake of pyruvate, propionate, and acetate, but it is not clear if it accepts glutarate (Jolkver et al., 2009).

Glutarate was observed as significant by-product. Unlike l-2HG, which likely is exported by SucE (s. above), glutarate is exported by YnfM (Fukui et al., 2019). Overexpression of ynfM improved production of glutarate, succinate, and 2-OG (Fukui et al., 2019; Han et al., 2020). Thus, deletion of ynfM is a suitable strategy to abolish export of glutarate as by-product of l-2HG production.

Alternative feedstocks are important to achieve sustainable biotechnological processes. In this respect, sidestreams are highly relevant. Glycerol is a sidestream of the biodiesel process and glycerol accumulates as stoichiometric by-product. C. glutamicum has been engineered to produce amino acids from pure and technical grade glycerol (Rittmann et al., 2008; Meiswinkel et al., 2013b). Access to nitrogenous sidestreams from the fishery industry such as glucosamine or N-acetyl-glucosamine is important if the target product is containing nitrogen atoms (Matano et al., 2014, 2016). For products lacking nitrogen atoms, access to lignocellulosics is pivotal (Gopinath et al., 2011; Buschke et al., 2013; Meiswinkel et al., 2013a; Hadiati et al., 2014; Matsuura et al., 2019). The focus of this study was a sidestream of industrial starch production. Although production was lower compared to glucose (30% for glutarate, 15% for l-2HG), media composition was easy and required only a buffer, a nitrogen source for growth and iron for glutarate hydroxylase. We envision improved production of l-2HG from wheat sidestream concentrate once access to sugars other than the native substrates glucose, fructose, maltose, and sucrose is achieved, e.g., to raffinose, xylose and arabinose, that are present in this feedstock (D'Appolonia and Rayas-Duarte, 1994). Strategies for access to the lignocellulosic pentoses xylose and arabinose have been established (Kawaguchi et al., 2008; Gopinath et al., 2011; Schneider et al., 2011; Meiswinkel et al., 2013a; Imao et al., 2017) and may prove useful for improved production of l-2HG from the alternative feedstock wheat sidestream concentrate. In this respect, provision of iron and/or iron chelators may prove essential as shown here for CsiD function.

The approach described here may be followed for production of the stereoisomer d-2HG. To this end, the glutarate pathway described here has to be extended with enzymes for conversion of glutarate to d-2HG. These enzymes may be sourced from lysine degrading bacteria as d-2HG occurs as intermediate in some of these pathways, e.g., in E. coli (Zhao and Winkler, 1996) and different Pseudomonaceace (Zhang et al., 2017; Guo et al., 2018; Thompson et al., 2019). Future work will reveal if this approach is suitable for fermentative production of d-2HG.

The original contributions presented in the study are included in the article/supplementary material, further inquiries can be directed to the corresponding author/s.

VFW wishes to dedicate this article in memoriam of scientific and personal friend Michael M. Goodin, Professor of Plant Pathology, University of Kentucky, Lexington, KY.

CP and VFW conceived and designed the experiments. CP constructed plasmids and strains, cultivated, and analyzed C. glutamicum strains and prepared a draft of the manuscript. CP and FM performed bioreactor experiments and evaluated data. AB conducted the growth experiment on wheat sidestream concentrate. CP, AB, FM, and VFW finalized the manuscript. VFW acquired funding and coordinated the study. All authors read and approved the final version of the manuscript.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.