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Systems Biocatalysis: Development and engineering of cell-free “artificial metabolisms” for preparative multi-enzymatic synthesis
New Biotechnology Volume 00, Number 00 January 2015
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Systems Biocatalysis: Development and engineering of cell-free ‘‘artificial metabolisms’’ for preparative multienzymatic synthesis Wolf-Dieter Fessner Q1 Technische Universita¨t Darmstadt, Institut fu¨r Organische Chemie und Biochemie, Alarich-Weiss-Str. 4, 64287 Darmstadt, Germany
Abstract
Systems Biocatalysis is an emerging concept of organizing enzymes in vitro to construct complex reaction cascades for an efficient, sustainable synthesis of valuable chemical products. The strategy merges the synthetic focus of chemistry with the modular design of biological systems, which is similar to metabolic engineering of cellular production systems but can be realized at a far lower level of complexity from a true reductionist approach. Such operations are free from material erosion by competing metabolic pathways, from kinetic restrictions by physical barriers and regulating circuits, and from toxicity problems with reactive foreign substrates, which are notorious problems in whole-cell systems. A particular advantage of cell-free concepts arises from the inherent opportunity to construct novel biocatalytic reaction systems for the efficient synthesis of non-natural products (‘‘artificial metabolisms’’) by using enzymes specifically chosen or engineered for non-natural substrate promiscuity. Examples illustrating the technology from our laboratory are discussed. Introduction Q4 Productive biosystems for industrial applications are in high demand because of their smaller ecological footprint as compared to chemical processes, and because of their higher sustainability in view of the looming end of the petrochemical age. In living organisms, chemical conversion of matter proceeds within metabolic flow systems that constitute a highly complex network in which metabolites are interconnected by nested and interdependent reactions. Enzymes have matured by evolution over billions of years into extremely efficient catalysts that can facilitate those reaction cascades with unmatched selectivity at ambient conditions. Despite advances in the genetic engineering of production strains (cell factories) to optimize natural product formation [1], metabolic streamlining is often hampered by an unpredictable impact on the entire metabolism, because cells have evolved intricate regulatory mechanisms that counteract the genetic mutation by employing alternative pathways [2]. More demanding goals, such as expanding the range of accessible products to Tel.:+49 6151 166666; fax: +49 6151 166636. Fessner, W.-D. ([email protected]) http://dx.doi.org/10.1016/j.nbt.2014.11.007 1871-6784/ß 2014 Published by Elsevier B.V.
structures with non-natural constitution or unusual chemical functionalization [3], have been a major challenge for cellular systems and will require new approaches. The design of cell-free biocatalytic systems is a promising heuristic approach to overcome such limitations [4]. This strategy, termed Systems Biocatalysis [5], focuses on the assembly of individual enzymes in vitro and benefits from the growing knowledge on genetic diversity of protein function and the ability of modern techniques in molecular biology and protein engineering to produce effective, stable and affordable enzymes for practical applications. The principle has been applied to the mimicking of various natural metabolic pathways in vitro such as for cell-free ethanol fermentation [6], nucleotide [7], vitamin [8] and oligosaccharide biosynthesis [9], or that of various microbial secondary metabolites. But also non-natural synthetic biocatalytic pathways can be assembled easily in vitro from purified enzymes and coenzymes, and they can be constructed by selecting natural enzymes having an appropriate substrate flexibility, or by choosing enzymes optimized by protein engineering for a desired substrate promiscuity or novel function. In combination with external man-made www.elsevier.com/locate/nbt
Please cite this article in press as: Fessner, W.-D., Systems Biocatalysis: Development and engineering of cell-free ‘‘artificial metabolisms’’ for preparative multi-enzymatic synthesis, New Biotechnol. (2014), http://dx.doi.org/10.1016/j.nbt.2014.11.007
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chemicals as starting substrates, non-natural products can thereby become accessible by efficient conversions along an ‘‘artificial metabolic pathway’’ [10]. This approach avoids complications with microorganisms that may suffer from reduced viability because of the potential toxicity of laboratory chemicals and their metabolites. Cell-free systems also avoid complications from the undesired consumption of substrates into deviating metabolic pathways of the cellular background, which can erode the product yield even with recombinant strains highly overexpressing the target genes. Such competition is particularly evident for desired reaction pathways involving substrates from the central carbon metabolism as the glycolytic pathway or the citrate cycle [11]. Examples are the triose phosphates or (phosphoenol) pyruvate, which qualify as highly attractive C3-building blocks for synthesis of valuable chiral products but are interrelated in an intricate metabolic network from which their concentration is depleted into undesired competing reaction pathways. Cell-free reaction systems offer some obvious advantages when compared to whole-cell bioproduction systems, such as the facilitated adjustment of defined process conditions, including broad variations to reaction parameters (temperature, pH, co-solvent etc.), easy reaction control and monitoring. The technology further profits from fast reaction rates by direct substrate supply to the catalyst in homogenous solution, without kinetic hurdles of substrate diffusion or active transport in and out of a cell across cellular membranes, and the option of high substrate loading that leads to high product titers with facilitated product isolation. Our group had pioneered the experimental development of laboratory ‘‘artificial metabolisms’’ with a particular focus on the synthesis of non-natural carbohydrates more than 20 years ago [10], and quantum leaps in molecular genetic tools as well as knowledge from systems biology have recently led to a rapid development for a range of diverse applications, spanning from simple few-steps laboratory scale preparations of fine chemicals [12,13] to complex cascades for bulk biofuel generation [14,15] from inexpensive sustainable resources. This article will deal with design considerations mostly derived from studies pursued in our own laboratories.
Synthesis of sugar phosphates by enzymatic phosphorylation Sugar phosphates play an important physiological role as intermediates and regulators in carbohydrate metabolism. Some simple sugar phosphates are accessible by fermentation or by isolation from waste biomass, but many specialty products are intrinsically unstable or based on expensive precursors and therefore cannot be produced using whole-cell catalysis but require specific synthesis. While the chemical synthesis of phosphorylated sugars constitutes a considerable challenge, enzymatic methods provide an attractive alternative. The following examples illustrate the cell-free synthesis of valuable sugar phosphates by implementation of complex synthetic reaction cascades without the need to isolate sensitive intermediates.
6-Phosphogluconate 6-Phosphogluconate (GA6P), an important food additive in the meat and beverage industries, can be obtained by enzymatic phosphorylation of glucose using hexokinase followed by oxidation of glucose 6-phosphate (G6P) by glucose 6-phosphate 2
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FIG. 2
Cell-free reconstitution of the glycolysis pathway in vitro for the generation of labile intermediates required as substrates for subsequent enzymatic conversions (black box). Starting from one mole of cheap saccharose, 4 mol of either dihydroxyacetone phosphate (DHAP) or D-glyceraldehyde 3phosphate (GAP) are produced and consumed in situ. ATP utilized in the phosphorylation steps is regenerated from phosphoenolpyruvate (blue box). The flow chart branches out into different artificial pathways that are highlighted in color. (A) Aldol synthesis of ketose 1-phosphates using DHAP as nucleophile (green box). (B) Aldol synthesis of 1-deoxy-D-fructose 1phosphate using glyceraldehyde 3-phosphate as electrophile (red box). (C) Synthesis of D-xylulose 5-phosphate from glyceraldehyde 3-phosphate as electrophile (pink box). Abbreviations used in the figure are: Glc, D-glucose; Fru, D-fructose; G6P, D-glucose 6-phosphate; F6P, D-fructose 6-phosphate; PEP, phosphoenolpyruvate; FBP, D-fructose 1,6-bisphosphate; DHAP, dihydroxyacetone phosphate; GAP, D-glyceraldehyde 3-phosphate; LiHP, lithium hydroxypyruvate; Xul5P, D-xylulose 5-phosphate. Enzymes are indicated in bold: Inv, invertase; HK, hexokinase; PGI, glucose-6-phosphate isomerase; PFK, phosphofructokinase; PK, pyruvate kinase; FruA, fructose1,6-bisphosphate aldolase; TPI, triosephosphate isomerase; TK, transketolase; FSA, fructose-6-phosphate aldolase.
dehydrogenase (Fig. 1A). To render the two cofactor-dependent enzymatic steps economical, the required costly ATP and NAD+ cosubstrates, respectively, must be recycled in situ [16,17]. A practical solution was found in a coupled two-step, one-pot conversion with use of phosphoenolpyruvate (PEP) as a sacrificial reagent for dualpurpose recycling in both cofactor-dependent steps [18]. PEP is a superior reagent for ATP regeneration in the enzymic preparation of sugar phosphates because of its kinetic stability to hydrolysis and strong phosphoryl donor capacity [19]. Integrated closed-loop cofactor regeneration was achieved by the conversion of the stoichiometrically formed byproduct pyruvate by L-lactate dehydrogenase, which also solves the practical problem of removing pyruvate, which may act as an inhibitor to other enzymes or cause side-product formation due to its high chemical reactivity, by enzymatic reduction to into stable L-lactate. Thermodynamic relations secure a complete overall conversion because both the phosphoryl transfer and the redox potentials are
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associated with a gain in free energy [16]. Indeed, substrates were effectively pulled through the pathway without accumulation of intermediates. The one-pot protocol for preparation of GA6P with closed-loop cofactor regeneration not only proved practical because no intermediate has to be isolated, but also because the overall conversion rate was considerably higher than for a separate two-step procedure owing to the fact that inhibitory intermediates are maintained at a minimum concentration level. A disadvantage is that the enzymatic phosphorylation relies on PEP as a stoichiometric reagent, which upon whole-cell reactions will be supplied intrinsically by cellular metabolism but is not a sustainable resource for in vitro applications. In fact, PEP is marketed as an expensive fine chemical. However, both chemical or enzymatic syntheses require a high-energy phosphorylation reagent, and PEP is readily obtained in high purity on a multi-molar scale in just two fairly simple manipulations from pyruvate [19]. As an alternative, less expensive phosphoryl sources such as acetyl phosphate or polyphosphate in combination with appropriate kinases can likewise be considered [17]. A different option is to avoid an ATP dependent kinase with need for cofactor recycling by applying an acid phosphatase for transphosphorylation from inexpensive pyrophosphate as donor. However, such methods often suffer from low regioselectivity and competing product hydrolysis [20]. D-Tagatose
1,6-bisphosphate
The degradation of lactose, D-galactose and galactitol proceeds in bacteria by a pathway capitalizing on the aldol cleavage of the intermediate D-tagatose 1,6-bisphosphate (TBP) [21]. When for assay purposes TBP was available only by a low yielding (2%), multistep synthesis from D-galacturonic acid [22], we had developed an alternative multi-enzymatic synthesis from dihydroxyacetone (DHA) [23,24]. After enzymatic phosphorylation of DHA using ATP and glycerol kinase, combined with cofactor regeneration based on PEP (Fig. 1B), the generated dihydroxyacetone phosphate (DHAP) first is partially isomerized in situ into D-glyceraldehyde 3-phosphate (GAP), then added to the latter in a stereoselective carboligation step catalyzed by tagatose-1,6bisphosphate aldolase (TagA) to furnish TBP. In principle, the synthesis strategy was based on a reversal of the catabolic route, which to become practical requires a high equilibrium concentration of TBP in the aldol reaction. However, as a consequence of its all-cis substitution in the furanose forms present in aqueous solution, the TBP product indeed is only barely favored (Keq = [DHAP] [GAP]/[TBP] = 103 M) [23]. For this reason the TBP yield cannot significantly exceed a level of 80%, even with reaction parameters optimized. As a consequence, high concentration of triose phosphate substrates is required to shift equilibrium conditions toward higher productivity.
Rare ketosugar 1-phosphates Microbial metabolisms comprise a remarkable repertoire for the catabolic utilization of carbohydrates as the sole source of carbon and energy. A common feature of most degradative sequences is the conversion of sugars and polyols into ketose L-phosphates that by breakdown of the carbon backbone into smaller units enter the central metabolism. Since the phosphorylated ketoses frequently function as the inducers of respective metabolic enzymes, they are
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thus primary targets for the investigation of mechanistic and regulatory aspects of sugar metabolism. We have developed a practical route to various ketose L-phosphates that starts from commercially available aldoses or ketoses and makes use of a ketose kinase in combination with an appropriate ketol isomerase (Fig. 1C) [25,26]. All enzymes are derived from L-rhamnose/L-fucose catabolism and are accessible in quantity by recombinant overexpression. The basis for this development was the observation that, unlike the functionally related hexokinases, the L-rhamnulokinase (RhuK) from Escherichia coli has a relatively broad tolerance for structural modifications of its natural substrate L-rhamnu1ose, except for the (3R)-configuration. The scheme is completed by two corresponding ketol isomerases, the L-rhamnose isomerase (RhaI) and the L-fucose isomerase (FucI) from E. coli. In common with RhuK, both isomerases require a fixed (3R)-configuration but are flexible toward structural variations at the C4–C6 segment. In the isomerization equilibria, the ketose isomers often turned out to be the less abundant constituent for structural reasons [27]. The thermodynamically favorable phosphoryl transfer from ATP, however, serves to constantly drain the ketose component from the equilibrium and to drive the reactions to completion. As an example, the RhaI/RhuK combination provides a straightforward and high-yielding access to rare ketose L-phosphates having a common (2R,3R)-configuration from a range of stereochemically related aldohexoses or aldopentoses that serve as starting materials [25]. The construction of cell-free biocatalytic systems according to the Systems Biocatalysis strategy offers the freedom to design the most appropriate sequence of reactions to the target product. To achieve maximum efficacy in one-pot conversions it is obviously important to balance endothermic or thermoneutral reactions by coupling to exothermic ones. However, it is also important to be able to choose preferentially the thermodynamically most effective combination of reagents for cofactor recycling, and particularly that sequence where this critical step is properly placed at the very end. Clearly, such operational freedom in vitro sets the Systems Biocatalysis approach apart from whole-cell technology where such is not, or at least not easily, attainable in vivo.
Glycolysis intermediates as substrates Triose phosphates (DHAP, GAP) are versatile small building blocks for asymmetric synthesis because of their dense, but differentiated functionality and their high chemical reactivity that can be readily exploited in a selective fashion. In particular, these structures can be utilized for the highly stereoselective enzymatic generation of a complete set of stereoisomeric vicinal diols by employing aldolase technology [28,29]. Although much research has been directed at their chemical or biochemical synthesis [30,31], both compounds remain very expensive fine chemicals. In addition, they are labile to phosphate elimination in aqueous solution with generation of cell-toxic methylglyoxal [32,33], which renders production by whole-cell approaches unfeasible.
Dihydroxyacetone phosphate In the context of a research program targeted at C–C bond forming aldolases for asymmetric synthesis we had been searching for means to make the essential substrate DHAP efficiently available. In a pioneering study [10] published in 1992 we were attracted by
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the potential of multi-enzyme systems in vitro that would allow by a series of specific serial or parallel conversions of shared substrates along the glycolysis pathway to exploit sugars as an economical source of DHAP. Investigations into these enzyme cascade reactions had been initiated, firstly because of the potential preparative benefit, and secondly to gain insight into the complex kinetic effects operative in vitro for such a multiply regulated multienzyme system. Starting with the central function of fructose-1,6-bisphosphate aldolase (FruA) in glycolysis to cleave and – in combination with triose phosphate isomerase – symmetrize fructose 1,6-bisphosphate (FBP) into C3 fragments, we have assembled additional steps in such a manner (Fig. 2) that several regenerative raw materials (glucose, fructose, or saccharose) can be utilized alternatively as possible precursors. The overall artificial reaction network comprises seven purified enzymes from commercial sources that, including ATP cofactor regeneration at several levels to drive overall synthetic equilibria to the FBP stage, jointly deliver DHAP in situ for consecutive preparative use. It became apparent that the phosphofructokinase (PFK) remained a limiting factor, in accord with the notion that PFK is the central regulatory element of glycolysis that is inhibited by numerous effectors (e.g., ATP,
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PEP, pyruvate). In metabolic engineering a critical element in a pathway is replaced by a homologous gene from another organism coding for an enzyme with superior properties [34]. Analogously, the artificial in vitro metabolism was engineered by replacing the mammalian PFK from rabbit muscle with the allosterically less regulated bacterial PFK-2 isoenzyme from E. coli [35,36], which significantly improved the flux to FBP production. Subsequently, to this scheme was combined an unnatural synthesis step by supplying an external aldehyde reactant to employ the FruA for a consecutive non-natural aldol reaction (Fig. 2A). Because aldol reactions are reversible, the overall equilibrium must be shifted toward product formation by using an excess of aldehyde. Relative to syntheses starting from pure FBP comparable yields were achieved with the multi-enzyme system. The method made available a selection of non-natural alkylated, spiro-annulated, deoxy and branched chain sugars that are difficult to prepare by classical chemical synthesis. Because this cell-free reaction scheme comprises an unusual accumulation of coupled reactions as a new combination of natural, catabolic processes with a nonphysiological, anabolic synthetic component by application of pure isolated enzymes in vitro, it was designated as the first artificial metabolism [10]. The reduced complexity as compared to living organisms makes in vitro biocatalytic systems readily amenable to pathway engineering and optimization. One further goal was to address the synthesis of stereoisomers different from those created by FruA catalysis, which requires supplementation of the metabolic scheme with a different aldolase such as fuculose-1-phosphate aldolase (FucA) having distinct stereospecificity [37]. Because FruA is mandatory for the aldol cleavage of FBP, the competing generation of stereoisomers was eliminated by replacing the class I FruA from rabbit muscle by its bacterial class II analog from E. coli, which has very narrow substrate specificity for GAP [38]. Indeed, the engineered in vitro metabolic pathway, including FucA from E. coli as an eighth enzyme in the cascade, produced diastereomerically pure products having the L-erythro rather than the D-threo configuration typical for FruA [38]. D-Glyceraldehyde
FIG. 1
One-pot multi-enzyme catalyzed synthesis of phosphorylated carbohydrate metabolites; in the flow charts starting materials (left) and products (right) are highlighted in blue. (A) Phosphoenolpyruvate (PEP) as a double-duty cofactor recycling reagent for in situ regeneration of ATP and NAD+. (B) Aldol synthesis of D-tagatose 1,6-bisphosphate (TagBP). (C) Coupled isomerization– phosphorylation for rare sugar 1-phosphate synthesis. Abbreviations used in the figure are: Glc6P, D-glucose 6-phosphate; GlcA6P, D-glucuronic acid 6phosphate; DHAP, dihydroxyacetone phosphate; GAP, D-glyceraldehyde 3phosphate; TagBP, D-tagatose 1,6-bisphosphate; PEP, phosphoenolpyruvate (PEP). Enzymes are indicated in bold: HK, hexokinase; PK, pyruvate kinase; GPDH, glucose-6-phosphate dehydrogenase; LDH, lactate dehydrogenase; GK, glycerol kinase; TPI, triosephosphate isomerase; TagA, tagatose-1,6bisphosphate aldolase; FucI, fucose isomerase; RhaI, rhamnose isomerase; RhuK, rhamnulokinase. 4
3-phosphate
For the preparation of 1-deoxyfructose 6-phosphate (dF6P) needed as an assay component for aldolase screening, we had envisioned a coupled enzymatic approach for its synthesis (Fig. 2B) based on the fructose 6-phosphate aldolase (FSA) catalyzed carboligation from GAP and hydroxyacetone [39]. FSA from E. coli is a member of the transaldolase class and specific for non-phosphorylated aldol donors [40,41]. Because of its low stability in solution, GAP is preferably generated and consumed in situ to avoid the build-up of high stationary concentrations, which is conveniently achieved by retro-aldol cleavage of stable FBP in the presence of FruA in concert with TPI. Both aldolases show a distinct and mutually exclusive, high substrate specificity (DHAP and ketoses 1-phosphates in the case of FruA, free ketoses in the case of FSA), which excludes any interference from substrates, intermediates, or products [40]. For the synthesis of dF6P, FBP was incubated with FruA, TPI and FSA in the presence of an excess of hydroxyacetone. The high concentration of the latter, necessary to propel the product formation in the all-equilibrium procedure, again required replacement of the conventional rabbit muscle FruA in the scheme
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by the corresponding recombinant FruA enzyme from Staphylococcus carnosus [42], which is highly stable and tolerant to enhanced levels of hydroxyacetone. The optimized method successfully delivered dF6P in 96% yield on a multi-gram scale [39]. Xylulose 5-phosphate (X5P), occurring as an intermediate in the pentose phosphate pathway, is a valuable substrate required for enzymatic assays, particularly for the screening of eucaryotic enzymes in anti-cancer drug design [43]. Because published protocols for the synthesis of X5P from free sugars or other phosphorylated precursors are tedious, micro-scale and/or low-yielding, we developed a practical and efficient procedure for the preparation of X5P by a multi-enzymatic route based on the stereospecific transketolase-catalyzed ketol transfer from hydroxypyruvate to GAP [44]. Again, retro-aldol fragmentation of FBP in the presence of FruA and TPI was considered as an in situ source of the triose phosphate electrophile (Fig. 2C). Although the equilibrium concentration of GAP is rather low, reaction rates were sufficiently high owing to the high affinity of transketolase for GAP and the release of CO2 resulting from decomposition of hydroxypyruvate, which renders the overall synthetic reaction practically irreversible. The product X5P could be isolated from this simple one-pot procedure in 82% overall yield on a multi-gram scale [44]. It ought to be noted that the above procedures all require the conversion of biogenic starting materials jointly with external non-natural chemicals (e.g., aldehydes, hydroxypyruvate), which are added in stoichiometric amounts and at high concentration. Owing to the high chemical reactivity of the latter components, this is readily tolerated by cell-free metabolic cascades that are composed of suitably selected stable and effective biocatalysts, but likewise would not be readily compatible with whole-cell technology because of toxicity issues.
Oxidative pathways for substrate generation Reactions involving elemental oxygen usually profit from a strong thermodynamic driving force by the high redox potential of oxygen reduction, which renders coupled reactions practically irreversible. In our quest for suitable DHAP sources, we have also investigated the potential of inexpensive flavin-dependent glycerol phosphate oxidase (GPO) from Streptococcus sp. for the air oxidation of L-glycerol 3-phosphate (Gly3P) as another method to generate DHAP in situ for consecutive use in aldol reactions (Fig. 3A) [45]. The reaction indeed proceeds smoothly and yields pure DHAP almost quantitatively. With GPO a separate cofactor regeneration step is obsolete because the reduced cofactor FAD (H2) remains tightly bound to the enzyme and is rapidly re-oxidized by elemental oxygen. Thereby, hydrogen peroxide is liberated, which must be decomposed by addition of catalase (Cat). In turn, as the flow chart suggests, hydrogen peroxide can be used alternatively as the oxidant [45,46]. In principle, Gly3P again could be obtained by coupled enzymatic phosphorylation of glycerol but this option had not been further pursued because of the low oxygen tolerance of available glycerol kinases [47]. However, oxidative DHAP generation could be smoothly coupled to its consumption in aldol reactions because the DHAP aldolases are not sensitive to oxygen. When compared to other established procedures, the GPO method usually furnished adducts of higher purity and in at least equal or higher yields. Interestingly, the GPO procedure proved also
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FIG. 3
One-pot multi-enzyme catalyzed carbohydrate synthesis by reversible aldol addition. The equilibrium of the carboligation reaction is driven by oxidative generation of the substrate(s) in situ using molecular oxygen. (A) Aldol synthesis of ketose 1-phosphates from glycerol phosphate. (B) Aldol synthesis of complex bicyclic 9-carbon sugar by carboligation from a galactoside precursor. Abbreviations used in the figure are: Gly3P, L-glycerol 3-phosphate; DHAP, dihydroxyacetone phosphate; GalOMe, b-methyl-D-galactoside; 6aldGalOMe, 6-aldehydo-b-methyl-D-galactoside. Enzymes are indicated in bold: GPO, glycerophosphate oxidase; Cat, catalase; GalO, galactose oxidase; RhuA, rhamnulose-1-phosphate aldolase.
adaptable to the synthesis of DHAP analogues, such as the corresponding bio-isosteric phosphonate and phosphorothioate, which are likewise accepted by the aldolases [48]. In a particularly interesting example (Fig. 3B), both the aldol donor and acceptor components for a subsequent carboligation were generated in situ by air oxidation using microbial oxidases [49]. For this purpose, the GPO technology for oxidative generation of DHAP as the nucleophilic component was coupled to an oxidative generation of the corresponding electrophilic aldehyde component. The latter was obtained by parallel oxidation of methyl b-D-galactopyranoside at the non-reducing end using the Cu(II)-dependent D-galactose oxidase (GalO) from Dactylium dendroides [50], and the system was completed by adding the highly substrate tolerant L-rhamnulose 1-phosphate aldolase (RhuA) from E. coli [37]. The ensuing aldol addition consumed both products in situ and thus obviated inhibition problems. This novel multi-enzymatic reaction system for the preparation of complex high-carbon sugar derivatives has the advantages of simplicity of operation, ready availability of starting materials at low cost, and high overall product yield [49].
Leloir-path sialoconjugation Sialic acid-containing glycoconjugates at the cell surface are of high importance in carbohydrate-mediated recognition phenomena in physiological and pathological events, including bacterial or viral infection [51]. Natural sialic acid derivatives typically incorporate various types of N- or O-modification but the resultant biological profiles are poorly understood because the individual compounds are not yet available in sufficient quantity and purity for testing [52]. The Leloir-type biosynthesis of sialic acid conjugates requires three dedicated enzymatic steps, which consist of a carboligation step to construct the extended sialic acid backbone, followed by nucleotide activation of the sugar and its regio- and stereospecific transfer to a suitable acceptor moiety (Fig. 4). Toward a generic method for the synthesis of sialoconjugate libraries [53], the catabolic neuraminic acid aldolase (NeuA) from
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FIG. 4
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One-pot, multi-enzyme catalyzed synthesis of sialoconjugates by Leloir-type sialyltransfer with in situ generation of natural and non-natural sialic acids by reversible carboligation. Abbreviations used in the figure are: Sia, sialic acid (N-acetylneuraminic acid and analogues); CMP-Sia,; GalOR, b-D-galactoside. Enzymes are indicated in bold: NeuA, neuraminic acid aldolase; CSS, CMPsialic acid synthase; 2,6SiaT, a2,6-sialyltransferase.
E. coli has been shown to offer broad substrate promiscuity for catalyzing the addition of pyruvate to N-acetyl-D-mannosamine and related aldoses to form various natural and non-natural sialic acids [29]. We have discovered that the CMP-sialic acid synthetase (CSS) from Neisseria meningitidis can activate an unusually wide range of sialic acid derivatives [54,55]. Allowable structural modifications included N-acyl variation as well as altered substitution pattern, backbone size, and inversion of stereoconfiguration. Recently, we have engineered the CSS by structure-guided sitespecific saturation mutagenesis to generate enzymes with significantly enhanced substrate promiscuity for sterically highly demanding N-acyl modified sialic acid analogues [56]. Top hits such as the F192S/F193Y variant display an improvement of up to 70-fold catalytic efficiency relative to wild-type CSS. CSS catalysis was then coupled in a cascade reaction system to an a2,6-sialyltransferase (2,6SiaT) conversion, which effects transfer of the sialyl moiety to a suitable galactoside acceptor. For this purpose, a novel highly substrate tolerant 2,6SiaT from Photobacterium leiognathi JTSHIZ-145 was applied, which best matches the requirements from the overall reaction system [57]. Rapid regio- and stereospecific sialyltransfer was achieved onto lactoside acceptors, and the coupled enzymatic system allowed preparing a library of
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neo-sialoconjugates (structural analogues of 60 -sialyllactosides) in a highly efficiently manner [55,56]. Cascade enzyme processes are especially important for glycosyltransferase-catalyzed reactions because they require nucleotideactivated substrates that are highly expensive and unstable but can be generated in situ from simpler starting materials using standard biosynthetic enzymes. Particularly, the development of promiscuous enzymes of the Leloir pathway is instrumental for the development of simplified, cascade-type synthetic procedures to facilitate rapid access to biologically important oligosaccharide structures for biomedical applications.
Summary and outlook This compilation is meant to illustrate the power of the Systems Biocatalysis approach, partly by highlighting early examples from the pioneer era in developing multi-step enzymatic reaction networks in vitro, partly from recent studies into more generic pathways using enzymes engineered for substrate promiscuity. It clearly demonstrates that cell-free artificial biocatalytic systems of high complexity can be designed and rapidly established, which are simple to operate and analyze for pathway engineering and process optimization, and which can be applied with high freedom to operate. Such is particularly evident for the synthesis of a wide range of non-natural fine chemicals that require reactive components that may not be compatible with living systems. The obvious drawback for creating in vitro biosystems currently still is the laborious purification of the parts, which usually makes exploratory laboratory developments unsustainable beyond the research scale. However, opportunities are emerging to significantly reduce costs such as with the use of recombinant enzymes from thermophilic organisms that can be purified by simple heat-shock treatment [58]. Together with the power of modern synthetic biology to identify replacement parts from nature, or to engineer improved parts by evolution [59], the concept of ‘‘artificial metabolisms’’ for sustainable synthesis in vitro is expected to make an important contribution to the development of productive biosystems of the future.
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[10] Fessner W-D, Walter C. Artificial metabolisms for the asymmetric one-pot synthesis of branched-chain saccharides. Angew Chem Int Ed Engl 1992;31: 614–6. [11] Bujara M, Schuemperli M, Pellaux R, Heinemann M, Panke S. Optimization of blueprint for in vitro glycolysis by metabolic real-time analysis. Nat Chem Biol 2011;7:271–7. [12] Bujara M, Schuemperli M, Billerbeck S, Heinemann M, Panke S. Exploiting cellfree systems: implementation and debugging of a system of biotransformations. Biotechnol Bioeng 2010;106:376–89. [13] Guterl J-K, Sieber V. Biosynthesis debugged: novel bioproduction strategies. Eng Life Sci 2013;13:4–18. [14] Zhang YHP, Sun J, Zhong J-J. Biofuel production by in vitro synthetic enzymatic pathway biotransformation. Curr Opin Biotechnol 2010;21:663–9. [15] Zhang YHP. Production of biocommodities and bioelectricity by cell-free synthetic enzymatic pathway biotransformations: challenges and opportunities. Biotechnol Bioeng 2010;105:663–77. [16] Chenault HK, Simon ES, Whitesides GM. Cofactor regeneration for enzymecatalyzed synthesis. Biotechnol Gen Eng Rev 1988;6:221–70. [17] Truppo MD. Cofactor recycling for enzyme catalyzed processes. Compre Chirality 2012;7:46–70. [18] Fessner W-D, Sinerius G. Phosphoenolpyruvate as a dual purpose reagent for integrated nucleotide/nicotinamide cofactor recycling. Bioorg Med Chem 1994;2:639–45. [19] Hirschbein BL, Mazenod FP, Whitesides GM. Synthesis of phosphoenolypyruvate and its use in ATP cofactor regeneration. J Org Chem 1982;47:3765–6.
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[20] Tanaka N, Hasan Z, Hartog AF, van Herk T, Wever R. Phosphorylation and dephosphorylation of polyhydroxy compounds by class A bacterial acid phosphatases. Org Biomol Chem 2003;1:2833–9. [21] LowKam C, Liotard B, Sygusch J. Structure of a class I tagatose-1,6-bisphosphate aldolase: investigation into an apparent loss of stereospecificity. J Biol Chem 2010;285:21143–52. [22] Anderson RL, Wenger WC, Bissett DL. D-Galactose 6-phosphate and D-tagatose 6-phosphate. Methods Enzymol 1982;89:93–8. [23] Fessner W-D, Eyrisch O. One-pot synthesis of tagatose 1,6-bisphosphate via diastereoselective enzymic aldol addition. Angew Chem Int Ed Engl 1992;31: 56–8. [24] Eyrisch O, Sinerius G, Fessner WD. Facile enzymic de novo synthesis and NMR spectroscopic characterization of D-tagatose 1,6-bisphosphate. Carbohydr Res 1993;238:287–306. [25] Fessner W-D, Badia J, Eyrisch O, Schneider A, Sinerius G. Enzymic syntheses of rare ketose 1-phosphates. Tetrahedron Lett 1992;33:5231–4. [26] Fessner W-D, Schneider A, Eyrisch O, Sinerius G, Badia J. 6-Deoxy-L-lyxo- and 6-deoxy-L-arabino-hexulose 1-phosphates. Enzymic syntheses by antagonistic metabolic pathways. Tetrahedron: Asymmetry 1993;4:1183–92. [27] Fessner W-D, Gosse C, Jaeschke G, Eyrisch O. Short enzymatic synthesis of L-fucose analogs. Eur J Org Chem 2000;125–32. Q5 [28] Fessner W-D, Helaine V. Biocatalytic synthesis of hydroxylated natural products using aldolases and related enzymes. Curr Opin Biotechnol 2001;12:574–86. [29] Fessner W-D. Aldol reactions. In: Drauz K, Groeger H, May O, editors. 3rd ed., Enzyme catalysis in organic synthesis, vol. 2, 3rd ed. Weinheim: Wiley-VCH; 2011. p. 857–917. [30] Schuemperli M, Pellaux R, Panke S. Chemical and enzymatic routes to dihydroxyacetone phosphate. Appl Microbiol Biotechnol 2007;75:33–45. [31] Gauss D, Schoenenberger B, Wohlgemuth R. Chemical and enzymatic methodologies for the synthesis of enantiomerically pure glyceraldehyde 3-phosphates. Carbohydr Res 2014;389:18–24. [32] Richard JP. Acid–base catalysis of the elimination and isomerization reactions of triose phosphates. J Am Chem Soc 1984;106:4926–36. [33] Richard JP. Kinetic parameters for the elimination reaction catalyzed by triosephosphate isomerase and an estimation of the reaction’s physiological significance. Biochemistry 1991;30:4581–5. [34] Stephanopoulos G. Synthetic biology and metabolic engineering. ACS Synth Biol 2012;1:514–25. [35] Babul J. Phosphofructokinases from Escherichia coli. Purification and characterization of the nonallosteric isozyme. J Biol Chem 1978;253:4350–5. [36] Daldal F. Molecular cloning of the gene for phosphofructokinase-2 of Escherichia coli and the nature of a mutation, pfkB1, causing a high level of the enzyme. J Mol Biol 1983;168:285–305. [37] Fessner W-D, Sinerius G, Schneider A, Dreyer M, Schulz GE, Badia J, et al. Diastereoselective enzymatic aldol additions: L-rhamnulose and L-fuculose 1-phosphate aldolases from E. coli. Angew Chem Int Ed Engl 1991;30: 555–8. [38] Fessner W-D, Walter C. Enzymatic C–C bond formation in asymmetric synthesis. Top Curr Chem 1996;184:97–194. [39] Fessner W-D, Heyl D, Rale M. Multi-enzymatic cascade synthesis of D-fructose 6-phosphate and deoxy analogs as substrates for high-throughput aldolase screening. Catal Sci Technol 2012;2:1596–601. [40] Samland AK, Rale M, Sprenger GA, Fessner W-D. The transaldolase family: new synthetic opportunities from an ancient enzyme scaffold. ChemBioChem 2011;12:1454–74.
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[41] Rale M, Schneider S, Sprenger GA, Samland AK, Fessner W-D. Broadening deoxysugar glycodiversity: natural and engineered transaldolases unlock a complementary substrate space. Chem Eur J 2011;17:2623–32. S/1–S/42. [42] Zannetti MT, Walter C, Knorst M, Fessner W-D. Fructose 1,6-bisphosphate aldolase from Staphylococcus carnosus: overexpression, structure prediction, stereoselectivity, and application in the synthesis of bicyclic sugars. Chem Eur J 1999;5:1882–90. [43] Du MX, Sim J, Fang L, Zheng Y, Koh S, Stratton J, et al. Identification of novel small-molecule inhibitors for human transketolase by high-throughput screening with fluorescent intensity (FLINT) assay. J Biomol Screening 2004;9:427–33. [44] Zimmermann FT, Schneider A, Schorken U, Sprenger GA, Fessner W-D. Efficient multi-enzymatic synthesis of D-xylulose 5-phosphate. Tetrahedron: Asymmetry 1999;10:1643–6. [45] Fessner W-D, Sinerius G. Synthesis of dihydroxyacetone phosphate (and isosteric analogs) by enzymatic oxidation: sugars from glycerol. Angew Chem Int Ed Engl 1994;33:209–12. [46] Hettwer J, Oldenburg H, Flaschel E. Enzymic routes to dihydroxyacetone phosphate or immediate precursors. J Mol Catal B: Enzym 2002;19–20:215–22. [47] Crans DC, Whitesides GM. Glycerol kinase: synthesis of dihydroxyacetone phosphate, sn-glycerol-3-phosphate, and chiral analogs. J Am Chem Soc 1985;107:7019–27. [48] Arth H-L, Fessner W-D. Practical synthesis of 4-hydroxy-3-oxobutylphosphonic acid and its evaluation as a bio-isosteric substrate of DHAP aldolase. Carbohydr Res 1998;305:313–21. [49] Eyrisch O, Keller M, Fessner W-D. Higher-carbon sugars by enzymatic chain extension. Oxidative generation of aldol precursors in situ. Tetrahedron Lett 1994;35:9013–6. [50] Whittaker JW. The radical chemistry of galactose oxidase. Arch Biochem Biophys 2005;433:227–39. [51] Chen X, Varki A. Advances in the biology and chemistry of sialic acids. ACS Chem Biol 2010;5:163–76. [52] Angata T, Varki A. Chemical diversity in the sialic acids and related alpha-keto acids: an evolutionary perspective. Chem Rev 2002;102:439–69. [53] Fessner WD, He N, Yi D, Unruh P, Knorst M. Enzymatic generation of sialoconjugate diversity. In: Riva S, Fessner WD, editors. Cascade biocatalysis. Weinheim: Wiley-VCH; 2014. p. 361–92. [54] Knorst M, Fessner W-D. CMP-sialate synthetase from Neisseria meningitidis— overexpression and application to the synthesis of oligosaccharides containing modified sialic acids. Adv Synth Catal 2001;343:698–710. [55] He N, Yi D, Fessner W-D. Flexibility of substrate binding of cytosine-50 -monophosphate-N-acetylneuraminate synthetase (CMP-sialate synthetase) from Neisseria meningitidis: an enabling catalyst for the synthesis of neo-sialoconjugates. Adv Synth Catal 2011;353:2384–98. [56] Yi D, He N, Kickstein M, Metzner J, Weiss M, Berry A, et al. Engineering of a cytidine 50 -monophosphate-sialic acid synthetase for improved tolerance to functional sialic acids. Adv Synth Catal 2013;355:3597–612. [57] Yamamoto T, Hamada Y, Ichikawa M, Kajiwara H, Mine T, Tsukamoto H, et al. A beta-galactoside alpha 2,6-sialyltransferase produced by a marine bacterium, Photobacterium leiognathi JT-SHIZ-145, is active at pH 8. Glycobiology 2007;17: 1167–74. [58] Abdoul-Zabar J, Sorel I, Helaine V, Charmantray F, Devamani T, Yi D, et al. Thermostable transketolase from Geobacillus stearothermophilus: characterization and catalytic properties. Adv Synth Catal 2013;355:116–28. [59] Kaul P, Asano Y. Strategies for discovery and improvement of enzyme function: state of the art and opportunities. Microbial Biotechnol 2012;5:18–33.
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Systems Biocatalysis: Development and engineering of cell-free ‘‘artificial metabolisms’’ for preparative multienzymatic synthesis Wolf-Dieter Fessner Q1 Technische Universita¨t Darmstadt, Institut fu¨r Organische Chemie und Biochemie, Alarich-Weiss-Str. 4, 64287 Darmstadt, Germany
Abstract
Systems Biocatalysis is an emerging concept of organizing enzymes in vitro to construct complex reaction cascades for an efficient, sustainable synthesis of valuable chemical products. The strategy merges the synthetic focus of chemistry with the modular design of biological systems, which is similar to metabolic engineering of cellular production systems but can be realized at a far lower level of complexity from a true reductionist approach. Such operations are free from material erosion by competing metabolic pathways, from kinetic restrictions by physical barriers and regulating circuits, and from toxicity problems with reactive foreign substrates, which are notorious problems in whole-cell systems. A particular advantage of cell-free concepts arises from the inherent opportunity to construct novel biocatalytic reaction systems for the efficient synthesis of non-natural products (‘‘artificial metabolisms’’) by using enzymes specifically chosen or engineered for non-natural substrate promiscuity. Examples illustrating the technology from our laboratory are discussed. Introduction Q4 Productive biosystems for industrial applications are in high demand because of their smaller ecological footprint as compared to chemical processes, and because of their higher sustainability in view of the looming end of the petrochemical age. In living organisms, chemical conversion of matter proceeds within metabolic flow systems that constitute a highly complex network in which metabolites are interconnected by nested and interdependent reactions. Enzymes have matured by evolution over billions of years into extremely efficient catalysts that can facilitate those reaction cascades with unmatched selectivity at ambient conditions. Despite advances in the genetic engineering of production strains (cell factories) to optimize natural product formation [1], metabolic streamlining is often hampered by an unpredictable impact on the entire metabolism, because cells have evolved intricate regulatory mechanisms that counteract the genetic mutation by employing alternative pathways [2]. More demanding goals, such as expanding the range of accessible products to Tel.:+49 6151 166666; fax: +49 6151 166636. Fessner, W.-D. ([email protected]) http://dx.doi.org/10.1016/j.nbt.2014.11.007 1871-6784/ß 2014 Published by Elsevier B.V.
structures with non-natural constitution or unusual chemical functionalization [3], have been a major challenge for cellular systems and will require new approaches. The design of cell-free biocatalytic systems is a promising heuristic approach to overcome such limitations [4]. This strategy, termed Systems Biocatalysis [5], focuses on the assembly of individual enzymes in vitro and benefits from the growing knowledge on genetic diversity of protein function and the ability of modern techniques in molecular biology and protein engineering to produce effective, stable and affordable enzymes for practical applications. The principle has been applied to the mimicking of various natural metabolic pathways in vitro such as for cell-free ethanol fermentation [6], nucleotide [7], vitamin [8] and oligosaccharide biosynthesis [9], or that of various microbial secondary metabolites. But also non-natural synthetic biocatalytic pathways can be assembled easily in vitro from purified enzymes and coenzymes, and they can be constructed by selecting natural enzymes having an appropriate substrate flexibility, or by choosing enzymes optimized by protein engineering for a desired substrate promiscuity or novel function. In combination with external man-made www.elsevier.com/locate/nbt
Please cite this article in press as: Fessner, W.-D., Systems Biocatalysis: Development and engineering of cell-free ‘‘artificial metabolisms’’ for preparative multi-enzymatic synthesis, New Biotechnol. (2014), http://dx.doi.org/10.1016/j.nbt.2014.11.007
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chemicals as starting substrates, non-natural products can thereby become accessible by efficient conversions along an ‘‘artificial metabolic pathway’’ [10]. This approach avoids complications with microorganisms that may suffer from reduced viability because of the potential toxicity of laboratory chemicals and their metabolites. Cell-free systems also avoid complications from the undesired consumption of substrates into deviating metabolic pathways of the cellular background, which can erode the product yield even with recombinant strains highly overexpressing the target genes. Such competition is particularly evident for desired reaction pathways involving substrates from the central carbon metabolism as the glycolytic pathway or the citrate cycle [11]. Examples are the triose phosphates or (phosphoenol) pyruvate, which qualify as highly attractive C3-building blocks for synthesis of valuable chiral products but are interrelated in an intricate metabolic network from which their concentration is depleted into undesired competing reaction pathways. Cell-free reaction systems offer some obvious advantages when compared to whole-cell bioproduction systems, such as the facilitated adjustment of defined process conditions, including broad variations to reaction parameters (temperature, pH, co-solvent etc.), easy reaction control and monitoring. The technology further profits from fast reaction rates by direct substrate supply to the catalyst in homogenous solution, without kinetic hurdles of substrate diffusion or active transport in and out of a cell across cellular membranes, and the option of high substrate loading that leads to high product titers with facilitated product isolation. Our group had pioneered the experimental development of laboratory ‘‘artificial metabolisms’’ with a particular focus on the synthesis of non-natural carbohydrates more than 20 years ago [10], and quantum leaps in molecular genetic tools as well as knowledge from systems biology have recently led to a rapid development for a range of diverse applications, spanning from simple few-steps laboratory scale preparations of fine chemicals [12,13] to complex cascades for bulk biofuel generation [14,15] from inexpensive sustainable resources. This article will deal with design considerations mostly derived from studies pursued in our own laboratories.
Synthesis of sugar phosphates by enzymatic phosphorylation Sugar phosphates play an important physiological role as intermediates and regulators in carbohydrate metabolism. Some simple sugar phosphates are accessible by fermentation or by isolation from waste biomass, but many specialty products are intrinsically unstable or based on expensive precursors and therefore cannot be produced using whole-cell catalysis but require specific synthesis. While the chemical synthesis of phosphorylated sugars constitutes a considerable challenge, enzymatic methods provide an attractive alternative. The following examples illustrate the cell-free synthesis of valuable sugar phosphates by implementation of complex synthetic reaction cascades without the need to isolate sensitive intermediates.
6-Phosphogluconate 6-Phosphogluconate (GA6P), an important food additive in the meat and beverage industries, can be obtained by enzymatic phosphorylation of glucose using hexokinase followed by oxidation of glucose 6-phosphate (G6P) by glucose 6-phosphate 2
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FIG. 2
Cell-free reconstitution of the glycolysis pathway in vitro for the generation of labile intermediates required as substrates for subsequent enzymatic conversions (black box). Starting from one mole of cheap saccharose, 4 mol of either dihydroxyacetone phosphate (DHAP) or D-glyceraldehyde 3phosphate (GAP) are produced and consumed in situ. ATP utilized in the phosphorylation steps is regenerated from phosphoenolpyruvate (blue box). The flow chart branches out into different artificial pathways that are highlighted in color. (A) Aldol synthesis of ketose 1-phosphates using DHAP as nucleophile (green box). (B) Aldol synthesis of 1-deoxy-D-fructose 1phosphate using glyceraldehyde 3-phosphate as electrophile (red box). (C) Synthesis of D-xylulose 5-phosphate from glyceraldehyde 3-phosphate as electrophile (pink box). Abbreviations used in the figure are: Glc, D-glucose; Fru, D-fructose; G6P, D-glucose 6-phosphate; F6P, D-fructose 6-phosphate; PEP, phosphoenolpyruvate; FBP, D-fructose 1,6-bisphosphate; DHAP, dihydroxyacetone phosphate; GAP, D-glyceraldehyde 3-phosphate; LiHP, lithium hydroxypyruvate; Xul5P, D-xylulose 5-phosphate. Enzymes are indicated in bold: Inv, invertase; HK, hexokinase; PGI, glucose-6-phosphate isomerase; PFK, phosphofructokinase; PK, pyruvate kinase; FruA, fructose1,6-bisphosphate aldolase; TPI, triosephosphate isomerase; TK, transketolase; FSA, fructose-6-phosphate aldolase.
dehydrogenase (Fig. 1A). To render the two cofactor-dependent enzymatic steps economical, the required costly ATP and NAD+ cosubstrates, respectively, must be recycled in situ [16,17]. A practical solution was found in a coupled two-step, one-pot conversion with use of phosphoenolpyruvate (PEP) as a sacrificial reagent for dualpurpose recycling in both cofactor-dependent steps [18]. PEP is a superior reagent for ATP regeneration in the enzymic preparation of sugar phosphates because of its kinetic stability to hydrolysis and strong phosphoryl donor capacity [19]. Integrated closed-loop cofactor regeneration was achieved by the conversion of the stoichiometrically formed byproduct pyruvate by L-lactate dehydrogenase, which also solves the practical problem of removing pyruvate, which may act as an inhibitor to other enzymes or cause side-product formation due to its high chemical reactivity, by enzymatic reduction to into stable L-lactate. Thermodynamic relations secure a complete overall conversion because both the phosphoryl transfer and the redox potentials are
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associated with a gain in free energy [16]. Indeed, substrates were effectively pulled through the pathway without accumulation of intermediates. The one-pot protocol for preparation of GA6P with closed-loop cofactor regeneration not only proved practical because no intermediate has to be isolated, but also because the overall conversion rate was considerably higher than for a separate two-step procedure owing to the fact that inhibitory intermediates are maintained at a minimum concentration level. A disadvantage is that the enzymatic phosphorylation relies on PEP as a stoichiometric reagent, which upon whole-cell reactions will be supplied intrinsically by cellular metabolism but is not a sustainable resource for in vitro applications. In fact, PEP is marketed as an expensive fine chemical. However, both chemical or enzymatic syntheses require a high-energy phosphorylation reagent, and PEP is readily obtained in high purity on a multi-molar scale in just two fairly simple manipulations from pyruvate [19]. As an alternative, less expensive phosphoryl sources such as acetyl phosphate or polyphosphate in combination with appropriate kinases can likewise be considered [17]. A different option is to avoid an ATP dependent kinase with need for cofactor recycling by applying an acid phosphatase for transphosphorylation from inexpensive pyrophosphate as donor. However, such methods often suffer from low regioselectivity and competing product hydrolysis [20]. D-Tagatose
1,6-bisphosphate
The degradation of lactose, D-galactose and galactitol proceeds in bacteria by a pathway capitalizing on the aldol cleavage of the intermediate D-tagatose 1,6-bisphosphate (TBP) [21]. When for assay purposes TBP was available only by a low yielding (2%), multistep synthesis from D-galacturonic acid [22], we had developed an alternative multi-enzymatic synthesis from dihydroxyacetone (DHA) [23,24]. After enzymatic phosphorylation of DHA using ATP and glycerol kinase, combined with cofactor regeneration based on PEP (Fig. 1B), the generated dihydroxyacetone phosphate (DHAP) first is partially isomerized in situ into D-glyceraldehyde 3-phosphate (GAP), then added to the latter in a stereoselective carboligation step catalyzed by tagatose-1,6bisphosphate aldolase (TagA) to furnish TBP. In principle, the synthesis strategy was based on a reversal of the catabolic route, which to become practical requires a high equilibrium concentration of TBP in the aldol reaction. However, as a consequence of its all-cis substitution in the furanose forms present in aqueous solution, the TBP product indeed is only barely favored (Keq = [DHAP] [GAP]/[TBP] = 103 M) [23]. For this reason the TBP yield cannot significantly exceed a level of 80%, even with reaction parameters optimized. As a consequence, high concentration of triose phosphate substrates is required to shift equilibrium conditions toward higher productivity.
Rare ketosugar 1-phosphates Microbial metabolisms comprise a remarkable repertoire for the catabolic utilization of carbohydrates as the sole source of carbon and energy. A common feature of most degradative sequences is the conversion of sugars and polyols into ketose L-phosphates that by breakdown of the carbon backbone into smaller units enter the central metabolism. Since the phosphorylated ketoses frequently function as the inducers of respective metabolic enzymes, they are
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thus primary targets for the investigation of mechanistic and regulatory aspects of sugar metabolism. We have developed a practical route to various ketose L-phosphates that starts from commercially available aldoses or ketoses and makes use of a ketose kinase in combination with an appropriate ketol isomerase (Fig. 1C) [25,26]. All enzymes are derived from L-rhamnose/L-fucose catabolism and are accessible in quantity by recombinant overexpression. The basis for this development was the observation that, unlike the functionally related hexokinases, the L-rhamnulokinase (RhuK) from Escherichia coli has a relatively broad tolerance for structural modifications of its natural substrate L-rhamnu1ose, except for the (3R)-configuration. The scheme is completed by two corresponding ketol isomerases, the L-rhamnose isomerase (RhaI) and the L-fucose isomerase (FucI) from E. coli. In common with RhuK, both isomerases require a fixed (3R)-configuration but are flexible toward structural variations at the C4–C6 segment. In the isomerization equilibria, the ketose isomers often turned out to be the less abundant constituent for structural reasons [27]. The thermodynamically favorable phosphoryl transfer from ATP, however, serves to constantly drain the ketose component from the equilibrium and to drive the reactions to completion. As an example, the RhaI/RhuK combination provides a straightforward and high-yielding access to rare ketose L-phosphates having a common (2R,3R)-configuration from a range of stereochemically related aldohexoses or aldopentoses that serve as starting materials [25]. The construction of cell-free biocatalytic systems according to the Systems Biocatalysis strategy offers the freedom to design the most appropriate sequence of reactions to the target product. To achieve maximum efficacy in one-pot conversions it is obviously important to balance endothermic or thermoneutral reactions by coupling to exothermic ones. However, it is also important to be able to choose preferentially the thermodynamically most effective combination of reagents for cofactor recycling, and particularly that sequence where this critical step is properly placed at the very end. Clearly, such operational freedom in vitro sets the Systems Biocatalysis approach apart from whole-cell technology where such is not, or at least not easily, attainable in vivo.
Glycolysis intermediates as substrates Triose phosphates (DHAP, GAP) are versatile small building blocks for asymmetric synthesis because of their dense, but differentiated functionality and their high chemical reactivity that can be readily exploited in a selective fashion. In particular, these structures can be utilized for the highly stereoselective enzymatic generation of a complete set of stereoisomeric vicinal diols by employing aldolase technology [28,29]. Although much research has been directed at their chemical or biochemical synthesis [30,31], both compounds remain very expensive fine chemicals. In addition, they are labile to phosphate elimination in aqueous solution with generation of cell-toxic methylglyoxal [32,33], which renders production by whole-cell approaches unfeasible.
Dihydroxyacetone phosphate In the context of a research program targeted at C–C bond forming aldolases for asymmetric synthesis we had been searching for means to make the essential substrate DHAP efficiently available. In a pioneering study [10] published in 1992 we were attracted by
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the potential of multi-enzyme systems in vitro that would allow by a series of specific serial or parallel conversions of shared substrates along the glycolysis pathway to exploit sugars as an economical source of DHAP. Investigations into these enzyme cascade reactions had been initiated, firstly because of the potential preparative benefit, and secondly to gain insight into the complex kinetic effects operative in vitro for such a multiply regulated multienzyme system. Starting with the central function of fructose-1,6-bisphosphate aldolase (FruA) in glycolysis to cleave and – in combination with triose phosphate isomerase – symmetrize fructose 1,6-bisphosphate (FBP) into C3 fragments, we have assembled additional steps in such a manner (Fig. 2) that several regenerative raw materials (glucose, fructose, or saccharose) can be utilized alternatively as possible precursors. The overall artificial reaction network comprises seven purified enzymes from commercial sources that, including ATP cofactor regeneration at several levels to drive overall synthetic equilibria to the FBP stage, jointly deliver DHAP in situ for consecutive preparative use. It became apparent that the phosphofructokinase (PFK) remained a limiting factor, in accord with the notion that PFK is the central regulatory element of glycolysis that is inhibited by numerous effectors (e.g., ATP,
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PEP, pyruvate). In metabolic engineering a critical element in a pathway is replaced by a homologous gene from another organism coding for an enzyme with superior properties [34]. Analogously, the artificial in vitro metabolism was engineered by replacing the mammalian PFK from rabbit muscle with the allosterically less regulated bacterial PFK-2 isoenzyme from E. coli [35,36], which significantly improved the flux to FBP production. Subsequently, to this scheme was combined an unnatural synthesis step by supplying an external aldehyde reactant to employ the FruA for a consecutive non-natural aldol reaction (Fig. 2A). Because aldol reactions are reversible, the overall equilibrium must be shifted toward product formation by using an excess of aldehyde. Relative to syntheses starting from pure FBP comparable yields were achieved with the multi-enzyme system. The method made available a selection of non-natural alkylated, spiro-annulated, deoxy and branched chain sugars that are difficult to prepare by classical chemical synthesis. Because this cell-free reaction scheme comprises an unusual accumulation of coupled reactions as a new combination of natural, catabolic processes with a nonphysiological, anabolic synthetic component by application of pure isolated enzymes in vitro, it was designated as the first artificial metabolism [10]. The reduced complexity as compared to living organisms makes in vitro biocatalytic systems readily amenable to pathway engineering and optimization. One further goal was to address the synthesis of stereoisomers different from those created by FruA catalysis, which requires supplementation of the metabolic scheme with a different aldolase such as fuculose-1-phosphate aldolase (FucA) having distinct stereospecificity [37]. Because FruA is mandatory for the aldol cleavage of FBP, the competing generation of stereoisomers was eliminated by replacing the class I FruA from rabbit muscle by its bacterial class II analog from E. coli, which has very narrow substrate specificity for GAP [38]. Indeed, the engineered in vitro metabolic pathway, including FucA from E. coli as an eighth enzyme in the cascade, produced diastereomerically pure products having the L-erythro rather than the D-threo configuration typical for FruA [38]. D-Glyceraldehyde
FIG. 1
One-pot multi-enzyme catalyzed synthesis of phosphorylated carbohydrate metabolites; in the flow charts starting materials (left) and products (right) are highlighted in blue. (A) Phosphoenolpyruvate (PEP) as a double-duty cofactor recycling reagent for in situ regeneration of ATP and NAD+. (B) Aldol synthesis of D-tagatose 1,6-bisphosphate (TagBP). (C) Coupled isomerization– phosphorylation for rare sugar 1-phosphate synthesis. Abbreviations used in the figure are: Glc6P, D-glucose 6-phosphate; GlcA6P, D-glucuronic acid 6phosphate; DHAP, dihydroxyacetone phosphate; GAP, D-glyceraldehyde 3phosphate; TagBP, D-tagatose 1,6-bisphosphate; PEP, phosphoenolpyruvate (PEP). Enzymes are indicated in bold: HK, hexokinase; PK, pyruvate kinase; GPDH, glucose-6-phosphate dehydrogenase; LDH, lactate dehydrogenase; GK, glycerol kinase; TPI, triosephosphate isomerase; TagA, tagatose-1,6bisphosphate aldolase; FucI, fucose isomerase; RhaI, rhamnose isomerase; RhuK, rhamnulokinase. 4
3-phosphate
For the preparation of 1-deoxyfructose 6-phosphate (dF6P) needed as an assay component for aldolase screening, we had envisioned a coupled enzymatic approach for its synthesis (Fig. 2B) based on the fructose 6-phosphate aldolase (FSA) catalyzed carboligation from GAP and hydroxyacetone [39]. FSA from E. coli is a member of the transaldolase class and specific for non-phosphorylated aldol donors [40,41]. Because of its low stability in solution, GAP is preferably generated and consumed in situ to avoid the build-up of high stationary concentrations, which is conveniently achieved by retro-aldol cleavage of stable FBP in the presence of FruA in concert with TPI. Both aldolases show a distinct and mutually exclusive, high substrate specificity (DHAP and ketoses 1-phosphates in the case of FruA, free ketoses in the case of FSA), which excludes any interference from substrates, intermediates, or products [40]. For the synthesis of dF6P, FBP was incubated with FruA, TPI and FSA in the presence of an excess of hydroxyacetone. The high concentration of the latter, necessary to propel the product formation in the all-equilibrium procedure, again required replacement of the conventional rabbit muscle FruA in the scheme
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by the corresponding recombinant FruA enzyme from Staphylococcus carnosus [42], which is highly stable and tolerant to enhanced levels of hydroxyacetone. The optimized method successfully delivered dF6P in 96% yield on a multi-gram scale [39]. Xylulose 5-phosphate (X5P), occurring as an intermediate in the pentose phosphate pathway, is a valuable substrate required for enzymatic assays, particularly for the screening of eucaryotic enzymes in anti-cancer drug design [43]. Because published protocols for the synthesis of X5P from free sugars or other phosphorylated precursors are tedious, micro-scale and/or low-yielding, we developed a practical and efficient procedure for the preparation of X5P by a multi-enzymatic route based on the stereospecific transketolase-catalyzed ketol transfer from hydroxypyruvate to GAP [44]. Again, retro-aldol fragmentation of FBP in the presence of FruA and TPI was considered as an in situ source of the triose phosphate electrophile (Fig. 2C). Although the equilibrium concentration of GAP is rather low, reaction rates were sufficiently high owing to the high affinity of transketolase for GAP and the release of CO2 resulting from decomposition of hydroxypyruvate, which renders the overall synthetic reaction practically irreversible. The product X5P could be isolated from this simple one-pot procedure in 82% overall yield on a multi-gram scale [44]. It ought to be noted that the above procedures all require the conversion of biogenic starting materials jointly with external non-natural chemicals (e.g., aldehydes, hydroxypyruvate), which are added in stoichiometric amounts and at high concentration. Owing to the high chemical reactivity of the latter components, this is readily tolerated by cell-free metabolic cascades that are composed of suitably selected stable and effective biocatalysts, but likewise would not be readily compatible with whole-cell technology because of toxicity issues.
Oxidative pathways for substrate generation Reactions involving elemental oxygen usually profit from a strong thermodynamic driving force by the high redox potential of oxygen reduction, which renders coupled reactions practically irreversible. In our quest for suitable DHAP sources, we have also investigated the potential of inexpensive flavin-dependent glycerol phosphate oxidase (GPO) from Streptococcus sp. for the air oxidation of L-glycerol 3-phosphate (Gly3P) as another method to generate DHAP in situ for consecutive use in aldol reactions (Fig. 3A) [45]. The reaction indeed proceeds smoothly and yields pure DHAP almost quantitatively. With GPO a separate cofactor regeneration step is obsolete because the reduced cofactor FAD (H2) remains tightly bound to the enzyme and is rapidly re-oxidized by elemental oxygen. Thereby, hydrogen peroxide is liberated, which must be decomposed by addition of catalase (Cat). In turn, as the flow chart suggests, hydrogen peroxide can be used alternatively as the oxidant [45,46]. In principle, Gly3P again could be obtained by coupled enzymatic phosphorylation of glycerol but this option had not been further pursued because of the low oxygen tolerance of available glycerol kinases [47]. However, oxidative DHAP generation could be smoothly coupled to its consumption in aldol reactions because the DHAP aldolases are not sensitive to oxygen. When compared to other established procedures, the GPO method usually furnished adducts of higher purity and in at least equal or higher yields. Interestingly, the GPO procedure proved also
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FIG. 3
One-pot multi-enzyme catalyzed carbohydrate synthesis by reversible aldol addition. The equilibrium of the carboligation reaction is driven by oxidative generation of the substrate(s) in situ using molecular oxygen. (A) Aldol synthesis of ketose 1-phosphates from glycerol phosphate. (B) Aldol synthesis of complex bicyclic 9-carbon sugar by carboligation from a galactoside precursor. Abbreviations used in the figure are: Gly3P, L-glycerol 3-phosphate; DHAP, dihydroxyacetone phosphate; GalOMe, b-methyl-D-galactoside; 6aldGalOMe, 6-aldehydo-b-methyl-D-galactoside. Enzymes are indicated in bold: GPO, glycerophosphate oxidase; Cat, catalase; GalO, galactose oxidase; RhuA, rhamnulose-1-phosphate aldolase.
adaptable to the synthesis of DHAP analogues, such as the corresponding bio-isosteric phosphonate and phosphorothioate, which are likewise accepted by the aldolases [48]. In a particularly interesting example (Fig. 3B), both the aldol donor and acceptor components for a subsequent carboligation were generated in situ by air oxidation using microbial oxidases [49]. For this purpose, the GPO technology for oxidative generation of DHAP as the nucleophilic component was coupled to an oxidative generation of the corresponding electrophilic aldehyde component. The latter was obtained by parallel oxidation of methyl b-D-galactopyranoside at the non-reducing end using the Cu(II)-dependent D-galactose oxidase (GalO) from Dactylium dendroides [50], and the system was completed by adding the highly substrate tolerant L-rhamnulose 1-phosphate aldolase (RhuA) from E. coli [37]. The ensuing aldol addition consumed both products in situ and thus obviated inhibition problems. This novel multi-enzymatic reaction system for the preparation of complex high-carbon sugar derivatives has the advantages of simplicity of operation, ready availability of starting materials at low cost, and high overall product yield [49].
Leloir-path sialoconjugation Sialic acid-containing glycoconjugates at the cell surface are of high importance in carbohydrate-mediated recognition phenomena in physiological and pathological events, including bacterial or viral infection [51]. Natural sialic acid derivatives typically incorporate various types of N- or O-modification but the resultant biological profiles are poorly understood because the individual compounds are not yet available in sufficient quantity and purity for testing [52]. The Leloir-type biosynthesis of sialic acid conjugates requires three dedicated enzymatic steps, which consist of a carboligation step to construct the extended sialic acid backbone, followed by nucleotide activation of the sugar and its regio- and stereospecific transfer to a suitable acceptor moiety (Fig. 4). Toward a generic method for the synthesis of sialoconjugate libraries [53], the catabolic neuraminic acid aldolase (NeuA) from
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FIG. 4
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One-pot, multi-enzyme catalyzed synthesis of sialoconjugates by Leloir-type sialyltransfer with in situ generation of natural and non-natural sialic acids by reversible carboligation. Abbreviations used in the figure are: Sia, sialic acid (N-acetylneuraminic acid and analogues); CMP-Sia,; GalOR, b-D-galactoside. Enzymes are indicated in bold: NeuA, neuraminic acid aldolase; CSS, CMPsialic acid synthase; 2,6SiaT, a2,6-sialyltransferase.
E. coli has been shown to offer broad substrate promiscuity for catalyzing the addition of pyruvate to N-acetyl-D-mannosamine and related aldoses to form various natural and non-natural sialic acids [29]. We have discovered that the CMP-sialic acid synthetase (CSS) from Neisseria meningitidis can activate an unusually wide range of sialic acid derivatives [54,55]. Allowable structural modifications included N-acyl variation as well as altered substitution pattern, backbone size, and inversion of stereoconfiguration. Recently, we have engineered the CSS by structure-guided sitespecific saturation mutagenesis to generate enzymes with significantly enhanced substrate promiscuity for sterically highly demanding N-acyl modified sialic acid analogues [56]. Top hits such as the F192S/F193Y variant display an improvement of up to 70-fold catalytic efficiency relative to wild-type CSS. CSS catalysis was then coupled in a cascade reaction system to an a2,6-sialyltransferase (2,6SiaT) conversion, which effects transfer of the sialyl moiety to a suitable galactoside acceptor. For this purpose, a novel highly substrate tolerant 2,6SiaT from Photobacterium leiognathi JTSHIZ-145 was applied, which best matches the requirements from the overall reaction system [57]. Rapid regio- and stereospecific sialyltransfer was achieved onto lactoside acceptors, and the coupled enzymatic system allowed preparing a library of
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neo-sialoconjugates (structural analogues of 60 -sialyllactosides) in a highly efficiently manner [55,56]. Cascade enzyme processes are especially important for glycosyltransferase-catalyzed reactions because they require nucleotideactivated substrates that are highly expensive and unstable but can be generated in situ from simpler starting materials using standard biosynthetic enzymes. Particularly, the development of promiscuous enzymes of the Leloir pathway is instrumental for the development of simplified, cascade-type synthetic procedures to facilitate rapid access to biologically important oligosaccharide structures for biomedical applications.
Summary and outlook This compilation is meant to illustrate the power of the Systems Biocatalysis approach, partly by highlighting early examples from the pioneer era in developing multi-step enzymatic reaction networks in vitro, partly from recent studies into more generic pathways using enzymes engineered for substrate promiscuity. It clearly demonstrates that cell-free artificial biocatalytic systems of high complexity can be designed and rapidly established, which are simple to operate and analyze for pathway engineering and process optimization, and which can be applied with high freedom to operate. Such is particularly evident for the synthesis of a wide range of non-natural fine chemicals that require reactive components that may not be compatible with living systems. The obvious drawback for creating in vitro biosystems currently still is the laborious purification of the parts, which usually makes exploratory laboratory developments unsustainable beyond the research scale. However, opportunities are emerging to significantly reduce costs such as with the use of recombinant enzymes from thermophilic organisms that can be purified by simple heat-shock treatment [58]. Together with the power of modern synthetic biology to identify replacement parts from nature, or to engineer improved parts by evolution [59], the concept of ‘‘artificial metabolisms’’ for sustainable synthesis in vitro is expected to make an important contribution to the development of productive biosystems of the future.
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