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Synthetic biology requires well-characterized biological parts that can be combined into functional modules. One type of biological parts are transcriptional regulators and their cognate operator elements, which enable to either generate an input-specific response or are used as actuator modules. A range of regulators has already been characterized and used for orthogonal gene expression engineering, however, previous efforts have mostly focused on bacterial regulators. This work aims to design and explore the use of an archaeal TetR family regulator, FadRSa from Sulfolobus acidocaldarius, in a bacterial system, namely Escherichia coli. This is a challenging objective given the fundamental difference between the bacterial and archaeal transcription machinery and the lack of a native TetR-like FadR regulatory system in E. coli. The synthetic σ70-dependent bacterial promoter proD was used as a starting point to design hybrid bacterial/archaeal promoter/operator regions, in combination with the mKate2 fluorescent reporter enabling a readout. Four variations of proD containing FadRSa binding sites were constructed and characterized. While expressional activity of the modified promoter proD was found to be severely diminished for two of the constructs, constructs in which the binding site was introduced adjacent to the -35 promoter element still displayed sufficient basal transcriptional activity and showed up to 7-fold repression upon expression of FadRSa. Addition of acyl-CoA has been shown to disrupt FadRSa binding to the DNA in vitro. However, extracellular concentrations of up to 2 mM dodecanoate, subsequently converted to acyl-CoA by the cell, did not have a significant effect on repression in the bacterial system. This work demonstrates that archaeal transcription regulators can be used to generate actuator elements for use in E. coli, although the lack of ligand response underscores the challenge of maintaining biological function when transferring parts to a phylogenetically divergent host.

Glycolipids are target molecules in biotechnology and biomedicine as biosurfactants, biomaterials and bioactive molecules. An engineered E. coli strain for the production of glycoglycerolipids (GGL) used the MG517 glycolipid synthase from M. genitalium for glucosyl transfer from UDPGlc to diacylglycerol acceptor (Mora-Buye et al., 2012). The intracellular diacylglycerol pool proved to be the limiting factor for GGL production. Here we designed different metabolic engineering strategies to enhance the availability of precursor substrates for the glycolipid synthase by modulating fatty acids, acyl donor and phosphatidic acid biosynthesis. Knockouts of tesA, fadE and fabR genes involved in fatty acids degradation, overexpression of the transcriptional regulator FadR, the acyltransferases PlsB and C, and the pyrophosphatase Cdh for phosphatidic acid biosynthesis, as well as the phosphatase PgpB for conversion to diacylglycerol were explored with the aim of improving GGL titers. Among the different engineered strains, the ΔtesA strain co-expressing MG517 and a fusion PlsCxPgpB protein was the best producer, with a 350% increase of GGL titer compared to the parental strain expressing MG517 alone. Attempts to boost UDPGlc availability by overexpressing the uridyltransferase GalU or knocking out the UDP-sugar diphosphatase encoding gene ushA did not further improve GGL titers. Most of the strains produced GGL containing a variable number of glucosyl units from mono-to tetra-saccharides. Interestingly, the strains co-expressing Cdh showed a shift in the GGL profile towards the diglucosylated lipid (up to 80% of total GGLs) whereas the strains with a fadR knockout presented a higher amount of unsaturated acyl chains. In all cases, GGL production altered the lipidic composition of the E. coli membrane, observing that GGL replace phosphatidylethanolamine to maintain the overall membrane charge balance.

Fine-tuning biosynthetic pathways is crucial for the development of economic feasible microbial cell factories. Therefore, the use of computational models able to predictably design regulatory sequences for pathway engineering proves to be a valuable tool, especially for modifying genes at the translational level. In this study we developed a computational approach for the de novo design of 5'-untranslated regions (5'UTRs) in Saccharomyces cerevisiae with a predictive outcome on translation initiation rate. On the basis of existing data, a partial least-squares (PLS) regression model was trained and showed good performance on predicting protein abundances of an independent test set. This model was further used for the construction of a "yUTR calculator" that can design 5'UTR sequences with a diverse range of desired translation efficiencies. The predictive power of our yUTR calculator was confirmed in vivo by different representative case studies. As such, these results show the great potential of data driven approaches for reliable pathway engineering in S. cerevisiae.

Synthetic genetic sensors and circuits enable programmable control over timing and conditions of gene expression and, as a result, are increasingly incorporated into the control of complex and multi-gene pathways. Size and complexity of genetic circuits are growing, but stay limited by a shortage of regulatory parts that can be used without interference. Therefore, orthogonal expression and regulation systems are needed to minimize undesired crosstalk and allow for dynamic control of separate modules. This work presents a set of orthogonal expression systems for use in Escherichia coli based on heterologous sigma factors from Bacillus subtilis that recognize specific promoter sequences. Up to four of the analyzed sigma factors can be combined to function orthogonally between each other and toward the host. Additionally, the toolbox is expanded by creating promoter libraries for three sigma factors without loss of their orthogonal nature. As this set covers a wide range of transcription initiation frequencies, it enables tuning of multiple outputs of the circuit in response to different sensory signals in an orthogonal manner. This sigma factor toolbox constitutes an interesting expansion of the synthetic biology toolbox and may contribute to the assembly of more complex synthetic genetic systems in the future.

As flavonoids have beneficial health effects on humans, they are gaining increasing interest from pharmaceutical and health industries. However, current production methods, such as plant extraction and chemical synthesis, are inadequate to meet the demand. Therefore, microbial production might offer a promising alternative. During recent years, microbial strains able to produce flavonoids to a certain extent have been developed. However production titers are limited to the mg l−1 range, hampering the industrial exploitation of these strains. The latter will not be achieved by simply introducing the heterologous pathway in the production host and optimizing the fermentation process, but will depend on the interaction of different aspects of metabolic engineering and process engineering to overcome the current limitations. Next to engineering the production strain to optimize the availability of precursors, the pathway itself also requires intensive engineering. Currently utilized strategies result in a wide variety of different production strains, requiring high-throughput screening methods to identify optimal performing strains. As more and more organisms are being characterized, each with their own specific properties which might be beneficial for the heterologous production of flavonoids, the choice of the production host is another important aspect. Finally, the use of co-cultures might offer an alternative in which different parts of the process are performed by different organisms. This review aims to provide an overview of the research that has been done on these separate aspects. The work presented here could be used as a framework for further research.

Monitoring cellular behavior and eventually properly adapting cellular processes is key to handle the enormous complexity of today’s metabolic engineering questions. Hence, transcriptional biosensors bear the potential to augment and accelerate current metabolic engineering strategies, catalyzing vital advances in industrial biotechnology. The development of such transcriptional biosensors typically starts with exploring nature’s richness. Hence, in a first part, the transcriptional biosensor architecture and the various modi operandi are briefly discussed, as well as experimental and computational methods and relevant ontologies to search for natural transcription factors and their corresponding binding sites. In the second part of this review, various engineering approaches are reviewed to tune the main characteristics of these (natural) transcriptional biosensors, i.e., the response curve and ligand specificity, in view of specific industrial biotechnology applications, which is illustrated using success stories of transcriptional biosensor engineering.

Taxadiene-5α-Hydroxylase (CYP725A4) is a membrane-bound plant cytochrome P450 that catalyzes the oxidation of taxadiene to taxadiene-5α-ol. This oxidation is a key step in the production of the valuable cancer therapeutic and natural plant product, taxol. In this work, we report the bacterial expression and purification of six different constructs of CYP725A4. All six of these constructs are N-terminally modified and three of them are fused to cytochrome P450 reductase to form a chimera construct. The construct with the highest yield of CYP725A4 protein was then selected for substrate binding and kinetic analysis. Taxadiene binding followed type-1 substrate patterns with an observed KD of 2.1 μM ± 0.4 μM. CYP725A4 was further incorporated into nanoscale lipid bilayers (nanodiscs) and taxadiene metabolism was measured. Taxadiene metabolism followed Michaelis-Menten kinetics with an observed Vmax of 30 ± 8 pmol/min/nmolCYP725A4 and a KM of 123 ± 52 μM. Additionally, molecular operating environment (MOE) modeling was performed in order to gain insight into the interactions of taxadiene with CYP725A4 active site. Taken together, we demonstrate the successful expression and purification of the functional membrane-bound plant CYP, CYP725A4, in E. coli.

Microorganisms possess a plethora of regulatory mechanisms to tightly control the flux through their metabolic network, allowing optimal behaviour in response to environmental conditions. However, these mechanisms typically counteract metabolic engineering efforts to rewire the metabolism with a view to overproduction. Hence, overcoming flux control is key in the development of microbial cell factories, illustrated in this contribution using the strictly controlled hexosamine biosynthesis pathway. The hexosamine biosynthesis pathway has recently garnered attention as gateway for the industrial biotechnological production of numerous mono-, oligo- and polysaccharidic compounds, composed of, i.a., glucosamine, N-acetylglucosamine, and neuraminic acid and with a vast application potential in the health, comsetics, and agricultural sector. First, the various alternative pathways in eukaryotes and prokaryotes are discussed. Second, the main regulatory mechanisms on transcriptional, translational and post-translational control, and the strategies to circumvent these pathway bottlenecks are highlighted. These efforts can serve as an inspiration to tackle regulatory control when optimizing any microbial cell factory.

Transcriptional biosensors enable key applications in both metabolic engineering and synthetic biology. Due to nature's immense variety of metabolites, these applications require biosensors with a ligand specificity profile customized to the researcher's needs. In this work, chimeric biosensors were created by introducing parts of a donor regulatory circuit from Sinorhizobium meliloti, delivering the desired luteolin-specific response, into a nonspecific biosensor chassis from Herbaspirillum seropedicae. Two strategies were evaluated for the development of chimeric LysR-type biosensors with customized ligand specificity profiles toward three closely related flavonoids, naringenin, apigenin, and luteolin. In the first strategy, chimeric promoter regions were constructed at the biosensor effector module, while in the second strategy, chimeric transcription factors were created at the biosensor detector module. Via both strategies, the biosensor repertoire was expanded with luteolin-specific chimeric biosensors demonstrating a variety of response curves and ligand specificity profiles. Starting from the nonspecific biosensor chassis, a shift from 27.5% to 95.3% luteolin specificity was achieved with the created chimeric biosensors. Both strategies provide a compelling, faster, and more accessible route for the customization of biosensor ligand specificity, compared to de novo design and construction of each biosensor circuit for every desired ligand specificity.

Altering gene expression regulation by promoter engineering is a very effective way to fine-tune heterologous pathways in eukaryotic hosts. Typically, pathway building approaches in yeast still use a limited set of long, native promoters. With the today’s introduction of longer and more complex pathways, an expansion of this synthetic biology toolbox is necessary. In this study we elucidated the core promoter structure of the well-characterized yeast TEF1 promoter and determined the minimal length needed for sufficient protein expression. Furthermore, this minimal core promoter sequence was used for the creation of a promoter library covering different expression strengths. This resulted in a group of short, 69 bp promoters with an 8.0-fold expression range. One exemplar had a two and four times higher expression compared to the native CYC1 and ADH1 promoter, respectively. Additionally, as it was described that the protein expression range could be broadened by upstream activating sequences (UASs), we integrated earlier described single and multiple short, synthetic UASs in front of the strongest yeast core promoter. This approach resulted to further variation in protein expression and an overall promoter library spanning a 20-fold activity range and covering a length from 69 bp to maximally 129 bp. Furthermore, the robustness of this library was assessed on three alternative carbon sources besides glucose. As such, the suitability of short yeast core promoters for metabolic engineering applications on different media, either in an individual context or combined with UAS elements, was demonstrated.

Combinatorial engineering approaches are becoming increasingly popular, yet they are hindered by the lack of specialized techniques for both efficient introduction of sequence variability and assembly of numerous DNA parts, required for the construction of lengthy multigene pathways. In this contribution, we introduce a new combinatorial multigene pathway assembly scheme based on Single Strand Assembly (SSA) methods and Golden Gate Assembly, exploiting the strengths of both assembly techniques. With a minimum of intermediary steps and an accompanying set of well-characterized and ready-to-use genetic parts, the developed workflow allows effective introduction of various libraries and efficient assembly of multigene pathways. It was put to the test by optimizing the lycopene pathway as proof-of-principle. The here constructed libraries yield ample variation in lycopene production. In addition, good-performing transformants with a significantly higher lycopene production were obtained as compared to previously published reference strains. The best selected producer yielded 3-fold improvement in lycopene titers up to 448 mg lycopene/g CDW. The proposed workflow in combination with the accompanying sets of ready-to-use expression and carrier plasmids, will allow the combinatorial assembly of increasingly lengthy product pathways with minimal effort.

Recent advances in metabolic engineering have demonstrated the potential to exploit biological chemistry for the synthesis of complex molecules. Much of the progress to date has leveraged increasingly precise genetic tools to control the transcription and translation of enzymes for superior biosynthetic pathway performance. However, applying these approaches and principles to the synthesis of more complex natural products will require a new set of tools for enabling various classes of metabolic chemistries (i.e., cyclization, oxygenation, glycosylation, and halogenation) in vivo. Of these diverse chemistries, oxygenation is one of the most challenging and pivotal for the synthesis of complex natural products. Here, using Taxol as a model system, we use nature's favored oxygenase, the cytochrome P450, to perform high-level oxygenation chemistry in Escherichia coli. An unexpected coupling of P450 expression and the expression of upstream pathway enzymes was discovered and identified as a key obstacle for functional oxidative chemistry. By optimizing P450 expression, reductase partner interactions, and N-terminal modifications, we achieved the highest reported titer of oxygenated taxanes (∼570 ± 45 mg/L) in E. coli. Altogether, this study establishes E. coli as a tractable host for P450 chemistry, highlights the potential magnitude of protein interdependency in the context of synthetic biology and metabolic engineering, and points to a promising future for the microbial synthesis of complex chemical entities.

Ethylene glycol (EG) is an important commodity chemical with broad industrial applications. It is presently produced from petroleum or natural gas feedstocks in processes requiring consumption of significant quantities of non-renewable resources. Here, we report a novel pathway for biosynthesis of EG from the renewable sugar glucose in metabolically engineered Escherichia coli. Serine-to-EG conversion was first achieved through a pathway comprising serine decarboxylase, ethanolamine oxidase, and glycolaldehyde reductase. Serine provision in E. coli was then enhanced by overexpression of the serine-biosynthesis pathway. The integration of these two parts into the complete EG-biosynthesis pathway in E. coli allowed for production of 4.1 g/L EG at a cumulative yield of 0.14 g-EG/g-glucose, establishing a foundation for a promising biotechnology. Biotechnol. Bioeng. 2016;113: 376–383. © 2015 Wiley Periodicals, Inc.

Glycosylation of small molecules can significantly alter their properties such as solubility, stability, and/or bioactivity, making glycosides attractive and highly demanded compounds. Consequently, many biotechnological glycosylation approaches have been developed, with enzymatic synthesis and whole-cell biocatalysis as the most prominent techniques. However, most processes still suffer from low yields, production rates and inefficient UDP-sugar formation. To this end, a novel metabolic engineering strategy is presented for the in vivo glucosylation of small molecules in Escherichia coli W. This strategy focuses on the introduction of an alternative sucrose metabolism using sucrose phosphorylase for the direct and efficient generation of glucose 1-phosphate as precursor for UDP-glucose formation and fructose, which serves as a carbon source for growth. By targeted gene deletions, a split metabolism is created whereby glucose 1-phosphate is rerouted from the glycolysis to product formation (i.e., glucosylation). Further, the production pathway was enhanced by increasing and preserving the intracellular UDP-glucose pool. Expression of a versatile glucosyltransferase from Vitis vinifera (VvGT2) enabled the strain to efficiently produce 14 glucose esters of various hydroxycinnamates and hydroxybenzoates with conversion yields up to 100%. To our knowledge, this fast growing (and simultaneously producing) E. coli mutant is the first versatile host described for the glucosylation of phenolic acids in a fermentative way using only sucrose as a cheap and sustainable carbon source.

Glycosylation of small molecules like specialized (secondary) metabolites has a profound impact on their solubility, stability or bioactivity, making glycosides attractive compounds as food additives, therapeutics or nutraceuticals. The subsequently growing market demand has fuelled the development of various biotechnological processes, which can be divided in the in vitro (using enzymes) or in vivo (using whole cells) production of glycosides. In this context, uridine glycosyltransferases (UGTs) have emerged as promising catalysts for the regio- and stereoselective glycosylation of various small molecules, hereby using uridine diphosphate (UDP) sugars as activated glycosyldonors. This review gives an extensive overview of the recently developed in vivo production processes using UGTs and discusses the major routes towards UDP-sugar formation. Furthermore, the use of interconverting enzymes and glycorandomization is highlighted for the production of unusual or new-to-nature glycosides. Finally, the technological challenges and future trends in UDP-sugar based glycosylation are critically evaluated and summarized.

Leaping DNA read-and-write technologies, and extensive automation and miniaturization are radically transforming the field of biological experimentation by providing the tools that enable the cost-effective high-throughput required to address the enormous complexity of biological systems. However, standardization of the synthetic biology workflow has not kept abreast with dwindling technical and resource constraints, leading, for example, to the collection of multi-level and multi-omics large data sets that end up disconnected or remain under- or even unexploited.In this contribution, we critically evaluate the various efforts, and the (limited) success thereof, in order to introduce standards for defining, designing, assembling, characterizing, and sharing synthetic biology parts. The causes for this success or the lack thereof, as well as possible solutions to overcome these, are discussed.Akin to other engineering disciplines, extensive standardization will undoubtedly speed-up and reduce the cost of bioprocess development. In this respect, further implementation of synthetic biology standards will be crucial for the field in order to redeem its promise, i.e. to enable predictable forward engineering.

A problem rarely tackled by current DNA assembly methods is the issue of cloning additional parts into an already assembled construct. Costly PCR workflows are often hindered by repeated sequences, and restriction based strategies impose design constraints for each enzyme used. Here we present Protected Oligonucleotide Duplex Assisted Cloning (PODAC), a novel technique that makes use of an oligonucleotide duplex for iterative Golden Gate cloning using only one restriction enzyme. Methylated bases confer protection from digestion during the assembly reaction and are removed during replication in vivo, unveiling a new cloning site in the process. We used this method to efficiently and accurately assemble a biosynthetic pathway and demonstrated its robustness toward sequence repeats by constructing artificial CRISPR arrays. As PODAC is readily amenable to standardization, it would make a useful addition to the synthetic biology toolkit.

The rapid and efficient assembly of multi-step metabolic pathways for generating microbial strains with desirable phenotypes is a critical procedure for metabolic engineering, and remains a significant challenge in synthetic biology. Although several DNA assembly methods have been developed and applied for metabolic pathway engineering, many of them are limited by their suitability for combinatorial pathway assembly. The introduction of transcriptional (promoters), translational (ribosome binding site (RBS)) and enzyme (mutant genes) variability to modulate pathway expression levels is essential for generating balanced metabolic pathways and maximizing the productivity of a strain. We report a novel, highly reliable and rapid single strand assembly (SSA) method for pathway engineering. The method was successfully optimized and applied to create constructs containing promoter, RBS and/or mutant enzyme libraries. To demonstrate its efficiency and reliability, the method was applied to fine-tune multi-gene pathways. Two promoter libraries were simultaneously introduced in front of two target genes, enabling orthogonal expression as demonstrated by principal component analysis. This shows that SSA will increase our ability to tune multi-gene pathways at all control levels for the biotechnological production of complex metabolites, achievable through the combinatorial modulation of transcription, translation and enzyme activity.

The increasing public awareness of adverse health impacts from excessive sugar consumption has created increasing interest in plant-derived, natural low-calorie or zero-calorie sweeteners. Two plant species which contain natural sweeteners, Stevia rebaudiana and Siraitia grosvenorii, have been extensively profiled to identify molecules with high intensity sweetening properties. However, sweetening ability does not necessarily make a product viable for commercial applications. Some criteria for product success are proposed to identify which targets are likely to be accepted by consumers. Limitations of plant-based production are discussed, and a case is put forward for the necessity of biotechnological production methods such as plant cell culture or microbial fermentation to meet needs for commercial-scale production of natural sweeteners.

The GNB/LNB (galacto- N -biose/lacto- N -biose) pathway plays a crucial role in bifidobacteria during growth on human milk or mucin from epithelial cells. It is thought to be the major route for galactose utilization in Bifidobacterium longum as it is an energy-saving variant of the Leloir pathway. Both pathways are present in B. bifidum , and galactose 1-phosphate (gal1P) is considered to play a key role. Due to its toxic nature, gal1P is further converted into its activated UDP-sugar through the action of poorly characterized uridylyltransferases. In this study, three uridylyltransferases ( galT1 , galT2 , and ugpA ) from Bifidobacterium bifidum were cloned in an Escherichia coli mutant and screened for activity on the key intermediate gal1P. GalT1 and GalT2 showed UDP-glucose-hexose-1-phosphate uridylyltransferase activity (EC 2.7.7.12), whereas UgpA showed promiscuous UTP-hexose-1-phosphate uridylyltransferase activity (EC 2.7.7.10). The activity of UgpA toward glucose 1-phosphate was about 33-fold higher than that toward gal1P. GalT1, as part of the bifidobacterial Leloir pathway, was about 357-fold more active than GalT2, the functional analog in the GNB/LNB pathway. These results suggest that GalT1 plays a more significant role than previously thought and predominates when B. bifidum grows on lactose and human milk oligosaccharides. GalT2 activity is required only during growth on substrates with a GNB core such as mucin glycans.

In natural 1,3-propanediol (PDO) producing microorganisms such as Klebsiella pneumoniae, Citrobacter freundii and Clostridium sp., the genes coding for PDO producing enzymes are grouped in a dha cluster. This article describes the dha cluster of a novel candidate for PDO production, Citrobacter werkmanii DSM17579 and compares the cluster to the currently known PDO clusters of Enterobacteriaceae and Clostridiaceae. Moreover, we attribute a putative function to two previously unannotated ORFs, OrfW and OrfY, both in C. freundii and in C. werkmanii: both proteins might form a complex and support the glycerol dehydratase by converting cob(I)alamin to the glycerol dehydratase cofactor coenzyme B12. Unraveling this biosynthesis cluster revealed high homology between the deduced amino acid sequence of the open reading frames of C. werkmanii DSM17579 and those of C. freundii DSM30040 and K. pneumoniae MGH78578, i.e., 96 and 87.5 % identity, respectively. On the other hand, major differences between the clusters have also been discovered. For example, only one dihydroxyacetone kinase (DHAK) is present in the dha cluster of C. werkmanii DSM17579, while two DHAK enzymes are present in the cluster of K. pneumoniae MGH78578 and Clostridium butyricum VPI1718.

The interactions between the intracellular metabolome, fluxome and growth rate of Escherichia coli after sudden glycolytic/gluconeogenic substrate shifts are studied based on pulses of different substrates to an aerobic glucose-limited steady-state (dilution rate=0.1h(-1)). After each added glycolytic (glucose) and gluconeogenic (pyruvate and succinate) substrate pulse, no by-products were secreted and a pseudo steady state in flux and metabolites was achieved in about 30-40s. In the pulse experiments a large oxygen uptake capacity of the cells was observed. The in vivo dynamic responses showed massive reorganization and flexibility (1/100-14-fold change) of extra/intracellular metabolic fluxes, matching with large changes in the concentrations of intracellular metabolites, including reversal of reaction rate for pseudo/near equilibrium reactions. The coupling of metabolome and fluxome could be described by Q-linear kinetics. Remarkably, the three different substrate pulses resulted in a very similar increase in growth rate (0.13-0.3h(-1)). Data analysis showed that there must exist as yet unknown mechanisms which couple the protein synthesis rate to changes in central metabolites.

Natural product metabolic engineering potentially offers sustainable and affordable access to numerous valuable molecules. However, challenges in characterizing and assembling complex biosynthetic pathways have prevented more rapid progress in this field. The anticancer agent Taxol represents an excellent case study. Assembly of a biosynthetic pathway for Taxol has long been stalled at its first functionalization, putatively an oxygenation performed by the cytochrome P450 CYP725A4, due to confounding characterizations. Here, through combined in vivo (Escherichia coli), in vitro (lipid nanodisc), and metabolite stability assays, we verify the presence and likely cause of this enzyme's inherent promiscuity. Thereby, we remove the possibility that promiscuity simply existed as an artifact of previous metabolic engineering approaches. Further, spontaneous rearrangement and the stabilizing effect of a hydrophobic overlay suggest a potential role for nonenzymatic chemistry in Taxol's biosynthesis. Taken together, this work confirms taxadiene-5α-ol as a primary enzymatic product of CYP725A4 and provides direction for future Taxol metabolic and protein engineering efforts.

Impressive advances in the field of synthetic biology go hand in hand with the discovery, design, and use of novel DNA and RNA elements. Efficient synthesis of large oligonucleotides and double-stranded DNA parts, chip-based synthesis of DNA libraries, and a detailed understanding of fundamental biological mechanisms and increased capacities in bioinformatics enable new findings and applications.

WOS: 000368884500001, PubMed ID: 26822953, Background: Imbalance in cofactors causing the accumulation of intermediates in biosynthesis pathways is a frequently occurring problem in metabolic engineering when optimizing a production pathway in a microorganism. In our previous study, a single knock-out Citrobacter werkmanii Delta dhaD was constructed for improved 1,3-propanediol (PDO) production. Instead of an enhanced PDO concentration on this strain, the gene knock-out led to the accumulation of the toxic intermediate 3-hydroxypropionaldehyde (3-HPA). The hypothesis was emerged that the accumulation of this toxic intermediate, 3-HPA, is due to a cofactor imbalance, i.e. to the limited supply of reducing equivalents (NADH). Here, this bottleneck is alleviated by rationally engineering cell metabolism to balance the cofactor supply. Results: By eliminating non-essential NADH consuming enzymes (such as lactate dehydrogenase coded by ldhA, and ethanol dehydrogenase coded by adhE) or by increasing NADH producing enzymes, the accumulation of 3-HPA is minimized. Combining the above modifications in C. werkmanii Delta dhaD resulted in the strain C. werkmanii Delta dhaD Delta ldhA.adhE, Promotion of Innovation through Science and Technology in Flanders (IWT-Vlaanderen)Institute for the Promotion of Innovation by Science and Technology in Flanders (IWT) [B/14045/07]; Multidisciplinary Research Partnership Ghent Bio-Economy, The authors wish to thank the Institute for the Promotion of Innovation through Science and Technology in Flanders (IWT-Vlaanderen) for financial support in the framework of the PhD grant (B/14045/07) of Maervoet V. The research was also supported by the Multidisciplinary Research Partnership Ghent Bio-Economy. The authors wish to thank dr. Ajikumar Parayil for valuable comments on and edits to the manuscript which was improved by his helpful suggestions.

Transcription is a complex and dynamic process representing the first step in gene expression that can be readily controlled through current tools in molecular biology. Elucidating and subsequently controlling transcriptional processes in various prokaryotic and eukaryotic organisms have been a key element in translational research, yielding a variety of new opportunities for scientists and engineers. This chapter aims to give an overview of how the fields of molecular and synthetic biology have contributed both historically and presently to the state of the art in transcriptional engineering. The described tools and techniques, as well as the emerging genetic circuit engineering discipline, open the door to new advances in the fields of medical and industrial biotechnology.

Industrial biotechnology promises to revolutionize conventional chemical manufacturing in the years ahead, largely owing to the excellent progress in our ability to re-engineer cellular metabolism. However, most successes of metabolic engineering have been confined to over-producing natively synthesized metabolites in E. coli and S. cerevisiae. A major reason for this development has been the descent of metabolic engineering, particularly secondary metabolic engineering, to a collection of demonstrations rather than a systematic practice with generalizable tools. Synthetic biology, a more recent development, faces similar criticisms. Herein, we attempt to lay down a framework around which bioreaction engineering can systematize itself just like chemical reaction engineering. Central to this undertaking is a new approach to engineering secondary metabolism known as ‘multivariate modular metabolic engineering’ (MMME), whose novelty lies in its assessment and elimination of regulatory and pathway bottlenecks by re-defining the metabolic network as a collection of distinct modules. After introducing the core principles of MMME, we shall then present a number of recent developments in secondary metabolic engineering that could potentially serve as its facilitators. It is hoped that the ever-declining costs of de novo gene synthesis; the improved use of bioinformatic tools to mine, sort and analyze biological data; and the increasing sensitivity and sophistication of investigational tools will make the maturation of microbial metabolic engineering an autocatalytic process. Encouraged by these advances, research groups across the world would take up the challenge of secondary metabolite production in simple hosts with renewed vigor, thereby adding to the range of products synthesized using metabolic engineering., National Institutes of Health (U.S.) (1-R01-GM085323-01A1), Special Research Funds BOF (BOF08/PDO/014), Research Foundation Flanders (FWO-Vlaandern V.4.174.10.N.01)

Deletion of both iclR and arcA in E. coli profoundly alters the central metabolic fluxes and decreases acetate excretion by 70%. In this study we investigate the metabolic consequences of both deletions in E. coli BL21 (DE3). No significant differences in biomass yields, acetate yields, CO(2) yields and metabolic fluxes could be observed between the wild type strain E. coli BL21 (DE3) and the double-knockout strain E. coli BL21 (DE3) ΔarcAΔiclR. This proves that arcA and iclR are poorly active in the BL21 wild type strain. Noteworthy, both strains co-assimilate glucose and acetate at high glucose concentrations (10-15 g l(-1)), while this was never observed in K12 strains. This implies that catabolite repression is less intense in BL21 strains compared to in E. coli K12.

In this study we validated the use of 24 square deepwell microtiterplates to screen large libraries of metabolically engineered strains by investigating the optimization of succinate production. Wild type E. coli MG1655 and 11 derived mutants were physiologically evaluated by growth in 24 deepwell MTPs and 2L benchtop bioreactors. Growth parameters, product yields and byproduct formation were determined for all mutants. The results show that similar average values and standard deviations for these parameters were obtained. Especially a high correlation was noticed for the acetate byproduct yield and the succinate production rate. For these parameters there was no significant difference for 8 out of 12 strains between MTPs and 2L bioreactors. However a lower maximum growth rate was observed in 2L reactors as opposed to 24 deepwell plates for 9 out of 12 mutants which could be linked to the higher amount of dead cells in the benchtop bioreactors (12% vs. 2% in MTPs). Finally, a cluster-based approach was used to select good producer strains, i.e. strains with a high succinate yield and succinate production rate. Bad, intermediate and good producer strains were clustered in the same groups for MTPs and benchtop bioreactors for 11 out of the 12 investigated strains.

1,3-Propanediol (PDO) is the starting point of a new-generation polymer with superior properties which is used in the textile and carpet industry. This product is mainly produced chemically, but high pressure, high temperature and expensive catalysts are required and toxic intermediates are released. Therefore, the biological production route has been studied intensively. DuPont and Genencor International, Inc. have modified Escherichia coli genetically so that this organism could produce PDO from glucose. However, due to the tremendous growth of the biofuel industry, a glycerol surplus has been created, so more and more researchers started to investigate the natural producing strains for the production of PDO from glycerol. Several metabolic engineering techniques have been used to enhance the production of PDO. This contribution gives an overview of the different strategies to increase the final titer, yield, and productivity of 1,3-propanediol in non-natural and natural producing strains.

To reduce the amount of consumables and number of pipetting steps in high-throughput screening, a constitutive expression system was developed that comprises four different promoters of varying strength. The system was validated by the expression of different sucrose phosphorylase enzymes from Leuconostoc mesenteroides, Lactobacillus acidophilus and Bifidobacterium adolescentis in 96-deep-and low-well plates at three temperatures. Drastically improved soluble expression in mini-cultures was observed for the enzymes from L. mesenteroides strains by reducing the promoter strength from strong to intermediate and by expressing the proteins at lower temperatures. In contrast, the enzymes from B. adolescentis and L. acidophilus were expressed most efficiently with a strong promoter. The constitutive expression of sucrose phosphorylases in low-well plates resulted in a level of activity that is equal or even better than what was achieved by inducible expression. Therefore, our plasmid set with varying constitutive promoters will be an indispensable tool to optimize enzyme expression for high-throughput screening.

The increased consciousness for environmental issues and the depletion of mineral oil reserves led to the search for alternative energy sources but also for alternative biochemical processes. One of these chemicals that is identified to have great economical potential in a biobased economy is succinic acid. This chemical is a precursor for various high value-added derivatives which have application in the detergent/surfactant market, the ion chelator market, the food market and the pharmaceutical market. This review investigates the goals and preconditions to have an economical viable bio succinic acid process. The different production hosts for bio succinic acid are examined and the metabolic engineering strategies and possibilities are discussed. Finally, the state of the art of bio succinic acid production processes is critically evaluated in function of the production host, media, fermentation strategy, titers and yields.

Escherichia coli is one of the most widely used hosts for the production of recombinant proteins, among other reasons because its genetics are far better characterized than those of any other microorganism. To improve the understanding of recombinant protein synthesis in E. coli, the production of a model recombinant protein, beta-galactosidase, was studied in response to the constitutive overexpression of the anaplerotic reaction afforded by PEP carboxylase. To this end, an IPTG wash-in experiment was performed starting from a well-defined steady-state condition for both the wild-type E. coli and a mutant with a defective acetate pathway and a constitutively overexpressed ppc. In order to compare the dynamics of the fluxes over time during the wash-in experiment, a method referred to as transient metabolic flux analysis, which is based on steady-state metabolic flux analysis, was used. This allowed us to track the intracellular changes/fluxes in both strains. It was observed that the flux towards fermentation products was 3.6 times lower in the ppc overexpression mutant compared to the wild-type E. coli. In the former on the other hand, the PPC flux is in general higher. In addition, the flux towards beta-galactosidase was higher (12.4 times), resulting in five times more protein activity. These results indicate that by constitutively overexpressing the anaplerotic ppc gene in E. coli, the TCA cycle intermediates are increasingly replenished. The additional supply of these protein precursors has a positive result on recombinant protein production.

Synthetic biology, in close concert with systems biology, is revolutionizing the field of metabolic engineering by providing novel tools and technologies to rationally, in a standardized way, reroute metabolism with a view to optimally converting renewable resources into a broad range of bio-products, bio-materials and bio-energy. Increasingly, these novel synthetic biology tools are exploiting the extensive programmable nature of RNA, vis-à-vis DNA- and protein-based devices, to rationally design standardized, composable, and orthogonal parts, which can be scaled and tuned promptly and at will. This review gives an extensive overview of the recently developed parts and tools for i) modulating gene expression ii) building genetic circuits iii) detecting molecules, iv) reporting cellular processes and v) building RNA nanostructures. These parts and tools are becoming necessary armamentarium for contemporary metabolic engineering. Furthermore, the design criteria, technological challenges, and recent metabolic engineering success stories of the use of RNA devices are highlighted. Finally, the future trends in transforming metabolism through RNA engineering are critically evaluated and summarized. publisher: Elsevier articletitle: Putting RNA to work: Translating RNA fundamentals into biotechnological engineering practice journaltitle: Biotechnology Advances articlelink: http://dx.doi.org/10.1016/j.biotechadv.2015.10.011 content_type: article copyright: Copyright © 2015 Elsevier Inc. All rights reserved. ispartof: Biotechnology Advances vol:3 issue:8 pages:1829-1844 ispartof: location:England status: published

Background Flavonoids are bio-active specialized plant metabolites which mainly occur as different glycosides. Due to the increasing market demand, various biotechnological approaches have been developed which use Escherichia coli as a microbial catalyst for the stereospecific glycosylation of flavonoids. Despite these efforts, most processes still display low production rates and titers, which render them unsuitable for large-scale applications. Results In this contribution, we expanded a previously developed in vivo glucosylation platform in E. coli W, into an efficient system for selective galactosylation and rhamnosylation. The rational of the novel metabolic engineering strategy constitutes of the introduction of an alternative sucrose metabolism in the form of a sucrose phosphorylase, which cleaves sucrose into fructose and glucose 1-phosphate as precursor for UDP-glucose. To preserve these intermediates for glycosylation purposes, metabolization reactions were knocked-out. Due to the pivotal role of UDP-glucose, overexpression of the interconverting enzymes galE and MUM4 ensured the formation of both UDP-galactose and UDP-rhamnose, respectively. By additionally supplying exogenously fed quercetin and overexpressing a flavonol galactosyltransferase (F3GT) or a rhamnosyltransferase (RhaGT), 0.94 g/L hyperoside (quercetin 3-O-galactoside) and 1.12 g/L quercitrin (quercetin 3-O-rhamnoside) could be produced, respectively. In addition, both strains showed activity towards other promising dietary flavonols like kaempferol, fisetin, morin and myricetin. Conclusions Two E. coli W mutants were engineered that could effectively produce the bio-active flavonol glycosides hyperoside and quercitrin starting from the cheap substrates sucrose and quercetin. This novel fermentation-based glycosylation strategy will allow the economically viable production of various glycosides. Electronic supplementary material The online version of this article (doi:10.1186/s12934-015-0326-1) contains supplementary material, which is available to authorized users.

A 2918 bp sequence coding for the orotidine-5'-monophosphate decarboxylase enzyme (OMPD) was isolated from the genome of Myrothecium gramineum. This sequence was analysed and, remarkably, it is the first OMPD gene of a Sordariomycete that has an intron. The gene codes for an enzyme of 282 amino acids. The nucleotide sequence and the amino acid sequence were compared with fungal OMPD sequences. They show the highest similarity to OMPD genes and enzymes of Aspergillus sp., Penicillium sp. and Cladosporium fulvum. The functionality of the gene as a selection marker was proven by complementation of the uracil auxotrophy of Aspergillus nidulans FGSC A722.

Despite the potential in utilizing microbial fermentation for chemical production, the field of industrial biotechnology still lacks a standard, universally applicable principle for strain optimization. A key challenge has been in finding and applying effective ways to address metabolic flux imbalances. Strategies based on rational design require significant a priori knowledge and often fail to take a holistic view of cellular metabolism. Combinatorial approaches enable more global searches but require a high-throughput screen. Here, we present the recent advances and promises of a novel approach to metabolic pathway and strain optimization called multivariate modular metabolic engineering (MMME). In this technique, key enzymes are organized into distinct modules and simultaneously varied based on expression to balance flux through a pathway. Because of its simplicity and broad applicability, MMME has the potential to systematize and revolutionize the field of metabolic engineering and industrial biotechnology.

Through microbial engineering, biosynthesis has the potential to produce thousands of chemicals used in everyday life. Metabolic engineering and synthetic biology are fields driven by the manipulation of genes, genetic regulatory systems, and enzymatic pathways for developing highly productive microbial strains. Fundamentally, it is the biochemical characteristics of the enzymes themselves that dictate flux through a biosynthetic pathway toward the product of interest. As metabolic engineers target sophisticated secondary metabolites, there has been little recognition of the reduced catalytic activity and increased substrate/product promiscuity of the corresponding enzymes compared to those of central metabolism. Thus, fine-tuning these enzymatic characteristics through protein engineering is paramount for developing high-productivity microbial strains for secondary metabolites. Here, we describe the importance of protein engineering for advancing metabolic engineering of secondary metabolism pathways. This pathway integrated enzyme optimization can enhance the collective toolkit of microbial engineering to shape the future of chemical manufacturing.

Since the development of recombinant DNA technology (Cohen et al., 1973), it became possible to express heterologous genes in proor eukaryotic hosts, i.e. genes which they naturally not express. This development enabled the production of all kinds of products of which the high-added value recombinant proteins, became increasingly important and as such boosted biopharmaceutical and industrial enzyme applications. Up to now, the FDA (Food and Drug Administration) and EMEA (European Medicines Agency) have licensed the application of more than 150 recombinant proteins to be used as a pharmaceutical (Ferrer-Miralles et al., 2009). Global sales of biopharmaceuticals are estimated to account for US$70–80 Billion today (Walsh, 2010). Industrial enzymes (e.g. proteases, amylases, lipases, cellulases, pullulanases, pectinases) are used in various industrial segments and the industrial enzyme market is still expanding, estimated to reach US$ 3.74 Billion by the year 2015 (Global Industry Analysts, 2011). To date, the majority of this industrial enzyme market value is generated by recombinant processes (Hodgson, 1994; Demain & Vaishnav, 2009).

Escherichia coli strains are widely used as host for the production of recombinant proteins. Compared to E. coli K12, E. coli BL21 (DE3) has several biotechnological advantages, such as a lower acetate yield and a higher biomass yield, which have a beneficial effect on protein production. In a previous study (BMC Microbiol. 2011, 11:70) we have altered the metabolic fluxes of a K12 strain (i.e. E. coli MG1655) by deleting the regulators ArcA and IclR in such a way that the biomass yield is remarkably increased, while the acetate production is decreased to a similar value as for BL21 (DE3). In this study we show that the increased biomass yield beneficially influences recombinant protein production as a higher GFP yield was observed for the double knockout strain compared to its wild type. However, at higher cell densities (>2 g L(-1) CDW), the GFP concentration decreases again, due to the activity of proteases which obstructs the application of the strain in high cell density cultivations. By further deleting the genes lon and ompT, which encode for proteases, this degradation could be reduced. Consequently, higher GFP yields were observed in the quadruple knockout strain as opposed to the double knockout strain and the MG1655 wild type and its yield approximates the GFP yield of E. coli BL21 (DE3), that is, 27±5 mg g(CDW)(-1) vs. 30±5 mg g(CDW)(-1), respectively.

Background Gene expression is regulated through a complex interplay of different transcription factors (TFs) which can enhance or inhibit gene transcription. ArcA is a global regulator that regulates genes involved in different metabolic pathways, while IclR as a local regulator, controls the transcription of the glyoxylate pathway genes of the aceBAK operon. This study investigates the physiological and metabolic consequences of arcA and iclR deletions on E. coli K12 MG1655 under glucose abundant and limiting conditions and compares the results with the metabolic characteristics of E. coli BL21 (DE3). Results The deletion of arcA and iclR results in an increase in the biomass yield both under glucose abundant and limiting conditions, approaching the maximum theoretical yield of 0.65 c-mole/c-mole glucose under glucose abundant conditions. This can be explained by the lower flux through several CO2 producing pathways in the E. coli K12 ΔarcAΔiclR double knockout strain. Due to iclR gene deletion, the glyoxylate pathway is activated resulting in a redirection of 30% of the isocitrate molecules directly to succinate and malate without CO2 production. Furthermore, a higher flux at the entrance of the TCA was noticed due to arcA gene deletion, resulting in a reduced production of acetate and less carbon loss. Under glucose limiting conditions the flux through the glyoxylate pathway is further increased in the ΔiclR knockout strain, but this effect was not observed in the double knockout strain. Also a striking correlation between the glyoxylate flux data and the isocitrate lyase activity was observed for almost all strains and under both growth conditions, illustrating the transcriptional control of this pathway. Finally, similar central metabolic fluxes were observed in E. coli K12 ΔarcA ΔiclR compared to the industrially relevant E. coli BL21 (DE3), especially with respect to the pentose pathway, the glyoxylate pathway, and the TCA fluxes. In addition, a comparison of the genome sequences of the two strains showed that BL21 possesses two mutations in the promoter region of iclR and rare codons are present in arcA implying a lower tRNA acceptance. Both phenomena presumably result in a reduced ArcA and IclR synthesis in BL21, which contributes to the similar physiology as observed in E. coli K12 ΔarcAΔiclR. Conclusions The deletion of arcA results in a decrease of repression on transcription of TCA cycle genes under glucose abundant conditions, without significantly affecting the glyoxylate pathway activity. IclR clearly represses transcription of glyoxylate pathway genes under glucose abundance, a condition in which Crp activation is absent. Under glucose limitation, Crp is responsible for the high glyoxylate flux, but IclR still represses transcription. Finally, in E. coli BL21 (DE3), ArcA and IclR are poorly expressed, explaining the similar fluxes observed compared to the ΔarcAΔiclR strain.

Background Metabolic engineering aims at channeling the metabolic fluxes towards a desired compound. An important strategy to achieve this is the modification of the expression level of specific genes. Several methods for the modification or the replacement of promoters have been proposed, but most of them involve time-consuming screening steps. We describe here a novel optimized method for the insertion of constitutive promoters (referred to as "promoter knock-in") whose strength can be compared with the native promoter by applying a promoter strength predictive (PSP) model. Results Our method was successfully applied to fine tune the ppc gene of Escherichia coli. While developing the promoter knock-in methodology, we showed the importance of conserving the natural leader region containing the ribosome binding site (RBS) of the gene of interest and of eliminating upstream regulatory elements (transcription factor binding sites). The gene expression was down regulated instead of up regulated when the natural RBS was not conserved and when the upstream regulatory elements were eliminated. Next, three different promoter knock-ins were created for the ppc gene selecting three different artificial promoters. The measured constitutive expression of the ppc gene in these knock-ins reflected the relative strength of the different promoters as predicted by the PSP model. The applicability of our PSP model and promoter knock-in methodology was further demonstrated by showing that the constitutivity and the relative levels of expression were independent of the genetic background (comparing wild-type and mutant E. coli strains). No differences were observed during scaling up from shake flask to bioreactor-scale, confirming that the obtained expression was independent of environmental conditions. Conclusion We are proposing a novel methodology for obtaining appropriate levels of expression of genes of interest, based on the prediction of the relative strength of selected synthetic promoters combined with an optimized promoter knock-in strategy. The obtained expression levels are independent of the genetic background and scale conditions. The method constitutes therefore a valuable addition to the genetic toolbox for the metabolic engineering of E. coli.

Three cytochrome P450 monooxygenases belonging to the CYP52 family were isolated from the genome of the sophorolipid-producing yeast Candida bombicola using degenerate PCR and genomic walking. One gene displayed high identity with the CYP52E members and was classified into this group (CYP52E3), whereas the other genes belonged to new groups: CYP52M and CYP52N. CYP52E3 and CYP52N1 turned out to be of no relevance for sophorolipid production, but show clear upregulation when the yeast cells are grown on alkanes as the sole carbon source. On the other hand, CYP52M1 is clearly upregulated during sophorolipid synthesis and very likely takes part in sophorolipid formation.

Many different extraction and analysis methods exist to determine the protein fraction of microbial cells. For metabolic engineering purposes it is important to have precise and accurate measurements. Therefore six different protein extraction protocols and seven protein quantification methods were tested and compared. Comparison was based on the reliability of the methods and boxplots of the normalized residuals. Some extraction techniques (SDS/chloroform and toluene) should never be used: the measurements are neither precise nor accurate. Bugbuster extraction combined with UV280 quantification gives the best results, followed by the combinations Sonication-UV280 and EasyLyse-UV280. However, if one does not want to use the quantification method UV280, one can opt to use Bugbuster, EasyLyse or sonication extraction combined with any quantification method with exception of the EasyLyse-BCA_P and Sonication-BCA_P combinations.

E. coli cells produce acetate as an extracellular coproduct of aerobic cultures. Acetate is undesirable because it retards growth and inhibits protein formation. Most process designs or genetic modifications to minimize acetate formation aim at balancing growth rate and oxygen consumption. In this research, three genetic approaches to reduce acetate formation were investigated: (1) direct reduction of the carbon flow to acetate (ackA-pta, poxB knock-out); (2) anticipation on the underlying metabolic and regulatory mechanisms that lead to acetate (constitutive ppc expression mutant); and (3) both (1) and (2). Initially, these mutants were compared to the wild-type E. coli via batch cultures under aerobic conditions. Subsequently, these mutants were further characterized using metabolic flux analysis on continuous cultures. It is concluded that a combination of directly reducing the carbon flow to acetate and anticipating on the underlying metabolic and regulatory mechanism that lead to acetate, is the most promising approach to overcome acetate formation and improve recombinant protein production. These genetic modifications have no significant influence on the metabolism when growing the micro-organisms under steady state at relatively low dilution rates (less than 0.4 h(-1)).

Escherichia coli remains the best established production organisms in industrial biotechnology. However, during aerobic fermentation runs at high growth rates, considerable amounts of acetate are accumulated as by-product. This by-product has negative effects on growth and protein production. Over the last 20 years, substantial research efforts have been spent to reduce acetate accumulation during aerobic growth of E. coli on glucose. From the onset it was clear that this quest should not be a simple nor uncomplicated one. Simple deletion of the acetate pathway, reduced the acetate accumulation, but instead other by-products were formed. This minireview gives a clear outline of these research efforts and the outcome of them, including bioprocess level approaches and genetic approaches. Recently, the latter seems to have some promising results.

Microbial metabolomics has received much attention in recent years mainly because it supports and complements a wide range of microbial research areas from new drug discovery efforts to metabolic engineering. Broadly, the term metabolomics refers to the comprehensive (qualitative and quantitative) analysis of the complete set of all low molecular weight metabolites present in and around growing cells at a given time during their growth or production cycle. This review focuses on the past, current and future development of various experimental protocols in the rapid developing area of metabolomics in the ongoing quest to reliably quantify microbial metabolites formed under defined physiological conditions. These developments range from rapid sample collection, instant quenching of microbial metabolic activity, extraction of the relevant intracellular metabolites as well as quantification of these metabolites using enzyme based and or modern high tech hyphenated analytical protocols, mainly chromatographic techniques coupled to mass spectrometry (LC-MS(n), GC-MS(n), CE-MS(n)), where n indicates the number of tandem mass spectrometry, and nuclear magnetic resonance spectroscopy (NMR).

L-Arabinose isomerase (E.C. 5.3.1.14) catalyzes the reversible isomerization between L-arabinose and L-ribulose and is highly selective towards L-arabinose. By using a directed evolution approach, enzyme variants with altered substrate specificity were created and screened in this research. More specifically, the screening was directed towards the identification of isomerase mutants with L-ribose isomerizing activity. Random mutagenesis was performed on the Escherichia coli L-arabinose isomerase gene (araA) by error-prone polymerase chain reaction to construct a mutant library. To enable screening of this library, a selection host was first constructed in which the mutant genes were transformed. In this selection host, the genes encoding for L-ribulokinase and L-ribulose-5-phosphate-4-epimerase were brought to constitutive expression and the gene encoding for the native L-arabinose isomerase was knocked out. L-Ribulokinase and L-ribulose-5-phosphate-4-epimerase are necessary to ensure the channeling of the formed product, L-ribulose, to the pentose phosphate pathway. Hence, the mutant clones could be screened on a minimal medium with L-ribose as the sole carbon source. Through the screening, two first-generation mutants were isolated, which expressed a small amount of L-ribose isomerase activity.