Microalgal products are an emerging class of food, feed, and nutraceuticals. They include dewatered or dried biomass, isolated pigments, and extracted fat. The oil, protein, and antioxidant-rich microalgal biomass is used as a feed and food supplement formulated as pastes, powders, tablets, capsules, or flakes designed for daily use. Pigments such as astaxanthin (red), lutein (yellow), chlorophyll (green), or phycocyanin (bright blue) are natural food dyes used as isolated pigments or pigment-rich biomass. Algal fat extracted from certain marine microalgae represents a vegetarian source of n-3-fatty acids (eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA), γ-linolenic acid (GLA)). Gaining an overview of the production of microalgal products is a time-consuming task. Here, requirements and options of microalgae cultivation are summarized in a concise manner, including light and nutrient requirements, growth conditions, and cultivation systems. The rentability of microalgal products remains the major obstacle in industrial application. Key challenges are the high costs of commercial-scale cultivation, harvesting (and dewatering), and product quality assurance (toxin analysis). High-value food ingredients are commonly regarded as profitable despite significant capital expenditures and energy inputs. Improvements in capital and operational costs shall enable economic production of low-value food products going down to fishmeal replacement in the future economy.

Access to chiral alcohols of high optical purity is today frequently provided by the enzymatic reduction of precursor ketones. However, bioreductions are complicated by the need for reducing equivalents in the form of NAD(P)H. The high price and molecular weight of NAD(P)H necessitate in situ recycling of catalytic quantities, which is mostly accomplished by enzymatic oxidation of a cheap co-substrate. The coupled oxidoreduction can be either performed by free enzymes in solution or by whole cells. Reductase selection, the decision between cell-free and whole cell reduction system, coenzyme recycling mode and reaction conditions represent design options that strongly affect bioreduction efficiency. In this paper, each option was critically scrutinized and decision rules formulated based on well-described literature examples. The development chain was visualized as a decision-tree that can be used to identify the most promising route towards the production of a specific chiral alcohol. General methods, applications and bottlenecks in the set-up are presented and key experiments required to "test" for decision-making attributes are defined. The reduction of o-chloroacetophenone to (S)-1-(2-chlorophenyl)ethanol was used as one example to demonstrate all the development steps. Detailed analysis of reported large scale bioreductions identified product isolation as a major bottleneck in process design.

Little is known about how substrates bind to CtXR (Candida tenuis xylose reductase; AKR2B5) and other members of the AKR (aldo–keto reductase) protein superfamily. Modelling of xylose into the active site of CtXR suggested that Trp23, Asp50 and Asn309 are the main components of pentose-specific substrate-binding recognition. Kinetic consequences of site-directed substitutions of these residues are reported. The mutants W23F and W23Y catalysed NADH-dependent reduction of xylose with only 4 and 1% of the wild-type efficiency (kcat/Km) respectively, but improved the wild-type selectivity for utilization of ketones, relative to xylose, by factors of 156 and 471 respectively. Comparison of multiple sequence alignment with reported specificities of AKR members emphasizes a conserved role of Trp23 in determining aldehyde-versus-ketone substrate selectivity. D50A showed 31 and 18% of the wild-type catalytic-centre activities for xylose reduction and xylitol oxidation respectively, consistent with a decrease in the rates of the chemical steps caused by the mutation, but no change in the apparent substrate binding constants and the pattern of substrate specificities. The 30-fold preference of the wild-type for D-galactose compared with 2-deoxy-D-galactose was lost completely in N309A and N309D mutants. Comparison of the 2.4 Å (1 Å=0.1 nm) X-ray crystal structure of mutant N309D bound to NAD+ with the previous structure of the wild-type holoenzyme reveals no major structural perturbations. The results suggest that replacement of Asn309 with alanine or aspartic acid disrupts the function of the original side chain in donating a hydrogen atom for bonding with the substrate C-2(R) hydroxy group, thus causing a loss of transition-state stabilization energy of 8–9 kJ/mol.

Aerobic, hydrogen oxidizing bacteria are capable of efficient, non-phototrophic CO₂ assimilation, using H₂ as a reducing agent. The presence of explosive gas mixtures requires strict safety measures for bioreactor and process design. Here, we report a simplified, reproducible, and safe cultivation method to produce Cupriavidus necator H16 on a gram scale. Conditions for long-term strain maintenance and mineral media composition were optimized. Cultivations on the gaseous substrates H₂, O₂, and CO₂ were accomplished in an explosion-proof bioreactor situated in a strong, grounded fume hood. Cells grew under O₂ control and H₂ and CO₂ excess. The starting gas mixture was H₂:CO₂:O₂ in a ratio of 85:10:2 (partial pressure of O₂ 0.02 atm). Dissolved oxygen was measured online and was kept below 1.6 mg/L by a stepwise increase of the O₂ supply. Use of gas compositions within the explosion limits of oxyhydrogen facilitated production of 13.1 ± 0.4 g/L total biomass (gram cell dry mass) with a content of 79 ± 2% poly-(R)-3-hydroxybutyrate in a simple cultivation set-up with dissolved oxygen as the single controlled parameter. Approximately 98% of the obtained PHB was formed from CO₂.

Aldo‐keto reductases (AKRs) constitute a large protein superfamily of mainly NAD(P)‐dependent oxidoreductases involved in carbonyl metabolism. Catalysis is promoted by a conserved tetrad of active site residues (Tyr, Lys, Asp and His). Recent results of structure‐function relationship studies for xylose reductase (AKR2B5) require an update of the proposed catalytic mechanism. Electrostatic stabilization by the ε‐NH3+ group of Lys is a key source of catalytic power of xylose reductase. A molecular‐level analysis of the substrate binding pocket of xylose reductase provides a case of how a very broadly specific AKR achieves the requisite selectivity for its physiological substrate and could serve as the basis for the design of novel reductases with improved specificities for biocatalytic applications. iubmb Life, 58: 499‐507, 2006

Background: Whole cell-catalyzed biotransformation is a clear process option for the production of chiral alcohols via enantioselective reduction of precursor ketones. A wide variety of synthetically useful reductases are expressed heterologously in Escherichia coli to a high level of activity. Therefore, this microbe has become a prime system for carrying out whole-cell bioreductions at different scales. The limited capacity of central metabolic pathways in E. coli usually requires that reductase coenzyme in the form of NADPH or NADH be regenerated through a suitable oxidation reaction catalyzed by a second NADP⁺ or NAD⁺ dependent dehydrogenase that is co-expressed. Candida tenuis xylose reductase (Ct XR) was previously shown to promote NADH dependent reduction of aromatic α-keto esters with high Prelog-type stereoselectivity. We describe here the development of a new whole-cell biocatalyst that is based on an E. coli strain co-expressing Ct XR and formate dehydrogenase from Candida boidinii (Cb FDH). The bacterial system was evaluated for the synthesis of ethyl R-4-cyanomandelate under different process conditions and benchmarked against a previously described catalyst derived from Saccharomyces cerevisiae expressing Ct XR. Results: Gene co-expression from a pETDuet-1 vector yielded about 260 and 90 units of intracellular Ct XR and Cb FDH activity per gram of dry E. coli cell mass (g_CDW). The maximum conversion rate (r _S) for ethyl 4-cyanobenzoylformate by intact or polymyxin B sulphate-permeabilized cells was similar (2 mmol/g_CDWh), suggesting that the activity of Cb FDH was partly rate-limiting overall. Uncatalyzed ester hydrolysis in substrate as well as inactivation of Ct XR and Cb FDH in the presence of the α-keto ester constituted major restrictions to the yield of alcohol product. Using optimized reaction conditions (100 mM substrate; 40 g_CDW/L), we obtained ethyl R-4-cyanomandelate with an enantiomeric excess (e.e.) of 97.2% in a yield of 82%. By increasing the substrate concentration to 500 mM, the e.e. could be enhanced to ≅100%, however, at the cost of a 3-fold decreased yield. A recombinant strain of S. cerevisiae converted 100 mM substrate to 45 mM ethyl R-4-cyanomandelate with an e.e. of ≥ 99.9%. Modifications to the recombinant E. coli (cell permeabilisation; addition of exogenous NAD⁺) and addition of a water immiscible solvent (e.g. hexane or 1-butyl-3-methylimidazolium hexafluorophosphate) were not useful. To enhance the overall capacity for NADH regeneration in the system, we supplemented the original biocatalyst after permeabilisation with also permeabilised E. coli cells that expressed solely Cb FDH (410 U/g_CDW). The positive effect on yield (18% → 62%; 100 mM substrate) caused by a change in the ratio of FDH to XR activity from 2 to 20 was invalidated by a corresponding loss in product enantiomeric purity from 86% to only 71%. Conclusion: A whole-cell system based on E. coli co-expressing Ct XR and Cb FDH is a powerful and surprisingly robust biocatalyst for the synthesis of ethyl R-4-cyanomandelate in high optical purity and yield. A clear requirement for further optimization of the specific productivity of the biocatalyst is to remove the kinetic bottleneck of NADH regeneration through enhancement (≥ 10-fold) of the intracellular level of FDH activity.