Chlamydomonas reinhardii cells, after a period of dark anaerobic adaptation, evolve H2 not only in the dark but also in the light. Our results show that high irradiances impair prolonged H2 evolution, while under low irradiances or darkness H2 evolution proceeds for more than 50 hours. NO3-and NO2-suppress H2 evolution both in the dark or under low irradiance. Apparently the cells prefer these oxidized nitrogen sources to protons as electron acceptors, since both N03-and NOTbecome reduced to NHW+, which is excreted to the culture medium in high amounts. H2 evolution started once these oxidized anions were largely depleted from the medium. Moreover, H2 evolution was consistently associated with NH4I excretion even if NH4' was already present in high amounts in the medium. This observation indicates that the cells utilize not only their carbohydrate but also their protein reserves as sources of reducing power for H2 evolution. This conclusion was supported by the observation that when nitrogen-starved cells were made anaerobic in a nitrogen-free medium, they not only evolved H2 at very high rates but excreted concomitantly NH4W up to concentrations in the millimolar range.After some controversy following the pioneering work of Gaffron and Rubin (1 1) on H2 evolution by green algae, it is now widely accepted that two different pathways exist in these organisms for H2 photoproduction with either water or carbohydrates as electron donors (1). Direct coupling between oxygenic photosynthetic activity and the H2 evolving system has been demonstrated to take place preferentially during initial periods of light exposure (7,12,22). Alternatively, simultaneous evolution of CO2 and H2 have been observed both in the dark (15) and in a light-dependent process involving solely PSI (18). In these two latter cases, cellular carbon reserves provide ultimately the reducing equivalents for H2 evolution, with H+ acting as final electron acceptors. Since H2 evolution implies such a simple redox reaction, it should provide an effective pathway for the disposal of excess internal reducing power, especially under low 02 tensions (18,26). A similar relief valve operates in anaerobic bacteria (1).In photosynthetic eukaryotes, the reductive utilization ofNO3-is carried out by two different enzymes: NAD(P)H-nitrate reductase which reduces N03 to N02 , and reduced ferredoxin-nitrite reductase which reduces N02-to NH4' (14 to the medium, most probably to unload excess photosynthetically generated reducing power (2, 3). Moreover, it was found that in this organism inorganic nitrogen metabolism was modulated by blue light (4).We report here the effect ofthe physiological electron acceptors N03 and N02 on H2 production by anaerobically adapted cells of C. reinhardii. When either of these oxidized nitrogen compounds were present, H2 production was suppressed, while, depending on the oxidized nitrogen source, NO2-and/or NH4' were released into the medium. However, with NH4' as the only nitrogen source, H2 evolution was enhanced. Furthermo...

During directed evolution to functionally express the high redox potential laccase from the PM1 basidiomycete in Saccharomyces cerevisiae (Mate et al. 2010), the characteristic maximum absorption at the T1 copper site (Abs610T1Cu) was quenched, switching the typical blue colour of the enzyme to yellow. To determine the molecular basis of this colour change, we characterized the original wild-type laccase and its evolved mutant. Peptide printing and Maldi-TOF analysis confirmed the absence of contaminating protein traces that could mask the Abs610T1Cu, while conservation of the redox potential at the T1 site was demonstrated by spectroelectrochemical redox titrations. Both wild-type and evolved laccases were capable of oxidizing a broad range of substrates (ABTS, guaiacol, DMP, synapic acid) and they displayed similar catalytic efficiencies. The laccase mutant could only oxidize high redox potential dyes (Poly R478, Reactive Black 5, Azure B) in the presence of exogenous mediators, indicating that the yellow enzyme behaves like a blue laccase. The main consequence of over-expressing the mutant laccase was the generation of a six-residue N-terminal acidic extension, which was associated with the failure of the STE13 protease in the Golgi compartment giving rise to alternative processing. Removal of the N-terminal tail had a negative effect on laccase stability, secretion and its kinetics, although the truncated mutant remained yellow. The results of CD spectra analysis suggested that polyproline helixes were formed during the directed evolution altering spectral properties. Moreover, introducing the A461T and S426N mutations in the T1 environment during the first cycles of laboratory evolution appeared to mediate the alterations to Abs610T1Cu by affecting its coordinating sphere. This laccase mutant is a valuable departure point for further protein engineering towards different fates.

We have developed a sustainable three-stage process for the revaluation of cheese whey permeate into D-tagatose, a rare sugar with functional properties used as sweetener. The experimental conditions (pH, temperature, cofactors, etc.) for each step were independently optimized. In the first step, concentrated whey containing 180–200 g/L of lactose was fully hydrolyzed by β-galactosidase from Bifidobacterium bifidum (Saphera®) in 3 h at 45 °C. Secondly, glucose was selectively removed by treatment with Pichia pastoris cells for 3 h at 30 °C. The best results were obtained with 350 mg of cells (previously grown for 16 h) per mL of solution. Finally, L-arabinose isomerase US100 from Bacillus stearothermophilus was employed to isomerize D-galactose into D-tagatose at pH 7.5 and 65 °C, in presence of 0.5 mM MnSO4. After 7 h, the concentration of D-tagatose was approximately 30 g/L (33.3% yield, referred to the initial D-galactose present in whey). The proposed integrated process takes place under mild conditions (neutral pH, moderate temperatures) in a short time (13 h), yielding a glucose-free syrup containing D-tagatose and galactose in a ratio 1:2 (w/w).

Glycosylation of polyphenols may increase their aqueous solubility, stability, bioavailability and pharmacological activity. Herein, we used a mutant of sucrose phosphorylase from Thermoanaerobacterium thermosaccharolyticum engineered to accept large polyphenols (variant TtSPP_R134A) to produce phloretin glucosides. The reaction was performed using 10% (v/v) acetone as cosolvent. The selective formation of a monoglucoside or a diglucoside (53% and 73% maximum conversion percentage, respectively) can be kinetically controlled. MS and 2D‐NMR determined that the monoglucoside was phloretin 4’‐O‐α‐D‐glucopyranoside and the diglucoside phloretin‐4’‐O‐[α‐D‐glucopyranosyl‐(1→3)‐O‐α‐D‐glucopyranoside], a novel compound. The molecular features that determine the specificity of this enzyme for 4’‐OH phenolic group were analysed by induced‐fit docking analysis of each putative derivative, using the crystal structure of TtSPP and changing the mutated residue. The mono‐ and diglucoside were, respectively, 71‐ and 1200‐fold more soluble in water than phloretin at room temperature. The α‐glucosylation decreased the antioxidant capacity of phloretin, measured by DPPH and ABTS assays; however, this loss was moderate and the activity could be recovered upon deglycosylation in vivo. Since phloretin attracts a great interest in dermocosmetic applications, we analyzed the percutaneous absorption of glucosides and the aglycon employing a pig skin model. Although the three compounds were detected in all skin layers (except the fluid receptor), the diglucoside was present mainly on superficial layers.

The transglycosylation activity of a novel commercial β-galactosidase from Bifidobacterium bifidum (Saphera) was evaluated. The optimal conditions of operation of this enzyme, measured with o-nitrophenyl-β-D-galactopyranoside, were 40 °C and pH around 6.0. Although at low lactose concentrations the character of this enzyme was basically hydrolytic, an increase of lactose concentration to 400 g/L resulted in a significant formation (107.2 g/L, 27% yield) of prebiotic galactooligosaccharides (GOS).The maximum amount of GOS was obtained at a lactose conversion of approximately 90%, which contrasts with other β-galactosidases, for which the highest GOS yield is achieved at 40-50% lactose conversion. Using HPAEC-PAD, semipreparative HPLC-HILIC, MS, 1D and 2D NMR, we determined the structure of most of the GOS synthesized by this enzyme.The main identified products were Gal-β(13)-Gal-β(14)-Glc (3´-O-Beta-galactosyllactose), Gal-β(1→6)-Glc (allolactose), Gal-β(13)-Glc (3-galactosyl-glucose), Galβ(1→3)-Gal (3-galactobiose) and the tetrasaccharide Gal-β(13)-Gal-β(13)-Galβ(14)-Glc. In general, the B. bifidum β-galactosidase showed a tendency to form β(13) linkages followed by β(16), and more scarcely β(14).