Characterization of erythrose reductases from filamentous fungi
© Jovanović et al.; licensee Springer. 2013
Received: 2 August 2013
Accepted: 3 August 2013
Published: 8 August 2013
Proteins with putative erythrose reductase activity have been identified in the filamentous fungi Trichoderma reesei, Aspergillus niger, and Fusarium graminearum by in silico analysis. The proteins found in T. reesei and A. niger had earlier been characterized as glycerol dehydrogenase and aldehyde reductase, respectively. Corresponding genes from all three fungi were cloned, heterologously expressed in Escherichia coli, and purified. Subsequently, they were used to establish optimal enzyme assay conditions. All three enzymes strictly require NADPH as cofactor, whereas with NADH no activity could be observed. The enzymatic characterization of the three enzymes using ten substrates revealed high substrate specificity and activity with D-erythrose and D-threose. The enzymes from T. reesei and A. niger herein showed comparable activities, whereas the one from F. graminearum reached only about a tenth of it for all tested substrates. In order to proof in vivo the proposed enzyme function, we overexpressed the erythrose reductase-encoding gene in T. reesei. An increased production of erythritol by the recombinant strain compared to the parental strain could be detected.
KeywordsTrichoderma reesei Aspergillus niger Fusarium graminearum Erythrose reductase Erythritol
Erythritol is a four-carbon sugar alcohol, which is applied as flavour enhancer, formulation aid, humectants, stabilizer, thickener, and as low-calorie sweetener, of which the latter is the main utilization. It has a natural occurrence in several foods including beer, sake, wine, soy sauce, water melon, pear and grape (O’Donnell and Kearsley ; Sreenath and Venkatesh ) and is well tolerated by the human body (Munro et al. ). Erythritol can be chemically synthesized from dialdehyde starch with a nickel catalyst at high temperatures, but this process is not stereospecific and low in yield, and therefore, not industrialized (Moon et al. ). Instead erythritol is produced in biotechnological processes using osmophilic yeasts obtained by random mutagenesis as Aureobasidium sp. (Ishizuka et al. ; Sasaki et al. ), Trichosporonoides sp. (Suh et al. ), (Torula sp. Oh et al. ), and Candida magnoliae (Koh et al. ; Ryu et al. ). As substrate a highly concentrated glucose solution (typically 40% (w/v)) is applied, which is gained from chemically and enzymatically hydrolyzed wheat- and cornstarch. It serves as carbon source and causes high osmotic pressure, which pushes the yeast to produce the osmolyte erythritol (reviewed by (Moon et al. )).
Even though the production of erythritol and the according enzyme, erythrose reductase, have been well studied in yeasts, no such enzymes have yet been identified in filamentous fungi. For this study the filamentous ascomycota Trichoderma reesei (telemorph Hypocrea jecorina, (Kuhls et al. )), Aspergillus niger, and Fusarium graminearum (telemorph Gibberella zeae) were chosen because of their great importance in biotechnology. The (hemi)cellulases of T. reesei are widely used in pulp and paper production (Buchert et al. ; Noé P. ; Welt ), food and feed industry (Galante ; Lanzarini ; Walsh et al. ), textile industry (Koo ; Kumar ; Pedersen ), and more recently, for 2nd generation biofuel (cellulose ethanol) production (Hahn-Hägerdal et al. ; Himmel et al. ; Ragauskas et al. ). A. niger is used for the production of organic acids, as citric acid and gluconic acid (Ruijter et al. ), for heterologous protein expression Archer and Turner (), as well as production of pectinases Bussink et al. (; Delgado et al. ; Parenicová et al. ) and hemicellulases, such as xylanases and arabinases Gielkens et al. (; van Peij et al. ). F. graminearum is a well studied filamentous fungus because of its relevance as plant pathogen that can infect numerous plants like cereals, but also dicotyledons (Pirgozliev et al. ; Urban et al. ). Additionally, it is also used in biotechnological applications such as heterologous protein expression (Royer et al. ).
The characterization of the enzyme performing this reduction, namely erythrose reductase, has been done for some yeasts e. g. by (Lee et al. (), (Lee et al. ), (Ookura et al. )), but until now no such enzyme has been identified for the above-mentioned filamentous fungi.
In this study, we identified by in silico analysis proteins in T. reesei, A. niger, and F. graminearum exhibiting a high sequence similarity to the erythrose reductase (ER1) from Trichosporonoides megachiliensis. Accordingly, in this manuscript the corresponding proteins from the three organisms are referred to the term Err1 (Erythrose reductase 1) for easier reading. The respective genes were cloned and their protein products were heterologously expressed and purified. All three putative Err1 proteins were characterized in enzymatic assays with respect to their substrate specificity to D-erythrose and nine other potential substrates. In order to do this, the optimal assay conditions (temperature and pH) for all three enzymes were determined before, and then their usages of the different substrates were tested. Finally, we aimed to prove the function of the putative erythrose reductase in vivo. Therefore, the corresponding T. reesei enzyme was overexpressed in this fungus and the production of erythritol in the recombinant strain was compared to the parental strain.
Materials and methods
Strains and cultivation conditions
The T. reesei strain QM6aΔtmus53 (Steiger et al. ), the A. niger strain N400 (CBS 120.49), and the F. graminearum strain PH1 (NRRL31084) were maintained on malt extract (MEX) agar, complete medium agar (Pontecorvo et al. ), and small nutrient agar (Brunner et al. ), respectively. The recombinant T. reesei strain PEC1, produced during this study, was maintained on MEX agar containing hygromycin B.
Cultivation in shakeflasks was performed in 1-l-Erlenmeyer flasks containing 250 ml (Mandels-Andreotti (MA) medium Mandels ) supplemented with 1% (w/v) D-xylose. For inoculation 109 conida per litre were used. Growth conditions were pH 5, 30°C, and 160 rpm shaking rate. For harvesting mycelia, samples of 60 ml were drawn after 24 h and 30 h. For short-term storage, mycelia were shock-frozen and kept in liquid nitrogen.
Oligonucleotides used during the study
Sequence (5′ - 3′) a
Vector construction for fungal transformation
For the construction of pGEX-err1T, pGEX-err1A, and pGEX-err1F the err1 gene was excised from pJET-1.2 by Eco RI/Bam HI digestion and inserted into pGEX-4T-2 (GE Healthcare Life sciences, Little Chalfont, Buckinghamshire, UK).
For the construction of pBJ-PEC1 the vector pRLMex30 Mach et al. (), which contains the hph gene flanked by the pki promoter and the cbh2 terminator, was used. The hph gene was removed by Nsi I/Xba I digestion and subsequently, err1, which was excised from JET-1.2 also by Nsi I/Xba I digestion, was inserted.
The protoplast transformation of T. reesei was performed as described by (Gruber et al. ). 5 μg of the plasmid pBJ-PEC1 and 1 μg pAN7, which confers hygromycin B resistance (Punt et al. ), were co-transformed into the fungal genome.
Fungal genomic DNA was isolated by phenol-chloroform extraction, using a FastPrep®-24 (MP Biomedicals, Santa Ana, CA, USA) for cell disruption. Therefore about 100 mg of mycelia was transfered to 400 μl DNA extraction buffer (0.1 M Tris–HCl pH 8.0, 1.2 M NaCl, 5 mM EDTA) and grounded with glass beads (0.37 g Ø 0.01 – 0.1 mm, 0.25 g Ø 1 mm, 1 piece Ø 3 mm) using the FastPrep. Afterwards, the mixture was immediately put on 65°C, supplemented with 9 μM RNase A, and incubated for 30 min. Then 200 μl of phenol (pH 7.9) and 200 μl of a chloroform-isoamyl alcohol-mixture (25:1) were added, and vigorous mixing followed each addition. Phases were separated by centrifugation (12000 g, 10 min, 4°C) and the aqueous phase was transferred into a new vial. DNA was precipitated by addition of the 0.7-fold volume of isopropanol. After 20 min incubation at room temperature the DNA was separated by centrifugation (20000 g, 20 min, 4°C) and washed with 500 μl ethanol (70%). The air-dried DNA pellet was solubilised in 50 μl Tris–HCl (10 mM, pH 7.5) at 60°C.
RNA isolation and cDNA synthesis
RNA extraction from fungal mycelia was performed with peqGOLD TriFast™ (peqlab, Erlangen, Germany) according to the manufacturer’s procedure, using a FastPrep®-24 (MP Biomedicals, Santa Ana, CA, USA) for cell disruption. RNA quantity and quality were determined with a NanoDrop 1000 (Thermo Scientific, Waltham, MA, USA). A 260 nm/280 nm ratio of at least 1.8 was stipulated for further sample processing.
cDNA synthesis was performed with RevertAid™ H Minus First Strand cDNA Synthesis Kit (Thermo Scientific, Waltham, MA, USA) according to the manufacturer’s procedure, using 0.5 μg of RNA.
where E is cycling efficiency, C is the threshold cycling number, r is the reference gene, t the target gene and o marks the sample which is taken for normalization Pfaffl ().
Glutathione S-transferase (GST): Err1 fusion proteins
GST fusion proteins of the erythrose reductases from T. reesei, A. niger, and F. graminearum were expressed using plasmids pGEX-err1T, pGEX-err1A, and pGEX-err1F, respectively, in E. coli BL21(DE3)pLysS (Promega, Madison, WI, USA). The protein expression was done in shakeflasks on lysogeny broth supplemented with 100 μg/ml ampicillin at 37°C and 200 rpm. For induction 0.1 mM isopropyl-β-D-thiogalactopyranoside (IPTG) was added when the culture reached an OD600 between 0.7 and 0.8. Cells were harvested 3 h after induction by centrifugation, resuspended in phosphate buffered saline supplemented with 1% Triton X-100, and sonicated using a Sonifier® 250 Cell Disruptor (Branson, Danbury, CT, USA) (power 70%, duty cycle 40%, power for 10 s, pause for 50 s, 10 cycles, on ice). Insoluble compounds were separated by centrifugation (2600 g, 10 min, 4°C). Purification of the proteins was performed using GSTrap™ FF (GE Healthcare Life sciences, Little Chalfont, Buckinghamshire, UK) according to standard procedures. The purified protein solutions were stored at 4°C. There was no considerable loss of activity observed within one month under these storage conditions. The addition of glycerol must be avoided because it has an influence on the enzymatic assay described later.
For the SDS-PAGE analysis a 10% polyacrylamide gel with a tris-glycine buffer (25 mM Trizma® base (Sigma Aldrich, St. Louis, MO, USA), 1.9 mM glycine, 0.5% SDS) was used. Gel casting and running the gel was done with the Mini-PROTEAN® Tetra Cell system (Bio-Rad Laboratories, Hercules, CA, USA). From all three protein expressions 2 μl of the crude extract, 2 μl of the flow-through, and 12 μl of the wash solution, respectively, were applied on the gel. Of the eluated protein from A. niger 2 μl, from F. graminearium 12 μl, and from T. reesei 1 μl were applied. All the samples were supplemented with 4 μl 4x Laemmli sample buffer (Bio-Rad Laboratories, Hercules, CA, USA), filled up with distilled water to a final volume of 16 μl, and incubated for 10 min at 95°C for denaturation. After denaturation, samples were kept on ice until application on the gel. For protein size estimation 2.5 μl of PageRuler™ Prestained ProteinLadder (Thermo Scientific, Waltham, MA, USA) were used. The electrophoresis was carried out at a constant voltage of 160 V. Staining of the gels was done with PageBlue Protein Staining Solution (Thermo Scientific, Waltham, MA, USA) according to the manufacturers protocol.
Enzymatic analysis was performed according to a slightly modified, previously by Lee et al. () described protocol. The reducing reaction was performed in a total volume of 1 ml containing 50 mM Sorenson’s phosphate buffer (pH 6.5), 160 μM NADPH or NADH, 100 μl purified GST::Err1 fusion protein, and 10 mM substrate. As substrates L-arabinose, dihydroxyacetone (DHA), D-erythrose, D-glucose, L-glyceraldehyde, glyoxal, methylglyoxal, D-threose, D-xylose, and D-xylulose were used. In a spectrophotometer the consumption of NADPH or NADH over time was followed at 340 nm at the indicated temperature. After 1 min incubation without substrate the reaction was started by adding 100 μl 100 mM substrate.
The oxidizing reaction was performed in a total volume of 1 ml containing 50 mM Tris/HCl (pH 9.0), 400 μM NADP+, 200 μl purified GST::Err1 fusion protein, and 10 mM erythritol. In a spectrophotometer the formation of NADPH over time was followed at 340 nm at a temperature of 40°C. After 1 min incubation without substrate the reaction was started by adding 100 μl 100 mM erythritol.
Enzymatic assays were performed in triplicates. Activity is defined in katal (kat), and 1 katal is the conversion of 1 mol substrate per second. The specific activity kcat is defined as 1 katal per mol enzyme and the catalytic efficacy is defined as kcat/Km.
Gas chromatography (GC) analysis
Mycelia were ground under liquid nitrogen. The powder was suspended in 3 ml distilled water and sonicated using a Sonifier® 250 Cell Disruptor (Branson, Danbury, CT, USA) (power 70%, duty cycle 40%, power for 3 min, on ice). Insoluble compounds were separated by centrifugation (20000 g, 10 min, 4°C). Sample preparation for GC was done in triplicates as follows: 300 μl of the supernatant, supplemented with 10 ng sorbitol as internal standard, was gently mixed with 1.2 ml ethanol (96%) and incubated for 30 min at room temperature for protein precipitation. The precipitant was separated by centrifugation (20000 g, 10 min, 4°C). Samples were dried under vacuum and thereafter silylated (50 μl pyridine, 250 μl hexamethyldisilazane, 120 μl trimethylsilyl chloride). For quantitative erythritol determination a GC equipment (Agilent Technologies, Santa Clara, CA, USA) with a HP-5-column (30 m, inner diameter 0.32 mm, film 0.26 μm) (Agilent Technologies, Santa Clara, CA, USA) was used. The mobile phase consisted of helium with a flow of 1.4 l/min, the column temperature was as follows: 150°C for 1 min, ramping 150 – 220°C (ΔT 4°C/min), ramping 220–320°C (ΔT 20°C/min), 320°C for 6.5 min. Detection was performed with FID at 300°C. The retention times were determined using pure standard substances.
Identification of putative erythrose reductase proteins by in silico analysis
Purification of heterologously expressed Err1 proteins
Optimal parameters for the erythrose reductase enzyme assay
Since neither of the proteins has yet been characterized using D-erythrose as a substrate, the optimal parameters for the enzymatic assay had to be determined. Enzyme assays were performed using the proteins heterologously expressed in E. coli.
For the reducing reaction, which converts D-erythrose to erythritol, a previous study reported a pH of 7.0 for the ER from C. magnoliae Lee et al. (). For the GLD1 from T. reesei Liepins et al. () also reported a pH optimum of 7.0, but this was determined with different substrates. Therefore, Sorenson’s phosphate buffers from pH 6.0 to 8.0 were tested in steps of 0.5 pH units. Online resource 2 depicts the measured progression of the absorption caused by NADPH consumption. We found that a pH of 6.5 is clearly favorable for the Err1 from T. reesei (Additional file 2a). The enzymes from A. niger and F. graminearum showed strongest decrease in absorbance at pH 7.0, but the differences between varying pH conditions were neglible for both (Additional file 2b and Additional file 2c). Therefore, the temperature optimization was carried out at pH 6.5 for all three enzymes from 10°C to 50°C (in steps of 10°C). For Err1 from T. reesei we found an increase in activity between 10°C and 40°C, whereas 40°C and 50°C already yielded almost identical activities (Additional file 2d). The enzyme from A. niger showed only slightly better performance at 50°C compared to 40°C (Additional file 2e). For the Err1 from F. graminearum enzyme denaturation occurred most probably at 50°C, which can be deduced from the early loss of activity at a still high NADPH concentration (Additional file 2f). Since the improvement in Err1 activity using 50°C instead of 40°C was negligibly anyway and with respect to better enzyme stability, 40°C was chosen for further measurements.
Testing the three enzymes under optimized conditions with NADH instead of NADPH as co-factor for neither of them yielded a detectable activity. This is in accordance with former reports on the T. reseei enzyme, which showed activity only under consumption of NADPH, but not with NADH Liepins et al. ().
For the oxidizing reaction, which converts erythritol to D-erythrose under consumption of NADP+, former studies proposed a pH of about 9 for similar reactions Colowick (). Consequently, Tris/HCl buffers of pH 8.0, 8.5, and 9.0 (equals the upper range of this buffer system) were tested at an assay temperature of 40°C. Only at pH 9 the oxidation of erythritol was the favored direction of the reaction, however, it proceeded much slower than the inverse reaction described before. At pH 8.5 an oscillating reaction was observed, whereas at pH 8.0 the equilibrium was completely on the reducing side of the reaction (data not shown).
Altogether, we suggest the usage of a buffers system at pH 6.5 and a temperature of 40°C for the erythrose reductase assay.
Substrate specificity and activity of Err1
Substrates were chosen in order to cover molecules from 2 to 6 carbon atoms (C2 – C6) on the one hand, and aldehydes and ketones on the other hand: the dialdehyde glyoxal (C2), the keto-aldehyde methylglyoxal (C3), the trioses DHA and L-glyceraldehyde, the aldotetroses D-erythrose and D-threose, the aldopentoses L-arabinose and D-xylose, the ketopentose D-xylulose, and the aldohexose D-glucose.
The three enzymes showed some differences in both, substrate specificity as well as in total activity. But for all of them the activity using DHA, D-glucose, D-xylose, and D-xylulose was too low to evaluate the kinetics parameters. Consequently, these substances will be neglected in the further discussion.
Substrate specificity of Err1 from T. reesei
k cat/K m[1/(mM·s)]
124.56 ± 9.78b
3.21 ± 0.22
25.80 ± 0.23
134.52 ± 9.34
36.51 ± 2.13
271.41 ± 3.00
158.04 ± 5.00
47.89 ± 1.86
303.02 ± 2.18
102.74 ± 9.76
18.84 ± 1.34
183.41 ± 4.40
131.86 ± 1.84
72.58 ± 0.28
550.41 ± 5.55
94.07 ± 2.46
29.03 ± 0.89
308.59 ± 1.36
Substrate specificity of Err1 from A. niger
k cat/K m[1/(mM·s)]
286.66 ± 27.06b
7.32 ± 0.51
25.55 ± 0.64
139.39 ± 6.45
24.95 ± 1.05
179.00 ± 0.76
319.28 ± 4.12
448.61 ± 2.29
330.95 ± 3.06
49.09 ± 0.68
148.34 ± 0.68
352.81 ± 24.42
196.04 ± 13.43
555.66 ± 0.39
279.50 ± 7.89
108.44 ± 1.98
387.97 ± 3.87
Substrate specificity of Err1 from F. graminearum
k cat/K m[1/(mM·s)]
227.61 ± 8.81c
3.72 ± 0.09
16.36 ± 0.23
298.72 ± 88.4
8.54 ± 2.28
28.57 ± 0.83
535.16 ± 6.42
2.90 ± 0.07
5.42 ± 0.06
214.32 ± 7.64
6.76 ± 0.24
31.55 ± 0.02
380.48 ± 18.65
2.91 ± 0.00d
7.64 ± 0.37
Overexpression of err1 in T. reesei proves its function in vivo
GC analysis of the intracellular erythritol concentration of both strains demonstrated that the err1 overexpression strain indeed was able to produce more erythritol than its parental strain. After 24 h, the erythritol concentration in the recombinant strain was 1.6-fold higher than in the parental strain, and after 30 h it was even 3.2-fold, respectively (Figure 3b).
Based on the protein sequences of the known erythrose reductases from Trichosporonoides megachiliensis SNG-42 (Ookura et al. ), we identified by in silico analysis candidate proteins for Err1 in T. reesei, A. niger, and F. graminearum. In vitro analysis of these proteins by an enzyme assay confirmed for all of them a high substrate specificity and turnover rate for D-erythrose. Out of ten tested aldehydes and ketones, ranging from C2 to C6, only methylglyoxal and L-glyceraldehyde partly showed better performance or substrate specificity than D-erythrose and its diastereomer D-threose. For the cell toxin methylglyoxal it is known that aldehyde reductases show considerable activity for it, and convert it to hydroxyacetone (95%) and D-lactaldehyde (5%) (Thornalley ). But the main detoxification of methylglyoxal is done by the glyoxalase system, consisting of glyoxalase I and II and catalytic amounts of reduced glutathione. These enzymes belong to superfamily cl14632, whereas Err1 belongs to superfamily cl00470 and utilizes NADPH as cofactor. Therefore, it is very unlikely that Err1 belongs to the glyoxalase system. Interestingly, the good performance of erythrose reductase with glyceraldehydes, which was observed in this study, was also reported by (Lee et al. ) for C. magnoliae.
Neither of the tested Err1 proteins from the three fungi has a clear specificity for D-erythrose over D-threose or vice versa. In case of the Err1 from T. reesei D-erythrose showed a higher turnover number than D-threose, but the differences in Km were not substantially. The Err1 from A. niger on the one hand clearly preferred D-erythrose considering Km, but on the other hand, the turnover number was considerably higher for D-threose. Only the enzyme from F. graminearum has a slight preference for D-erythrose, which is reflected by both characteristic numbers, Km and kcat. Since Err1 takes various short-chained aldehydes as substrate it is not surprising that it utilizes the diastereomers D-erythrose and D-threose in a similar manner.
Aside from D-erythrose (C4), D-threose (C4), L-glyceraldehyde (C3) and methylglyoxal (C3) also glyoxal (C2) caused distinct activity. The enzymes from T. reesei and A. niger also showed measurable activity with the C5-sugar L-arabinose, but it was much lower than the activity of the substrates mentioned before. With D-xylose, the other C5-aldehyde tested, only a poor activity of these two enzymes was detected, which turned out to be too low to calculate kinetic parameters. The C6-sugar D-glucose showed no activity at all. It can therefore be proposed that Err1 is limited to substrates with a chain length ≤ 5 C-atoms, with best performance for 3 and 4 C-atoms. The two ketones analyzed, DHA and D-xylulose, showed no measurable activity. This leads to the assumption that only aldehydes are suitable substrates, which is in accordance with the previous general assignment of the A. niger enzyme as aldehyde reductase.
The Err1 from T. reesei and A. niger performed quite similar (activity is in the same order of magnitude), whereas the enzyme from F. graminearum showed much lower activity (about one tenth) for all substrates. Also, the latter was found to be less temperature-stable than the other ones, as the loss of activity was visible within minutes if kept at 50°C.
Comparing the kinetic parameters using D-erythrose as substrate and NADPH as co-factor, a ten times higher Km was observed for the Err1 proteins from T. reesei and A. niger characterized in this study than for ER1 and ER2 from C. magnoliae (Lee et al. ). The kcat of Err1 from T. reesei and A. niger is in the same order of magnitude as ER2, resulting in a 10-fold higher catalytic efficacy of ER2. The strict requirement of NADPH as cofactor is in accordance with results for C. magnoliae (Lee et al. ). However, the presence of erythrose reductase activity in these filamentous fungi is an important prerequisite for the possibility of developing production strategies using non-food plant biomass. Notably, the enhanced err1 expression in a recombinant T. reesei strain led to an increased formation of erythritol. Even if the yield is not at the level of the yeast production strains, it should be considered that these strains have already undergone extensive mutagenesis and were screened for erythritol production. Any kind of engineering steps are still open in order to increase erythritol production in filamentous fungi. As this is an attractive alternative that would use cheap and sustainable starting materials an according patent was issued (Mach-Aigner et al. ).
Finally, the recombinant T. reesei strain, which overexpressed err1, and its parental strain demonstrated functionality of the erythrose reductase in vivo. This emphasizes that the earlier characterizations of the enzyme from T. reesei as Gld1 (Liepins et al. ) and the one from A. niger as Alr1 missed an important biological function of the enzyme. In summary, all three levels of investigation (in silico, in vitro, and in vivo) have provided evidence that the proteins identified are catalyzing the side reaction of the PPP, in which D-erythrose is converted to erythritol and vice versa. Altogether, this supports their capability to function as erythrose reductases.
This study was supported by Annikki GmbH, by two grants from the Austrian Science Fund (FWF): [P20192, P24851] given to RLM and ARMA, respectively, and by a doctoral program of Vienna University of Technology (AB-Tec).
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