Utilizing Alcohol for Alkane Biosynthesis by Introducing a Fatty Alcohol Dehydrogenase
ABSTRACT
Alkanes produced by microorganisms are expected to be an alternative to fossil fuels as an energy source. Microbial synthesis of alkanes involves the formation of fatty aldehydes via fatty acyl coenzyme A (acyl-CoA) intermediates derived from fatty acid metabolism, followed by aldehyde decarbonylation to generate alkanes. Advancements in metabolic engineering have enabled the construction of such pathways in various microorganisms, including Escherichia coli. However, endogenous aldehyde reductases in the host microorganisms are highly active in converting fatty aldehydes to fatty alcohols, limiting the substrate pool for alkane production. To reuse the alcohol by-product, a screening of fatty alcohol-assimilating microorganisms was conducted, and a bacterial strain, Pantoea sp. strain 7-4, was found to convert 1-tetradecanol to tetradecanal. From this strain, an alcohol dehydrogenase, PsADH, was purified and found to be involved in 1-tetradecanol-oxidizing reaction. Subsequent heterologous expression of the PsADH gene in E. coli was conducted, and recombinant PsADH was purified for a series of biochemical characterizations, including cofactors, optimal reaction conditions, and kinetic parameters. Furthermore, direct alkane production from alcohol was achieved in E. coli by coexpressing PsADH with a cyanobacterial aldehyde-deformylating oxygenase and a reducing system, including ferredoxin and ferredoxin reductase, from Nostoc punctiforme PCC73102. The alcohol-aldehyde-alkane synthetic route established in this study will provide a new approach to utilizing fatty alcohols for the production of alkane biofuel.
IMPORTANCE Alcohol dehydrogenases are a group of enzymes found in many organisms. Unfortunately, studies on these enzymes mainly focus on their activities toward short-chain alcohols. In this study, we discovered an alcohol dehydrogenase, PsADH, from the bacterium Pantoea sp. 7-4, which can oxidize 1-tetradecanol to tetradecanal. The medium-chain aldehyde products generated by this enzyme can serve as the substrate of aldehyde-deformylating oxygenase to produce alkanes. The enzyme found in this study can be applied to the biosynthetic pathway involving the formation of medium-chain aldehydes to produce alkanes and other valuable compounds.
INTRODUCTION
Biofuels produced from renewable biomass are promising alternatives to fossil fuels (1, 2). Recently, biobased hydrocarbons, such as alkanes and alkenes, have attracted particularly considerable attention since they have a chemical composition similar to that of petroleum-based fuels and show better performance than other biofuels due to their higher energy content (3). Alkanes are produced by many organisms in nature, including plants, insects, and microorganisms, where in most cases, fatty aldehydes derived from fatty acid metabolism are converted to alkanes through a decarbonylation reaction mechanism (4–9). To date, there have been numerous reports on the production of medium-chain (C13 to C17) alkanes from glucose using Escherichia coli, Saccharomyces cerevisiae, and other metabolically engineered microorganisms (10–14). For example, by introducing cyanobacterial acyl-acyl carrier protein (acyl-ACP) reductase and aldehyde-deformylating oxygenase (ADO), the alkane titer in E. coli reached 300 mg L−1 (15).
A bottleneck in alkane production using E. coli as the host is caused by the endogenous aldehyde reductase, which converts intracellular aldehyde to alcohol (16, 17). The substrate pool for ADO was competed by aldehyde reductase, resulting in poor alkane production. It has been reported that the alkane titer was increased nearly 2-fold by removing yqhD, a gene encoding aldehyde reductase found in E. coli (18). Also, it has been reported that up to 13 genes encoding aldehyde reductase are found in E. coli, and the deletion of these genes results in a 90% reduction in endogenous alcohol accumulation (19). However, it takes tremendous effort to conduct gene deletion, and the resulting aldehyde accumulation is toxic to the cell (19, 20). To avoid these concerns and to efficiently drive metabolic flow to alkane production, a better approach to utilize alcohol by-products in E. coli remains necessary.
We aimed to construct a pathway linking alcohol oxidation with alkane biosynthesis. Coupling an alcohol dehydrogenase capable of converting 1-tetradecanol to the corresponding medium-chain fatty aldehyde with a cyanobacterial ADO (15, 21) could improve alkane production by utilizing the alcohol as the substrate again. Here, we report the discovery of an enzyme, PsADH, originated from a soil isolate of Pantoea sp. suitable for this purpose. Biochemical characterization of PsADH revealed its potential as a tool to facilitate alkane production. Regarding the lack of research about utilizing fatty alcohol for alkane biosynthesis (22), this study provides a new point of view for the development of biofuel.
RESULTS
Screening and identification of 1-tetradecanol-oxidizing microorganisms.
A total of 350 microbial strains were isolated from the soil as 1-tetradecanol-assimilating microorganisms. Among them, 60 strains showed the ability to oxidize 1-tetradecanol, including 19 strains that produced tetradecanal, 27 strains that produced tetradecanoic acid, and 14 strains that produced both tetradecanal and tetradecanoic acid. Since we aimed to find an enzyme that can generate tetradecanal for alkane production, we chose strain 7-4, with the highest tetradecanal production among all strains. The tetradecanal produced by strain 7-4 was confirmed using gas chromatography-mass spectrometry (GC-MS; see Fig. S1 in the supplemental material), and the 16S rRNA sequence of strain 7-4 was found to exhibit 99% homology with that of Pantoea. To further evaluate Pantoea sp. strain 7-4, we conducted a reaction using its cell extracts and found that 1-tetradecanol is converted to tetradecanal only when NAD(P)+ was present (Fig. 1). Also, tetradecanal production remained at the same level when the reaction was performed under anaerobic conditions. These results indicated that Pantoea sp. 7-4 might possess a dehydrogenase, requiring NAD(P)+ as an electron acceptor, rather than an oxidase, which requires molecular oxygen as an electron acceptor.
FIG 1
Purification and identification of an alcohol dehydrogenase from Pantoea.
To identify the enzyme involved in alcohol oxidation, we conducted enzyme purification from the cell lysate of Pantoea sp. 7-4 via a fast protein liquid chromatography (FPLC) system and successfully purified an enzyme shown as a single protein band in Fig. S2. The detailed activity and yield during the purification process are shown in Table 1. The NH2-terminal amino acid sequence of the purified protein was identified as MNMKIKTTMKAAVVKSFGEP.
TABLE 1
| Step | Total protein (mg) | Total activity (U) | Sp act (U/mg) | Yield (%) | Fold |
|---|---|---|---|---|---|
| Cell-free extracts | 511 | 162 | 0.32 | 100 | 1.00 |
| Ammonium sulfate | 347 | 145 | 0.42 | 89.0 | 1.31 |
| HiTrap Phenyl HP | 25.2 | 18.4 | 0.73 | 11.3 | 2.30 |
| Superdex 200 | 3.89 | 8.68 | 2.23 | 5.30 | 7.02 |
| Q Sepharose XL | 0.0556 | 0.183 | 3.29 | 0.11 | 10.4 |
The above amino acid sequence was subjected to a BLAST search, and the result showed that it was 100% homologous with the sequence of an alcohol dehydrogenase from Pantoea sp. strain MSR2 (23). Based on these results, we named the purified enzyme PsADH.
Cloning, expression, and purification of alcohol dehydrogenase PsADH.
To elucidate the catalytic properties of PsADH, we cloned the PsADH gene from the isolated Pantoea sp. strain 7-4 into the pET-21b(+) vector and transformed it into E. coli Rosetta 2 (DE3). The cell lysate of isopropyl-β-d-thiogalactopyranoside (IPTG)-induced E. coli was then subjected to a column chromatography series using an FPLC system, and the recombinant PsADH was purified (Fig. S3). The purified enzyme was used for further characterization.
Characterization of recombinant PsADH.
First, the effect of the cofactors was evaluated in the alcohol-oxidizing reaction using purified recombinant PsADH (Fig. 2a). We found that PsADH catalyzed the reaction when beta-NAD, oxidized (NAD+), was added but not when beta-NADP, oxidized (NADP+), was added. Based on this result, we assumed that PsADH is a NAD+-dependent alcohol dehydrogenase. Next, the optimal reaction pH of PsADH was evaluated for both oxidation and reduction reactions (Fig. 2b). PsADH showed its highest activity at pH 9.0 with Tris-HCl buffer when catalyzing an oxidation reaction, while its highest activity occurred at pH 7.0 with potassium phosphate buffer (KPB) when catalyzing a reduction reaction. Furthermore, the optimal reaction temperature and the thermal stability of PsADH were evaluated for the oxidation reaction. PsADH showed the highest activity when it catalyzed the reaction at 40°C (Fig. 2c). For thermal stability, we found that PsADH retained 90% of its original activity when incubated at temperatures between 20°C and 50°C, while its activity dropped to <20% when incubated at 60°C (Fig. 2d). Finally, a kinetic analysis of PsADH was conducted using either 1-tetradenacol with NAD+ as the substrate (oxidation reaction) or tetradecanal with NADH as the substrate (reduction reaction) (Table 2). The PsADH follows the Michaelis-Menten-type reaction mechanism, where its kcat/Km values for oxidation and reduction reactions were 171 s−1·mM−1 and 586 s−1·mM−1, respectively.
FIG 2
TABLE 2
| Substrate | Km (mM) | kcat (s−1) | kcat/Km (s−1·mM−1) |
|---|---|---|---|
| 1-Tetradecanol | 0.168 | 28.7 | 171 |
| Tetradecanal | 0.129 | 75.6 | 586 |
Coexpression of PsADH and cyanobacterial ADO results in alkane production from alcohol.
To evaluate the potential use of PsADH in alkane production, we conducted PsADH coexpression with an ADO using E. coli as the host. We chose NpAD, the ADO from the cyanobacterium Nostoc punctiforme PCC73102, as well as ferredoxin (Fd) and ferredoxin-NADP+ reductase (FNR) from the same species, a system reported to convert aldehydes to alkanes (15). The expression of enzymes in the transformed E. coli strains was confirmed using SDS-PAGE (Fig. S4), and the alkane production was analyzed using GC-MS (Fig. S5). The alkane productivity assay results revealed that the E. coli strain harboring NpAD-Fd-FNR gene clusters produced no tridecane from 1-tetradecanol, whereas the strain harboring PsADH-NpAD-Fd-FNR gene clusters produced 0.15 mM tetradecanal and 0.12 mM tridecane from 2 mM 1-tetradecanol (Fig. 3).
FIG 3
DISCUSSION
In this study, we aimed to develop a novel pathway to utilize alcohol by-products generated in E. coli cells during alkane biosynthesis. The concept is to couple an oxidase with ADO where fatty alcohols can be reoxidized to fatty aldehydes for generating alkanes. Various enzymes found in nature perform alcohol oxidation; however, limited information has been reported regarding their activity toward medium-chain fatty alcohols (24–28). Therefore, we conducted a screening of 1-tetradecanol-assimilating microorganisms. 1-Tetradecanol has a carbon length of 14. Its corresponding product, tetradecanal, is a substrate favored by cyanobacterial ADOs to generate alkanes (15). As a result, we discovered that Pantoea sp. strain 7-4 is a good candidate since it produced the largest amount of tetradecanal among all strains. During the screening, we also found other positive strains that can utilize 1-tetradecanol. For example, Pseudomonas sp. strain 1-2 showed the ability to convert 1-tetradecanol to a relatively large amount of both tetradecanal and tetradecanoic acid (data not shown). However, since fatty acid is the undesired substrate for ADO in the downstream alkane biosynthetic process, we chose Pantoea sp. 7-4 for further analysis.
Observed throughout nature, enzymes that perform alcohol oxidation can be divided into two groups, the alcohol dehydrogenase group (EC 1.1.1) and the alcohol oxidase group (EC 1.1.3). An alcohol dehydrogenase normally requires NAD(P)+ or FAD as an electron acceptor, whereas an alcohol oxidase requires molecular oxygen as an electron acceptor (23, 29). Our initial results indicated that Pantoea sp. 7-4 might possess a dehydrogenase (Fig. 1). Subsequent sequence analysis of the enzyme PsADH purified from this strain also revealed that it belongs to the medium-chain dehydrogenase/reductase (MDR) superfamily. The enzymes in this superfamily were reported to have a size of around 350 residues (38.5 kDa) and function as dimers or tetramers (30, 31). Interestingly, when we tried estimating the molecular weight of PsADH, the results of gel filtration chromatography indicated its molecular weight to be 296 kDa (see Fig. S6 in the supplemental material), while the results of SDS-PAGE as well as its sequence information indicated 36.8 kDa (Fig. S3 and Table S1). Considering the denaturation process during SDS-PAGE, we assumed that PsADH functions as an octamer, which is an unusual character in the MDR superfamily.
To evaluate the synergy between PsADH and ADO, a series of biochemical parameters of PsADH were characterized using its recombinant protein. Kinetic analysis of PsADH (Table 2) showed a higher kcat/Km toward the reduction reaction (aldehyde to alcohol) than the oxidation reaction (alcohol to aldehyde), a result also found for many alcohol dehydrogenases where short-chain alcohols were used as substrates (24–28). Thus, it will be important to maintain the concentration of substrate and cofactor in the cell when applying this enzyme to alkane production. Conversely, PsADH was found to be an NAD+-dependent alcohol dehydrogenase, based on our results (Fig. 2a), making PsADH a suitable enzyme to be coupled with ADOs. It has been reported that the reaction catalyzed by cyanobacterial ADO requires oxygen as the cofactor and a reducing system consisting of ferredoxin, ferredoxin reductase, and NADPH, which provides four electrons per turnover (15, 32–36). PsADH needs neither oxygen nor NADP+ to catalyze alcohol oxidation. In the reduction of aldehyde, PsADH uses NADH but not NADPH. Based on these cofactor preferences, there will be no competition among these cofactors when PsADH is combined with ADO for alkane production.
Furthermore, we demonstrated the feasibility of our strategy by constructing an E. coli transformant expressing PsADH-NpAD-Fd-FNR, resulting in direct alkane production from alcohol. The conversion rate was approximately 6%, with 0.12 mM tridecane being produced from 2 mM 1-tetradecanol (Fig. 3). We performed the alkane productivity assay at 40°C under pH 9.0, which is favored by PsADH. However, such conditions may be unsuitable for NpAD. Since PsADH can catalyze alcohol-oxidizing reactions under a wide range of pHs and temperatures (Fig. 2b to d), we believe that the efficiency of this system can be improved by adjusting these parameters. Another way to improve alkane production may be adding a system to transport extracellular 1-tetradecanol into the cytoplasm. However, when we conducted the alkane productivity assay using the cell extracts of E. coli expressing PsADH-NpAD-Fd-FNR, we found that the alkane titer did not increase comparing to the data using the resting cells (data not shown). Hence, we assumed that the transportation of the substrate would not affect the yield.
It has been reported that the hydrocarbons produced by cyanobacterial ADOs are mainly C13 to C17 alkanes or alkenes (15), while our data also demonstrated the production of C13 alkane by coexpressing PsADH and cyanobacterial ADO. Nevertheless, further research on the substrate specificity of PsADH is required to evaluate its potential for alkane production. Similar to other alcohol dehydrogenases found in nature (24–28), PsADH may also show activity toward short-chain alcohols. An ADH with a broad substrate spectrum would be favored since the petroleum-based fuels currently used are composed of alkanes and alkenes ranging from short to medium (C8 to C16) chain. Unfortunately, there has been limited information on ADOs with substrate specificity toward short-chain aldehydes, and those well-studied cyanobacterial ADOs have shown a low turnover rate when catalyzing decarbonylation reactions (32, 36). To improve the alkane titer to a competitive level with other biofuels, such as bioethanol (37), an enzyme with better alkane-producing activity is required.
Conclusions.
This work reveals a new approach to reuse alcohol by-products for alkane production. We discovered an alcohol dehydrogenase PsADH from a Pantoea sp. strain isolated from the soil and characterized its optimal reaction conditions and kinetic parameters. By coexpressing PsADH with a cyanobacterial ADO in E. coli, the direct production of alkane from alcohol was reported for the first time. Our data suggest that PsADH is NAD+ dependent and does not require oxygen to catalyze the alcohol-oxidizing reaction. These characteristics provide PsADH with a great potential to be coupled with ADOs and other enzymes when constructing alkane biosynthetic pathways. By expanding the possible routes for metabolic engineering, this study will contribute to the future development of biobased sustainable fuels.
MATERIALS AND METHODS
Chemicals, microbial strains, primers, and plasmids.
1-Tetradecanol and tetradecanal were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan). The NAD+ disodium salt and NADP+ tetrasodium salt were purchased from Oriental Yeast Co., Ltd. (Tokyo, Japan). Yeast extract was purchased from BD Biosciences (Franklin Lakes, NJ, USA). Other reagents were purchased from Fujifilm Wako Pure Chemical Corporation (Osaka, Japan). E. coli DH5α and E. coli Rosetta 2 (DE3) (Merck, Darmstadt, Germany) were used as the hosts for gene cloning and expression, respectively. The primers used in this study are shown in Table 3. Plasmids pET-21b (+), pRSFDuet-1, and pCDFDuet-1 (Merck) were used as the expression vectors. Other chemicals used in this study were of analytical grade and commercially available.
TABLE 3
| Primer | Sequence |
|---|---|
| 1492R | 5′-GNTACCTTGTTACGACTT-3′ |
| 1100R | 5′-AGGGTTGCGCTCGTTG-3′ |
| 802R | 5′-TACCAGGGTATCTAATCC-3′ |
| 341F | 5′-CCTACGGGNGGCWGCAG-3′ |
| T7-F | 5′-TAATACGACTCACTATAGGG-3′ |
| T7-R | 5′-TAGTTATTGCTCAGCGGTGG-3′ |
| Adh-AS SalF | 5′-GGGGTCGACATGAACATGAAGATCAAAACCACGATGAAAGCG-3′ |
| Adh-AS NotR | 5′-AAAGCGGCCGCTTACAGCGACATATCAACAACAATACGGCCTTC-3′ |
| Adhseq1F | 5′-ATGAAGGTGTTGGTCAGGTT-3′ |
| Adhseq1R | 5′-TTGCTCAAAGGCTTTCGGTG-3′ |
| NpAD NdeF | 5′-GGGCATATGATGCAGCAGCTTACAGACCAATCTAAA-3′ |
| NpAD XhoIR | 5′-GGGCTCGAGTTAAGCACCTATGAGTCCGTAGGCTGA-3′ |
Isolation of 1-tetradecanol-assimilating microorganisms.
Soil samples were collected from places all over Japan and used as the source for screening and isolation of 1-tetradecanol-assimilating microorganisms. First, soil samples were suspended in saline water and incubated at room temperature for 1 h. Approximately 20 to 30 μL of the supernatant of the solution was then inoculated into 3 mL of enrichment medium that consisted of 0.5% (wt/vol) (NH4)2SO4, 0.1% (wt/vol) NH4Cl, 0.1% (wt/vol) KH2PO4, 0.3% (wt/vol) K2HPO4, 0.05% (wt/vol) MgSO4, 0.001% (wt/vol) FeSO4, 0.1% (wt/vol) yeast extract, 1% (wt/vol) 1-tetradecanol, and 0.1% (wt/vol) Triton X-100. The culture medium was incubated aerobically with shaking (300 strokes/min) at 28°C for 24 to 48 h. Next, an aliquot (20 to 50 μL) of the medium, in which microorganism growth was observed, was transferred to 3 mL of fresh enrichment medium and incubated with shaking (300 strokes/min) at 28°C for another 24 to 48 h. Finally, an aliquot (10 μL) of the medium containing microorganisms was streaked onto a 2% (wt/vol) agar plate with the same composition as the isolation medium and cultivated at 28°C for 24 to 48 h. Following cultivation, single colonies on the agar plate were isolated and used for subsequent experiments.
Screening of microorganisms isolated from soil samples.
The 1-tetradecanol-assimilating microorganisms were inoculated into 10 mL of isolation medium and cultivated at 28°C for 48 h. After cultivation, the cells were collected by centrifugation at room temperature and 1,500 × g for 10 min and added to a 1-mL reaction mixture consisting of 10 mM 1-tetradecanol, 5 mM NAD+, 5 mM NADP+, and 1% (wt/vol) Triton X-100 in 100 mM Tris-HCl buffer (pH 8.5). The reaction mixture was incubated with shaking (300 strokes/min) at 28°C for 24 h. After the reaction, the mixture was acidified with 5 N HCl, extracted with 1 mL of toluene containing 1 mM 1-pentadecanol as an internal standard, and subjected to GC and GC-MS analyses.
GC and GC-MS analyses.
GC analysis was conducted using a Shimadzu GC-2010 Plus gas chromatograph (Shimadzu, Kyoto, Japan) equipped with a flame ionization detector and a Nukol capillary column (30 m by 0.25 mm by 0.25 μm; Supelco, Bellefonte, PA, USA). Helium at a flow rate of 1.12 mL/min was used as the carrier gas. Samples were injected with a split ratio of 50:1. The injector and detector were operated at 200°C. The heating program of the column consisted of the following steps: the initial column temperature of 80°C was raised to 112°C at a rate of 5°C/min, then raised to 190°C at a rate of 40°C/min, and held at 190°C for 45 min. GC-MS was conducted using a Shimadzu QP2010 GC-MS (Shimadzu, Kyoto, Japan) equipped with a Nukol capillary column (30 m by 0.25 mm by 0.25 μm; Supelco) to identify the structure of products produced by microbial strains. The flow rate and the heating program were the same as those described above. A mass spectrometer was operated in the electron impact mode at 70 eV, with an ion source temperature of 200°C. Quantification was conducted by comparing peak areas with those from standard compounds.
Identification of selected strains via 16S rRNA sequencing.
Genomic DNA of a selected bacterial strain was extracted with a DNeasy blood and tissue kit (Qiagen, Hilden, Germany) and used as the template for PCR. The gene encoding the 16S rRNA sequence was amplified using PCR with the primers 1492R, 1100R, 802R, and 341F, listed in Table 3. The PCR was performed using KOD-Plus-Neo DNA polymerase (Toyobo, Osaka, Japan) under the following reaction conditions: 94°C for 2 min and 30 cycles of 98°C for 10 s, 55°C for 30 s, and 68°C for an appropriate time (30 s/kbp). The PCR products were purified using the QIAquick gel extraction kit (Qiagen) and subjected to genetic analysis using the GenomeLab DTCS Quick Start kit and a Beckman Coulter CEQ8000 analyzer (Beckman Coulter, Brea, CA, USA).
Purification of the alcohol-oxidizing enzyme from Pantoea sp. 7-4.
All protein purification processes in this study were conducted at 0°C to 5°C using the AKTA pure chromatography system (GE Healthcare, Chicago, IL, USA). The activity of all protein fractions was measured using the spectrophotometric method mentioned in the next section, and all active fractions were subjected to SDS-PAGE analysis. First, cell pellets of Pantoea sp. 7-4 were suspended in 20 mM KPB (pH 7.0) and disrupted with an Insonator 201M ultrasonic oscillator (KUBOTA, Tokyo, Japan) for 5 min (6 cycles) followed by centrifugation at 12,000 × g for 20 min and ultracentrifugation at 100,000 × g for 60 min. The supernatant was recovered as a soluble protein fraction and was mixed with an equal volume of 20 mM KPB (pH 7.0) with 3 M ammonium sulfate. After mixing and centrifugation at 6,500 × g for 20 min, the supernatant was recovered and subjected to a chromatography series. Protein was purified using hydrophobic interaction chromatography in a HiTrap Phenyl HP 5-mL column (GE Healthcare) and eluted with 10 column volumes (CV) of 20 mM KPB (pH 7.0). The active protein fractions were desalted and further purified via gel filtration chromatography in a Superdex 200 Increase 100/300 GL column (GE Healthcare) and eluted with 1.3 CV of 20 mM KPB (pH 7.0), followed by anion-exchange chromatography in a Q Sepharose XL 1-mL column (GE Healthcare) with a NaCl linear gradient (0 to 1 M in 20 CV) in 20 mM KPB (pH 7.0).
Determination of enzyme activity.
Enzyme activity was determined by measuring the absorbance change at 340 nm, which represented the NAD(P)H generated from NAD(P)+ during the oxidation reaction catalyzed by alcohol dehydrogenase. The reaction mixture (1 mL) consisted of 100 mM Tris-HCl (pH 8.5), 1 mM 1-tetradecanol, 3 mM NAD+, 3 mM NADP+, 0.01% (wt/vol) Triton X-100, and 10 μL of the enzyme fraction. The reaction components were mixed in a cuvette and incubated at 25°C for 3 min before adding the substrate 1-tetradecanol. After adding the substrate, the absorbance at 340 nm of the reaction mixture was measured.
Protein analysis.
The protein concentration was determined using a Bradford protein assay with a Bio-Rad protein assay kit (Bio-Rad Laboratories, Inc., Hercules, CA, USA). SDS-PAGE was conducted on a 12.5% polyacrylamide gel with a Tris-glycine buffer system. Protein bands were stained using Coomassie brilliant blue R250. Protein bands were cut from the gel and transferred onto a polyvinylidene difluoride (PVDF) membrane for sequencing. The NH2-terminal amino acid sequence of the target protein was determined using automated Edman degradation with a PPSQ-31A protein sequencer (Shimadzu, Kyoto, Japan).
Cloning and expression of recombinant alcohol dehydrogenase PsADH.
The PsADH gene (DDBJ accession number LC727629 [Table S1]) was amplified via PCR using the genomic DNA of Pantoea sp. strain 7-4 as a template and the primer of Adh-AS SalF and Adh-AS NotR listed in Table 3. The PCR was performed using PrimeSTAR MAX DNA polymerase (TaKaRa Bio, Shiga, Japan) under the following reaction conditions: 94°C for 2 min and 35 cycles of 98°C for 10 s, 55°C for 5 s, and 72°C for 1.5 min. The PCR product was mixed with the pET-21b(+) vector, which was digested with NotI and SalI and ligated using ClonTech ligation mix (TaKaRa Bio), resulting in the plasmid pET-21b(+)-PsADH. The plasmid was then transformed into E. coli competent cells. E. coli DH5α was used as the cloning host and E. coli Rosetta 2 (DE3) was used as the expression host. The resulting strain, E. coli Rosetta 2 (DE3)/pET-21b(+)-PsADH, was inoculated into an LB medium containing ampicillin and chloramphenicol and cultivated at 37°C with shaking. When the optical density at 600 nm reached 0.8, culture was induced with 1 mM IPTG and cultivated at 16°C for 16 h with shaking. Protein expression was monitored by SDS-PAGE. After cultivation, cells were harvested via centrifugation, washed with 0.85% NaCl solution, and stored at −20°C for further use.
Purification of recombinant PsADH.
For the purification of recombinant PsADH, cells of E. coli Rosetta 2 (DE3)/pET21b(+)-PsADH induced with IPTG were disrupted using the method mentioned in the previous section. Recombinant PsADH was then purified via anion-exchange chromatography in a Q Sepharose XL (1-mL) column with a NaCl linear gradient (0 to 1 M in 20 CV) in 20 mM KPB (pH 7.0), followed by gel filtration chromatography in a Superdex 200 Increase 100/300 GL column, eluted with 1.3 CV of 20 mM KPB (pH 7.0). The native molecular weight of PsADH was determined via gel filtration chromatography using gel filtration calibration kits (low molecular weight [LMW] and high molecular weight [HMW]; GE Healthcare).
Effect of electron acceptor.
The effect of NAD+ or NADP+ on the purified PsADH was evaluated by measuring the absorbance change at 340 nm. The reaction mixture (1 mL), which consisted of 100 mM Tris-HCl buffer (pH 8.5), 0.5 mM 1-tetradecanol, 1 mM NAD+ or NADP+, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH, was incubated at 25°C, and the absorbance at 340 nm was measured. The amount of NAD(P)H generated was calculated using its molar extinction coefficient (6.22 mmol−1 1 cm−1).
Effect of reaction pH.
The pH-dependent enzyme assay was conducted for either an oxidation or reduction reaction catalyzed by purified PsADH. For the oxidation reaction, the reaction mixture (1 mL) consisted of 0.5 mM 1-tetradecanol, 1 mM NAD+, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH in different types of buffer. For the reduction reaction, the mixture (1 mL) consisted of 0.5 mM tetradecanal, 1 mM NADH, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH in different kinds of buffer. The buffers used were all at a concentration of 100 mM, with pHs as follows: sodium acetate buffer, pH 5.0 to 5.5; KPB, pH 6.0 to 8.0; Tris-HCl buffer, pH 7.5 to 9.0; and sodium carbonate buffer, pH 9.0 to 10.0. The reaction mixture was incubated at 25°C and the absorbance at 340 nm was measured.
Effect of reaction temperature.
The temperature-dependent enzyme assay was conducted for an oxidation reaction catalyzed by purified PsADH. The reaction mixture (1 mL), which consisted of 100 mM Tris-HCl buffer (pH 9.0), 0.5 mM 1-tetradecanol, 1 mM NAD+, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH, was incubated at different temperatures (20°C to 60°C) for 10 min for each trial, and the absorbance at 340 nm was measured.
Thermal stability.
The thermal stability of PsADH was evaluated for the oxidation reaction. Before mixing with other reaction components, the purified PsADH was incubated at different temperatures (20°C to 70°C) for 1 h in 100 mM Tris-HCl buffer (pH 9.0) for each trial. The reaction mixture (1 mL) consisted of 100 mM Tris-HCl buffer (pH 9.0), 0.5 mM 1-tetradecanol, 1 mM NAD+, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH. After mixing, the reaction mixture was incubated at 25°C for 5 min, and the absorbance at 340 nm was measured.
Kinetic analysis.
The kinetics parameters, including the Michaelis-Menten constant (Km and kcat values) against 1-tetradecanol (oxidation reaction) or tetradecanal (reduction reaction), were calculated using the Hanes-Woolf plot. The molar extinction coefficient for NADH at 340 nm is 6.22 mmol−1 1 cm−1. For the oxidation reaction, the mixture (1 mL) consisted of 100 mM Tris-HCl buffer (pH 9.0), 0.5 mM 1-tetradecanol, 0.5 mM NAD+, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH. For the reduction reaction, the mixture (1 mL) consisted of 100 mM KPB (pH 7.0), 0.5 mM tetradecanal, 0.24 mM NADH, 0.1% (wt/vol) Triton X-100, and 3.0 μg of purified PsADH. After mixing, the reaction mixture was incubated at 25°C for 3 min and the absorbance at 340 nm was monitored.
Coexpression of PsADH, NpAD, Fd, and FNR.
The methods of vector construction are the same as those described in the previous section. The genes encoding NpAD, Fd, and FNR are listed in Table S1 (NCBI:protein accession numbers WP_012408400, WP_012408585, and WP_012409282, respectively). For constructing the pRSF-PsADH-NpAD plasmid, the PsADH gene was amplified using PCR, digested with SalI and NotI, and cloned into the pRSF Duet-1 vector, creating the pRSF-PsADH plasmid. Then the pRSF-PsADH was cut with NdeI and XhoI and ligated with the NdeI/XhoI-digested PCR product of the NpAD gene, resulting in pRSF-PsADH-NpAD. The same procedure was applied when constructing the pCDF-Fd-FNR plasmid, where codon-optimized Fd and FNR genes were cloned into the pCDFDuet-1 vector using the restriction enzymes NcoI/NotI and NdeI/PacI, respectively. The two plasmids were transformed into E. coli Rosetta 2 (DE3) competent cells, resulting in E. coli Rosetta 2 (DE3)/pRSF-PsADH-NpAD/pCDF-Fd-FNR. For expression, the transformed E. coli cells were inoculated into LB medium containing chloramphenicol, kanamycin, and streptomycin and cultivated at 37°C with shaking. When the optical density at 600 nm reached 0.8, culture was induced with 0.5 mM IPTG and cultivated at 16°C for 16 h with shaking. After cultivation, cells were harvested via centrifugation, washed with 0.85% NaCl solution, and stored at −20°C for further use.
Alkane productivity assay.
Alkane production by E. coli Rosetta 2 (DE3)/pRSF-PsADH-NpAD/pCDF-Fd-FNR was evaluated using its resting cells. Another strain of E. coli, Rosetta 2 (DE3)/pRSF-NpAD/pCDF-Fd-FNR, which does not harbor the PsADH gene, was used as the control. The reaction mixture (1 mL) consisted of 100 mM Tris-HCl buffer (pH 9.0), 2 mM 1-tetradecanol, 5 mM NAD+, 5 mM NADPH, 0.1% (wt/vol) Triton X-100, and approximately 20 mg (wet weight) of induced E. coli cells. The reaction mixture was prepared in a 15-mL screw-cap test tube and incubated with shaking (120 strokes/min) at 40°C for 24 h. After the reaction, the mixture was acidified with 5 N HCl and extracted with 1 mL of toluene containing 1 mM 1-pentadecanol as the internal standard. The extracted sample was then subjected to GC and GC-MS to analyze the products.
Data availability.
All data are included here and in the supplemental material.
ACKNOWLEDGMENTS
This work was partially supported by the New Energy and Industrial Technology Development Organization (NEDO). The results were obtained by using microbial resources developed in the NEDO project (J.O.).
Y.-A.S., S.K., S.M., and J.O. designed the experiments. Y.-A.S. and S.M. conducted the experiments. Y.-A.S., S.K., S.M., M.I., M.M., S.O., and J.O. analyzed the data. Y.-A.S., S.K., and J.O. wrote the manuscript. M.I., M.M., S.O., and J.O. supervised the project. All authors read and approved the manuscript.
We declare no conflict of interest.
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Information & Contributors
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Published In
Applied and Environmental Microbiology
Volume 88 • Number 23 • 13 December 2022
eLocator: e01264-22
Editor: Arpita Bose, Washington University in St. Louis
PubMed: 36416567
Copyright
© 2022 American Society for Microbiology. All Rights Reserved.
History
Received: 2 August 2022
Accepted: 24 October 2022
Published online: 23 November 2022
Keywords
Contributors
Editor
Arpita Bose
Editor
Washington University in St. Louis
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2022.
Utilizing Alcohol for Alkane Biosynthesis by Introducing a Fatty Alcohol Dehydrogenase. Appl Environ Microbiol
88:e01264-22.
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