Improved n-butanol production via co-expression of membrane-targeted tilapia metallothionein and the clostridial metabolic pathway in Escherichia coli
© The Author(s). 2017
Received: 27 October 2016
Accepted: 22 March 2017
Published: 11 April 2017
N-Butanol has favorable characteristics for use as either an alternative fuel or platform chemical. Bio-based n-butanol production using microbes is an emerging technology that requires further development. Although bio-industrial microbes such as Escherichia coli have been engineered to produce n-butanol, reactive oxygen species (ROS)-mediated toxicity may limit productivity. Previously, we show that outer-membrane-targeted tilapia metallothionein (OmpC-TMT) is more effective as an ROS scavenger than human and mouse metallothioneins to reduce oxidative stress in the host cell.
The host strain (BUT1-DE) containing the clostridial n-butanol pathway displayed a decreased growth rate and limited n-butanol productivity, likely due to ROS accumulation. The clostridial n-butanol pathway was co-engineered with inducible OmpC-TMT in E. coli (BUT3-DE) for simultaneous ROS removal, and its effect on n-butanol productivity was examined. The ROS scavenging ability of cells overexpressing OmpC-TMT was examined and showed an approximately twofold increase in capacity. The modified strain improved n-butanol productivity to 320 mg/L, whereas the control strain produced only 95.1 mg/L. Transcriptomic analysis revealed three major KEGG pathways that were significantly differentially expressed in the BUT3-DE strain compared with their expression in the BUT1-DE strain, including genes involved in oxidative phosphorylation, fructose and mannose metabolism and glycolysis/gluconeogenesis.
These results indicate that OmpC-TMT can increase n-butanol production by scavenging ROS. The transcriptomic analysis suggested that n-butanol causes quinone malfunction, resulting in oxidative-phosphorylation-related nuo operon downregulation, which would diminish the ability to convert NADH to NAD+ and generate proton motive force. However, fructose and mannose metabolism-related genes (fucA, srlE and srlA) were upregulated, and glycolysis/gluconeogenesis-related genes (pfkB, pgm) were downregulated, which further assisted in regulating NADH/NAD+ redox and preventing additional ATP depletion. These results indicated that more NADH and ATP were required in the n-butanol synthetic pathway. Our study demonstrates a potential approach to increase the robustness of microorganisms and the production of toxic chemicals through the ability to reduce oxidative stress.
KeywordsTilapia metallothionein OmpC n-butanol E. coli Oxidative stress Transcriptomic analysis
n-Butanol has many advantages over ethanol, including a higher energy density due to two extra carbons, and can be used in gasoline engines without modification. n-Butanol is less hygroscopic and volatile than ethanol and has been recently regarded as a more viable transportation biofuel than ethanol [1–3]. n-Butanol is also a versatile platform chemical that can be produced from a variety of biomass sources. In industry, many important chemicals derived from n-butanol are used extensively as solvents or as intermediates in the production of acrylates, n-butyl acetate, esters, and glycol ethers .
Several industrial microbes, including Escherichia coli, have been engineered to produce n-butanol. In the initial stage of engineering E. coli for n-butanol production, the whole n-butanol pathway from Clostridium was transferred to E. coli, including the thil, hbd, crt, bcd, etfA, etfB, and adhe (or adhel) genes, catalyzing the six-step conversion of two molecules of acetyl-CoA into one molecule of n-butanol. However, when the clostridial n-butanol pathway was first transferred to E. coli using plasmids in 2007, the engineered strain produced less than 1 g/L n-butanol (vs. a clostridial n-butanol titer of 10–20 g/L) [5, 6]. The results indicated that engineering efficient n-butanol-producing E. coli is not as simple as expressing several clostridial n-butanol pathway genes. Over the following years, the best heterologous n-butanol-producing strains derived from E. coli were able to produce 14–15 g/L n-butanol [7, 8], providing industrial advantages compared to clostridial strains . However, the toxicity of n-butanol to both natural producers and engineered hosts increases with accumulation [10, 11]. This toxicity complicates the economically efficient production of large titers of n-butanol; therefore, high-titer n-butanol (30 g/L) production currently relies on in situ product removal by continuous gas stripping .
Furthermore, although E. coli can convert sugars into n-butanol at relatively high levels, the cells cannot tolerate 2% (v/v) n-butanol  and produce n-butanol at insufficient levels. Considering the relationship between n-butanol tolerance and n-butanol production in Clostridial strains [12, 13], the toxicity of n-butanol to E. coli can be considered a production bottleneck. In fact, product toxicity is a widespread problem in the production of biorenewables. Therefore, the development of a stress-tolerant host strain for the bio-production of titer-dependent toxic chemicals is important.
To understand the effect of n-butanol toxicity on the host, cell-wide studies have been conducted to obtain a global view of the n-butanol stress-response at the transcript, protein, and metabolite levels. Transcriptomic analysis in Clostridium acetobutylicum indicated that the primary response involved the accumulation of transcripts encoding chaperones, proteases, and other heat shock-related proteins .
In E. coli, several transcriptional analyses have been performed to investigate the stress caused by alcohols, including ethanol, n-butanol, and isobutanol [15–17]. Additionally, observations from fluorescent dye-staining indicated a large increase in reactive oxygen species (ROS) during n-butanol stress . Increased oxidative stress is a common cellular response to extracellular xenobiotics, which may mediate macromolecular damage. These free radicals can directly attack the membrane via lipid peroxidation or cause DNA mutations, protein misfolding and fragmentation, and apoptosis [18–20].
To decrease ROS-induced oxidative damage, microorganisms synthesize many antioxidant enzymes, including catalases, superoxide dismutases and glutathione peroxidase [21, 22]. Recently, metallothioneins (MTs), beneficial antioxidant enzymes that occur widely in mammals, plants and fungi, have been identified . MTs are heat-stable, low-molecular-weight and cysteine-rich intracellular proteins [23–25]. MTs also act as a defense system against oxidative stress through their ROS scavenging ability . Furthermore, purified tilapia MT (TMT) has been shown to outperform glutathione (GSH) in scavenging both 2-diphenyl-1-picrylhydrazyl (DPPH●) and 2,2-azinobis (3-ethylbenzothiazoline- 6-sulfonic acid) diammonium salt (ABTS●+) . To reduce oxidative stress in the host cell, we previously engineered MTs into E. coli hosts for both cytosolic and outer-membrane-targeted (osmoregulatory membrane protein, OmpC, fused) expression. The ability of these engineered E. coli to scavenge intracellular or extracellular ROS was examined, and OmpC-fused TMT performed the best, growing in medium containing 1.5% n-butanol. n-Butanol tolerance was increased through the scavenging of intracellular and extracellular free radicals, and the fusion protein still contributed to osmotic tolerance via either glycine betaine or glucose uptake .
Intracellular ROS increase levels in E. coli after exposure to n-butanol . We therefore considered whether introducing OmpC-fused TMT into butanologenic E. coli would improve n-butanol tolerance and productivity. In the present study, we engineered the C. acetobutylicum n-butanol synthetic pathway into E. coli, along with OmpC-TMT co-expression to increase n-butanol tolerance. A synthetic biological technical platform (Ordered Gene Assembly in Bacillus subtilis – OGAB method) was employed for the genetic manipulation of the n-butanol production and tolerance gene clusters . Our results demonstrate that co-expressing OmpC-fused TMT in a butanologenic E. coli strain enhances n-butanol production.
Results and discussion
Construction of butanologenic E. coli strains
Strains and plasmids used in this study
Strains and plasmids
Genotype and descriptiona
Reference or source
E. coli strains
E. coli BL21/PET30a, T7 promoter, f1 origin; ompC-tmt, Kmr
endA1, recA1, gyrA96, thi, hsdR17 (rk–, mk+), relA1, supE44, Δ(lac-proAB), [F’ traD36, proAB, laqIqZΔM15].
E. coli JM109/pGETS118, CATr
E. coli JM109/pBUT, CATr
E. coli JM109/pBUT-rO-tMT, CATr
E. coli JM109/pBUT-T7-rO-tMT, CATr
JM109 + λ(DE3),Contains an IPTG-inducible gene for T7 RNA polymerase
E. coli JM109 (DE3)/pGETS118, CATr
E. coli JM109 (DE3)/pBUT, CATr
E. coli JM109 (DE3)/pBUT-rO-tMT, CATr
E. coli JM109 (DE3)/pBUT-T7-rO-tMT, CATr
pGETS118SfiI-Pr, Tcr, CATr
pGETS118::thil-crt-bcd-etfAB-hbd-adhe, Tcr, CATr
pBUT::rbs-ompC-tmt, Tcr, CATr
pBUT::T7 promoter-ompC-tmt, Tcr, CATr
Growth profile and n-butanol production
Recently, many groups have reported successful n-butanol production in E. coli [5, 6, 8]; a similar approach was previously applied to n-butanol production in E. coli carrying the CoA-dependent synthetic pathway, yielding final n-butanol titers of 552 mg/L to 1.2 g/L [5, 6]. In addition, Shen et al.  further improved the strain by using NADH- and acetyl-CoA-driving forces; the best strain in their study showed high-titer production of n-butanol through in situ product removal. The production of n-butanol has been confirmed to affect host growth.
Improved tolerance and n-butanol production
A clostridial strain overexpressing GSH was reported to exhibit improved tolerance and increased n-butanol productivity . Boyarskiy et al.  reported increased n-butanol tolerance and production (approximately 35% increases) through transcriptional feedback regulation of efflux protein expression.
In the present study, we attempted to improve the n-butanol tolerance of butanologenic E. coli to further enhance n-butanol production by expressing OmpC-TMT. The concentrations of n-butanol found to be inhibitory to E. coli in n-butanol production strains (Fig. 2) were consistent with the trends noted in the literature for alcohol toxicity in E. coli . Hence, we examined the normal expression of OmpC-TMT (BUT2, BUT3) in butanologenic E. coli strains, and the results showed that cell growth was still affected by n-butanol. This result is similar to that obtained for the BUT1 strain (Fig. 2a).
However, the BUT2 and BUT3 strains showed similar n-butanol production (118 mg/L and 121 mg/L) after 20 h of incubation (Fig. 3a). After incubation for 60 h, the butanologenic E. coli strains BUT2 and BUT3 showed 1.6-fold and 2.7-fold higher n-butanol production, respectively, than BUT1 (94.6 mg/L). Notably, BUT3-DE achieved a 1.7-fold increase in n-butanol production compared to that achieved by BUT2-DE after 20 h of incubation (Fig. 3b). Furthermore, cell growth in the OmpC-TMT-overexpressing butanologenic E. coli (BUT3-DE) was nearly the same as in the control strain (PGETS118-DE) after 60 h of incubation (Fig. 2b). More importantly, n-butanol production increased nearly 3.4-fold compared with that of BUT1-DE (320 mg/L) and showed a significantly improved conversion rate from glucose to n-butanol (4.97 mg/L n-butanol/1 g glucose) (Fig. 3b). These results suggested that OmpC-TMT overexpression rescued the growth of the butanologenic E. coli strains, improved host robustness and increased n-butanol productivity. Our results are consistent with previous reports [28, 30, 31].
Engineering strategies to improve butanol tolerance and production in E. coli
Overexpression of ompC-tmt
From 1.50 to 2.00%
thil, crt, bcd, etfA, etfB, hbd, adhe, ompC-tmt
atoB, crt, bcd, etfAB, hbd, adhe, ΔadhE, ΔldhA, ΔfrdBC, Δfnr, Δpta
thil, hbd, crt, bcd, etfA, etfB, adhe, high cell density
Overexpression of ATF, fabD, feoA and srpABC
From 1.50 to 2.00%
double disruptions of proV and marR Mutation of lon
From 1.00 to 2.00%
Overexpression of groESL
From 0.75 to 1.00%
Mutation of cyclic AMP receptor protein (CRP)
From 0.80 to 1.20%
Aside from n-butanol, carboxylic acids are attractive biorenewable chemicals, such as fatty acid and aromatic carboxylic acids [39–41]. However, product toxicity is frequently encountered in metabolic engineering, and the toxicity of these carboxylic acids to the microbial biocatalyst appears to limit biocatalyst performance. Strong evidence suggests that membrane damage is the main mechanism of toxicity [42, 43], and previous metabolic engineering efforts successfully increased membrane integrity by modulating membrane composition to alter free fatty acid tolerance in E. coli [44–46].
Free radical scavenging ability and membrane integrity test
Since the cellular membrane is a vital factor that allows cells to acclimate to external stresses and is also one of the components that is strongly affected by organic solvents , many studies have proposed that the plasma membrane is the most affected target of organic solvents and plays a significant role in adapting to stress. Additionally, the length of the carbon backbone of the organic solvent can alter the toxicity mechanism; increasing the hydrophobicity of the solvent increases the level of toxicity . Long- and short-chain alcohols cause stress during biofuel production by altering membrane fluidity (also known as Overton’s Rule). Ethanol and n-butanol, respectively, decrease and increase membrane fluidity [47, 49, 50].
Toxicity that causes membrane damage is likely a key limitation for butanologenic microbes to produce butanol, suggesting that our engineering strategy of expressing OmpC-TMT is effective in protecting membrane integrity and scavenging free radicals to improve n-butanol tolerance and production in butanologenic E. coli strains.
Expression profiles of n-butanol and tolerance genes in E. coli
Real-time RT-PCR analysis
Gene expression log2 fold change a [log2 (2-ΔΔCT (Exp-Control))]
-7.79 ± 0.02
-8.64 ± 0.08
-7.72 ± 0.06
-1.40 ± 0.02
-3.09 ± 0.12
-4.14 ± 0.69
-0.76 ± 0.10
2.84 ± 0.09
4.70 ± 0.11
4.50 ± 0.68
6.96 ± 0.16
4.24 ± 0.08
Transcriptomic analysis in engineered E. coli
KEGG biological pathways for significantly expressed genes a
NADH:ubiquinone oxidoreductase, chain I
NADH:ubiquinone oxidoreductase, chain G
NADH:ubiquinone oxidoreductase, chain F
NADH:ubiquinone oxidoreductase, fused CD subunit
Fructose and mannose metabolism
glucitol/sorbitol-specific enzyme IIB component of PTS
glucitol/sorbitol-specific enzyme IIC component of PTS
50S ribosomal subunit protein L1
30S ribosomal subunit protein S6
50S ribosomal subunit protein L32
50S ribosomal subunit protein L9
Nicotinate and nicotinamide metabolism
pyridine nucleotide transhydrogenase, alpha subunit
However, the past results showed that isobutanol disrupts the cell membrane, leading to quinone malfunction, which results in the release of quinone inhibition on ArcB and the subsequent autophosphorylation of ArcB and activation of ArcA, which adapts cellular metabolism for growth with decreased respiratory efficiency, such as downregulation of NADH dehydrogenase I (nuo operon) or upregulation of cytochrome d oxidase (cydAB operon) . This hypothesis coincides with our results. In addition, microbial fermentation pathways involve many redox reactions, which usually require NADH and NAD+ as cofactors. However, in the clostridial synthetic pathway, the synthesis of n-butanol from glucose can cause an NADH/NAD+ redox imbalance because more NADH is required in the synthetic pathway than is generated in the glycolytic pathway . Therefore, compared with the control strain, in the modified strain, a number of upregulated genes, such as fucA, srlE and srlA, may help supplement metabolic intermediates for fructose and mannose metabolism to generate NADH and assist in regulating NADH/NAD+ redox [66, 67]. These results suggested that maintaining the redox balance of NADH and NAD+ is a key to ensure the continued operation of cellular metabolism under n-butanol fermentation. The same strategy was used in previous research [8, 65]. Moreover, isobutanol disrupts quinone/quinol function, resulting in the malfunction of NADH ubiquinone oxidoreductase (complex I), which would diminish the ability to convert NADH to NAD+ and generate the proton motive force (PMF) . This process may affect the biosynthesis of ATP in the cell. We also found that two genes (pfkB, pgm) involved in glycolysis/gluconeogenesis were downregulated. One possible explanation is that the downregulation of pfkB, which encodes 6-phosphofructokinase II, results in the decreased conversion of fructose-6-phosphate (F6P) to fructose-1, 6-biphosphate, thereby reducing the consumption of ATP. Additionally, pgm encodes phosphoglucomutase and facilitates the interconversion of glucose-1-phosphate and glucose-6-phosphate (G6P). One possible explanation for the downregulation of pgm is that glucose-1-phosphate is further converted to ADP-glucose by ATP depletion. This result indicated that BUT3-DE tried to reduce ATP consumption following NADH dehydrogenase I malfunction. Moreover, both the pfkB and pgm genes were downregulated, which may result in the accumulation of G6P and F6P. Both G6P and F6P, as starting metabolites for the pentose phosphate pathway, produce precursors and provide a major source of NADPH for biosynthesis, which is involved in most of reductive pathways [68, 69]. However, previous studies show the energy-dependent reduction of NADP+ with NADH by PntAB, but increased NADPH formation will downregulate pntA for genetic and environmental manipulation .
In general, cells exposed to n-butanol show elevated levels of oxidative stress, and many oxidative-related genes have been identified as being significantly upregulated . However, in our case, although BUT3-DE showed elevated butanol production and higher radical scavenging capacity than BUT1-DE (Figs. 3b and 5b), we did not detect significant expression of oxidative stress-related genes. In fact, our analysis found that genes related to oxidative stress (sodA) and cell envelope stress (cpxP) were expressed (Additional file 1: Table S3). Meanwhile, KEGG pathway analysis revealed no significant differences between BUT1-DE and BUT3-DE with respect to these genes, and thus was not included in Table 4. Given that BUT1-DE experienced higher oxidative stress than BUT3-DE (Fig. 5b), these results suggest that OmpC-TMT overexpression decreased ROS production, as well as the expression of oxidative stress-related genes, in BUT3-DE.
This study uses a novel approach to develop E. coli strains that express both the n-butanol synthesis pathway and membrane-targeted MTs to improve cell fermentation via ROS scavenging. OmpC-TMT was able to decrease the production of free radicals and to improve host robustness during fermentation. Furthermore, a balanced intracellular redox state in microbes is recognized as essential to ensure the efficient production of fermentation products. Unfortunately, our studies suggest that although OmpC-TMT expression increased n-butanol production compared to that of the control strain, the accumulation of fermented products (e.g., n-butanol) in the cell or external environment cause malfunctions in the NADH and ATP synthesis system. Therefore, this fermentation system requires further improvement. In principle, the expression of membrane-targeted TMT has the potential to improve the alcohol fermentation bioprocess and may also serve as a practical strategy for the construction of platform E. coli strains for biofuel production.
All chemicals and reagents were purchased from Sigma-Aldrich Co., USA, unless otherwise noted. The reagents, when available, were molecular biology grade. All solutions were prepared using these reagents and sterile distilled water.
Construction of plasmids via the OGAB method
The OGAB method was described previously . The primer sequences used in this study are listed in Additional file 2: Table S1. To construct an expression plasmid containing the clostridial n-butanol synthesis pathway, thil (1.2 kb) and the crt-bcd-etfB-etfA-hbd cluster (4.8 kb) were first amplified from the genomic DNA of C. acetobutylicum ATCC824, and the adhel (2.7 kb) gene was amplified from C. acetobutylicum ATCC824 megaplasmid pSOL1. ompC-tmt (1.3 kb) was amplified from pET-OmTmt . The PCR products were subcloned into the yT&A cloning vector (YEASTERN) and confirmed by DNA sequencing. All PCR products were digested and then assembled as one transcriptional unit into pGETS118 in a polycistronic manner using the OGAB method, resulting in pBUT, pBUT-rO-tMT and pBUT-T7-rO-tMT (Fig. 1). All genes and an assembly vector were prepared as DNA fragments with arbitrary 3’-nucleotide protrusions that allowed the genes to be linked together in a specific order and direction. The protrusions were generated by type II restriction endonucleases that recognize two separate recognition sites, such as SfiI, which recognizes 5’-GGCCANNNAGGCC-3’, where N is any nucleotide in a random sequence. SfiI and BglI were chosen in this study since SfiI and BglI sites are absent in the relevant genes and vectors. The constructed plasmids were transferred into E. coli JM109 and JM109 (DE3) to obtain the recombinant E. coli strains BUT1, BUT2, BUT3, BUT1-DE, BUT2-DE and BUT3-DE (Table 1) and to assay the production of n-butanol.
Bacterial strains, culture media and growth conditions
MT-expressing engineered strains, protein expression and their locations in recombinants E. coli hosts were confirmed in our previous study . All batch cultures were grown at 37 °C in a rotary shaker at 200 rpm. Batch cultures were grown in 150 mL PYG medium under anaerobic conditions (per liter: 5 g of peptone, 10 g of yeast extract, 60 g of glucose, 5 g of tryptone, 40 mg of K2HPO4, 19.2 mg of MgSO4.7 (H2O), 8 mg of CaCl2, 40 mg of KH2PO4, 0.4 g of NaHCO3, 80 mg of NaCl and 1.1 mg of FeSO4.7 H2O) or in M9 (AMRESCO-J863) media. Each engineered E. coli strain, including PGETS118, BUT1, BUT2, BUT3, PGETS118-DE, BUT1-DE, BUT2-DE, and BUT3-DE (Table 1), was grown in medium supplemented with 50 μg/mL chloramphenicol. When the culture density reached OD 0.6, isopropyl-β-D-thiogalactopyranoside (IPTG) was added to a final culture concentration of 0.6 mM. After 12 h of incubation, cells were harvested for ROS and n-butanol production experiments. All solvent concentrations in media are reported as mg/L.
Cell density was measured at 600 nm using a UV–vis spectrophotometer (GENESYS 10S, Thermo Scientific, USA). Fermentation samples were analyzed by gas chromatography (GC) on an Agilent 7890A equipped with a split/splitless injector, flame ionization detector, and a DP-FFAP capillary column (30 m, 0.32 mm i.d., 0.25 μm film thickness). The temperatures of the injector and detector were both maintained at 225 °C. The temperature profile of the column oven was as follows: (i) initial hold at 50 °C for 4 min, increase to 100 °C at 20 °C/min and hold for 1 min; (ii) increase to 170 °C at 30 °C/min and hold for 2.5 min; and (iii) increase to 220 °C at 20 °C/min and hold for 4 min. Nitrogen gas was used as the carrier gas. Split injection mode was used, with a split ratio of 1:10.
Reactive oxygen species detected by carboxy-H2DCFDA under n-butanol stress
The engineered E. coli strains were cultured in PYG medium, and different concentrations of n-butanol were produced after 0, 20, 36 and 60 h of incubation. Aliquots of 100 μL of the cultured strains were re-suspended in 5 mL of M9 medium, and 140 μL of each diluted sample was transferred to a 96-well plate, followed by incubation at 37 °C. The assay method was adapted from a previous study . All samples were treated with 10 μL of 25 mM carboxy-H2DCFDA (Invitrogen, Co., Carlsbad, CA) and incubated at 37 °C for 15 min. The optical density at 600 nm and the fluorescence excitation/emission at 535/600 nm of each sample were measured on a plate reader. Tert-butyl hydroperoxide (TBHP) (Invitrogen, Carlsbad, CA) is a known stressor that produces intracellular H2O2; a set of positive controls for the ROS assay were prepared with strains cultured after inoculation and were treated as above, except with an initial dilution step with 10 μL of 7.78 M TBHP.
Staining of bacterial suspensions with DAPI and SYTOX Green
Bacterial suspensions containing either 1.0 × 106 or 1.2 × 106 organisms/mL were stained with 0.5 μM SYTOX Green and 1 μg/ml DAPI. The fluorescence emissions from E. coli were compared with the background fluorescence of each stain alone at the optimal excitation wavelength for each nucleic acid stain (DAPI, 358 nm; SYTOX Green, 504 nm). The mixture of cells and DAPI was incubated at 37 °C for 10 min, and the cells were washed twice with TBS buffer (pH 7.6). Next, the DAPI-stained cells were stained with SYTOX Green nucleic acid again, incubated at 37 °C for 10 min, and washed three times with TBS buffer (pH 7.6). Fluorescence was observed on a Nikon ECLIPSE 80i (Nikon, Tokyo, Japan).
Next-generation sequencing and analysis
After 60 h of incubation, total RNA was isolated from both butanologenic strains (BUT1-DE and BUT3-DE) for next-generation sequencing (RNA-Seq), which was performed by Welgene Biotech Co., Ltd. (Taipei, Taiwan). Total RNA was extracted using the Trizol® Reagent (Invitrogen, USA) according to the manufacturer’s instructions. Purified RNA was quantified at 260 nm using an ND-1000 spectrophotometer (Nanodrop Technology, USA) and was qualitatively analyzed using a Bioanalyzer 2100 (Agilent Technology, USA) with an RNA 6000 labchip kit (Agilent Technologies, USA). All procedures were performed according to the Illumina protocol. For all samples, library construction was performed using TruSeq RNA Sample Prep Kits v2 for 75 bp (Single-End) sequencing and the Solexa platform (Illumina Inc.). Sequences were directly determined using sequencing-by-synthesis technology via the TruSeq SBS Kit. Raw sequences were obtained from the Illumina Pipeline software bcl2fastq v2.0 and were expected to generate 10 M (million reads or Gb) per sample. The generated sequences were filtered to obtain qualified reads. Trimmomatics was implemented to trim or remove reads according to the quality score . Qualified reads after filtering low-quality data were analyzed using TopHat/Cufflinks for gene expression estimation . The gene expression level was calculated as FPKM (fragments per kilobase of transcript per million mapped reads). For differential expression analysis, CummeRbund was employed to perform statistical analysis of the gene expression profiles. The reference genome and gene annotations were retrieved from the Ensembl database for E. coli str. K-12 substr. W3110. The cuffdiff tool from the cufflinks package was run to calculate expression changes and the associated P-values for each gene between strain BUT1-DE (control) and strain BUT3-DE (Table 1) after 60 h of incubation. The output files of cuffdiff were annotated by adding gene functional descriptions. KEGG term enrichment analysis and fold-change enrichment for the gene lists of significantly upregulated and downregulated genes in two butanologenic strains were performed. KEGG pathways were analyzed using NIH DAVID Bioinformatics Resources 6.7  to identify the regulated biological themes.
Quantitative Real-Time PCR Analysis
Total RNA for quantitative reverse transcriptase PCR (qPCR) was extracted as described for next-generation sequencing and analysis. Two-step quantitative real-time PCR (qPCR) was conducted using LightCycler ® FastStart DNA Master PLUS SYBR Green I (Roche Life Science) on a Light Cycler 480 Real-Time PCR 3System (Roche Co., Germany) and was performed by Welgene Biotech Co., Ltd. (Taipei, Taiwan). The four genetic regions involved in n-butanol production pathway (thil, crt-bcd-etfB-etfA-hbd, adhe) and ROS-targeted scavenging (ompC-tmt) were analyzed. R16s (rrsA gene) was used as a reference in all analyses. The primers are provided in Additional file 3: Table S2 in the supplemental material. For the relative quantification of gene expression, the comparative CT method was employed. The averaged CT was subtracted from the corresponding averaged r16S value for each sample; resulting in ∆CT. ∆∆CT was obtained by subtracting the average control ∆CT value from the average experimental ∆CT. The fold increase was determined by calculating log2 (2−∆∆CT) for the experimental vs. control samples.
Medium containing peptone, yeast extract and glucose.
Reactive oxygen species
We thank Jian-Rong Li and Tzu-Hang Yuan for help with the next-generation sequencing data analysis and in gathering information.
This work was supported by grants (MOST 104-2621-M-005-003-MY3) from the National Science Council, Taiwan.
Availability of data and materials
The datasets supporting the conclusions of this article are included within the manuscript and its additional files. The data set supporting the results of this article are available in the NCBI Sequence Read Archive (http://www.ncbi.nlm.nih.gov/sra/) repositories, SRR5352318, and SRR5352319.
WCC performed the experiments, analyzed the data and drafted the manuscript. KHL performed some of the experimental work. KHL contributed to data interpretation. WCC and KHL wrote and revised the manuscript. WCC and CCH designed the study. CCH coordinated the study. All authors read and approved the final manuscript.
The authors declare no competing interests.
Consent for publication
Ethics approval and consent to participate
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Ezeji TC, Qureshi N, Blaschek HP. Bioproduction of butanol from biomass: from genes to bioreactors. Curr Opin Biotechnol. 2007;18(3):220–7.View ArticleGoogle Scholar
- Green EM. Fermentative production of butanol--the industrial perspective. Curr Opin Biotechnol. 2011;22(3):337–43.View ArticleGoogle Scholar
- Procentese A, Raganati F, Olivieri G, Russo ME, De La Feld M, Marzocchella A. Renewable feedstocks for biobutanol production by fermentation. N Biotechnol. 2016. doi: 10.1016/j.nbt.2016.10.010.
- Harvey BG, Meylemans HA. The role of butanol in the development of sustainable fuel technologies. J Chem Technol Biotechnol. 2011;86(1):2–9.View ArticleGoogle Scholar
- Atsumi S, Cann AF, Connor MR, Shen CR, Smith KM, Brynildsen MP, et al. Metabolic engineering of Escherichia coli for 1-butanol production. Metab Eng. 2008;10(6):305–11.View ArticleGoogle Scholar
- Inui M, Suda M, Kimura S, Yasuda K, Suzuki H, Toda H, et al. Expression of Clostridium acetobutylicum butanol synthetic genes in Escherichia coli. Appl Microbiol Biotechnol. 2008;77(6):1305–16.View ArticleGoogle Scholar
- Dellomonaco C, Clomburg JM, Miller EN, Gonzalez R. Engineered reversal of the beta-oxidation cycle for the synthesis of fuels and chemicals. Nature. 2011;476(7360):355–9.View ArticleGoogle Scholar
- Shen CR, Lan EI, Dekishima Y, Baez A, Cho KM, Liao JC. Driving forces enable high-titer anaerobic 1-butanol synthesis in Escherichia coli. Appl Environ Microbiol. 2011;77(9):2905–15.View ArticleGoogle Scholar
- Ye Q, Bao J, Zhong J-J. Bioreactor Engineering Research and Industrial Applications I: Cell Factories. 1st ed. Springer: Berlin; 2016. https://link.springer.com/book/10.1007%2F978-3-662-49161-4.
- Bowles LK, Ellefson WL. Effects of butanol on Clostridium acetobutylicum. Appl Environ Microbiol. 1985;50(5):1165–70.Google Scholar
- Knoshaug EP, Zhang M. Butanol tolerance in a selection of microorganisms. Appl Biochem Biotechnol. 2009;153(1-3):13–20.View ArticleGoogle Scholar
- Mao S, Luo Y, Zhang T, Li J, Bao G, Zhu Y, et al. Proteome reference map and comparative proteomic analysis between a wild type Clostridium acetobutylicum DSM 1731 and its mutant with enhanced butanol tolerance and butanol yield. J Proteome Res. 2010;9(6):3046–61.View ArticleGoogle Scholar
- Liu XB, Gu QY, Yu XB. Repetitive domestication to enhance butanol tolerance and production in Clostridium acetobutylicum through artificial simulation of bio-evolution. Bioresour Technol. 2013;130:638–43.View ArticleGoogle Scholar
- Tomas CA, Beamish J, Papoutsakis ET. Transcriptional analysis of butanol stress and tolerance in Clostridium acetobutylicum. J Bacteriol. 2004;186(7):2006–18.View ArticleGoogle Scholar
- Gonzalez R, Tao H, Purvis JE, York SW, Shanmugam KT, Ingram LO. Gene array-based identification of changes that contribute to ethanol tolerance in ethanologenic Escherichia coli: comparison of KO11 (parent) to LY01 (resistant mutant). Biotechnol Prog. 2003;19(2):612–23.View ArticleGoogle Scholar
- Brynildsen MP, Liao JC. An integrated network approach identifies the isobutanol response network of Escherichia coli. Mol Syst Biol. 2009;5:277.View ArticleGoogle Scholar
- Rutherford BJ, Dahl RH, Price RE, Szmidt HL, Benke PI, Mukhopadhyay A, et al. Functional genomic study of exogenous n-butanol stress in Escherichia coli. Appl Environ Microbiol. 2010;76(6):1935–45.View ArticleGoogle Scholar
- Sigler K, Chaloupka J, Brozmanova J, Stadler N, Hofer M. Oxidative stress in microorganisms--I. Microbial vs. higher cells--damage and defenses in relation to cell aging and death. Folia Microbiol (Praha). 1999;44(6):587–624.View ArticleGoogle Scholar
- Perrone GG, Tan SX, Dawes IW. Reactive oxygen species and yeast apoptosis. Biochim Biophys Acta. 2008;1783(7):1354–68.View ArticleGoogle Scholar
- Jones DP. Radical-free biology of oxidative stress. Am J Physiol Cell Physiol. 2008;295(4):C849–68.View ArticleGoogle Scholar
- Fridovich I. Superoxide radical and superoxide dismutases. Annu Rev Biochem. 1995;64:97–112.View ArticleGoogle Scholar
- Imlay JA. Pathways of oxidative damage. Annu Rev Microbiol. 2003;57:395–418.View ArticleGoogle Scholar
- Coyle P, Philcox JC, Carey LC, Rofe AM. Metallothionein: the multipurpose protein. Cell Mol Life Sci. 2002;59(4):627–47.View ArticleGoogle Scholar
- Vallee BL. The function of metallothionein. Neurochem Int. 1995;27(1):23–33.View ArticleGoogle Scholar
- Park JD, Liu Y, Klaassen CD. Protective effect of metallothionein against the toxicity of cadmium and other metals(1). Toxicology. 2001;163(2-3):93–100.View ArticleGoogle Scholar
- Sato M, Kondoh M. Recent studies on metallothionein: protection against toxicity of heavy metals and oxygen free radicals. Tohoku J Exp Med. 2002;196(1):9–22.View ArticleGoogle Scholar
- Lin KH, Chien MF, Hsieh JL, Huang CC. Mercury resistance and accumulation in Escherichia coli with cell surface expression of fish metallothionein. Appl Microbiol Biotechnol. 2010;87(2):561-69.
- Chin WC, Lin KH, Chang JJ, Huang CC. Improvement of n-butanol tolerance in Escherichia coli by membrane-targeted tilapia metallothionein. Biotechnol Biofuels. 2013;6(1):130.View ArticleGoogle Scholar
- Tsuge K, Matsui K, Itaya M. One step assembly of multiple DNA fragments with a designed order and orientation in Bacillus subtilis plasmid. Nucleic Acids Res. 2003;31(21):e133.View ArticleGoogle Scholar
- Zhu L, Dong H, Zhang Y, Li Y. Engineering the robustness of Clostridium acetobutylicum by introducing glutathione biosynthetic capability. Metab Eng. 2011;13(4):426–34.View ArticleGoogle Scholar
- Boyarskiy S, Davis Lopez S, Kong N, Tullman-Ercek D. Transcriptional feedback regulation of efflux protein expression for increased tolerance to and production of n-butanol. Metab Eng. 2016;33:130–7.View ArticleGoogle Scholar
- Kempf B, Bremer E. Uptake and synthesis of compatible solutes as microbial stress responses to high-osmolality environments. Arch Microbiol. 1998;170(5):319–30.View ArticleGoogle Scholar
- Zhang H, Chong H, Ching CB, Song H, Jiang R. Engineering global transcription factor cyclic AMP receptor protein of Escherichia coli for improved 1-butanol tolerance. Appl Microbiol Biotechnol. 2012;94(4):1107–17.View ArticleGoogle Scholar
- Zingaro KA, Terry PE. GroESL overexpression imparts Escherichia coli tolerance to i-, n-, and 2-butanol, 1,2,4-butanetriol and ethanol with complex and unpredictable patterns. Metab Eng. 2013;15:196–205.View ArticleGoogle Scholar
- Yuan Y, Bi C, Nicolaou SA, Zingaro KA, Ralston M, Papoutsakis ET. Overexpression of the Lactobacillus plantarum peptidoglycan biosynthesis murA2 gene increases the tolerance of Escherichia coli to alcohols and enhances ethanol production. Appl Microbiol Biotechnol. 2014;98(19):8399–411.View ArticleGoogle Scholar
- Zingaro KA, Nicolaou SA, Yuan Y, Papoutsakis ET. Exploring the heterologous genomic space for building, stepwise, complex, multicomponent tolerance to toxic chemicals. ACS Synth Biol. 2014;3(7):476–86.View ArticleGoogle Scholar
- Watanabe R, Doukyu N. Improvement of organic solvent tolerance by disruption of the lon gene in Escherichia coli. J Biosci Bioeng. 2014;118(2):139–44.View ArticleGoogle Scholar
- le Bui M, Lee JY, Geraldi A, Rahman Z, Lee JH, Kim SC. Improved n-butanol tolerance in Escherichia coli by controlling membrane related functions. J Biotechnol. 2015;204:33–44.View ArticleGoogle Scholar
- Si HM, Zhang F, Wu AN, Han RZ, Xu GC, Ni Y. DNA microarray of global transcription factor mutant reveals membrane-related proteins involved in n-butanol tolerance in Escherichia coli. Biotechnol Biofuels. 2016;9:114.View ArticleGoogle Scholar
- Mialon L, Vanderhenst R, Pemba AG, Miller SA. Polyalkylenehydroxybenzoates (PAHBs): biorenewable aromatic/aliphatic polyesters from lignin. Macromol Rapid Commun. 2011;32(17):1386–92.View ArticleGoogle Scholar
- Polen T, Spelberg M, Bott M. Toward biotechnological production of adipic acid and precursors from biorenewables. J Biotechnol. 2013;167(2):75–84.View ArticleGoogle Scholar
- Steen EJ, Kang Y, Bokinsky G, Hu Z, Schirmer A, McClure A, et al. Microbial production of fatty-acid-derived fuels and chemicals from plant biomass. Nature. 2010;463(7280):559–62.View ArticleGoogle Scholar
- Lennen RM, Kruziki MA, Kumar K, Zinkel RA, Burnum KE, Lipton MS, et al. Membrane stresses induced by overproduction of free fatty acids in Escherichia coli. Appl Environ Microbiol. 2011;77(22):8114–28.View ArticleGoogle Scholar
- Jarboe LR, Royce LA, Liu P. Understanding biocatalyst inhibition by carboxylic acids. Front Microbiol. 2013;4:272.View ArticleGoogle Scholar
- Lennen RM, Pfleger BF. Modulating membrane composition alters free fatty acid tolerance in Escherichia coli. PLoS One. 2013;8(1):e54031.View ArticleGoogle Scholar
- Royce LA, Yoon JM, Chen Y, Rickenbach E, Shanks JV, Jarboe LR. Evolution for exogenous octanoic acid tolerance improves carboxylic acid production and membrane integrity. Metab Eng. 2015;29:180–8.View ArticleGoogle Scholar
- Tan Z, Yoon JM, Nielsen DR, Shanks JV, Jarboe LR. Membrane engineering via trans unsaturated fatty acids production improves Escherichia coli robustness and production of biorenewables. Metab Eng. 2016;35:105–13.View ArticleGoogle Scholar
- Ingram L. Microbial tolerance to alcohols: role of the cell membrane. Trends Biotechnol. 1986;4(2):40–4.View ArticleGoogle Scholar
- Aono R, Nakajima H. Organic solvent tolerance in Escherichia coli. Tanpakushitsu Kakusan Koso. 1997;42(15):2532–41.Google Scholar
- Ingram LO, Buttke TM. Effects of alcohols on micro-organisms. Adv Microb Physiol. 1984;25:253–300.View ArticleGoogle Scholar
- Kabelitz N, Santos PM, Heipieper HJ. Effect of aliphatic alcohols on growth and degree of saturation of membrane lipids in Acinetobacter calcoaceticus. FEMS Microbiol Lett. 2003;220(2):223–7.View ArticleGoogle Scholar
- Trautwein K, Kuhner S, Wohlbrand L, Halder T, Kuchta K, Steinbuchel A, et al. Solvent stress response of the denitrifying bacterium “Aromatoleum aromaticum” strain EbN1. Appl Environ Microbiol. 2008;74(8):2267–74.View ArticleGoogle Scholar
- Minty JJ, Lesnefsky AA, Lin F, Chen Y, Zaroff TA, Veloso AB, et al. Evolution combined with genomic study elucidates genetic bases of isobutanol tolerance in Escherichia coli. Microb Cell Fact. 2011;10:18.View ArticleGoogle Scholar
- Ingram LO. Ethanol tolerance in bacteria. Crit Rev Biotechnol. 1990;9(4):305–19.View ArticleGoogle Scholar
- Goodarzi H, Bennett BD, Amini S, Reaves ML, Hottes AK, Rabinowitz JD, et al. Regulatory and metabolic rewiring during laboratory evolution of ethanol tolerance in E. coli. Mol Syst Biol. 2010;6:378.View ArticleGoogle Scholar
- Woodruff LB, Pandhal J, Ow SY, Karimpour-Fard A, Weiss SJ, Wright PC, et al. Genome-scale identification and characterization of ethanol tolerance genes in Escherichia coli. Metab Eng. 2013;15:124–33.View ArticleGoogle Scholar
- Asako H, Nakajima H, Kobayashi K, Kobayashi M, Aono R. Organic solvent tolerance and antibiotic resistance increased by overexpression of marA in Escherichia coli. Appl Environ Microbiol. 1997;63(4):1428–33.Google Scholar
- Dien BS, Cotta MA, Jeffries TW. Bacteria engineered for fuel ethanol production: current status. Appl Microbiol Biotechnol. 2003;63(3):258–66.View ArticleGoogle Scholar
- Luo LH, Seo PS, Seo JW, Heo SY, Kim DH, Kim CH. Improved ethanol tolerance in Escherichia coli by changing the cellular fatty acids composition through genetic manipulation. Biotechnol Lett. 2009;31(12):1867–71.View ArticleGoogle Scholar
- Jarboe LR, Liu P, Royce LA. Engineering inhibitor tolerance for the production of biorenewable fuels and chemicals. Curr Opin Chem Eng. 2011;1(1):38–42.View ArticleGoogle Scholar
- Ramskold D, Wang ET, Burge CB, Sandberg R. An abundance of ubiquitously expressed genes revealed by tissue transcriptome sequence data. PLoS Comput Biol. 2009;5(12):e1000598.View ArticleGoogle Scholar
- Ingledew WJ, Poole RK. The respiratory chains of Escherichia coli. Microbiol Rev. 1984;48(3):222–71.Google Scholar
- Georgellis D, Kwon O, Lin EC. Quinones as the redox signal for the arc two-component system of bacteria. Science. 2001;292(5525):2314–6.View ArticleGoogle Scholar
- Malpica R, Franco B, Rodriguez C, Kwon O, Georgellis D. Identification of a quinone-sensitive redox switch in the ArcB sensor kinase. Proc Natl Acad Sci U S A. 2004;101(36):13318–23.View ArticleGoogle Scholar
- Malpica R, Sandoval GR, Rodriguez C, Franco B, Georgellis D. Signaling by the arc two-component system provides a link between the redox state of the quinone pool and gene expression. Antioxid Redox Signal. 2006;8(5-6):781–95.View ArticleGoogle Scholar
- Saini M, Li SY, Wang ZW, Chiang CJ, Chao YP. Systematic engineering of the central metabolism in Escherichia coli for effective production of n-butanol. Biotechnol Biofuels. 2016;9:69.View ArticleGoogle Scholar
- Du Toit PJ, Kotze JP. The isolation and characterization of sorbitol-6-phosphate dehydrogenase from Clostridium pasteurianum. Biochim Biophys Acta. 1970;206(3):333–42.View ArticleGoogle Scholar
- Hacking AJ, Lin EC. Disruption of the fucose pathway as a consequence of genetic adaptation to propanediol as a carbon source in Escherichia coli. J Bacteriol. 1976;126(3):1166–72.Google Scholar
- White D. The physiology and biochemistry of prokaryotes. 3rd ed. New York: Oxford University Press; 2007.Google Scholar
- Sundara Sekar B, Seol E, Mohan Raj S, Park S. Co-production of hydrogen and ethanol by pfkA-deficient Escherichia coli with activated pentose-phosphate pathway: reduction of pyruvate accumulation. Biotechnol Biofuels. 2016;9:95.View ArticleGoogle Scholar
- Sauer U, Canonaco F, Heri S, Perrenoud A, Fischer E. The soluble and membrane-bound transhydrogenases UdhA and PntAB have divergent functions in NADPH metabolism of Escherichia coli. J Biol Chem. 2004;279(8):6613–9.View ArticleGoogle Scholar
- Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30(15):2114–20.View ArticleGoogle Scholar
- Trapnell C, Roberts A, Goff L, Pertea G, Kim D, Kelley DR, et al. Differential gene and transcript expression analysis of RNA-seq experiments with TopHat and Cufflinks. Nat Protoc. 2012;7(3):562–78.View ArticleGoogle Scholar
- NIH DAVID Bioinformatics Resources 6.7. Available online: http://david.abcc.ncifcrf.gov/(accessed on 28 Dec 2014).
- Nishizaki T, Tsuge K, Itaya M, Doi N, Yanagawa H. Metabolic engineering of carotenoid biosynthesis in Escherichia coli by ordered gene assembly in Bacillus subtilis. Appl Environ Microbiol. 2007;73(4):1355–61.View ArticleGoogle Scholar