Metabolic engineering of Clostridium autoethanogenum for ethyl acetate production from CO
Microbial Cell Factories volume 21, Article number: 243 (2022)
Ethyl acetate is a bulk chemical traditionally produced via energy intensive chemical esterification. Microbial production of this compound offers promise as a more sustainable alternative process. So far, efforts have focused on using sugar-based feedstocks for microbial ester production, but extension to one-carbon substrates, such as CO and CO2/H2, is desirable. Acetogens present a promising microbial platform for the production of ethyl esters from these one-carbon substrates.
We engineered the acetogen C. autoethanogenum to produce ethyl acetate from CO by heterologous expression of an alcohol acetyltransferase (AAT), which catalyzes the formation of ethyl acetate from acetyl-CoA and ethanol. Two AATs, Eat1 from Kluyveromyces marxianus and Atf1 from Saccharomyces cerevisiae, were expressed in C. autoethanogenum. Strains expressing Atf1 produced up to 0.2 mM ethyl acetate. Ethyl acetate production was barely detectable (< 0.01 mM) for strains expressing Eat1. Supplementation of ethanol was investigated as potential boost for ethyl acetate production but resulted only in a 1.5-fold increase (0.3 mM ethyl acetate). Besides ethyl acetate, C. autoethanogenum expressing Atf1 could produce 4.5 mM of butyl acetate when 20 mM butanol was supplemented to the growth medium.
This work offers for the first time a proof-of-principle that autotrophic short chain ester production from C1-carbon feedstocks is possible and offers leads on how this approach can be optimized in the future.
Short-chain esters, such as ethyl acetate, are important bulk chemicals used as organic solvents for the production of paints, coatings and resins [1,2,3]. The global ethyl acetate market was valued at $4.6 billion in 2021 and forecasted to grow to $8.3 billion by 2028 . Traditionally, esters are produced by Fischer-Speier esterification of an organic acid and an alcohol . This process relies on equilibrium reactions and demands high temperature and acidic conditions . On the other hand, microbial biosynthesis of ethyl acetate from renewable or waste feedstocks offers a promising sustainable alternative .
Certain microorganisms are naturally capable of producing ethyl acetate, of which yeasts are well studied examples . For instance, production of ethyl acetate by Kluyveromyces marxianus has been shown at pilot-scale with product yields up to around 50% . The biosynthesis pathway for ethyl acetate formation in yeast utilizes alcohol acetyltransferases (AATs), which catalyze the condensation of acetyl-CoA and ethanol to ethyl acetate . Atf1 and Eat1 are the two AATs that have been described to play a major role in the natural ethyl acetate formation [7, 9, 10].
Both Atf1 and Eat1 have been harnessed for metabolic engineering of various microbes to produce esters from sugars. Expression of either the Wickerhamomyces anomalus (Wan) Eat1 or the Kluyveromyces marxianus (Kma) Eat1 in Escherichia coli BW25113 ΔackAΔldhA (DE3) both enabled up to 50% of the theoretical maximum yield of ethyl acetate from glucose . Furthermore, truncation of Eat1 by removal of the mitochondrial pre-sequences resulted in an improved ethyl acetate yield and titer, i.e., 72% (C-mol ethyl acetate/C-mol glucose) and 43 mM, respectively, for the Wan Eat1 . By contrast, expression of Atf1 in E. coli seems to result in the production of low amounts of ethyl acetate. For example, 0.1 mM ethyl acetate was produced with supplementation of 100 mM ethanol by E. coli TOPO expressing Saccharomyces cerevisiae (Sce) Atf1 grown in LB . Furthermore, a yield of 5% was reported for ethyl acetate production from glucose by E. coli BW25113 ΔackAΔldhA (DE3) expressing Sce Atf1 . On the other hand, use of the Sce Atf1 is associated with high-level production of other short-chain esters such as butyl acetate . For example, up to ca. 175 mM butyl acetate was produced from glucose by an engineered Clostridium saccharoperbutylacetonicum, representing 80% of the maximum theoretical yield .
So far only sugar-based microbial production platforms have been explored for short-chain ester production. Utilization of more sustainable (waste) feedstocks and avoiding competition with food supply is therefore desirable. An alternative to sugar fermentation is gas fermentation, which utilizes gaseous carbon oxides (CO and/or CO2). These carbon oxides are abundantly available from industrial off-gases or from gasification of carbonaceous waste material (e.g., municipal solid waste) to syngas (CO, CO2 and H2) [15,16,17]. Anaerobic acetogens, such as Clostridium autoethanogenum, are amongst the best studied autotrophic bacteria for gas fermentation and make use of the Wood-Ljungdahl (or reductive acetyl-CoA) pathway for CO and CO2 (and H2) conversion [16, 18,19,20,21]. In C. autoethanogenum, CO and CO2/H2 conversion results in the formation of acetyl-CoA and subsequently ethanol; both precursors for a potential AAT-catalyzed ethyl acetate biosynthesis pathway (Fig. 1). Here, we report on the engineering of C. autoethanogenum for autotrophic ethyl acetate production from CO by expressing the AATs Kma Eat1 and Sce Atf1. By screening the two AATs in combination with two promoters and several production hosts, Atf1 was found to outperform Eat1 for ethyl acetate production. Additionally, we also established a proof of principle for production of other acetate esters from CO.
Bacterial strains and growth conditions
The bacterial strains used in this study are listed in Table 1. E. coli species were cultured aerobically in LB medium at 37 °C in the presence of the appropriate antibiotics (25 mg/L chloramphenicol with or without 50 mg/L kanamycin). Chemically competent E. coli DH5α (NEB, Ipswich, MA, United States) was used for general cloning, and electrocompetent E. coli sExpress (NEB Express carrying the conjugative plasmid R702) was used as donor for conjugation . C. autoethanogenum JA1-1 (DSM 10061) was purchased from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ) GmbH (Braunschweig, Germany) and cultivated under strict anaerobic conditions.
For all reported growth experiments, C. autoethanogenum was grown in modified ATCC medium 1754 PETC (ATCC, Manassas, VA, USA) at 37 °C. Modified ATCC medium 1754 PETC contained (per liter): 1.0 g NH4Cl, 0.1 g KCL, 0.2 g MgSO4, 0.8 g NaCl, 0.1 g KH2PO4, 0.02 g CaCl2 ⋅ 2H2O, 0.25 g sodium acetate, 1 g yeast extract, 20 g 2-(N-morpholino) ethanesulfonic acid (MES), 0.5 mg resazurin, 0.75 g L-cysteine ⋅ HCl, 40 mM fructose (only for heterotrophic growth), 10 mL trace element solution, and 10 mL Wolfe’s vitamin solution. The composition of the trace element solution (per liter) was 2 g nitrilotriacetic acid, 1 g MnSO4 ⋅ H2O, 0.8 g Fe(SO4)2(NH4)2 ⋅ 6H2O, 0.2 g CoCl2 ⋅ 6H2O, 0.2 mg ZnSO4 ⋅ 7H2O, 0.02 g CuCl2 ⋅ 2H2O, 0.02 g NiCl2 ⋅ 6H2O, 0.02 g NaMoO4 ⋅ 2H2O, 0.02 g Na2SeO4, 0.02 g Na2WO4 ⋅ 2H2O. The vitamin solution composition (per liter) was 2 mg biotin, 2 mg folic acid, 10 mg pyridoxine hydrochloride, 5 mg thiamine ⋅ HCl, 5 mg riboflavin, 5 mg nicotinic acid, 5 mg calcium pantothenate, 0.1 mg vitamin B12, 5 mg p-aminobenzoic acid, and 5 mg thioctic acid. Antibiotics were supplemented when required (6 mg/L or 15 mg/L thiamphenicol for autotrophic or heterotrophic growth, respectively).
For routine cultivation or growth on solid medium of C. autoethanogenum YTF medium was used that contained (per liter): 10 g yeast extract, 16 g tryptone, 10 g fructose, 0.2 g sodium chloride, 0.75 g L-cysteine ⋅ HCl, 0.5 mg resazurin, 10 mL ATCC medium 1754 trace element solution, 10 mL Wolfe’s vitamin solution, and 15 g agar (only for solid medium). When required, medium was supplemented with antibiotics (15 mg/L thiamphenicol and/or 250 mg/L D-cycloserine).
The growth experiments with C. autoethanogenum strains were carried out in 250 mL serum bottles containing 50 mL of modified ATCC medium 1754 PETC medium. The serum bottles contained either a gas phase of 170 kPa CO, or 40 mM fructose with a gas phase of 150 kPa N2. Strains were cultured in YTF before being transferred at approximately 5% (v/v) to precultures in ATCC 1754 PETC medium with CO or fructose as main carbon source. Precultures were transferred at least twice in ATCC 1754 PETC medium before used for growth experiments. For precultures with CO as carbon source, strains were incubated stationary until a decrease in total headspace pressure was measured, after which cultures were transferred to incubation at 150 rpm. Growth experiments were inoculated from these precultures at approximately 5% (v/v) and shaken at 150 rpm. For growth experiments involving supplementation of alcohols (ethanol, butanol, or hexanol) or butyl acetate, C. autoethanogenum strains were incubated one day until growth was observed prior to addition of the alcohols or butyl acetate. All growth experiments were carried out in biological triplicates. Sampling of cultures for analysis of final metabolite production was generally performed after depletion of substrate and/or growth arrest.
All mutagenesis work was performed in a Coy anaerobic chamber (Coy Laboratory Products, Inc., Ann.
Arbor, MI) with an atmosphere of approximately 1% H2 and 99% N2. Agar plates with C. autoethanogenum were incubated outside the anaerobic chamber at 37 °C in Mitsubishi™ AnaeroPack™ rectangular jars (Thermo Fisher, Waltham, MA) that contained an AnaeroGen™ sachet (Thermo Fisher). C. autoethanogenum freezer stocks were prepared by combining an equal volume of culture and 50% (v/v) glycerol in anaerobic vials reduced with a two to three drops of 100 mM titanium(III) citrate, and were stored at − 80 °C.
Genomic DNA from C. autoethanogenum was isolated with the MasterPure™ Gram Positive DNA Purification Kit (Epicenter, Madison, WI) according to manufacturer’s protocol. PCR was carried out using either Q5® High-Fidelity DNA Polymerase (New England Biolabs, Ipswich, MA) for cloning purposes or OneTaq® DNA Polymerase (NEB) for diagnostic purposes according to the manufacturers’ protocols.
DNA fragments were isolated and purified from agarose gel using the Zymoclean™ Gel DNA Recovery Kit (Zymo Research Corporation) after gel electrophoresis or from PCR reaction mixture using the Thermo Scientific™ GeneJET PCR Purification Kit (Thermo Fisher), according to the manufacturer’s protocols. Plasmid DNA was isolated using the Thermo Scientific™ GeneJET Plasmid Miniprep Kit (Thermo Fisher, Waltham, MA) according to manufacturer’s protocol.
Plasmids were assembled either by using NEBuilder® HiFi DNA Assembly Master Mix (NEB), for which the protocol was modified by downscaling the total reaction mixture volume to 5 μL, or by using standard restriction ligation cloning techniques for which necessary restriction and ligation enzymes were purchased from NEB and used according to the manufacturer’s protocols.
All primers used in this study are listed in Table S1 (see Additional File 1). Primers were synthesized by IDT (Integrated DNA Technologies, Coralville, IA) or Sigma-Aldrich (Saint Louis, MO). Sanger sequencing of plasmids and amplicons was carried out by Macrogen (Macrogen Europe, Amsterdam, Netherlands).
All plasmids used in this study are listed in Table 1. All plasmids are based on the E. coli-Clostridium shuttle vectors pMTL83151 or pMTL83122  with pCB102, catP, traJ and ColE1 or p15a, respectively. The genes eat1 (KMAR_10772, Kluyveromyces marxianus KM1777_03) and atf1 (Saccharomyces cerevisiae CEN.PK113-7D) were codon harmonized with the algorithm described previously  for expression in C. autoethanogenum DSM 10061. The codon harmonized genes were synthesized by Twist Bioscience (San Francisco, CA). Sequences of codon harmonized genes eat1 and atf1 are available in Table S2 (see Additional File 1).
For construction of the AAT expression plasmids, the genes eat1 and atf1 and the truncated version of eat1 were cloned into the multiple cloning site (MCS) of either pMTL83151 together with promoter PPta-AckA  or pMTL83122 downstream of PThl. The aforementioned plasmids were linearized prior to cloning of the AAT genes via PCR amplification. The promoter PPta-ackA was amplified from C. autoethanogenum genomic DNA.
The vector pMTL-SIBR-Cas-pta for targeting of the pta (CAETHG_3358) gene was assembled as specified before  and described in more detail below. Plasmids maps for the plasmids used in this study can be found in the Additional File 2.
Conjugation of C. autoethanogenum
Plasmids were transferred into the conjugative donor E. coli strain sExpress and subsequently conjugated into C. autoethanogenum according to previously established methods (Woods et al., ). Thiamphenicol (15 mg/L) was used to select for the catP-based plasmids. D-cycloserine (250 mg/L) was used to counter select against E. coli sExpress after conjugation.
Generation of C. autoethanogenum knockout strains
The Ald domain of adhE1 (CAETHG_3747), designated adhE1a as described before , was targeted using CRISPR Cas9/RiboCas gene editing techniques based on previously established methods . The homology arms were composed of ca. 500 bp and 1 kb, upstream and downstream of the gene of interest, respectively. The synthetic guide sequence was designed using the software Benchling (https://www.benchling.com/crispr/). Assembly of the targeting vector pRECas1-adhE1a was achieved through traditional restriction ligation cloning methods of the sgRNA module and editing template, as described before .
The pta gene (CAETHG_3358) was targeted using the recently described SIBR-Cas gene editing technique . The targeting vector was based on the backbone pMTL83151 with expression of the FnCas12a variant with Int3  under control of the fdx promoter from Clostridium sporogenes , a crRNA under control of the araE promoter from Clostridium acetobutylicum  and homology arms of ca. 1 kb up- and downstream of pta. To insert the homology arms, the vector backbone was first digested with restriction enzymes KpnI and NcoI. The synthetic guide sequence for the crRNA was designed using the software Benchling.
The vectors pRECas-adhE1a and pMTL-SIBR-Cas-pta were sequence verified using Sanger sequencing before and after transformation to the conjugative E. coli donor strain sExpress, and subsequently conjugated into C. autoethanogenum as described previously . Resultant colonies resistant to thiamphenicol and D-cycloserine were inoculated into liquid medium with thiamphenicol, and subsequently subcultured into liquid medium with thiamphenicol and 5 mM theophylline. After growth was observed, single colonies were isolated on solid media and were screened by PCR for the desired in-frame deletion in the pta or adhE1 locus. The PCR product was sequenced to confirm the nature of the deletion (see Additional file 1, Figure S1, 2). Plasmid loss was achieved through serial passage in non-selective medium and confirmed through replica plating on medium with or without thiamphenicol as well as lack of amplification of the CRISPR-Cas plasmid in C. autoethanogenum knockout strain.
Cell growth in liquid medium was monitored spectrophotometrically at 600 nm (OD600) using the spectrophotometer UV-1800 (Shimadzu, Tokyo, Japan). Headspace pressure was measured with a Greisinger GMH 3151 pressure reader (GSG Geologie-Service GmbH, Würzburg, Germany).
For analysis of metabolites, samples from the cultures were spun down by centrifugation (21,000 × g for 3 min, room temperature) after which the supernatant was stored at − 20 °C until analysis or analyzed immediately. Fructose, acetate, and ethanol were analyzed by HPLC with a LC-2030C Plus (Shimadzu) equipped with a Shodex SH1821 column operated at a temperature of 45 °C. The eluent used was 0.01 N H2SO4 at a flow rate of 1 ml/min. Peaks were detected with a refractive index detector. Relevant standards were used for identification and quantification. Ethyl acetate, butyl acetate, and hexyl acetate were analyzed by gas chromatography on a GC-2010 equipped with headspace sampler HS-20 (Shimadzu) and flame ionization detector. Supernatant (0.1 mL) from centrifuged culture samples (21,000 × g for 3 min, room temperature) was mixed with 0.1 mL internal standard (10 mM 1-propanol) in gas-tight glass vials. The vials were heated at 60 °C for 5 min and pressurized (100 kPa) with N2. The vial headspace was injected on an Agilent DB-WAX Ultra Inert column (30 m, 0.53 mm, 1.0 μm) with a split ration of 1 to 20. The column was operated at a constant pressure (30 kPa) with N2 as carrier gas and with a temperature gradient. The starting temperature was 70 °C which was held for 1 min followed by a temperature ramp of 50 °C/min to 125 °C and finally by a temperature ramp of 70 °C/min to 230 °C at which the temperature was kept until a total run time of 6 min was reached. Ethyl acetate was also measured by GC–MS with the GC Trace 1300 coupled with an EI Mass spectrometer ISQ 7000 and TriPlus RSH (Thermo Fisher). The GC–MS was equipped with an Agilent Porabond Q column (25 m, 0.25 mm, 3 μm) and operated with a flow of 1 ml/min of helium as carrier gas and a split of 50. Liquid samples of 0.2 mL were prepared in gas-tight glass vials, which were heated at 50 °C for 5 min after which 0.5 mL of the headspace was injected into the GC–MS. The column temperature was set to 100 °C for one min, followed by a ramp of 25 °C/min to 200 °C at which the temperature was kept for 3 min. Relevant standards were used for identification and quantification.
CO was analyzed by gas chromatography by injecting 0.2 mL of culture headspace into a CompactGC4.0 (Global Analyzer Solutions, The Netherlands). The CompactGC4.0 was equipped with a Carboxen 110 column (3 m, 0.32 mm) followed by a Molsieve 5A column, (30 m, 0.32 mm). The column oven temperature was set to 140 °C. The GC was operated with argon as carrier gas at a flowrate of 5 mL/min. A standard with a known amount of CO was used for identification and quantification of CO in culture samples.
Heterologous expression of an AAT enables ethyl acetate production from CO
To investigate the potential of producing ethyl acetate from CO by acetogenic bacteria, we initially screened different combinations of AATs, promoters, and expression hosts. The genes encoding the two AATs, Kma Eat1 and Sce Atf1, were each expressed from two different promoters: the C. autoethanogenum Pta-AckA promoter (PPta-AckA)  and the C. acetobutylicum thiolase promoter (Pthl) . Besides the wild type Eat1, the truncated version of Eat1  was included as well. The availability of the acetyl-CoA and ethanol precursors may also be important for the AAT-catalyzed formation of ethyl acetate. For instance, ethanol supplementation has shown to improve ethyl acetate production using Sce Atf1 . Therefore, next to wild-type C. autoethanogenum, we also included C. autoethanogenum strains in which knockouts were made of either the phosphotransacetylase (pta) or the aldehyde dehydrogenase (Ald) subunit of the adhE1 gene. Previously, disruption of the Ald subunit of adhE1 has already been shown to enhance ethanol production from CO by nearly threefold . Similarly, knockout of pta may also influence acetyl-CoA or ethanol metabolism (Fig. 1), and therefore ethyl acetate formation. The different C. autoethanogenum strains harboring the resulting plasmids for AAT expression or an empty plasmid control (pMT83151) were cultivated under autotrophic conditions with CO as main carbon source. The final ethyl acetate concentration was the highest for C. autoethanogenum [PThl-Atf1] at approximately 0.2 mM ethyl acetate (Fig. 2). This corresponded to a final ethyl acetate yield of ca. 0.7% of the theoretical maximum on CO (see Additional file 1: Figure S3). Ethyl acetate formation was approximately fourfold lower for C. autoethanogenum [PPta-AckA-Atf1] using the PPta-AckA rather than the PThl promoter for Atf1 expression. Only trace presence of ethyl acetate (< 0.01 mM) was detected for the C. autoethanogenum strains expressing Eat1, regardless of the promoter or Eat1 type (full length or truncated) used. No improvement in ethyl acetate production was observed for the Δpta or ΔadhE1a strains, in fact, deletion of the Ald subunit of adhE1 resulted in an unexpected decrease in ethyl acetate formation. Knockout of pta or adhE1a did not cause a substantial effect on growth or product spectrum under these conditions compared to wild-type C. autoethanogenum (data not shown). Screening of C. autoethanogenum strains harboring the Eat1 and Atf1 expression plasmids under heterotrophic conditions with 40 mM fructose as main carbon source showed a similar outcome to autotrophic growth; only Atf1-expressing strains produced ethyl acetate at low final concentrations (0.1 mM) (see Additional file 1: Figure S4). Furthermore, we also examined the potential detrimental ability of the production host to naturally degrade ethyl acetate esters by incubating ethyl acetate with and without C. autoethanogenum, but no ethyl acetate degradation was observed (see Additional file 1: Figure S5-6).
Next, we investigated the lack of ethyl acetate production by Eat1-expressing C. autoethanogenum strains by assaying these strains for Eat1 activity. Because measurement of Eat1 AAT activity in cell-free extracts has been proven to be difficult , Eat1 activity was assayed using in vivo alcoholysis, a secondary activity of Eat1 in which the alcohol moiety of an ester is replaced by another alcohol . Together with exogenously supplied butyl acetate and endogenously produced ethanol, Eat1-catalyzed alcoholysis can be easily measured via ethyl acetate production. Varying concentrations of butyl acetate were added to cultures of C. autoethanogenum strains harboring plasmids PPta-AckA-Eat1, PThl-Eat1 or empty plasmid pMTL83151. However, neither of the Eat1-expressing C. autoethanogenum strains showed alcoholysis activity (see Additional file 1: Figure S7), suggesting that Eat1 may not be functionally expressed in C. autoethanogenum.
As another control, varying concentrations of butyl acetate were also added to cultures of E. coli strains harboring the same Eat1 expression plasmids as the C. autoethanogenum strains. In contrast to what was observed for C. autoethanogenum, E. coli strains showed Eat1-dependent alcoholysis activity in presence of butyl acetate (see Additional file 1: Figure S7). The Eat1-expressing E. coli strains also produced ethyl acetate in absence of butyl acetate, confirming Eat1-dependent AAT activity from the same plasmids used for the expression of Eat1 in C. autoethanogenum.
Ethanol supplementation boosts ethyl acetate production
To gain further insights into autotrophic ethyl acetate production, potential metabolic bottlenecks in the currently engineered C. autoethanogenum strains were investigated. First, the best performing strain C. autoethanogenum [PThl-Atf1] was characterized in more detail. For both C. autoethanogenum [pMTL83151] and C. autoethanogenum [PThl-Atf1] the CO fermentation pattern was characterized by initial formation of ethanol, followed by acetate formation and ethanol consumption (Fig. 3a). Ethyl acetate did not show degradation over time. Because ethyl acetate production appeared to coincide with ethanol formation during part of the fermentation, ethanol was investigated as possible limiting precursor for ethyl acetate formation. Ethanol was supplemented to cultures of C. autoethanogenum [PThl-Atf1] to final concentrations of 25 – 200 mM; these cultures were incubated with CO in the headspace for 7 days. Compared to the control (0 mM ethanol), cultures with 50 mM ethanol supplementation showed 1.5-fold increase in ethyl acetate production to approximately 0.35 mM (Fig. 3b). Higher ethanol concentrations were detrimental to growth and final ethyl acetate titers but appeared to enhance biomass-specific ethyl acetate production (see Additional file 1: Figure S8-9).
Alcohol doping shows potential of Atf1 for ester production from CO
Besides formation of ethyl acetate, Atf1 is known to have broad alcohol specificity and is able to catalyze formation of other acetate esters, including butyl acetate and hexyl acetate [13, 14, 32, 33]. To investigate whether an acetogen expressing Atf1 also has potential for biosynthesis of other acetate esters from CO, C. autoethanogenum [PThl-Atf1] was grown on CO supplemented with either butanol or hexanol and was compared to a reference condition with and without ethanol supplementation (Fig. 4a). Strikingly, up to 4.5 mM of butyl acetate was formed in the presence of butanol. Only trace amounts (< 0.05 mM) of hexyl acetate could be detected in the presence of hexanol (Fig. 4b).
Compared to supplementation with ethanol, tenfold more total ester production was achieved with butanol supplementation. Although ethyl acetate production with and without the presence of butanol remained similar, 20-fold more butyl acetate than ethyl acetate was produced in the presence of butanol, representing a 95% selectivity for butyl acetate production.
We describe a successful proof-of-concept of engineering acetogens to produce ethyl acetate from CO as main carbon source. However, more efforts are needed before the envisioned autotrophic ethyl acetate production can compete with the conventional chemical catalysis or sugar-based microbial ethyl acetate production. Out of the two AATs tested, only expression of the Sce Atf1 resulted in ethyl acetate formation in C. autoethanogenum, regardless of the carbon source, background hosts, and promoters tested. The trace amounts of ethyl acetate observed for the empty plasmid control and Eat1-expression strains (but not for wild-type C. autoethanogenum) likely originated from the chloramphenicol acetyltransferase (CatP) encoded on the plasmids as antibiotic resistance marker . The higher ethyl acetate production with the promoter PThl compared to PPta-AckA suggests that the PThl is stronger or more suitable for Atf1 expression on both CO and fructose. This may be specific to the microorganism and genes expressed. For instance, a study on autotrophic 2-propanol production from syngas in Clostridium ljungdahlii comparing the native C. ljungdahlii PPta-AckA and the PThl promoters showed that only the PPta-AckA enabled isopropanol production . On the other hand, acetone production levels were similar comparing the PThl and the C. ljungdahlii PPta-AckA for pathway expression in Acetobacterium woodii grown on H2/CO2 .
Eat1 was a promising catalyst for ethyl acetate production based on previous work in E. coli . However, Eat1 did not facilitate ethyl acetate production in C. autoethanogenum and was likely not functionally expressed based on lack of AAT and alcoholysis activity. Differences in AAT activity between Eat1 and Atf1 in C. autoethanogenum could be a result of differences in expression, enzyme characteristics or other factors. For instance, Atf1 was successfully expressed in C. autoethanogenum using the same promoters as for Eat1. This indicates that Eat1 might require other expression conditions than Atf1 for functional activity in C. autoethanogenum. Choice of inducible promoter and level of induction have previously been shown to strongly influence Eat1-catalyzed ethyl acetate production in E. coli, with Eat1 activity correlating positively with promoter strength . Achieving functional expression of Eat1 in C. autoethanogenum might therefore benefit from a more extensive promoter library testing. Concerning enzyme kinetics, the affinity of purified Wan Eat1 for acetyl-CoA is relatively low (apparent Km = 2.43 mM) compared to that of purified Sce Atf (apparent Km = 45 μM) [10, 37]. This might indicate that Eat1 requires higher intracellular acetyl-CoA concentrations than Atf1. However, concentrations of intracellular metabolites, including acetyl-CoA, have been described to be similar between E. coli and C. autoethanogenum , and Eat1 can successfully catalyze ethyl acetate formation in E. coli . Metabolic pathway optimization in E. coli lysates has also been shown to be representative for C. autoethanogenum [39, 40]. Yet there still may be physiological differences that affect activity of certain enzymes. For instance, the intracellular pH of the yeast mitochondria, where Eat1 is localized , is tightly regulated near pH 7.5 . Atf1 appears to be located in the endoplasmic reticulum or cytosol [43,44,45], which have a pH near 7.2 . Correspondingly, the intracellular pH of E. coli is near pH 7.6 . However, the intracellular pH of clostridia like C. autoethanogenum and Clostridium acetobutylicum is significantly lower, near pH 6 [47,48,49]. Purified Sce Atf1 has a pH optimum near pH 7 but retains high activity at pH 6 . For Eat1, the effect of pH on enzyme activity has not yet been investigated but it might be less suited for the acidic intracellular environment of clostridia. A previous study expressing the Wan Eat1 in another clostridial host, Clostridium beijerinckii, using the strong constitutive C. beijerinkcii Pthl also showed low AAT activity . Curiously, C. beijerinckii expressing Wan Eat1 still showed high alcoholysis activity, in contrast to C. autoethanogenum expressing Kma Eat1 in this study. Host-specific functionality is further supported by the fact that Kma Eat1 was successfully expressed in E. coli using the same expression plasmids as for C. autoethanogenum in this study. However, host-specific expression elements, such as host-specific origins of replication, may contribute to this observation. Nevertheless, this also confirms previous findings that the promoters used are also active in E. coli [27, 51]. Overall, these results indicate that expression of Eat1 in Clostridia is troublesome and requires further investigation.
For Sce Atf1, only low final ethyl acetate titers were formed, despite ethanol supplementation, in line with previous observations [11, 12]. The inefficient ethyl acetate formation by Sce Atf1 might be due to low ethanol specificity, or other factors, such as suboptimal expression of Atf1. The first option is supported by our observations that, using the same Atf1 expression plasmid, nearly 20-fold more acetate esters could be produced with 20 mM butanol addition compared to a similar amount of ethanol addition. Therefore, this study indicates that the potential of Sce Atf1 may be limited for ethyl acetate production in acetogens under these conditions.
On the other hand, these experiments indicate that Atf1 might be an effective AAT to produce other acetate esters with acetogens from C1 substrates, such as butyl acetate or hexyl acetate. Surprisingly, the amount of butyl acetate produced in this study (4.5 mM) indicated that approximately 10% of the acetyl-CoA formed out of CO could be readily directed to this non-native product without further optimization. Hexyl acetate was only produced in trace amounts, comparable to earlier findings in E. coli expressing Sce Atf1 . Besides butyl acetate and hexyl acetate, Atf1 is known to be able to catalyze formation of many other acetate esters from different alcohols such as isobutyl acetate or propyl acetate . Research on acetogens has already focused on autotrophic production of a variety of alcohols such as butanol, hexanol, isobutanol, and isopropanol [39, 40, 52,53,54,55,56,57,58,59]. For efficient autotrophic production of non-native products using acetogens, the limited energy availability during autotrophic growth needs to be taken into account [60,61,62,63]. Importantly, production of acetate esters is ATP positive in acetogens on both CO and H2/CO2 assuming the bioenergetics for acetogens like C. autoethanogenum [49, 63]. Besides several alcohols, certain acetogens like Clostridium carboxidivorans can natively also produce longer-chain acyl-CoAs, such as butyryl-CoA, which expand the possibilities of autotrophic ester biosynthesis even further.
We have shown that production of ethyl acetate from the gaseous C1 substrate CO can be successfully achieved by heterologous expression of an AAT in C. autoethanogenum. The highest ethyl acetate production was achieved using the Sce Atf1, which reached 0.2 mM ethyl acetate on CO, or approximately 0.7% of the theoretical yield. C. autoethanogenum expressing Atf1 was also able to produce other acetate esters from CO, including butyl acetate and hexyl acetate, when supplied with their respective alcohols. Although there is ample room for improvement, we provide a foundation for autotrophic production of acetate esters from one-carbon substrates.
Availability of data and materials
The datasets supporting the conclusions of this article are included within the article and its additional file.
Löser C, Urit T, Bley T. Perspectives for the biotechnological production of ethyl acetate by yeasts. Appl Microbiol Biotechnol. 2014;98:5397–415.
Kam S-K, Kim J-K, Lee M-G. Removal characteristics of mixed gas of ethyl acetate and 2-butanol by a biofilter packed with Jeju scoria. Korean J Chem Eng. 2011;28:1019–22.
Chan W-C, Su M-Q. Biofiltration of ethyl acetate and amyl acetate using a composite bead biofilter. Bioresour Technol. 2008;99:8016–21.
GVR. Ethyl Acetate Market Size, Share & Trends Analysis Report By End Use (Food & Beverages, Pharmaceutical, Automotive, Artificial Leather, Packaging), By Region, And Segment Forecasts, 2021–2028. 2021.
Fischer E, Speier A. Darstellung der Ester. Berichte der Dtsch Chem Gesellschaft. 1895;28:3252–8.
Liu Y, Lotero E, Goodwin JG. Effect of water on sulfuric acid catalyzed esterification. J Mol Catal A Chem. 2006;245:132–40.
Kruis AJ, Bohnenkamp AC, Patinios C, van Nuland YM, Levisson M, Mars AE, et al. Microbial production of short and medium chain esters: enzymes, pathways, and applications. Biotechnol Adv. 2019;37:107407.
Löser C, Urit T, Stukert A, Bley T. Formation of ethyl acetate from whey by Kluyveromyces marxianus on a pilot scale. J Biotechnol. 2013;163:17–23.
Minetoki T, Bogaki T, Iwamatsu A, Fujii T, Hamachi M. The purification, properties and internal peptide sequences of alcohol acetyltransferase isolated from Saccharomyces cerevisiae Kyokai No 7. Biosci Biotechnol Biochem. 1993;57:2094–8.
Kruis AJ, Levisson M, Mars AE, van der Ploeg M, Garcés Daza F, Ellena V, et al. Ethyl acetate production by the elusive alcohol acetyltransferase from yeast. Metab Eng. 2017;41:92–101.
Bohnenkamp AC, Kruis AJ, Mars AE, Wijffels RH, van der Oost J, Kengen SWM, et al. Multilevel optimisation of anaerobic ethyl acetate production in engineered Escherichia coli. Biotechnol Biofuels. 2020;13:65.
Horton CE, Bennett GN. Ester production in E. coli and C. acetobutylicum. Enzyme Microb Technol. 2006;38:937–43.
Wang Q, Al Makishah NH, Li Q, Li Y, Liu W, Sun X-M, et al. Developing clostridia as cell factories for short- and medium-chain ester production. Front Bioeng Biotechnol. 2021;9:386.
Feng J, Zhang J, Ma Y, Feng Y, Wang SS, Guo N, et al. Renewable fatty acid ester production in Clostridium. Nat Commun. 2021;12:1–13.
Redl S, Diender M, Jensen TØ, Sousa DZ, Nielsen AT. Exploiting the potential of gas fermentation. Ind Crops Prod. 2017;106:21–30.
Fackler N, Heijstra BD, Rasor BJ, Brown H, Martin J, Ni Z, et al. Stepping on the gas to a circular economy: accelerating development of carbon-negative chemical production from gas fermentation. Annu Rev Chem Biomol Eng. 2021;12:439–70.
Köpke M, Simpson SD. Pollution to products: recycling of ‘above ground’ carbon by gas fermentation. Curr Opin Biotechnol. 2020;65:180–9.
Ragsdale SW, Pierce E. Acetogenesis and the Wood-Ljungdahl pathway of CO2 fixation. Biochim Biophys Acta - Proteins Proteomics. 2008;1784:1873–98.
Wood HG. Life with CO or CO2 and H2 as a source of carbon and energy. FASEB J. 1991;5:156–63.
Diender M, Stams AJM, Sousa DZ. Pathways and bioenergetics of anaerobic carbon monoxide fermentation. Front Microbiol. 2015;6:1–18.
Pavan M, Reinmets K, Garg S, Mueller AP, Marcellin E, Köpke M, et al. Advances in systems metabolic engineering of autotrophic carbon oxide-fixing biocatalysts towards a circular economy. Metab Eng. 2022;71:117–41.
Woods C, Humphreys CM, Rodrigues RM, Ingle P, Rowe P, Henstra AM, et al. A novel conjugal donor strain for improved DNA transfer into Clostridium spp. Anaerobe. 2019;59:184–91.
Heap JT, Pennington OJ, Cartman ST, Minton NP. A modular system for Clostridium shuttle plasmids. J Microbiol Methods. 2009;78:79–85.
Cañadas IC, Groothuis D, Zygouropoulou M, Rodrigues R, Minton NP. RiboCas: a universal CRISPR-based editing tool for Clostridium. ACS Synth Biol. 2019. https://doi.org/10.1021/acssynbio.9b00075.
Patinios C, Creutzburg SCA, Arifah AQ, Adiego-Pérez B, Gyimah EA, Ingham CJ, et al. Streamlined CRISPR genome engineering in wild-type bacteria using SIBR-Cas. Nucleic Acids Res. 2021;49:11392–404.
Claassens NJ, Siliakus MF, Spaans SK, Creutzburg SCA, Nijsse B, Schaap PJ, et al. Improving heterologous membrane protein production in Escherichia coli by combining transcriptional tuning and codon usage algorithms. PLoS One. 2017;12:e0184355.
Köpke M, Gerth ML, Maddock DJ, Mueller AP, Liew F, Simpson SD, et al. Reconstruction of an acetogenic 2,3-butanediol pathway involving a novel NADPH-dependent primary-secondary alcohol dehydrogenase. Appl Environ Microbiol. 2014;80:3394–403.
Liew F, Henstra AM, Kӧpke M, Winzer K, Simpson SD, Minton NP. Metabolic engineering of Clostridium autoethanogenum for selective alcohol production. Metab Eng. 2017;40:104–14.
Huang H, Chai C, Li N, Rowe P, Minton NP, Yang S, et al. CRISPR/Cas9-based efficient genome editing in Clostridium ljungdahlii, an autotrophic gas-fermenting bacterium. ACS Synth Biol. 2016;5:1355–61.
Kruis AJ, Bohnenkamp AC, Nap B, Nielsen J, Mars AE, Wijffels RH, et al. From eat to treat: engineering the mitochondrial eat1 enzyme for enhanced ethyl acetate production in Escherichia coli. Biotechnol Biofuels BioMed Central. 2020;13:1–10.
Patinios C, Lanza L, Corino I, Franssen MCR, Van der Oost J, Weusthuis RA, et al. Eat1-like alcohol acyl transferases from yeasts have high alcoholysis and thiolysis activity. Front Microbiol. 2020;11:1.
Rodriguez GM, Tashiro Y, Atsumi S. Expanding ester biosynthesis in Escherichia coli. Nat Chem Biol. 2014;10:259–65.
Nancolas B, Bull ID, Stenner R, Dufour V, Curnow P. Saccharomyces cerevisiae Atf1p is an alcohol acetyltransferase and a thioesterase in vitro. Yeast. 2017;34:239–51.
Seo H, Lee JW, Giannone RJ, Dunlap NJ, Trinh CT. Engineering promiscuity of chloramphenicol acetyltransferase for microbial designer ester biosynthesis. Metab Eng. 2021;66:179–90.
Bengelsdorf FR, Poehlein A, Linder S, Erz C, Hummel T, Hoffmeister S, et al. Industrial acetogenic biocatalysts: a comparative metabolic and genomic analysis. Front Microbiol. 2016;7:1–15.
Hoffmeister S, Gerdom M, Bengelsdorf FR, Linder S, Flüchter S, Öztürk H, et al. Acetone production with metabolically engineered strains of Acetobacterium woodii. Metab Eng Elsevier. 2016;36:37–47.
Malcrops P, Dufour J-P. Short-chain and medium-chain aliphatic-ester synthesis in Saccharomyces cerevisiae. Eur J Biochem. 1992;210:1015–22.
Valgepea K, de Souza PLR, Meaghan K, Palfreyman RW, Abdalla T, Heijstra BD, et al. Maintenance of ATP homeostasis triggers metabolic shifts in gas-fermenting acetogens. Cell Syst. 2017;4:505-515.e5.
Karim AS, Dudley QM, Juminaga A, Yuan Y, Crowe SA, Heggestad JT, et al. In vitro prototyping and rapid optimization of biosynthetic enzymes for cell design. Nat Chem Biol. 2020;16:912–9.
Vögeli B, Schulz L, Garg S, Tarasava K, Clomburg JM, Lee SH, et al. Cell-free prototyping enables implementation of optimized reverse β-oxidation pathways in heterotrophic and autotrophic bacteria. Nat Commun. 2022;13:3058.
Kruis AJ, Mars AE, Kengen SWM, Borst JW, van der Oost J, Weusthuis RA. Alcohol Acetyltransferase Eat1 Is Located in Yeast Mitochondria. Appl Environ Microbiol. 2018;84:1–11.
Orij R, Postmus J, Ter Beek A, Brul S, Smits GJ. In vivo measurement of cytosolic and mitochondrial pH using a pH-sensitive GFP derivative in Saccharomyces cerevisiae reveals a relation between intracellular pH and growth. Microbiology. 2009;155:268–78.
Verstrepen KJ, Van Laere SDM, Vercammen J, Derdelinckx G, Dufour J-P, Pretorius IS, et al. The Saccharomyces cerevisiae alcohol acetyl transferase Atf1p is localized in lipid particles. Yeast. 2004;21:367–77.
Lin J-L, Wheeldon I. Dual N- and C-terminal helices are required for endoplasmic reticulum and lipid droplet association of alcohol acetyltransferases in Saccharomyces cerevisiae. PLoS ONE. 2014;9:e104141.
Zhu J, Lin J-L, Palomec L, Wheeldon I. Microbial host selection affects intracellular localization and activity of alcohol-O-acetyltransferase. Microb Cell Fact. 2015;14:35.
Slonczewski JL, Rosen BP, Alger JR, Macnab RM. pH homeostasis in Escherichia coli: measurement by 31P nuclear magnetic resonance of methylphosphonate and phosphate. Proc Natl Acad Sci. 1981;78:6271–5.
Huang L, Forsberg CW, Gibbins LN. Influence of external pH and fermentation products on Clostridium acetobutylicum Intracellular pH and cellular distribution of fermentation products. Appl Environ Microbiol. 1986;51:1230–4.
Terracciano JS, Kashket ER. Intracellular conditions required for initiation of solvent production by Clostridium acetobutylicum. Appl Environ Microbiol. 1986;52:86–91.
Mock J, Zheng Y, Mueller AP, Ly S, Tran L, Segovia S, et al. Energy conservation associated with ethanol formation from H2 and CO2 in Clostridium autoethanogenum involving electron bifurcation. J Bacteriol. 2015;197:2965–80.
Akita O, Suzuki S, Obata T, Hara S. Purification and some properties of alcohol acetyltransferase from sake Yeast. Agric Biol Chem. 1990;54:1485–90.
Mordaka PM, Heap JT. Stringency of synthetic promoter sequences in clostridium revealed and circumvented by tuning promoter library mutation rates. ACS Synth Biol. 2018;7:672–81.
Köpke M, Held C, Hujer S, Liesegang H, Wiezer A, Wollherr A, et al. Clostridium ljungdahlii represents a microbial production platform based on syngas. Proc Natl Acad Sci. 2010;107:13087–92.
Banerjee A, Leang C, Ueki T, Nevin KP, Lovley DR. Lactose-inducible system for metabolic engineering of Clostridium ljungdahlii. Appl Environ Microbiol. 2014;80:2410–6.
Dürre P. Butanol formation from gaseous substrates. FEMS Microbiol Lett. 2016;363:fnw040.
Liou JSC, Balkwill DL, Drake GR, Tanner RS. Clostridium carboxidivorans sp. nov., a solvent-producing clostridium isolated from an agricultural settling lagoon, and reclassification of the acetogen Clostridium scatologenes strain SL1 as Clostridium drakei sp. nov. Int J Syst Evol Microbiol. 2005;55:2085–91.
Lauer I, Philipps G, Jennewein S. Metabolic engineering of Clostridium ljungdahlii for the production of hexanol and butanol from CO2 and H2. Microb Cell Fact. 2022;21:85.
Weitz S, Hermann M, Linder S, Bengelsdorf FR, Takors R, Dürre P. Isobutanol production by autotrophic acetogenic bacteria. Front Bioeng Biotechnol. 2021;9:1–12.
Jia D, He M, Tian Y, Shen S, Zhu X, Wang Y, et al. Metabolic engineering of gas-fermenting Clostridium ljungdahlii for efficient co-production of isopropanol, 3-hydroxybutyrate, and ethanol. ACS Synth Biol. 2021;10:2628–38.
Liew FE, Nogle R, Abdalla T, Rasor BJ, Canter C, Jensen RO, et al. Carbon-negative production of acetone and isopropanol by gas fermentation at industrial pilot scale. Nat Biotechnol. 2022;40:335–44.
Schuchmann K, Müller V. Autotrophy at the thermodynamic limit of life: a model for energy conservation in acetogenic bacteria. Nat Rev Microbiol. 2014;12:809–21.
Bertsch J, Müller V. Bioenergetic constraints for conversion of syngas to biofuels in acetogenic bacteria. Biotechnol biofuels. BioMed Central. 2015;8:210.
Molitor B, Marcellin E, Angenent LT. Overcoming the energetic limitations of syngas fermentation. Curr Opin Chem Biol. 2017;41:84–92.
Katsyv A, Müller V. Overcoming energetic barriers in acetogenic C1 conversion. Front Bioeng Biotechnol. 2020;8:1–23.
We thank Nigel Minton (University of Nottingham, United Kingdom) for kindly providing the RiboCas pRECas1 plasmid. Also, we would like to thank Ton van Gelder, Iame Alves Guedes and Rob Joosten (Wageningen University and Research) for their technical support.
This work is part of the research program MicroSynC with project number P16-10/6 financed by the NWO Domain Applied and Engineering Sciences (AES) of the Netherlands Organization for Scientific Research (NWO).
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List of primers used in this study. Table S2. Sequences of genes expressed in this study. Figure S1. Image of Sanger sequencing results showed the nature of the in-frame deletion in the pta gene (CAETHG_3358). Figure S2. Image of Sanger sequencing results showed the nature of the in-frame deletion in the Ald subunit of the adhE1 gene (CAETHG_3747). Figure S3. Change in CO headspace pressure for C. autoethanogenum strains carrying plasmids for AAT expression on CO as main carbon source. Figure S4. Screening of ethyl acetate production by C. autoethanogenum strains carrying plasmids for AAT expression on 40 mM fructose. Figure S4. Investigating growth of C. autoethanogenum on CO and ethyl acetate. Figure S5. Investigating ethyl acetate degradation by C. autoethanogenum grown on CO. Figure S6. Eat1 in vivo alcoholysis assay for C. autoethanogenum and E. coli. Figure S7. Investigating the effects of ethanol supplementation on growth of C. autoethanogenum [PThl-Atf1] grown on CO. Figure S8. Investigating the effects of ethanol supplementation on ethyl acetate production by C. autoethanogenum [PThl-Atf1] grown on CO
: zip. GenBank files of plasmids used in this study, including: ppta-acka-atf1.gb, ppta-acka-eat1.gb, ppta-acka-treat1.gb, precas1-ald.gb, pthl-atf1.gb, SIBR-Cas pta.gb.
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Dykstra, J.C., van Oort, J., Yazdi, A.T. et al. Metabolic engineering of Clostridium autoethanogenum for ethyl acetate production from CO. Microb Cell Fact 21, 243 (2022). https://doi.org/10.1186/s12934-022-01964-5
- Syngas fermentation
- Ethyl acetate
- Butyl acetate
- Alcohol acetyl transferase