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Adding the Propionyl-CoA Pathway to E.coli

Adding the Propionyl-CoA Pathway to E.coli


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So I'm trying to simulate the production of 3-Hydroxypropionic Acid with E.coli via the Propionyl-CoA Pathway. However I'm not really sure how the production in the pathway goes. I got the following picture:

If I understand it correctly the pathway goes: Acetyl-Coa--> Propionate --> Propionyl-CoA -->… --> 3-HP When adding these reactions in Cobra are the stoichiometric coefficients always just 1?


Source of the figure: Luo H, Zhou D, Liu X, Nie Z, Quiroga- Sánchez DL, Chang Y (2016) Production of 3- Hydroxypropionic Acid via the Propionyl-CoA Pathway Using Recombinant Escherichia coli Strains. PLoS ONE 11(5): e0156286. doi:10.1371/journal. pone.0156286


If I understand it correctly the pathway goes: Acetyl-CoA --> Propionate --> Proprionyl-CoA -->… --> 3-HP

Not exactly. The first reaction is: $propionate + acetyl extrm-CoA ightarrow propionyl extrm-CoA + acetate$ The CoA group is transferred from acetate to propionate. The following reactions are an oxidation to acryloyl-CoA and a subsequent hydration to 3-HP-CoA. Lastly the CoA group is transferred back to acetate.

Propionate is a 3 carbon molecule and so is 3-HP. So stoechiometric coefficients of 1 seem logical.


Propionyl-CoA:succinate CoA transferase

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Abstract

We have previously reported in vivo biosynthesis of 2-hydroxyacid containing polyesters including polylactic acid (PLA), poly(3-hydroxybutyrate-co-lactate) [P(3HB-co-LA)], and poly(3-hydroxybutyrate-co-2-hydroxybutyrate-co-lactate) [P(3HB-co-2HB-co-LA)] employing metabolically engineered Escherichia coli strains by the introduction of evolved Clostridium propionicum propionyl-CoA transferase (PctCp) and Pseudomonas sp. MBEL 6–19 polyhydroxyalkanoate (PHA) synthase 1 (PhaC1Ps6–19). In this study, we further engineered in vivo PLA biosynthesis system in E. coli to synthesize 2HB-containing PHA, in which propionyl-CoA was used as precursor for 2-ketobutyrate that was converted into 2HB-CoA by the sequential actions of Lactococcus lactis ( d )-2-hydroxybutyrate dehydrogenase (PanE) and PctCp and then 2HB-CoA was polymerized by PhaC1Ps6–19. The recombinant E. coli XL1-blue expressing the phaC1437 gene, the pct540 gene, and the Ralstonia eutropha prpE gene together with the panE gene could be grown to 0.66 g/L and successfully produced P(70 mol%3HB-co-18 mol%2HB-co-12 mol%LA) up to the PHA content of 66 wt% from 20 g/L of glucose, 2 g/L of 3HB and 1 g/L of sodium propionate. Removal of the prpC gene in the chromosome of E. coli XL1-blue could increase the mole fraction of 2HB in copolymer, but the PHA content was decreased.

The metabolic engineering strategy reported here suggests that propionyl-CoA can be successfully used as the precursor to provide PHA synthase with 2HB-CoA for the production of PHAs containing 2HB monomer.


Poly(3-hydroxypropionate) [poly(3HP)] is a promising monomer of the polyhydroxyalkanoate (PHA) family of material properties similar to other short chain lengths (SCL) PHAs like poly(3-hydroxybutyrate) [poly(3HB)]. It is a well-characterized polyester, biodegradable, biocompatible, and has excellent mechanical properties such as high flexibility, high tensile strength, and high rigidity but more stable than polylactic acid (Andree෾n et al., 2010, 2014b). Poly(3HP) homopolymer biosynthesis came to the limelight in 2010, using glycerol as a carbon source in a two-step fed-batch fermentation and proved to be a promising alternative petrochemical-derived plastic.

Within half a century, glycerol, a by-product of biodiesel production, has become a common carbon source for several biotechnological research in the world. Due to environmental pollution associated with the petrochemical industry resulting in climate change and overexploited fossil reserves, there needs to use biological approaches through metabolic engineering to produce many chemicals from non-renewable sources using inexpensive and abundant raw materials (Keasling, 2010).

The effectiveness and success of the biological production of a platform chemical like 3-hydroxypropoic acid (3-HP) are dependent on the development of microbial Cell Factories thus prompting vigorous and extensive researches conducted in diverse fields such as biosynthesis, gene amplification, engineering of enzymes, design of the genetic circuit, genome editing, bioinformatics, adaptive laboratory evolution, multiomics using suitable hosts, and biosynthetic pathways of microorganisms through the process called �sign-Build-Test-Learn (DBTL)” cycles (Li et al., 2012 Abatemarco et al., 2013 Cheong et al., 2015 Wu et al., 2016). Furthermore, to commercialize these processes through effective biosynthetic pathways to achieve economically viable products, biosensors must be constructed in a suitable metabolic route, optimized to produce desirable results (Liu et al., 2015a).

In order to increase yields of PHAs through synthetic biotechnology and metabolic engineering, there has been the use of metabolite biosensors to detect and monitor a target metabolite in vivo by high-throughput screening (HTS) (Eggeling et al., 2015). Although previous review papers have discussed some biosynthetic pathways, material and chemical characteristics of poly(3HP) (Andree෾n et al., 2014b Chang et al., 2018), however, this review extensively discusses up to date the metabolic pathways that can be used to produce poly(3HP) from glycerol and glucose. The paper also focuses on the production host, the use of malonyl-CoA biosensors to optimize yield, life cycle assessment for poly(3HP) production using corn oil as a carbon source, and the physical and chemical properties. Finally, we will highlight some applications of poly(3HP).

Polyhydroxyalkanoates are classes of intracellular, linear high molecular aliphatic bacterial storage polyesters, which are biocompatible thermoplastics and biodegradable into carbon dioxide and water by many microorganisms (Zhang et al., 2018). Physical or material properties of PHAs are similar to petroleum-derived plastics but exhibit some different characteristics ranging from brittleness to flexibility and elasticity, as shown in Table 1 (Muhammadi et al., 2015). PHA polymers exhibition of features such as elasticity, flexibility, biodegradability, and renewability are highly dependent on these factors, namely (1) their biosynthesis pathways of production (Masood et al., 2014), (2) monomeric composition (Luzi et al., 2019), and (3) chemical structure (Bugnicourt et al., 2014).

Table 1. Comparison of thermodynamic properties of poly (3-HP) compared to other PHAs copolymers.

Furthermore, the production of PHAs is currently not economical in comparison to that of synthetic plastics. To make bacterial PHAs production cost-effective, a few critical factors need to be addressed, such as screening and selection of potential bacterial strains (Kumar et al., 2016, 2020), synthesis pathways, advanced tools and technologies (Mo៎jko-Ciesielska and Mostek, 2019), carbon and nitrogen source (Zahari et al., 2014), and cost-efficient down-stream pro-cesses (Koller and Braunegg, 2018). The purposeful application of the PHAs is also a crucial factor determining its importance and economy (Chen et al., 2017 Cao et al., 2019).

Many researchers have channeled their interest in the production of PHAs recently due to inexpensive carbon sources as substrate and the dangerous problem associated with single-use plastic waste disposal (Chen and Patel, 2012).

Previously, ring-opening polymerization (ROP) of β-propiolactone and 3-HP ester condensation was the chemical processes for synthesizing poly(3HP). There has been no evidence of a natural organism capable of synthesizing it (Wang et al., 2012, 2013 Heinrich et al., 2013).

However, poly(3HP) was not commercially feasible to produce in large quantities because β-propiolactone as a substrate has carcinogenic properties (Dunn, 2012 Dunn et al., 2016). 3-HP, which has low crystallinity, is an ideal precursor that forms a constituent for several copolymers like poly(3-hydroxybutyrate-co-3-hydroxypropionate) [poly(3HB-co-3HP)] or poly(3-hydroxypropionate-co-4-hydroxybutyrate) [Poly(3HP-co-4HB)] (Zhou et al., 2011 Wang et al., 2013).

3-hydroxypropoic acid, acrylate, and 1,3-propanediol (1,3-PDO) were previously utilized as dependent precursors for the biosynthesis of poly(3HP) and 3-HP-containing copolymer (Gao et al., 2014). However, these expensive precursors and Vitamin B12 increased poly(3HP) production cost making it economically not feasible.

Artificial pathways were constructed for poly(3HP) biosynthesis from inexpensive carbon sources such as glucose and glycerol without the addition of precursors to remedy the challenges arising from high production costs (Andree෾n et al., 2010). Nevertheless, only 13 mg/L poly(3HP) was produced by recombinant Escherichia coli strain using glucose as a substrate. Andree෾n et al. (2010) produced 1.42 mg/L poly(3HP) by glycerol conversion using a recombinant E. coli strain carrying glycerol dehydratase of Clostridium buytricum DhaB1Cb, propionaldehyde dehydrogenase of Salmonella enterica serovar Typhimurium LT2 (PduPSe), and PHA synthase gene of Ralstonia eutropha HI6 (PhaC1Re) in a two-step fed-batch fermentation process (Andree෾n et al., 2010).


RESULTS

Identification of the putative mesaconyl-CoA hydratase gene.

Previous work on the novel pathway of acetate assimilation in R. sphaeroides identified an essential gene that codes for an enzyme of the (R)-enoyl-CoA hydratase family (2). Similar genes were found in other bacteria that have in common with R. sphaeroides the absence of a functional glyoxylate cycle for acetate assimilation. However, the overall sequence identity of these proteins to the characterized member of this so-called MaoC or FkbR2 family, an R-specific 3-hydroxyacyl-CoA dehydratase from Aeromonas caviae (18), is only in the range of 20%. Furthermore, the expected molecular mass of the encoded enzyme is about 38 kDa, which is larger than the masses of normal enoyl-CoA hydratases (14 to 28 kDa). A similar gene was found in C. aurantiacus next to the gene coding for l -malyl-CoA/β-methylmalyl-CoA lyase. The gene products from C. aurantiacus and R. sphaeroides had 58% identical amino acid sequences. They showed two hydratase domains, which exhibited only 25% identity on the amino acid level. Normally, enoyl-CoA hydratases contain only one such domain. This suggests gene duplication and also explains the larger size of the subunits compared to other enoyl-CoA hydratases.

Cloning and expression.

The putative 1.06-kb Chloroflexus mesaconyl-CoA hydratase gene coded for a 38-kDa protein (352 amino acids), and the 1.02-kb Rhodobacter gene coded for a 37-kDa protein (343 amino acids). The genes from both bacteria were amplified, and the expected PCR products were cloned into the expression vector pET16b, resulting in coding for N-terminal His10-tagged proteins with altered molecular masses of about 40 kDa. Both plasmids were transformed into E. coli DH5α and then transformed into E. coli BL-21(DE3) for expression. As a control, the expression vector without an insert was also transformed. Both genes were heterologously expressed, and the enzymes were soluble, as deduced from an induced protein band around 40 kDa in SDS-PAGE of the soluble cell fraction (Fig. ​ (Fig.2 2 ).

Denaturing PAGE (12.5%) of various purification steps of heterologously expressed mesaconyl-CoA hydratase of C. aurantiacus and R. sphaeroides from extracts of 3 g E. coli cells. Lanes 1 and 7, molecular mass standards lanes 2 to 4, Chloroflexus enzyme lane 2, extract (100,000 × g supernatant) of induced E. coli cells (45 μg protein) lane 3, heat precipitation fraction (20 μg protein) lane 4, affinity chromatography fraction (8 μg protein) lanes 5 and 6, Rhodobacter enzyme lane 5, extract (100,000 × g supernatant) of induced E. coli cells (30 μg protein) lane 6, affinity chromatography fraction (8 μg protein). The gel was stained with Coomassie brilliant blue R-250.

Spectrophotometric enzyme assay.

The recombinant l - malyl-CoA/β-methylmalyl-CoA lyase from C. aurantiacus was used in a coupled spectrophotometric assay to enzymatically convert propionyl-CoA and glyoxylate to β-methylmalyl-CoA, the substrate of the postulated mesaconyl-CoA hydratase/β-methylmalyl-CoA dehydratase. The further transformation of β-methylmalyl-CoA to other coenzyme A thioesters was tested with E. coli extracts containing overproduced putative mesaconyl-CoA hydratase. Both extracts, which contained either the Chloroflexus or the Rhodobacter gene product, catalyzed a reaction that was associated with an increase in absorbance above 260 nm, with a maximum at around 284 nm. Because protein absorption interfered at this wavelength, we used 290 nm to follow the reaction spectrophotometrically (Fig. ​ (Fig.3). 3 ). The reaction was dependent on propionyl-CoA, glyoxylate, and l -malyl-CoA/β-methylmalyl-CoA lyase. In contrast, extracts of wild-type E. coli or of E. coli carrying the plasmid without an insert were inactive.

UV spectra of coenzyme A ( .._.. ), propionyl-CoA (—), β-methylmalyl-CoA (—), and the product of the hydratase reaction, mesaconyl-CoA ( ……. ). The spectra were recorded in 40 mM K2HPO4/formic acid buffer, pH 4.2, and were normalized to the same absorption at 260 nm.

Confirmation of a β-methylmalyl-CoA-transforming enzyme activity.

HPLC analysis revealed that β-methylmalyl-CoA was consumed almost completely and a new product was formed (Fig. ​ (Fig.4). 4 ). At equilibrium, the ratio of the concentrations of β-methylmalyl-CoA to the product was near 1:10. The product still had the 260-nm adenine nucleotide absorbance maximum characteristic of CoA, propionyl-CoA, or β-methylmalyl-CoA. However, it had an additional absorbance shoulder at longer wavelengths (Fig. ​ (Fig.3). 3 ). The estimated molar 290-nm absorption coefficient of this product (most likely mesaconyl-CoA) was 2,350 M 𢄡 cm 𢄡 , that of the substrate β-methylmalyl-CoA was 200 M 𢄡 cm 𢄡 , and that of propionyl-CoA was approximately 200 M 𢄡 cm 𢄡 . This estimation is based on the assumption that the molar absorption coefficients at 260 nm (ɛ260) of the substrate and the product are very similar to that of normal CoA esters, such as acetyl-CoA (16,400 M 𢄡 cm 𢄡 for comparison, the ɛ260 of CoA is 16,800 M 𢄡 cm 𢄡 ) (11). The Δɛ290 (mesaconyl-CoA minus β-methylmalyl-CoA) of 2,150 M 𢄡 cm 𢄡 was used to quantify the β-methylmalyl-CoA consumption and product formation reaction in the spectrophotometric assay.

HPLC separation of [ 14 C]mesaconyl-CoA as the product of the mesaconyl-CoA hydratase from [ 14 C]β-methylmalyl-CoA and [ 14 C]propionyl-CoA. [ 14 C]β-methylmalyl-CoA is enzymatically formed from [ 14 C]propionyl-CoA and glyoxylate with recombinant l -malyl-CoA/β-methylmalyl-CoA lyase. CoA and its derivatives were detected at 260 nm. Free acids of the corresponding CoA thioesters were eluted between 2 and 5 min. (A) Before addition of glyoxylate to the assay mixture. (B) Ten minutes after addition of glyoxylate. (C) Formation of mesaconyl-CoA after another 10 minutes of incubation with recombinant mesaconyl-CoA hydratase. The sensitivities of 14 C detection by solid-phase scintillation counting in the different runs were the same, and therefore, the signals can be directly compared. For the conditions, see Materials and Methods.

Enzyme activity in cell extract of C. aurantiacus.

Using excess amounts of recombinant l -malyl-CoA/β-methylmalyl-CoA lyase from C. aurantiacus, saturating concentrations of [ 14 C]propionyl-CoA, and an excess of glyoxylate, the specific activity of the enzyme system that produced labeled mesaconyl-CoA from labeled propionyl-CoA in cell extract of C. aurantiacus could be estimated (55ଌ). The specific enzyme activity in extracts of autotrophically grown cells amounted to 20 to 30 nmol min 𢄡 mg protein 𢄡 , depending on the batch of cells. The specific enzyme activity was 10- to 15-fold lower in heterotrophically grown cells.

Purification of recombinant enzymes.

Most of the protein overproduced in E. coli was in the soluble cell fraction, and purification started from the 100,000 × g supernatant. For the recombinant Chloroflexus enzyme, E. coli extract was heated for 10 min at 70ଌ the enzyme from this moderate thermophile remained active and in the supernatant (Fig. ​ (Fig.2). 2 ). The supernatant (or cell extract in the case of the recombinant Rhodobacter enzyme) was then chromatographed on an Ni-Sepharose high-performance affinity column. From 3 g E. coli cells (wet weight), 6.4 mg of Chloroflexus enzyme with a yield of 87% or 4 mg of Rhodobacter enzyme with a yield of 52% was obtained (Table ​ (Table1 1 ).

TABLE 1.

Purification of recombinant His-tagged mesaconyl-CoA hydratase of C. aurantiacus and R. sphaeroides from 3 g of E. coli cells a

Purification stepTotal enzyme activity (μmol min 𢄡 )Total protein (mg)Sp act (μmol min 𢄡 mg protein 𢄡 )Recovery (%)Purification (fold)
Cell extract−/10,500106−/99−/100−/1
Heat precipitation9,300/−16.5/−560/−100/−1−
Ni-Sepharose high performance8,100/5,5006.4/4.01,300/1,40087/522.3/14.0

Properties of mesaconyl-CoA hydratases.

The specific activity of the Chloroflexus enzyme at 55ଌ was 1,300 μmol min 𢄡 mg 𢄡 , and that of the Rhodobacter enzyme at 30ଌ was 1,400 μmol min 𢄡 mg 𢄡 . The enzyme turnover per dimeric enzyme based on specific activities for the Chloroflexus enzyme was 1,700 s 𢄡 , and that for the Rhodobacter enzyme was 1,900 s 𢄡 . Both enzymes exhibited optimal activity at pH 7.5 and half-maximal activity at pH 6.5 and 8.5 (data not shown). The UV spectra of the enzymes showed absorptions only at 280 nm. SDS-PAGE revealed that the enzymes were composed of approximately 40-kDa subunits. Gel filtration indicated a molecular mass of the native enzymes of about 80 kDa, suggesting a homodimeric structure.

Isolation and structure elucidation of the product of the transformation of 13 C-labeled β-methylmalyl-CoA as mesaconyl-CoA.

The 13 C-enriched product of the conversion of [1,2, 3- 13 C]propionyl-CoA and glyoxylate by recombinant l -malyl-CoA/β-methylmalyl-CoA lyase and recombinant mesaconyl-CoA hydratase of C. aurantiacus or R. sphaeroides was analyzed by HPLC-MS. The product had a virtual molecular mass of 877.9 Da, as determined by ESI-MS (negative ion mode). This corresponds to a molecular mass of 878.9 Da, which is close to the expected value of 878.6 Da for mesaconyl-CoA.

The product was isolated by HPLC and analyzed by one- and two-dimensional NMR spectroscopy. Mesaconyl-CoA was characterized through proton NMR signals at 0.87 ppm (s, 3H, 10″ CH3), 1.10 (s, 3H, 11″ CH3), 2.26 (ddd, J = 130.0 Hz, 5.8 Hz, 1.5 Hz, 3H, ʬH3 [mesaconyl]), 2.46 (m, 2H, H6″), 3.13 (q, J = 6.5 Hz, 2H, H9″), 3.40 (t, J = 6.4 Hz, 2H, H8″), 3.51 (t, J = 6.8 Hz, 2H, H5″), 3.64 (m, 1H, H1𠌺), 4.06 (m, 1H, H1𠌻), 4.11 (s, 1H, H3″), 4.32 (s, br, 2H, H5′), 4.51 (s, br, 1H, H4′), 4.91 (m, 1H, H2′), 5.01 (s, br, 1H, H3′), 6.16 (d, J = 5.8 Hz, 1H, H1′), 6.70 (“t,” br, J = 8.0 Hz, 1H [mesaconyl]), 8.24 (s, 1H, H2), 8.61 (s, 1H, H8). The 13 C-enriched carbons (C1, C2, and α-methyl) of mesaconyl-CoA gave signals at 14.3 ppm (dd, J = 42.6 Hz, 2.3 Hz), 149.4 (dd, J = 55.5 Hz, 42.6 Hz), and 195.3 (dd, J = 55.5Hz, 2.3 Hz), indicating the incorporation of the intact propionyl 13 C-enriched carbon chain. The corresponding signal in the proton NMR at 2.26 ppm of the 13 C-enriched α-methyl group ( 13 C NMR, 14.3 ppm) was identified through a gradient heteronuclear single quantum coherence experiment. Moreover, the signal at 6.70 ppm (proton NMR, β-CH) showed a cross-peak to the three signals at 14.3, 149.4, and 195.3 ppm ( 13 C NMR) in the heteronuclear multiple-bond correlation experiment, verifying the mesaconyl-CoA structure. The CH2 group at 3.13 ppm (proton NMR) gave a cross-peak with the 13 C-enriched carbonyl-C at 195.3 ppm ( 13 C NMR) in the heteronuclear multiple-bond correlation experiment, verifying that the CoA residue was attached at C-1 of mesaconyl-CoA.


Abstract

The Escherichia coli genome encodes seven paralogues of the crotonase (enoyl CoA hydratase) superfamily. Four of these have unknown or uncertain functions their existence was unknown prior to the completion of the E. coli genome sequencing project. The gene encoding one of these, YgfG, is located in a four-gene operon that encodes homologues of methylmalonyl CoA mutases (Sbm) and acyl CoA transferases (YgfH) as well as a putative protein kinase (YgfD/ArgK). We have determined that YgfG is methylmalonyl CoA decarboxylase, YgfH is propionyl CoA:succinate CoA transferase, and Sbm is methylmalonyl CoA mutase. These reactions are sufficient to form a metabolic cycle by which E. coli can catalyze the decarboxylation of succinate to propionate, although the metabolic context of this cycle is unknown. The identification of YgfG as methylmalonyl CoA decarboxylase expands the range of reactions catalyzed by members of the crotonase superfamily.

This research was supported by the Universität Karlsruhe, the Deutsche Forschungsgemeinschaft, and the Fonds der Chemischen Industrie (to J.R.) and NIH Grants GM-40570 and GM-52594 (to J.A.G.).

Address correspondence to this author. Telephone: 217-333-3945. Fax: 217-265-0385. E-mail: [email protected]


Conclusions and Future Perspectives

The commercialisation of production methods for biologically-sourced gaseous fuels is crucial to support the global challenges of realising renewable energy supplies and reducing the carbon footprint and other pollutants. Further research is needed to develop tuneable alkane production across the spectrum of short to very long chain hydrocarbons, effectively converting microorganisms into the ‘oilfields of the future’. This will satisfy the demand for blending with or even replacing the current dependence on petroleum-based fuels and synthetic precursors.

The transition from ‘proof-of-principle’ research to successful commercialisation requires a detailed understanding of the techno-economic factors associated with scaling biological processes. A recent study into the commercial potential of fermentative alkane gas production identified key parameters that needed optimisation to enable cost-effective fuel production, and proposed mitigations to overcome these barriers [6]. These mitigations included the transition towards robust industrial microorganisms requiring drastically reduced capital and running costs, sourcing low cost and renewable energy sources, and increasing gas production titres. The latter is particularly important for biological alkane production as the terminal ADO / CvFAP-dependent deformylation / decarboxylation step is thought to be the rate limiting step.

Identification of the important barriers to commercial success can help focus further research, for example to improve In vivo biocatalytic efficiencies, by applying enzyme evolution or redesign strategies to increase reaction rates, stability and expression within a chosen chassis. The advent of synthetic biology techniques enables more in-depth optimisation of process development beyond traditional enzyme redesign. Enhancements in productivity can be obtained by optimising DNA regulatory parts [74], both on a transcriptional and translational level [75, 76], metabolic engineering of auxiliary supply pathways to relieve bottlenecks, and the elimination or downregulation of competing side-reactions. These are important areas of process optimisation realised through bioengineering but overall process optimisation will be essential beyond the need to improve microbial cell factories for bio alkane gas production.

The unprecedented curtailment of global economic activity and mobility during early 2020 due to the Covid 19 pandemic has reduced global energy demand by 3.8% relative to the same time period in 2019 [77]. In spite of this, fossil fuel supplies remain limited and non-renewable, with demand still at high levels. The development of (ultimately) sustainable bio-manufacturing of gaseous hydrocarbons is therefore timely, with success measured by the ability to compete on price and abundancy with existing non-renewable and commercial synthesis routes.


Materials and Methods

Strains and Media

Media and growth conditions for E. coli were previously described by Sambrook and Russell (2001) those for Y. lipolytica were previously described by Barth and Gaillardin (1997). Rich medium (YPD) and minimal glucose medium (YNB) were prepared as described elsewhere (Milckova et al., 2004). The YNB contained 0.17% (w/v) yeast nitrogen base (without amino acids and ammonium sulfate, YNBww), 0.5% (w/v) NH4Cl, 50 mM KH2PO4-Na2HPO4 (pH 6.8), and 2% (w/v) glucose. To complement strain auxotrophies, 0.1 g/L of uracil or leucine was added as necessary. To screen for hygromycin resistance, 250 μg/ml of hygromycin was added to the YPD. Solid media were prepared by adding 1.5% (w/v) agar.

Construction of Plasmids and Strains (E. coli and Y. lipolytica)

We used standard molecular genetic techniques (Sambrook and Russell, 2001). Restriction enzymes were obtained from New England Biolabs (Ipswich, MA, USA). PCR amplification was performed in an Eppendorf 2720 Thermal Cycler with either Q5 High-Fidelity DNA Polymerase (New England Biolabs) or GoTaq DNA polymerases (Progema, WI, USA). PCR fragments were purified using a PCR Purification Kit (Macherey-Nagel, Duren, Germany), and plasmids were purified with a Plasmid Miniprep Kit (Macherey-Nagel).

The plasmids used in this study were constructed using Golden Gate assembly, as described in Celinska et al. (2017). The genes in the A, T, and H module were obtained via PCR using the genomic DNA of Y. lipolytica W29. Internal BsaI recognition sites were removed via PCR using the primers listed in Supplementary Table 1. The plasmids for each module included the Zeta sequence, the URA3 ex marker, and gene expression cassettes containing the TEF1 promoter and LIP2 terminator.

For the cytosolic PDH complex, all the genes were synthesized and cloned in the plasmid pUC57 by GenScript Biotech (New Jersey, US). Cytosolic PDX1 was cloned into the expression plasmid (JME2563) using the BamHI and AvrII restriction sites. The other four genes were cloned into two plasmids (JME4774 and JME4775) using Golden Gate assembly.

To disrupt PHD1, the cassettes were constructed to include a promoter (pPHD1), a marker (URA3 or LEU2), and a terminator (TPHD1), which allowed the ORF gene to be removed via homologous recombination, as described in Papanikolaou et al. (2013).

Gene expression and disruption cassettes were prepared by NotI digestion and transformed into Y. lipolytica strains using the lithium acetate method, as described previously (Barth and Gaillardin, 1997). Gene integration and disruption were verified via colony PCR using the primers listed in Supplementary Table 1. The replicative plasmid harboring the Cre gene (JME547 Table 1) was used for marker rescue (Fickers et al., 2003). After transformation with the Cre expression plasmid, the loss of the marker gene was verified on YNB with/without uracil. The loss of the replicative plasmid was checked using replica plating on YPD with/without hygromycin after culturing on YPD for 24 h. To construct the prototrophic strain, a LEU2 fragment from plasmid JMP2563 was transformed. All the strains and plasmids used in this study are listed in Table 1.

Table 1. The plasmids and strains used in this study.

Culture Conditions for the Lipid Biosynthesis Experiments

The lipid biosynthesis experiments were carried out in minimal media, and the cultures were prepared as follows: an initial pre-culture was established by inoculating 10 mL of YPD medium in 50 mL Erlenmeyer flasks. Then, the pre-culture was incubated overnight at 28ଌ and 180 rpm. The resulting cell suspension was washed with sterile distilled water and used to inoculate 50 mL of minimal medium YNBD6 containing 0.17% (w/v) yeast nitrogen base (without amino acids and ammonium sulfate, YNBww, Difco), 0.15% (w/v) NH4Cl, 50 mM KH2PO4-Na2HPO4 buffer (pH 6.8), and 6% (w/v) glucose. This medium had been placed in 250 mL Erlenmeyer flasks. The cultures were then incubated at 28ଌ and 180 rpm.

Lipid Determination

Lipids were extracted from 10 to 20 mg of freeze-dried cells and converted into FA methyl esters (FAMEs) using the procedure described by Browse et al. (1986). The FAMEs were then analyzed using gas chromatography (GC), which was carried out with a Varian 3900 instrument equipped with a flame ionization detector and a Varian FactorFour vf-23ms column, where the bleed specification at 260ଌ is 3 pA (30 m, 0.25 mm, 0.25 μm). The FAMEs were identified via comparisons with commercial standards (FAME32, Supelco) and quantified using the internal standard method, which involves the addition of 100 μg of commercial dodecanoic acid (Sigma-Aldrich). Commercial odd-chain FAs (Odd Carbon Straight Chains Kit containing 9 FAs, OC9, Supelco) were converted into their FAMEs using the same method employed with the yeast samples. They were then identified using GC and compared with the odd-chain FAs from the yeast samples.

To determine dry cell weight (DCW), 2 mL of the culture was taken from the flasks, washed, and lysophilized in a pre-weighed tube. The differences in mass corresponded to the mg of cells found in 2 mL of culture.


Abstract

The Escherichia coli genome encodes seven paralogues of the crotonase (enoyl CoA hydratase) superfamily. Four of these have unknown or uncertain functions their existence was unknown prior to the completion of the E. coli genome sequencing project. The gene encoding one of these, YgfG, is located in a four-gene operon that encodes homologues of methylmalonyl CoA mutases (Sbm) and acyl CoA transferases (YgfH) as well as a putative protein kinase (YgfD/ArgK). We have determined that YgfG is methylmalonyl CoA decarboxylase, YgfH is propionyl CoA:succinate CoA transferase, and Sbm is methylmalonyl CoA mutase. These reactions are sufficient to form a metabolic cycle by which E. coli can catalyze the decarboxylation of succinate to propionate, although the metabolic context of this cycle is unknown. The identification of YgfG as methylmalonyl CoA decarboxylase expands the range of reactions catalyzed by members of the crotonase superfamily.

This research was supported by the Universität Karlsruhe, the Deutsche Forschungsgemeinschaft, and the Fonds der Chemischen Industrie (to J.R.) and NIH Grants GM-40570 and GM-52594 (to J.A.G.).

Address correspondence to this author. Telephone: 217-333-3945. Fax: 217-265-0385. E-mail: [email protected]


Discussion

Several examples of unusual type I modular PKS extender units have been reported over the past half-decade. In all cases, these are assembled via a common final biosynthetic step, involving reductive carboxylation of an α, β-unsaturated CoA thioester by a CCRC homologue. The data reported here reveal an alternative mechanism for assembly of unusual type I modular PKS extender units. This employs a unique YCC β-subunit (MccB) that, in partnership with the primary metabolic YCC α-subunit, is able to directly carboxylate medium chain acyl-CoA thioesters. X-ray crystallographic analysis revealed the structural basis for substrate recognition by MccB, allowing a homologue likely involved in the biosynthesis of butylmalonyl-CoA and other unusual extender units incorporated into the primycin complex of clinically-used antibiotics to be identified.

These findings inspired the precursor-directed biosynthesis of novel azide- and alkyne-containing stambomycin analogues. Several biosynthetic engineering approaches have recently been utilized to introduce alkyne-terminated extender units containing 5–8 carbons into erythromycin and antimycin analogues 26,27,28 . Feeding of 8-nonynoic acid to S. ambofaciens resulted in the production of a stambomycin analogue containing a 9-carbon alkyne-terminated extender unit in comparable quantities to the natural products. Thus, MccB, the AT domain from module 12 of the stambomycin PKS and the medium chain acyl-CoA synthetase encoded by samR0482 are useful additions to the synthetic biology toolkit for bio-orthogonal tagging of polyketides.


Methods

Microbial strains and media

Table 1 lists the various strains and plasmids used in this study. The E. coli LS5218 strain [fadR, atoC(Con)], which constitutively expresses the enzymes of fatty acid β-oxidation pathway, allows expression of various pathway genes cloned into pTrc99a and pBBR1MCS2 vectors upon induction with isopropyl-β- d -1-thiogalactopyranoside (IPTG) [38]. The IPTG was added when the cells had reached an optical density at a wavelength of 600 nm (OD600) of 0.6. The PHA-producing strain LZ05 with deletion of the genes ptsG, tesB and yciA was described previously [23]. The E. coli DH5α strain served as the host strain for subsequent construction and propagation of various PHA-producing plasmids. During the recombinant plasmid construction, strains were cultivated in Luria–Bertani (LB) medium (10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl). For gene knockout, SOB medium (20 g/L tryptone, 5 g/L yeast extract, 0.5 g/L NaCl, 10 mM MgCl2 and 2.5 mM KCl) was utilized.

Plasmid construction

For the even-chain monomer supply, the construction of plasmid pQQ05 has been previously described [23]. Briefly, the genes yqeF, fadB, phaJ1Pa, ter and phaC2Pa were all cloned and ligated into the corresponding sites of pTrc99a which were cut with the same restriction enzymes stepwise to generate plasmid pQQ05.

The construction of odd-chain monomer generation pathway was as follows. The codon-optimized prpP gene was cloned into the pBBR1MCS2 vector between the KpnI and BamHI sites to construct the plasmid of pZQ01. Later, in order to form the plasmid pBBR1MCS2-acs, namely pZQ02, the acs gene amplified via polymerase chain reaction (PCR) using E. coli MG1655 genomic DNA (gDNA) as template was also inserted into the pBBR1MCS2. The prpE and pct fragments amplified from R. eutropha H16 gDNA with primers prpE-F/prpE-R and pct-F/pct-R were separately ligated into the pBBR1MCS2 to yield the plasmids pZQ03 and pZQ04. Subsequently, co-expression of two genes prpP and acs, prpP and prpE, prpP and pct in the pBBR1MCS2 was utilized to form the plasmids pZQ05, pZQ06 and pZQ07, respectively. All of the genes were under the control of the lac promoter with separated ribosomal binding site located upstream of each gene to facilitate the translation. The R. eutropha H16 template used for these PCR reactions was isolated using the TIANamp Bacterial DNA Kit (TIANGEN BIOTECH, China). The primers used to amplify different fragments for cloning reactions are listed in Additional file 1: Table S1.

In all cases, PCR was performed using an S1000 Thermal Cycler (Bio-Rad, USA). PrimeSTAR HS DNA polymerase was purchased from Takara (Tokyo, Japan), restriction endonucleases were from Fermentas/Thermo Scientific (Pittsburgh, USA), and T4 DNA ligase was from New England Biolabs (Ipswich, USA). Propagated plasmids were prepared by TIANGEN Plasmid Mini Extraction Kit (TIANGEN BIOTECH, China), and restriction enzyme-digested products were purified using an E.Z.N.A.™ Gel Extraction Kit (Omega, USA). DNA sequencing of all constructed plasmids were performed by Liuhe BGI Tech Co. Ltd (Beijing, China). All of the constructed plasmids were transformed into the strain LZ05 and the optimum double plasmids were then transformed into the strain LZ08 according to standard procedures [39].

Gene knockout

The gene pflB which encodes pyruvate formate lyase was knocked out by the one-step inactivation method as described previously [40] and poxB encoding pyruvate oxidase was knocked out by linearized DNA fragments with extending homologous sequence [41]. First, the linerized DNA fragments with the FLP recognition target sites and 39 bp homologous sequences were obtained via PCR using pKD4 (Km R ) as a template and pflB-F/pflB-R as primers. After the DNA gel extraction, the purified PCR product was electroporated into the host cells which carried the plasmid pKD46, and then E. coli LZ05 was induced by 0.3% (w/v) l -arabinose to express the λ Red system. The positive transformants were selected and identified by colony PCR using the primers pflB-test-F/pflB-test-R. Regarding the poxB deletion, primers poxB-F/poxB-R and chromosomal DNA of the strain QZ1111 were applied to amplify the linearized DNA fragments for poxB. The deletion procedure of poxB gene was as follows. After DpnI digestion, the PCR products were then purified and electroporated into the competent strain E. coli LZ05 containing the plasmid pKD46. Transformant cells were selected in solid LB medium (1% tryptone, 0.5% yeast extract, 1% NaCl, and 1.5% agar powder) containing chloramphenicol (Cm R ). Candidate clones were screened by PCR employing primers poxB-F/poxB-R. The PCR products were ultimately sequenced in Liuhe BGI Tech Co. Ltd (Beijing, China) if necessary. After removing pKD46, the corresponding Km R or Cm R cassette was removed with the helper plasmid pCP20. The plasmids pKD46 and pCP20 were eliminated by overnight cultivation at 42 °C. Finally, the strain LZ08 with the above two gene inactivation was generated.

Cultivation condition

The medium of shake flask study contains 10 g/L tryptone, 5 g/L yeast extract, 30 mM NH4Cl, 5 mM (NH4)2SO4, 1.48 mM Na2HPO4, and 100 μM FeSO4 supplemented with 125 mM MOPS.

For all shake flask experiments, single colony was inoculated into 5 mL LB broth and grown at 37 °C overnight. 0.5 mL pre-culture was inoculated to 300 mL Erlenmeyer flask containing 50 mL LB and cultivated for 8 to 10 h and then 1% (v/v) seed inoculum for shake flask cultivation was incubated in 50 mL fermentation medium. When all liquid fermentation medium (50 mL) was incubated in 300 mL conical flasks at 37 °C with an agitation of 250 rpm to an optical density at 600 nm (OD600) of 0.6–0.8, 1 mM IPTG was added to the culture broth as an inducer. After induction, 30 g/L glucose was supplied as the sole carbon source at the appropriate time and then fermented for 72 h at 30 °C with shaking at 250 rpm. When necessary, ampicillin (100 μg/mL), kanamycin (50 μg/mL) or chloramphenicol (25 μg/mL) was added to the medium to maintain the stability of the plasmids. After cultivation, cells were gathered by centrifugation at 12,000 rpm for 15 min, washed with water twice and treated with ethanol once and then lyophilized.

PHA production analysis

The content and monomer compositions of intracellular accumulated PHA were analyzed by gas chromatography (GC) as described previously [42]. PHA content was defined as the percent ratio of PHA concentration to CDW. Liquid culture was centrifuged to obtain the supernatant and cellular biomass. 15 mg lyophilized cells were subjected to methanolysis in the presence of 1 mL of chloroform and 1 mL of 3% (v/v) sulfuric acid in methanol for 1 h at 100 °C. The samples were cooled to room temperature and then 1 mL of distilled water was added in order to extract the cell debris that is soluble in the aqueous phase. 10 mg/mL pentadecanoic acid in ethanol was added as an internal standard. The mixture was vortexed and centrifuged at 12,000 rpm for 10 min. After the layer separation, the organic (chloroform) phase (500 μL) was transferred to another new vial and analyzed using a Shimadzu GC2010 gas chromatograph (Kyoto, Japan) equipped with an AOC-20i auto-injector and a RestekRxi ® -5 column. PHA standard samples were dissolved in chloroform and also analyzed according to the method above by GC. The temperature program used was as follows: 80 °C hold for 1 min, ramp from 60 to 230 °C at 10 °C per min and a final hold at 230 °C for 10 min [23].

Cell growth, glucose consumption and acetate assimilation analyses

Cell growth was monitored by measuring OD600 utilizing a spectrophotometer (Shimazu, Japan). Glucose and acetate were quantitatively analyzed by high-performance liquid chromatography (HPLC) (Shimazu, Japan) which equipped with a refractive index detector (RID-10A) and an Ion Exclusion column (Bio-Rad, HPX-87H). The samples were first centrifuged at 12,000 rpm for 10 min, and then the supernatant was filtrated with a 0.22 μm filter membrane. 5 mM sulfuric acid was utilized as the mobile phase of HPLC with the flow rate of 0.6 mL/min and the utilized column temperature was 65 °C.

Statistical analyses

All data examined were expressed as mean ± SD. Statistical analyses of the data were carried out using two-tailed Student’s t-test between two groups, and one-way ANOVA followed by the post hoc Tukey’s test for multiple groups. P < 0.05 was considered significant. The * denotes P < 0.05, the *** denotes P < 0.001.