Engineering Corynebacterium glutamicum for methanol-dependent growth and glutamate production

Author’s Accepted Manuscript Engineering Corynebacterium glutamicum for methanol-dependent growth and glutamate production Philibert Tuyishime, Yu Wang, Liwen Fan, Qiongqiong Zhang, Qinggang Li, Ping Zheng, Jibin Sun, Yanhe Ma www.elsevier.com/locate/ymben

PII: DOI: Reference:

S1096-7176(18)30252-0 https://doi.org/10.1016/j.ymben.2018.07.011 YMBEN1440

To appear in: Metabolic Engineering Received date: 19 June 2018 Accepted date: 19 July 2018 Cite this article as: Philibert Tuyishime, Yu Wang, Liwen Fan, Qiongqiong Zhang, Qinggang Li, Ping Zheng, Jibin Sun and Yanhe Ma, Engineering Corynebacterium glutamicum for methanol-dependent growth and glutamate production, Metabolic Engineering, https://doi.org/10.1016/j.ymben.2018.07.011 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Engineering Corynebacterium glutamicum for methanol-dependent growth and glutamate production Philibert Tuyishimea,b,c,1, Yu Wanga,b,1, Liwen Fana,b,d, Qiongqiong Zhanga,b, Qinggang Lia,b, Ping Zhenga,b,*, Jibin Suna,b,c,*, Yanhe Mab a

Key Laboratory of Systems Microbial Biotechnology, Chinese Academy of Sciences,

Tianjin 300308, China b

Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences, Tianjin

300308, China c

University of Chinese Academy of Sciences, Beijing 100049, China

d

School of Life Science, University of Science and Technology of China, Hefei 230026,

China [email protected] [email protected]). *

Correspondence to: Address: Tianjin Institute of Industrial Biotechnology, Chinese

Academy of Sciences, Tianjin 300308, China. Fax/Phone: +86-02284861943.

Abstract Methanol is a promising feedstock for bioproduction of fuels and chemicals, thus massive efforts have been devoted to engineering non-native methylotrophic platform microorganisms to utilize methanol. Herein, we rationally designed and experimentally engineered the industrial workhorse Corynebacterium glutamicum to serve as a methanol-dependent synthetic methylotroph. The cell growth of the methanol-dependent

1

These authors contributed equally to this work. 1

strain relies on co-utilization of methanol and xylose, and most notably methanol is an indispensable carbon source. Due to the methanol-dependent characteristic, adaptive laboratory evolution was successfully applied to improving methanol utilization. The evolved mutant showed a 20-fold increase in cell growth on methanol-xylose minimal medium and utilized methanol and xylose with a high mole ratio of 3.83:1. 13C-labeling experiments demonstrated that the carbon derived from methanol was assimilated into intracellular building blocks, high-energy carriers, cofactors, and biomass (up to 63% 13

C-labeling). By inhibiting cell wall biosynthesis, methanol-dependent glutamate

production was also achieved, demonstrating the potential application in bioconversion of methanol into useful chemicals. Genetic mutations detected in the evolved strains indicate the importance of intracellular NAD+/NADH ratio, substrate uptake, and methanol tolerance on methanol utilization. This study reports significant improvement in the area of developing fully synthetic methylotrophs.

Keywords:

Synthetic

methylotrophy;

Methanol-dependent;

Corynebacterium

glutamicum; Adaptive laboratory evolution; Ribose phosphate isomerase

1. Introduction The growing global population and food demand necessitate the sustainable production of chemicals and fuels from one-carbon compounds (Clomburg et al., 2017). Recently, abundant natural gas supplies (composed primarily of methane) and feasible conversion of methane to methanol have made methanol a promising alternative substrate with low cost and good availability for biomanufacturing (Bennett et al., 2018b).

2

Methanol is more reduced and energy richer than traditional carbon sources, such as glucose and glycerol. In consequence, the use of methanol as a sole substrate or a cosubstrate is expected to improve the product titer and yield (Whitaker et al., 2015). Although native methylotrophs possess the ability to utilize one-carbon compounds including methanol as sole carbon and energy sources, they are mostly obligate aerobes, produce few metabolites at high levels, and lack advanced genetic tools (Whitaker et al., 2015). Therefore, synthetic methylotrophs derived from well-characterized platform organisms that have rapid growth rates, modern genetic manipulation techniques, and extensive metabolic knowledge are preferred for bioconversion of methanol into chemicals and fuels (Zhang et al., 2017). In the last few years, massive efforts have been devoted to enable platform organisms such as Corynebacterium glutamicum, Escherichia coli, and Saccharomyces cerevisiae to incorporate methanol for building up cellular constituents and producing chemicals (Bennett et al., 2018a; Dai et al., 2017; Gonzalez et al., 2018; Leßmeier et al., 2015; Müller et al., 2015a; Meyer et al., 2018; Price et al., 2016; Rohlhill et al., 2017; Wang et al., 2017; Whitaker et al., 2017; Witthoff et al., 2015; Woolston et al., 2018). The NAD-dependent methanol dehydrogenase (Mdh) and ribulose monophosphate (RuMP) pathway, consisting of 3-hexulose-6-phosphate synthase (Hps) and 6-phospho-3hexuloisomerase (Phi), are usually selected to construct methanol assimilation pathway. With the function of Mdh and RuMP pathway, methanol assimilation in the presence of yeast extract, glucose, ribose, or gluconate was achieved (Müller et al., 2015a; Meyer et al., 2018; Whitaker et al., 2017; Witthoff et al., 2015). The efficiency of methanol utilization was further improved by expressing heterologous non-oxidative pentose

3

phosphate pathway (PPP) from native methylotroph Bacillus methanolicus, deactivating phosphoglucose isomerase (Bennett et al., 2018a), using formaldehyde-inducible promoters (Rohlhill et al., 2017; Woolston et al., 2018), or using engineered enzyme assemblies (Price et al., 2016). It is worth nothing that a very recently reported methylotrophic E. coli achieved methanol-essential growth by coupling methanol assimilation to modified gluconate catabolism (Meyer et al., 2018). By integrating methanol utilization pathway and biosynthesis pathway of specific chemicals in C. glutamicum or E. coli, methanol was successfully used as a co-substrate to produce chemicals such as cadaverine (Leßmeier et al., 2015), naringenin (Whitaker et al., 2017), acetone, and butanol (Bennett et al., 2018a). These recent studies provide evidence that methylotrophic pathways can be transferred to platform organisms and the resulting synthetic methylotrophs can assimilate methanol into biomass and extracellular metabolites. Adaptive laboratory evolution (ALE), a powerful strategy for metabolic engineering, has been shown to be a promising approach for improving microorganisms’ capacity to utilize nonnative carbon sources (Antonovsky et al., 2016; Lee and Palsson, 2010; Meyer et al., 2018; Portnoy et al., 2011). Development of a methanol-dependent synthetic methylotroph will facilitate direct improvement in methanol utilization by ALE. Besides, formaldehyde, the intermediate of methanol utilization, is highly toxic to cells. Therefore, oxidation of methanol, assimilation of formaldehyde, and generation of formaldehyde accepters should be carefully enhanced and fine-tuned in a coordinated way to improve the methanol utilization, which is very challenging to be realized by direct genetic manipulation. ALE, in this case, will be particularly useful. This prompted us to design

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and construct a synthetic methylotroph based on platform organisms whose cell growth is dependent on methanol assimilation. In this study, xylose utilization pathway was engineered in C. glutamicum to enable conversion of xylose into ribulose 5-phosphate (Ru5P) but deactivate the conversion of Ru5P to ribose 5-phosphate (R5P). In consequence, the engineered strain can utilize xylose as a precursor for generating formaldehyde accepter Ru5P but not the sole carbon source. The first round of ALE was conducted using ribose and xylose as carbon sources to adapt the strain’s metabolism to xylose utilization. Furthermore, by introducing Mdh and RuMP genes with the best performance in C. glutamicum, a methanol-dependent synthetic methylotroph whose cell growth was dependent on co-utilization of methanol and xylose was successfully constructed (Fig. 1). The second round of ALE was then applied to accelerate its cell growth and methanol utilization. A mutant with significantly improved performance was screened and used for glutamate production from methanol and xylose. The

13

C-labeling experiments were conducted to analyze the integration of

methanol into intracellular metabolites and cell biomass. Several mutations that were supposed to be responsible for the improved performance on methanol utilization were also detected. We demonstrated a detailed metabolic engineering strategy to construct a methanol-dependent strain and achieved amino acid production using methanol as substrate. This work represents significant improvements in construction of synthetic methylotrophs and bioconversion of methanol into valuable chemicals.

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2. Materials and methods 2.1. Bacterial strains and growth conditions The bacterial strains used in this study are listed in Table 1. E. coli DH5α was used for general cloning and cultivated at 37°C and with shaking at 220 rpm in Luria–Bertani (LB) broth. Kanamycin (Km, 50 μg/mL) or chloramphenicol (Cm, 20 µg/mL) was added to LB broth as required. C. glutamicum ATCC 13032 and its derivatives were cultivated at 30°C and with shaking at 220 rpm in LBG medium or CGXII minimal medium (Keilhauer et al., 1993). The LBG medium is LB broth supplemented with 4 g/L glucose. When CGXII minimal medium was used, 4 g/L of each carbon source (glucose, ribose, xylose, and/or methanol) was added. To avoid evaporation of methanol, the shake flasks were covered with a sealing membrane. Km (25 μg/mL) or Cm (5 µg/mL) was added to the LBG medium or CGXII minimal medium as required.

2.2. Gene expression in C. glutamicum E. coli-C. glutamicum shuttle vectors pXMJ19 and pEC-XK99E were used for gene expression in C. glutamicum (Jakoby et al., 1999; Kirchner and Tauch, 2003). The plasmids used in this study are listed in Table 1. To express xylA from E. coli in C. glutamicum, xylA was amplified from the genomic DNA of E. coli MG1655 using the primer pair xylA-F/xylA-R (Table S1). The PCR product was inserted into the pXMJ19 vector between HindIII and EcoRI sites under the control of the isopropyl-β-Dthiogalactopyranoside (IPTG)-inducible promoter Ptac using the ClonExpress II One Step Cloning Kit (Vazyme Biotech, Nanjing, China). The resultant plasmid pXMJ19-xylA was transformed into C. glutamicum by electroporation according to the protocol described

6

previously (Ruan et al., 2015). IPTG of 1 mM was added into the culture to induce xylA expression. The methanol assimilation genes were expressed in C. glutamicum using the pEC-XK99E vector. The genes encoding Mdh3 and its activator from B. methanolicus MGA3 (Mdh3-ActBm) and Mdh from B. stearothermophilus DSM 2334 (MdhBs2334) were amplified from the genomic DNAs of B. methanolicus MGA3 and B. stearothermophilus DSM 2334 using the primer pairs mdh3-actBm-F/mdh3-actBm-R and mdhBs2334F/mdhBs2334-R, respectively. The gene encoding Mdh2 variant (A26V, A31V, A169V) from Cupriavidus necator N-1 (Mdh2Cn) (Wu et al., 2016) was synthesized by GENEWIZ (Suzhou, China) and amplified using the primer pair mdh2Cn-F/mdh2Cn-R. The pEC-XK99E vector was linearized using PCR and the primer pair pEC-XK99EF/pEC-XK99E-R. The PCR products of Mdh encoding genes were ligated with the linearized pEC-XK99E under the control of the IPTG-inducible promoter Ptrc. The resultant plasmids pEC-XK99E-mdh3-actBm, pEC-XK99E-mdhBs2334, and pEC-XK99Emdh2Cn

were

transformed

into

C.

glutamicum

by electroporation.

Different

concentrations of IPTG (0.1, 0.5, and 1.0 mM) were tested for inducing Mdh expression. To express RuMP genes in C. glutamicum, hps-phiBm and hps-phiBs168 were first amplified from the genomic DNAs of B. methanolicus MGA3 and B. subtilis 168 using the primer pairs hps-phiBm-F/hps-phiBm-R and hps-phiBs168-F/hps-phiBs168-R, respectively. A constitutive promoter PP5 was added upstream of the hps-phiBm and hps-phiBs168 genes via the first round of PCR. A second round of PCR was then performed to add 20 bp overlaps at the two terminals of hps-phiBm and hps-phiBs168 PCR products for ligation using

the

primer

pairs

hps-phi-F2/hps-phiBm-R

and

hps-phi-F2/hps-phiBs168-R,

respectively. The PCR products were ligated with the XmaI linearized pEC-XK99E-

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mdhBs2334, producing pEC-XK99E-mdhBs2334-hps-phiBm and pEC-XK99E-mdhBs2334-hpsphiBs168, respectively. The plasmids were transformed into C. glutamicum by electroporation. IPTG of 1.0 mM were used to induce gene expression. The oligonucleotides for PCR fragment generation used in this study are listed in Table S1.

2.3. Gene knockout in C. glutamicum The suicide plasmid pK18mobsacB was used for gene knockout via allele exchange in C. glutamicum (Schäfer et al., 1994). pK18mobsacB-ΔadhE containing a mutant allele of adhE was constructed to knock out the adhE gene. The mutant allele of adhE was generated by connecting a left and a right homologous flank of adhE. First, the left and right flanks were amplified from the genomic DNA of C. glutamicum ATCC 13032 using the primer pairs ΔadhE-F1/ΔadhE-R1 and ΔadhE-F2/ΔadhE-R2, respectively (Table S1). The two fragments were ligated with the BamHI linearized pK18mobsacB to construct pK18mobsacB-ΔadhE.

Mutant

MX-1

was

then

constructed

by

transforming

pK18mobsacB-ΔadhE into C. glutamicum Δald via electroporation and according to procedures described previously (Ruan et al., 2015; Schäfer et al., 1994). To knock out rpiB gene, pK18mobsacB-ΔrpiB was constructed using the primer pairs ΔrpiB-F1/rpiBR1 and ΔrpiB-F2/ΔrpiB-R2 according to the aforementioned procedure. pK18mobsacBΔrpiB was then transformed into strain MX-2 and the similar procedure was used. Ribose and xylose were used as carbon sources during the whole gene knockout process to avoid growth defect caused by rpiB disruption.

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2.4. Enzyme activity assay Cells of C. glutamicum at exponential phase were harvested and then washed twice with 50 mM potassium phosphate buffer (pH 7.4). For crude cell extract preparation, mechanical lysis of cells was performed with glass beads using FastPrep®-24 Classic Instrument (MP Biomedicals, CA, USA). The crude cell extract was centrifuged at 12,000 × g for 30 min at 4°C to remove the cell debris. The obtained supernatant was used for enzyme assay. Mdh activity was assayed using the method described previously (Müller et al., 2015a) with minor modifications. The reaction mixture contained 50 mM potassium phosphate buffer (pH 7.4), 5 mM MgSO4, 0.5 mM NAD+ and 1 M methanol preheated at 30ºC. The reaction was started by adding crude extracts. Specific Mdh activity (U/mg) was defined as the amount of enzyme producing 1 µmol NADH per minute per mg of total protein. The protein concentration of the cell extracts was determined by the method of Bradford using bovine serum albumin as the standard. The coupled Hps-Phi activity assay was performed as described previously (Arfman et al., 1990) with minor modifications. The reaction mixture contained 50 mM potassium phosphate buffer (pH 7.4), 5 mM MgCl2, 5 mM R5P, 2.5 mM NADP+, 5 U phosphoriboisomerase from spinach (type I, partially purified powder, Sigma-Aldrich, MO, USA), 5 U phosphoglucose isomerase from baker's yeast (Type III, ammonium sulfate suspension, Sigma-Aldrich, MO, USA), 5 U glucose-6-phosphate dehydrogenase from baker's yeast (Type XV, lyophilized powder, Sigma-Aldrich, MO, USA), and 40 µL cell extract at different dilutions. The reaction mixture was incubated for 10 min at 30°C to ensure generation of enough Ru5P from R5P. Subsequently, formaldehyde (methanolfree) was added to the mixture with a final concentration of 5mM to start the reaction and

9

OD340nm was monitored. Specific coupled Hps-Phi activity (U/mg) was defined as the amount of enzyme producing 1 µmol NADPH per minute per mg of total protein.

2.5. Adaptive laboratory evolution (ALE) ALE of C. glutamicum MX-3 was performed using CGXII minimal medium supplemented with ribose and xylose (4 g/L for each carbon source). C. glutamicum MX3 was first cultivated in the aforementioned medium at 30°C and with shaking at 220 rpm. After cultivated for 24 h, the culture was used as a seed to inoculate fresh medium with an initial optical density at 600 nm (OD600nm) of 0.1, which was then incubated under the same conditions. At the certain passage that showed the best cell growth, the culture was diluted, spread on CGXII solid medium supplemented with ribose and xylose, and incubated at 30°C. The colonies that grew fast were cultivated in LB medium supplemented with ribose and xylose and stored for further experimental analysis. ALE of C. glutamicum MX-10 was performed using CGXII minimal medium supplemented with methanol and xylose (4 g/L for each carbon source). To avoid evaporation of methanol, the shake flasks were covered with a sealing membrane. C. glutamicum MX10 was first cultivated in the aforementioned medium at 30°C and with shaking at 220 rpm. When OD600nm of the culture reached over 1.0, the culture was used as a seed to inoculate fresh medium with an initial OD600nm of 0.5, which was then incubated under the same conditions. At the certain passage that showed the best cell growth, the culture was diluted, spread on CGXII solid medium supplemented with methanol and xylose, and incubated at 30°C. The colonies that grew fast were cultivated in LB medium supplemented with methanol and xylose and stored for further experimental analysis.

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2.6. Determination of methanol and xylose The quantitative measurement of methanol and xylose was performed using SBA-40 biosensor analyzer (Institute of Biology of Shandong Province Academy of Sciences, Shandong, China) and Prominence Ultra-Fast Liquid Chromatography (UFLC, Shimadzu, Kyoto, Japan), respectively. Briefly, the culture was harvested and centrifuged at 12,000 × g for 10 min at room temperature and the supernatant was used for methanol and xylose determination. For methanol analysis, a convenient and simple method was adopted. The SBA-40 biosensor analyzer (Institute of Biology of Shandong Province Academy of Sciences, Shandong, China) equipped with an alcohol oxidase membrane was used. The analytical signal was given by quantifying the production of H2O2, which was generated by methanol oxidation catalyzed by alcohol oxidase. Xylose was measured by using Prominence UFLC (Shimadzu, Kyoto, Japan) equipped with a refractive index detector and a Bio-Rad Aminex HPX-87H column (300 × 7.8 mm). The analysis was performed with a mobile phase of 5 mM H2SO4 at 55°C with a flow rate of 0.5 mL/min. The injection volume was 10 μL. The substrate uptake rate (mM/h) was calculated using Eq. (1). The specific methanol uptake rate qM (mmol/gCDW·h) was calculated according to Eq. (2). t, X0, and μ represent the time in hours, the initial biomass concentration, and the specific growth rate in h-1, respectively. Cellular dry weight (CDW) was determined using a conversion factor of 0.30 gCDW/L·OD600nm. The biomass yield on substrate YMS (gCDW/gS) was calculated by employing Eq. (3), where X and S represent the concentrations of biomass and substrate, respectively. The specific growth rate μ for strain MX-11 was obtained using exponential regression on growth data. The specific

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growth rate for each passage of ALE was calculated using the OD600nm values at the initial and final time points. (1) (2) (3)

2.7. Determination of 13C-labeled intracellular metabolites For

13

C-labeled intracellular metabolite analysis, C. glutamicum MX-11 was

cultivated in CGXII minimal medium supplemented with

13

C-methanol (99% atom

enrichment, Sigma-Aldrich, MO, USA) and xylose (4 g/L for each carbon source) for 120 h. For quenching of the cellular metabolism, the culture was quickly injected into -20°C 40% methanol and mixed by vortex (1 s). The mixture was centrifuged at 12,000 × g for 30 s at 0°C and the supernatant was removed. The resulting cell pellet was re-suspended in 5 mL 75% hot ethanol (100°C) and incubated in boiling water bath for 15 min. The cell extract was centrifuged at 5,000 × g for 5 min at 0°C. The supernatant was transferred into a pre-cooled tube. Additionally, 2 mL 50% acidic acetonitrile (0.1% formic acid, -20°C) was added into cell pellet and incubated in ice water bath for 15 min to further extract the intracellular metabolites. The mixture was centrifuged at 12,000 × g for 15 min at 0°C, and the supernatant was transferred into the same tube containing the metabolites extracted using 75% hot ethanol (100°C). The mixture was centrifuged at 12,000 × g for 30 min at 0°C. The supernatant was freeze dried and stored at -80°C.

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The freeze-dried samples were re-suspended in 50 μL 50% acetonitrile and centrifuged at 12,000 × g for 30 min at 0°C. The intracellular metabolites were detected and analyzed by LC-MS/MS using a Shimadzu Nexera Ultra-Performance Liquid Chromatography (UPLC) 30A (Shimadzu, Kyoto, Japan) equipped with a SeQuant ZICHILIC column (100 mm × 2.1 mm, 3.5 μm, Merck, Germany) and coupled with an Applied Biosystem TripleTOFTM 5600 mass spectrometer at a resolution of 30,000 FWHM (Applied Biosystem, USA) at the negative electrospray ionization (ESI) mode. The mobile phase included A phase (10 mmol/L (NH4)2COOH) and B phase (100% acetonitrile) and the flow rate was 0.2 mL/min. The LC gradient was 0-3 min, 90% B; 36 min, 90%-60% B; 6-25 min, 60%-50% B; 25-30 min, 50% B; 30-30.5 min, 50%-90% B; 30.5-38 min, 90% B. The flow rate was 0.2 mL/min. The mass spectra were obtained from ESI negative mode of -35 eV with a scan range of 30-1200 m/z. Moreover, mass accuracy was calibrated by automated calibrant delivery system (AB Sciex, Concord, Canada) interfaced to the second inlet of the DuoSpray source. The injection volume was 5 μL.

13

C-labeling was determined from the measured mass isotopomer data. First, the

mass isotopomer distributions were corrected for natural isotope abundance using the method described previously (Millard et al., 2012). Furthermore, average

13

C-labeling

was determined using the method by Whitaker and colleagues (Whitaker et al., 2017). Average

13

C-labeling (%)=sum(Mi*i)/n, where n is the number of carbon atoms for the

measured fragment and Mi is the corrected mass isotopomer abundance. Taking glutamate as an example, n is the maximum number of carbon atoms that can be labeled (5 carbons).

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2.8. Determination of 13C-labeled proteinogenic amino acids For

13

C-labeled proteinogenic amino acids analysis, C. glutamicum MX-11 was

cultivated in CGXII minimal medium supplemented with

13

C-methanol (99% atom

enrichment, Sigma-Aldrich, MO, USA) and xylose (4 g/L for each carbon source) for 120 h. Cells were then harvested and washed twice with 50 mM potassium phosphate buffer (pH 7.4). Next, cells were resuspended in 6 M HCl and transferred to glass screw-top GC vials. The vials were placed in a 105°C oven for 24 h to hydrolyze the biomass proteins into amino acids. The hydrolysates were centrifuged at 12,000 × g for 10 min to remove solid particles in the hydrolysis solution and the supernatants were transferred into new centrifuge tubes for desiccation. For derivatization, 100 μL of 10 mg/mL methoxylamine hydrochloride in pyridine was added to the samples, which were then vortexed occasionally and incubated at 30°C for 90 min. Then, the samples were mixed with 100 μL of N-methyl-N-trimethylsilyltrifluoroacetamide, vortexed occasionally and incubated at 37°C for 60 min. After derivatization, the samples were centrifuged at 12,000 × g for 10 min, and the supernatant was transferred to new GC vials. The derivatized amino acid samples were analyzed by GC/Q-TOF-MS using an Agilent 7890A GC coupled with a 7200 Accurate-Mass Q-TOF (Agilent Technologies, Germany). A DB-5MS Ultra Inert column (30 m × 0.25 mm, 0.25 μm film thickness) was used (Agilent Technologies, USA). The sample (1 μL) was injected into the GC in splitless mode. The oven temperature was programmed as follows: 60°C for 1 min, 8°C/min to 132°C, 2°C/min to 150°C, 5°C/min to 185°C, 10°C/min to 325°C, 5 min hold. Mass spectra of the amino acids were in the mass range of 50-650 m/z at an acquisition rate of 5 spectra/s. The temperatures of the ion source and transfer line were 250°C and

14

290°C, respectively. The electron ionization was carried out at 70 eV. Agilent Mass Hunter Qualitative Analysis Software was used for peak detection and mass spectral deconvolution. The annotation of amino acids was performed via matching their mass fragmentation patterns with those in the National Institute of Standards and Technology mass spectral library (match factor > 80%). The

13

C-labeling was determined from the

measured mass isotopomer data as described above.

2.9. Glutamate production and determination C. glutamicum MX-11 was cultivated in modified CGXII minimal medium supplemented with methanol and xylose (4 g/L for each carbon source) at 30°C and with shaking at 220 rpm. Biotin was added to the medium to a final concentration of 0.5 μg/L. When the OD600nm of the culture reached 2.0-2.5, penicillin G was added to a final concentration of 60 U/ml to induce glutamate production. Samples were taken periodically and extracellular glutamate concentrations were quantified using an SBA40D biosensor analyzer (Institute of Biology of Shandong Province Academy of Sciences, Shandong, China) as described previously (Wang et al., 2018a).

2.10. Whole genome sequencing Genomic DNAs of evolved methanol-dependent C. glutamicum strains were extracted using the Promega Wizard Genomic DNA Purification Kit (Madison, WI, USA). Library construction and whole genome sequencing were conducted by Berry Genomics (Beijing, China) by using Illumina Hiseq2500 sequencing platform (San Diego, CA, USA). The FastQC software (v.0.10.1) and NGSQC Toolkit software (v.2.3.3) were

15

used to analyze the output for quality assurance. The BWA alignment software (v.0.7.15r1140) and SAM tools software (v1.2) were used for alignment and variant calling, respectively. Annotation of the variations were performed using the SnpEff software (v.4.3i).

2.11. Determination of relative transcriptional levels C. glutamicum strains MX-3 and MX-4 were cultivated in CGXII minimal medium supplemented with ribose and xylose (4 g/L for each carbon source) at 30°C and with shaking at 220 rpm. Cells were collected in the mid-exponential phase. RNA was isolated from the cell pellet using RNAprep Pure Cell/Bacteria Kit (Tiangen Biotech, Beijing, China). After treatment with DNase I (Tiangen Biotech, Beijing, China), the RNA was used to synthesize cDNA using random primers and Fast Quant RT Kit (Tiangen Biotech, Beijing, China). The resultant cDNA was used as a template for quantitative PCR (qPCR) analysis. The RNA samples were also used as templates for PCR to confirm that genomic DNA contamination during RNA extraction was minimal. Specific primers for qPCR were designed using Beacon Designer software (Table S1). qPCR was performed by using SuperReal Premix SYBR Green kit (Tiangen Biotech, Beijing, China) and the Applied Biosystems® 7500 Real-Time PCR System (Thermo Fisher Scientific, USA) according to the manufacturer's instructions.

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3. Results 3.1. Design of methanol-dependent C. glutamicum For synthetic methylotrophs, methanol oxidation and Ru5P regeneration are suggested to be crucial for methanol utilization (Whitaker et al., 2015; Zhang et al., 2017). Ru5P is a PPP intermediate of platform organisms such as C. glutamicum and E. coli. Therefore, the existing synthetic methylotrophs usually preserve intact PPP to regenerate Ru5P for formaldehyde assimilation (Bennett et al., 2018a; Gonzalez et al., 2018; Leßmeier et al., 2015; Müller et al., 2015a; Whitaker et al., 2017; Witthoff et al., 2015). However, problems arise when cells are cultivated using methanol and another cosubstrate such as yeast extract, glucose, or ribose. The intact PPP allows cells to grow on the co-substrate even methanol is not utilized. Therefore, engineering PPP is necessary to construct a methanol-dependent strain. C. glutamicum can grow on ribose as the sole carbon source through PPP. By introducing a heterologous gene encoding xylose isomerase (xylA), the recombinant strain is capable of utilizing xylose as the sole carbon source (Blombach and Seibold, 2010; Kawaguchi et al., 2006). Pentose metabolism is dependent on two key enzymes of PPP, ribose phosphate isomerase (RpiB) and ribulose phosphate epimerase (Rpe), which mediate the conversion among xylulose 5-phosphate (X5P), Ru5P and R5P for carbon rearrangement and nucleotide synthesis (Fig. 1A and Fig. 1B). Knockout of rpiB could block the conversion between Ru5P and R5P and consequently abrogate the capacity of cells to grow on ribose or xylose individually. Nevertheless, growing on ribose and xylose as co-substrates is feasible (Fig. 1C). Noteworthily, if methanol oxidation and RuMP genes are introduced into the rpiB-deficient background, methanol can be co-

17

utilized with xylose by cells. Under this condition, methanol and xylose can provide C1 (formaldehyde) and C5 (Ru5P) intermediates, respectively, for generation of C6 intermediate, fructose 6-phosphate (F6P). The resulting F6P will enter the lower glycolysis or be rearranged through PPP to generate R5P for nucleotide synthesis and Ru5P for formaldehyde assimilation (Fig. 1D). Since methanol utilization is indispensable for cell growth in this case, construction of the methanol-dependent synthetic methylotroph is accomplished and ALE can be applied to evolve the methanol utilization capacity.

3.2. Engineering and evolution of xylose utilization pathway Prior to engineering xylose utilization pathway, the formaldehyde detoxification pathway was blocked in C. glutamicum by deleting genes encoding mycothiol-dependent formaldehyde dehydrogenase (AdhE) and acetaldehyde dehydrogenase (Ald) that catalyze oxidation of formaldehyde (Witthoff et al., 2013). The formaldehyde detoxification pathway deficient C. glutamicum strain MX-1 was used as a background in the subsequent experiments (Table 1). Then we attempted to knock out rpiB in strain MX-1, whereas no positive mutants were obtained. It was speculated that rpiB knockout caused growth defect to cells, making screening of positive mutants very difficult. According to our design, the rpiBdeleted mutant can grow on ribose and xylose mixture if a heterologous xylA gene is introduced (Fig. 1C). The xylA from E. coli was then inserted into pXMJ19 vector under the control of the IPTG-inducible promoter Ptac and transformed into strain MX-1, resulting in strain MX-2 that could grow on xylose (Fig. 2A). To knock out rpiB in strain

18

MX-2 and avoid growth defect, ribose and xylose were used as carbon sources during the whole gene knockout process. The rpiB gene was successfully deleted in strain MX-2, producing strain MX-3. As expected, strain MX-3 did not grow on ribose or xylose as the sole carbon source but could co-utilize ribose and xylose to grow (Fig. 2A). However, its cell growth on ribose and xylose was much slower compared to strain MX-2 on the same carbon sources, suggesting its xylose utilization pathway or PPP was not optimized for pentose utilization. To prepare the rpiB-deleted strain for methanol-dependent cell growth, it is necessary to adapt its metabolism to xylose utilization for more efficient generation of R5P and Ru5P. Therefore, ALE was applied to accelerate the cell growth of strain MX-3 on ribose and xylose. Strain MX-3 was first inoculated into CGXII minimal medium supplemented with 4 g/L ribose and 4 g/L xylose and cultivated aerobically. Cells were transferred into fresh medium with the same carbon sources every 24 h to continue the cultivation. As shown in Fig. 2B, the specific growth rate of strain MX-3 on ribose and xylose gradually increased from 0.03 h-1 to 0.15 h-1 after 10 passages of ALE (≈ 32 generations). Finally, an evolved strain that grew fast on ribose and xylose was screened and designated as strain MX-4.

3.3. Introduction of methanol utilization pathway The next essential step for constructing a methanol-dependent C. glutamicum strain is to introduce methanol utilization pathway in strain MX-4. NAD-dependent Mdh and RuMP pathway were selected to construct methanol utilization pathway in this study since they have been widely used in synthetic methylotrophs (Bennett et al., 2018a;

19

Gonzalez et al., 2018; Leßmeier et al., 2015; Müller et al., 2015a; Meyer et al., 2018; Whitaker et al., 2017; Witthoff et al., 2015). In addition to the activator-dependent Mdhs from native methylotroph B. methanolicus (Krog et al., 2013), two activator-independent Mdhs, Mdh from B. stearothermophilus DSM 2334 (MdhBs2334) and Mdh2 variant (A26V, A31V, A169V) from C. necator N-1 (Mdh2Cn), have been reported and expressed in E. coli for methanol oxidation (Sheehan et al., 1988; Whitaker et al., 2017; Wu et al., 2016). Therefore, we aimed to identify a more suitable Mdh to engineer methanol-dependent C. glutamicum. MdhBs2334, Mdh2Cn, and Mdh3 and its activator from B. methanolicus MGA3 (Mdh3-ActBm) were expressed and compared in the formaldehyde detoxification pathway deficient strain MX-1 using pEC-XK99E vector with the IPTG-inducible promoter Ptrc. Obviously, the recombinant MX-6 expressing MdhBs2334 showed the highest methanol oxidation activity in crude extract compared to strains MX-5 and MX-7 expressing Mdh3-ActBm and Mdh2Cn, respectively (Fig. 2C). Different IPTG concentrations were then tested and 1 mM IPTG was chosen in the subsequent experiments for the best performance on inducing Mdh expression. Since wild-type C. glutamicum strains do not possess RuMP enzymes, heterologous enzymes from methylotroph B. methanolicus MGA3 (Hps-PhiBm) and non-methylotroph B. subtilis 168 (Hps-PhiBs168), which were proven to be functional for formaldehyde assimilation (Leßmeier et al., 2015; Yasueda et al., 1999), were introduced into C. glutamicum, respectively. The native hps-phi cassettes were cloned from B. methanolicus MGA3 and B. subtilis 168 genomic DNAs and inserted into pEC-XK99E-mdhBs2334 under the control of a constitutive promoter PP5, respectively. The recombinant strain MX-8 expressing Hps-PhiBm showed higher formaldehyde assimilation activity in crude extract,

20

which was approximately 2-fold of MX-9 expressing Hps-PhiBs168 (Fig. 2D). Considering the enzyme assay results, Mdh from B. stearothermophilus DSM 2334 (MdhBs2334) and RuMP enzymes from B. methanolicus MGA3 (Hps-PhiBm) were selected and expressed in strain MX-4, generating strain MX-10.

3.4. ALE to accelerate cell growth of methanol-dependent C. glutamicum Strain MX-10 that was deficient in Ru5P-R5P conversion and equipped with methanol utilization enzymes was suggested to be the methanol-dependent C. glutamicum strain according to our design. Its cell growth on xylose- and methanolxylose-containing CGXII minimal medium was then tested. Consistent with our prediction, cell growth was only observed when methanol was present in the medium (Fig. S1). Although the methanol-dependent cell growth rate of strain MX-10 was very low, this promoted us to further accelerate its methanol utilization and cell growth. ALE is a useful metabolic engineering strategy for enabling the development of novel characteristics in microorganisms. Recently, this strategy has been successfully applied to enhance the cell growth performance of C. glutamicum on xylose (Radek et al., 2017). Therefore, we hypothesized that ALE could promote the acquisition of beneficial mutations that will improve cell fitness of the methanol-dependent C. glutamicum. Continuous passage cultivation of strain MX-10 using CGXII minimal medium supplemented with methanol and xylose as carbon sources was then conducted. The ALE experiment continued for 206 days and total 14 passages were cultivated (≈ 27 generations). The OD600nm values at the initial and final time points of each passage of

21

ALE were used to determine the specific growth rate on methanol and xylose, which was improved from 0.0006 h-1 to 0.0121 h-1, demonstrating a 20-fold increase (Fig. 3A). Then an evolved strain with improved cell growth on methanol and xylose was isolated and designated as MX-11. Compared to the unevolved strain MX-10, the evolved strain MX-11 showed significantly improved cell growth when methanol and xylose was used as co-substrates. A final OD600nm of 3.82 ± 0.08 (1.17 ± 0.03 gCDW/L) was obtained by strain MX-11 with a specific growth rate of 0.03 h-1 during exponential growth (Fig. 3B), which is within the same range as very recently reported methanolgluconate co-metabolizing E. coli (0.017-0.081 h-1) (Meyer et al., 2018). Given that no cell growth was observed using xylose as the sole carbon source (Fig. 3B), methanol was an indispensable carbon source for strain MX-11. The results support our hypothesis that the engineered C. glutamicum is a methanol-dependent strain and its cell growth on methanol can be enhanced through ALE. It is also supposed that the enhanced cell growth was caused by improved methanol utilization. During the cultivation, strain MX11 consumed 3.10 g/L methanol (96.90 mM) and 3.80 g/L xylose (25.32 mM), demonstrating that methanol and xylose were co-metabolized at an average mole ratio of 3.83:1 (Fig. 3C). The specific methanol uptake rate during exponential growth reached 0.86 mmol/gCDW·h, which is above that of previously published methanol-yeast extract co-metabolizing E. coli (0.019 g/gCDW·h corresponding to 0.59 mmol/gCDW·h) (Whitaker et al., 2017). However, there is still room for improvement considering that native methylotrophs and the very recently reported methanol-gluconate co-metabolizing E. coli utilize methanol at approximately 15 mmol/gCDW·h during exponential growth (Meyer et al., 2018; Pluschkell and Flickinger, 2002). In addition, we determined the

22

biomass yield of strain MX-11 from methanol and xylose to be 0.16 gCDW/gS, which is within the same range as previously reported methanol-yeast extract co-metabolizing E. coli (0.10-0.37 gCDW/gS) (Whitaker et al., 2017). It is worth noting that xylose was almost used up at 72 h by strain MX-11, thus the subsequent cell growth seemed to be mainly based on methanol utilization (Fig. 3C). To test the possibility of pure methylotrophic growth, strain MX-11 cultivated in methanolxylose-containing CGXII minimal medium was used as a seed culture to inoculate methanol-containing CGXII minimal medium. Unfortunately, no growth was observed after 144 hours’ cultivation (Fig. S2). By comparing the relative abundance of intracellular R5P/Ru5P/X5P of strain MX-11 at 72 h and 120 h, we observed higher R5P/Ru5P/X5P level when xylose was almost exhausted (Fig. S3). The results suggested that xylose was converted to C5 intermediates and accumulated in cells, which might support the successive methanol assimilation and growth.

3.5. Incorporation of 13C-methanol into intracellular metabolites and biomass 13

C-methanol labeling approach has been chosen for examining whether synthetic

methylotrophs could assimilate methanol as a carbon source (Bennett et al., 2018a; Gonzalez et al., 2018; Leßmeier et al., 2015; Müller et al., 2015a; Meyer et al., 2018; Wang et al., 2017; Whitaker et al., 2017; Witthoff et al., 2015). In this study, strain MX11 was cultivated in CGXII minimal medium supplemented with

13

C-methanol and

unlabeled xylose for 120 h and intracellular metabolites were analyzed (Fig. 4A).

13

C-

labeled PPP metabolites, including R5P/Ru5P/X5P and erythrose 4-phosphate (E4P) were detected. We observed that 19.8% R5P/Ru5P/X5P pool contained M+1 labeling and

23

15.3% contained M+2 labeling, suggesting 10.1% average carbon labeling (Fig. 4B and Fig. 4C). Regarding E4P, 14.1% of the pool contained M+1 labeling, 52.2% contained M+2 labeling, 10.1% contained M+3 labeling, and 8.0% was completely labeled (Fig. 4B). Compared to R5P/Ru5P/X5P, a much higher average carbon labeling percentage was observed for E4P (45.9%) (Fig. 4C). Regarding to hexulose 6-phosphate (H6P) and F6P, the downstream intermediates of methanol assimilation, on average 26.9% of carbons were derived from

13

C-methanol (Fig. 4C), which was similar to the 23.9%

average carbon labeling of H6P obtained by the methanol-gluconate co-metabolizing E. coli (Meyer et al., 2018). This value was above the theoretical expected stoichiometric value of 16.7%, which was consistent with the exceeded equimolar consumption of methanol and xylose mentioned above. Production of multiple carbons labeled R5P/Ru5P/X5P, E4P, and H6P/F6P suggest that the RuMP pathway is actually cycling and the formaldehyde accepter is provided not only through xylose utilization but also through RuMP cycle. Glycolysis and TCA cycle intermediates were also labeled by carbons derived from 13

C-methanol. We measured that 43.6% and 12.4% of 3-phosphoglycerate (3PG) pool

contain

M+1

and

M+2

labeling,

respectively.

Similarly,

37.5%

of

the

phosphoenolpyruvate (PEP) pool contained M+1 labeling and 14.1% contained M+2 labeling. Interestingly, pyruvate, the downstream intermediate of 3PG and PEP, showed a significantly increase in 13C labeling level. Up to 24.2% of pyruvate pool contained M+1 labeling, 28.2% contained M+2 labeling, and 16.5% was completely labeled (Fig. 4B). The average carbon labeling percentage of pyruvate reached 43.4% (Fig. 4C). The C6 TCA intermediate citrate/isocitrate was labeled by up to four

24

13

C-methanol-derived

carbons and its average carbon labeling percentage was 17.3%. Similar average carbon labeling levels were observed for α-ketoglutarate, succinate, fumarate, and malate, which ranged from 27.5% to 31.9%. Since carbons derived from methanol passed through RuMP pathway and glycolysis and entered the TCA cycle, they could be used to produce carbon skeleton and energy for biosynthesis. Detection of

13

C-labeled amino acids and cofactors confirmed this

hypothesis. Total fourteen amino acids including structural isomers isoleucine and leucine were found to be labeled by 13C-methanol-derived carbons. Among them, alanine, glutamine, proline, and histidine had completely labeled isotopomers. Glutamine and histidine showed higher average carbon labeling levels (54.5% and 63.9%, respectively) than other amino acids (Fig. 4C). Glutamate is an important product of C. glutamicum fermentation. M+1, M+2, M+3, and M+4 mass isotopomers of glutamate were detected and exhibited relative abundances of 37.3%, 29.9%, 12.2%, and 2.7%, respectively (Fig. 4B). These demonstrate that the methanol-dependent strain MX-11 possesses the potential for producing glutamate from methanol. In addition to the aforementioned metabolites, adenylates (ATP, ADP, and AMP), NAD(P)H, and FADH2 were also labeled by

13

C-methanol-derived carbons. For example, 23.8% of AMP pool contained

M+1 labeling, 39.7% contained M+2 labeling, 19.2% contained M+3 labeling, 6.9% contained M+4 labeling, and 2.1% contained M+5 labeling, suggesting 19.9% average carbon labeling (Fig. 4B and Fig. 4C). Besides the aforementioned difference in labeling between pyruvate and 3PG, different labeling patterns were also observed for glutamate/glutamine, aspartate/asparagine, and NADH/NADPH, which were expected to share similar labeling pattern (Fig. 4B). In a previous study evaluating different co-

25

substrates

for

methanol

assimilation,

such

different

labeling

patterns

of

glutamate/glutamine and aspartate/asparagine were also detected (Gonzalez et al., 2018). To further confirm that methanol was utilized by methanol-dependent strain for cell growth, biomass samples were hydrolyzed and 13C-labeling of proteinogenic amino acids were analyzed. All the detected amino acids were

13

C-labeled, including completely

labeled alanine (Fig. 5A). The average carbon labeling levels of these amino acids were between 15% and 25% (Fig. 5B). Taken together, these results again suggest that methanol was assimilated by the engineered and evolved methanol-dependent C. glutamicum to produce building blocks, high-energy carriers, and cofactors necessary for cellular metabolism.

3.6. Glutamate production by methanol-dependent C. glutamicum To demonstrate that the methanol-dependent strain has the capability to produce industrial relevance chemicals using methanol as a carbon source, glutamate that represents the largest product segment within the amino acid market was selected as a proof-of-concept example. Glutamate is a native product of C. glutamicum ATCC 13032 but its production requires inducing treatments such as biotin limitation or penicillin G addition (Wang et al., 2018a). The methanol-dependent C. glutamicum MX-11 was cultivated in CGXII minimal medium supplemented with methanol and xylose (4 g/L for each carbon source). Penicillin G was added into the culture at the mid-exponential phase when the OD600nm reached approximately 2.3. After the inducing treatment, cell growth was hindered, whereas extracellular glutamate accumulation was observed. At the end of fermentation, 90 mg/mL extracellular glutamate was produced. In absence of the inducer

26

penicillin G or the crucial substrate methanol, no extracellular glutamate was detected (Fig. 6). The results demonstrate the methanol-dependent biosynthesis of useful chemicals.

3.7. Genome sequencing of evolved methanol-dependent strains To identify the genetic mutations responsible for the improved growth and methanol utilization of evolved strains, whole-genome sequencing of strain MX-4 from the first round of ALE and three mutants with similar phenotypes from the second round of ALE (including strain MX-11) was conducted. Three missense mutations were detected in strain MX-4, which are altRI146S (encoding multi-function regulator of carbohydrate metabolism), cgl2030P179S (encoding predicted ATPase with chaperone activity), and ctaET145A (encoding cytochrome c oxidase subunit III) (Table 2). Additional, six missense mutations were detected in all the three evolved strains from the second round of ALE (including strain MX-11) (Table 2). No mutations were found in the xylA, mdh, hps, and phi expressing plasmids in these strains. AltR serves as a multi-function regulator of carbohydrate metabolism, which controls the expression of xylulokinase gene xylB, alcohol dehydrogenase gene adhA, succinyl-CoA synthetase operon sucCD, etc. (Auchter et al., 2011; Cho et al., 2010; Laslo et al., 2012). XylB catalyzes phosphorylation of xylulose and is necessary for xylose utilization (Blombach and Seibold, 2010). AdhA is not only involved in ethanol metabolism but also functions as a native methanol dehydrogenase that catalyzes oxidation of methanol to formaldehyde in C. glutamicum (Witthoff et al., 2013). It is possible that mutation of AltR affected expression of adhA and xylB and improved co-

27

utilization of methanol and xylose consequently. To validate this hypothesis, qPCR was conducted to determine the relative transcription levels of xylB and adhA in strains MX-3 and MX-4. As expected, the transcription levels of xylB and adhA in strain MX-4 are 7.7fold and 5.8-fold higher, respectively, than those in MX-3 (Fig. S4). SucCD is a TCA cycle enzyme and catalyzes the interconversion of succinyl-CoA and succinate. Low TCA cycle activity or even interrupted TCA cycle was observed during methylotrophic growth (Müller et al., 2015b; Meyer et al., 2018). It is thus speculated that regulation of sucCD operon by mutated AltR alters TCA cycle activity and benefits methanoldependent growth. Another two transcriptional regulator encoding genes, mtrA (cgl0754) and uriR (cgl1367), were mutated during the second round of ALE. MtrA is the response regulator of the two-component signal transduction system MtrAB, which influences the expression of genes involved in cell wall metabolism, cell division, and osmoregulation. MtrAB also regulates expression of glutaredoxin-like protein NrdH and NAD+ synthetase NadE that participate in maintaining intracellular redox state (Brocker et al., 2011). Compared with glucose catabolism, more NADH is generated by methanol catabolism. Balancing the NADH/NAD+ level has been proven to be beneficial for methanol oxidation in vitro (Price et al., 2016). MtrA mutation may favor methanol utilization by changing the regulation pattern of MtrAB. The third mutated transcriptional regulator is the repressor of uridine and ribose utilization genes, UriR (Brinkrolf et al., 2008; Nentwich et al., 2009). Since it directly controls uridine and ribose transport and catabolism, its function in methanol and xylose co-utilization was unknown.

28

Finally, we identified a mutation in O-acetylhomoserine sulfhydrylase gene cgl0653 (metY) that is responsible for methionine and cysteine metabolism. Recently, it has been reported that the enzymatic side reaction of MetY contributes to methanol toxicity and mutation of MetY increases methanol tolerance of C. glutamicum (Leßmeier and Wendisch, 2015). The mutation in MetY detected here may benefit methanol-dependent growth by improving cell adaptability in methanol-containing medium. The remaining three missense mutations happened in hypothetical or uncharacterized proteins with unknown function, making interpretation of these mutations difficult (Table 2).

4. Discussion Considering that methanol is a promising one-carbon substrate for biomanufacturing but native methylotrophs have limitations in industrial applications, great effects have been devoted to develop synthetic methylotrophs for converting methanol to fuels and chemicals (Bennett et al., 2018b; Whitaker et al., 2015). Theoretically, introduction of only three heterologous enzymes (i.e. Mdh, Hps, and Phi) is required to achieve synthetic methylotrophy (Zhang et al., 2017). However, according to recent studies, it is truly not the case since autonomous growth on methanol as the sole carbon source cannot be achieved by introducing Mdh, Hps, and Phi into E. coli or C. glutamicum (Müller et al., 2015a; Whitaker et al., 2017; Witthoff et al., 2015). Therefore, it is important to identify the limiting factors of methanol utilization in synthetic methylotrophs. We propose that combination of ALE and inverse metabolic engineering will help to identify the mutations responsible for improved methanol utilization and cell growth, which in turn

29

guides the optimization of synthetic methylotrophs. For this purpose, construction of a methanol-dependent strain is essential. In this study, the industrial workhorse C. glutamicum was engineered by modifying its PPP and introducing RuMP-based methanol utilization pathway to serve as a methanol-dependent strain. Although co-utilization of methanol and xylose was still required for cell growth, methanol is an indispensable carbon source for this strain. Notably, ALE was successfully applied to improve the strain’s performance on methanol utilization, which led to a 20-fold increase in cell growth rate on the mixed carbon sources of methanol and xylose. The engineered and evolved methanol-dependent strain MX-11 showed excellent methanol utilization capability. Methanol and xylose were coutilized by strain MX-11 at an average mole ratio of 3.83:1 (96.90 mM methanol and 25.32 mM xylose) in minimal medium. In the previously engineered methylotrophic C. glutamicum, methanol and glucose were co-utilized at an average mole ratio of 1.45:1 (approximately 80 mM methanol and 55 mM glucose) in minimal medium (Witthoff et al., 2015). For synthetically methylotrophic E. coli, co-utilization of methanol and glucose was reported with an average mole ratio of 0.14:1 (38.3 mM methanol and approximately 275 mM glucose) (Bennett et al., 2018a). The excellent methanol utilization capability of strain MX-11 was further validated by 13C-labeling experiments. Strain MX-11 incorporated up to 63% methanol into intracellular metabolites (such as building blocks, high-energy carriers, and cofactors) and biomass components (such as proteinogenic amino acids), which was above recently reported synthetically methylotrophs (Bennett et al., 2018a; Meyer et al., 2018; Witthoff et al., 2015).

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Two rounds of ALE were conducted to obtain the best mutant MX-11, which we believe both are important. It has been well characterized that PPP plays a crucial role in methanol utilization by providing Ru5P for formaldehyde assimilation (Bennett et al., 2018a). The rpiB-deficient and xylA-expressing strain MX-3 grew slowly on ribose and xylose, suggesting that the xylose utilization pathway and PPP were non-optimized. To strengthen these pathways, the first round of ALE using ribose and xylose as cosubstrates was performed. This step was supposed to enhance Ru5P regeneration. The second round of ALE using methanol and xylose as co-substrates significantly accelerated the methanol utilization rate of the methanol-dependence strain. Based on these results, it is reasonable to speculate that some bottlenecks have been resolved. Genome sequencing of evolved methanol-dependent strains sheds light on identifying the cryptic limiting factors. So far, the notorious Mdh kinetics, NADH excess, and the aforementioned inefficient Ru5P regeneration are proven to restrain methanol utilization in synthetic methylotrophs (Bennett et al., 2018a; Meyer et al., 2018; Price et al., 2016; Whitaker et al., 2017; Wu et al., 2016). It is also suggested that regulation plays an important role in methylotrophy but mechanisms need to be uncovered (Gonzalez et al., 2018). The detected mutations in this study seem to be in correspondence with the existing knowledge and hypothesis. One third of the mutations happened in transcriptional regulator genes. The regulation network of the three regulators covers methanol oxidation, pentose uptake, TCA activity, and intracellular redox balance (Auchter et al., 2011; Brinkrolf et al., 2008; Brocker et al., 2011; Cho et al., 2010; Laslo et al., 2012; Nentwich et al., 2009). Additional mutations involved in energy metabolism and methanol tolerance were also detected (Leßmeier and Wendisch, 2015). Further

31

inverse metabolic engineering towards the methanol-dependent strain will assist elucidation of the underlying mechanisms and guide the optimization of synthetic methylotrophs. ALE of strain MX-11 is still ongoing. Screening better mutants and reintegrating the PPP will likely lead to synthetic methylotrophs capable of growing on methanol as the sole carbon source. The methanol-dependent strain presented in this study can also serve as a screening platform for mutagenesis library of Mdh, since higher Mdh activity is supposed to result in higher cell growth rate on methanol. In conclusion, a methanol-dependent C. glutamicum was rationally designed and experimentally constructed in this study. Subsequent ALE significantly accelerated the strain’s methanol consumption rate. The methanol-dependent strain is capable of converting methanol to biomass and useful chemicals. Furthermore, the approach used in this study is highly feasible and provides access to study the mutations accumulated during ALE, which will help to identify the limiting factors of methanol utilization and guide the construction of synthetic methylotrophs for autonomous growth on methanol as the sole carbon source. In the time this manuscript was written, a study describing design and construction of methanol-dependent E. coli appeared (Meyer et al., 2018), employing a similar idea but a distinct strategy. In that study, methanol assimilation to biomass was coupled to modified gluconate catabolism. Gluconate was channeled to Ru5P via decarboxylation but its entry into the Entner–Doudoroff pathway is blocked by the edd knockout. Coutilization of methanol and gluconate was then improved by ALE with the help of another co-substrate pyruvate (Meyer et al., 2018). Both the similarities (i.e. the carbon source co-utilization strategy) and differences (i.e. the choice of co-substrate for

32

methanol assimilation, genes to knock out, co-substrates for ALE, and bacterial hosts) report significant improvement in the area of developing synthetic methylotrophs. Notably, completely different mutations were accumulated in the chromosomes of methanol-dependent E. coli (Meyer et al., 2018) and methanol-dependent C. glutamicum during ALE. However, mutations detected in the two independent studies both indicate the importance of the intracellular NAD+/NADH ratio and substrate uptake on methanol utilization. As mentioned above, our study reports methanol-dependent production of amino acids in addition to methanol-dependent cell growth.

Competing Interests The authors declare no competing financial interests.

Acknowledgements This work was supported by the National Natural Science Foundation of China (31700044), the Key Research Program of Chinese Academy of Sciences (ZDRW-ZS2016-2), the International Partnership Program of Chinese Academy of Sciences (153D31KYSB20170121), the Special Program of Talents Development for Excellent Youth Scholars in Tianjin, and the first Special Support Plan for Talents Development and High-level Innovation and Entrepreneurship Team of the Tianjin Municipal City.

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Fig. 1. Strategies to construct the methanol-dependent C. glutamicum. (A) C. glutamicum ATCC 13032 utilizes ribose as the sole carbon source. (B) C. glutamicum MX-2 utilizes xylose as the sole carbon source. (C) C. glutamicum MX-3 co-utilizes ribose and xylose as the carbon sources. (D) C. glutamicum MX-10 co-utilizes methanol and xylose as the carbon sources. Enzymes: ribose phosphate isomerase (RpiB), ribulose phosphate epimerase (Rpe), transketolase (Tkt), transaldolase (Tal), xylose isomerase (XylA), xylulokinase (XylB), methanol dehydrogenase (Mdh), 3-hexulose-6-phosphate synthase (Hps),

6-phospho-3-hexuloisomerase

(Phi),

mycothiol-dependent

formaldehyde

dehydrogenase (AdhE), acetaldehyde dehydrogenase (Ald). Metabolites: ribose 5phosphate

(R5P),

ribulose

5-phosphate

(Ru5P),

xylulose

5-phosphate

(X5P),

glyceraldehyde 3-phosphate (G3P), erythrose 4-phosphate (E4P), sedoheptulose 7phosphate (S7P), fructose 6-phosphate (F6P), hexulose 6-phosphate (H6P).

Fig. 2. Engineering of xylose and methanol utilization pathway in C. glutamicum. (A) Cell growth of C. glutamicum strains using different carbon sources (4 g/L for each carbon source). Glucose (Glc), ribose (Rib), xylose (Xyl). Error bars indicate standard deviations from three parallel experiments. (B) ALE of C. glutamicum MX-3 in CGXII minimal medium supplemented with ribose (Rib) and xylose (Xyl) as the carbon sources. C. glutamicum MX-3 was first cultivated in CGXII minimal medium supplemented with

40

ribose and xylose (4 g/L for each carbon source). After cultivated for 24 h at 30°C and with shaking at 220 rpm, the culture was used as a seed to inoculate fresh medium with an initial OD600nm of 0.1, which was then incubated under the same conditions. (C) Optimization of Mdh from different resources in C. glutamicum. Mdh3 and Act from B. methanolicus MGA3 (Mdh3-ActBm), Mdh from B. stearothermophilus DSM 2334 (MdhBs2334), Mdh2 variant (A26V, A31V, A169V) from C. necator N-1 (Mdh2Cn). Error bars indicate standard deviations from three parallel experiments. (D) Optimization of RuMP enzymes from different resources in C. glutamicum. Hps and Phi from B. methanolicus MGA3 (Hps-PhiBm), Hps and Phi from B. subtilis 168 (Hps-PhiBs168). Error bars indicate standard deviations from three parallel experiments.

Fig. 3. ALE for improving methanol uptake and growth of methanol-dependent C. glutamicum. (A) ALE of strain MX-10 in CGXII minimal medium supplemented with methanol and xylose. The specific growth rate for each passage of ALE (the black curve) was calculated using the OD600nm values at the initial and final time points. C. glutamicum MX-10 was first cultivated in CGXII minimal medium supplemented with methanol and xylose (4 g/L for each carbon source). To avoid evaporation of methanol, the shake flasks were covered with a sealing membrane. After cultivation at 30°C with shaking at 220 rpm until the OD600nm reached over 1.0, the culture was used as a seed to inoculate fresh medium with an initial OD600nm of 0.5, which was then incubated under the same conditions. (B) Growth curve of methanol-dependent C. glutamicum MX-11. Strain MX-11 was cultivated using CGXII minimal medium supplemented with methanol and xylose (Xyl) or only xylose as the carbon source(s). (C) Substrate uptake of

41

methanol-dependent C. glutamicum MX-11. Strain MX-11 was cultivated using CGXII minimal medium supplemented with methanol and xylose (Xyl) as the carbon sources. An evaporation control without inoculation of strain MX-11 was conducted simultaneously. Error bars indicate standard deviations from three parallel experiments.

Fig. 4.

13

C-labeling in intracellular metabolites from the methanol-dependent C.

glutamicum MX-11. Strain MX-11 was cultivated in CGXII minimal medium supplemented with 13C-methanol and xylose (4 g/L for each carbon source). (A) Central metabolism map of strain MX-11. (B) Relative abundance of intracellular metabolite and amino acid mass isotopomers at 120 h. (C) Average carbon labeling of intracellular metabolites and amino acids at 120 h. Intracellular metabolites: hexulose 6-phosphate (H6P), fructose 6-phosphate (F6P), fructose 1,6-diphosphate (F16dP), ribulose 5phosphate (Ru5P), ribose 5-phosphate (R5P), xylulose 5-phosphate (X5P), erythrose 4phosphate (E4P), glyceraldehyde 3-phosphate (G3P), dihydroxyacetone phosphate (DHAP), 3-phosphoglycerate (3PG), 2-phosphoglycerate (2PG), phosphoenolpyruvate (PEP), pyruvate (Pyr), citrate (Cit), isocitrate (Ici), α-ketoglutarate (α-KG), succinate (Suc), fumarate (Fum), malate (Mal), oxaloacetate (Oxa). Error bars indicate standard deviations from three parallel experiments.

Fig. 5.

13

C-labeling in proteinogenic amino acids from the methanol-dependent C.

glutamicum MX-11. Strain MX-11 was cultivated in CGXII minimal medium supplemented with 13C-methanol and xylose (4 g/L for each carbon source). (A) Relative abundance of proteinogenic amino acid mass isotopomers at 120 h. (B) Average carbon

42

labeling of proteinogenic amino acids at 120 h. Error bars indicate standard deviations from three parallel experiments.

Fig. 6. Glutamate production from methanol and xylose by C. glutamicum MX-11. C. glutamicum MX-11 was cultivated in CGXII minimal medium supplemented with only xylose or mixture of methanol and xylose (4 g/L for each carbon source) at 30°C and with shaking at 220 rpm. To induce glutamate production, penicillin G was added to a final concentration of 60 U/ml when the OD600nm of the culture reached approximately 2.3. Error bars indicate standard deviations from three parallel experiments.

Table 1. Strains and plasmids used in this study Strain or plasmid

Descriptiona

Reference or source

Strain E. coli DH5α MG1655

General cloning host Wild-type strain, source for xylA gene

TaKaRa Lab collection

Wild-type strain Derivative of strain ATCC 13032 with its ald gene deleted Derivative of strain Δald with its adhE gene deleted Derivative of strain MX-1 harboring pXMJ19xylA Derivative of strain MX-2 with its rpiB gene deleted Mutant of strain MX-3 that grows fast using ribose and xylose as carbon sources Derivative of strain MX-1 harboring pECXK99E-mdh3-actBm Derivative of strain MX-1 harboring pECXK99E-mdhBs2334

ATCC (Wang et al., 2018b) This study

C. glutamicum ATCC 13032 Δald MX-1 MX-2 MX-3 MX-4 MX-5 MX-6

43

This study This study This study This study This study

MX-7 MX-8 MX-9 MX-10 MX-11 B. methanolicus MGA3 B. stearothermophilus DSM 2334 B. subtilis 168 Plasmid pK18mobsacB pXMJ19 pEC-XK99E

pK18mobsacB-ΔadhE pK18mobsacB-ΔrpiB pXMJ19-xylA pEC-XK99E-mdh3-actBm pEC-XK99E-mdhBs2334 pEC-XK99E-mdh2Cn pEC-XK99E-mdhBs2334hps-phiBm

pEC-XK99E-mdhBs2334hps-phiBs168

Derivative of strain MX-1 harboring pECXK99E-mdh2Cn Derivative of strain MX-1 harboring pECXK99E-mdhBs2334-hps-phiBm Derivative of strain MX-1 harboring pECXK99E-mdhBs2334-hps-phiBs168 Derivative of strain MX-4 harboring pECXK99E-mdhBs2334-hps-phiBm Mutant of strain MX-10 that grows fast using methanol and xylose as carbon sources Wild-type strain, source for mdh3-actBm and hpsphiBm Wild-type strain, source for mdhBs2334

This study

Wild-type strain, source for hps-phiBs168

Lab collection

Gene deletion vector, mob, sacB, KmR

(Schäfer et al., 1994) (Jakoby et al., 1999) (Kirchner and Tauch, 2003) This study This study This study

Expression vector, IPTG-inducible promoter Ptac, CmR Expression vector, IPTG-inducible promoter Ptrc, KmR pK18mobsacB derivative for deleting adhE pK18mobsacB derivative for deleting rpiB pXMJ19 derivative harboring xylA from E. coli, under the control of Ptac pEC-XK99E derivative harboring mdh3 and act genes from B. methanolicus MGA3, under the control of Ptrc pEC-XK99E derivative harboring mdh gene from B. stearothermophilus DSM 2334, under the control of Ptrc pEC-XK99E derivative harboring mdh2 variant (A26V, A31V, A169V) from C. necator N-1, under the control of Ptrc pEC-XK99E derivative harboring mdh gene from B. stearothermophilus DSM 2334, under the control of Ptrc, and hps and phi genes from B. methanolicus MGA3, under the control of constitutive promoter PP5 pEC-XK99E derivative harboring mdh gene from B. stearothermophilus DSM 2334, under the control of Ptrc, and hps and phi genes from B. subtilis 168, under the control of constitutive 44

This study This study This study This study Lab collection DSM

This study This study This study This study

This study

a

promoter PP5 Km and Cm represent resistance to kanamycin and chloramphenicol, respectively. R

R

Table 2. Mutations of evolved strains identified by genome sequencing Gene ID

Nucleotide Amino acid alteration alteration T437G I146S

cgl2192

Gene Gene product name atlR Multi-function regulator of carbohydrate metabolism – Predicted ATPase with chaperone activity ctaE Cytochrome c oxidase subunit III

cgl0653

metY

cgl0754

mtrA

cgl1367

uriR

cgl1520 cgl2424

– –

cgl2998

–

cgl0111 cgl2030

O-acetylhomoserine sulfhydrylase Dual regulator of genes involved in cell morphology, antibiotics susceptibility and osmoprotection Transcriptional repressor of uridine utilization and ribose uptake genes Hypothetical protein Uncharacterized membrane protein Hypothetical protein

Strain

C535T

P179S

A433G

T145A

G1256A

G419D

MX-4 & MX-11 MX-4 & MX-11 MX-4 & MX-11 MX-11

C582A

H194Q

MX-11

C584T

T195I

MX-11

A574G C446A

M192V A149E

MX-11 MX-11

G104T

G35V

MX-11

Highlights 

Methanol-dependent strain was rationally designed and experimentally constructed.



Adaptive laboratory evolution improved methanol-dependent cell growth by 20fold.



Methanol was co-utilized with xylose at a high mole ratio of 3.83:1.



Up to 63% 13C-labeling was detected in intracellular metabolites and biomass.

45



Methanol-dependent glutamate production was achieved by engineered C. glutamicum.

46

47

48

49

50