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. 2025 Sep 24;122(12):3393–3405. doi: 10.1002/bit.70072

Succinic Acid Production by Engineered Mannheimia succiniciproducens and Its Use in Chemoenzymatic Poly(Butylene Succinate) Synthesis

Ji Yeon Kim 1,2, Jong An Lee 1,2, Gi Bae Kim 1,2, Youngjoon Lee 1,2, Sang Yup Lee 1,2,3,4,5,6,✉
PMCID: PMC12599488  PMID: 40990211

ABSTRACT

Amid growing concerns about climate change, environmental pollution, and fossil resource depletion, bio‐based chemicals and materials have garnered significant attention. Succinic acid (SA), a four‐carbon dicarboxylic acid, is a key platform chemical for producing industrially important compounds such as poly(butylene succinate) (PBS), a biodegradable polymer. In this paper, we demonstrate the enhanced production of SA using metabolically engineered Mannheimia succiniciproducens PALK (pMS3‐gltA) strain, which achieves high SA yields through enhanced biomass production and optimized metabolic pathways. High‐purity SA, purified from the fermentation broth, was used in an environmentally friendly two‐step polymerization process under mild conditions. In the first step, diethyl succinate was synthesized from SA and ethanol using Amberlyst as a catalyst. This was followed by polymerization with 1,4‐butanediol, catalyzed by Candida antarctica lipase B. The resulting PBS exhibited comparable thermal stability and molecular weight distribution to its petrochemical counterpart, with enhanced biodegradability. By integrating metabolic engineering with sustainable polymer synthesis, this study underscores the potential for an eco‐friendly approach to developing bio‐based polymers, offering a promising solution to the environmental challenges posed by conventional plastics.

Keywords: biodegradability, metabolic engineering, poly(butylene succinate), succinic acid, sustainability

Succinic acid is produced by the fermentation of metabolically engineered Mannheimia succiniciproducens, purified, and used to synthesize poly(butylene succinate) using a two‐step process that involves the esterification of succinic acid and ethanol, followed by enzymatic transesterification of diethyl succinate and 1,4‐butanediol.

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1. Introduction

Growing awareness of climate change, environmental pollution, and fossil resource depletion has intensified the demand for sustainable alternatives to petrochemical‐based chemicals and materials. In response, the bio‐based production of platform chemicals has emerged as a promising solution, offering the potential to reduce greenhouse gas emissions and reliance on nonrenewable fossil resources (Cho et al. 2022; Kim et al. 2023). Among these chemicals, succinic acid (SA), a four‐carbon dicarboxylic acid, is recognized as a versatile precursor for the synthesis of various industrially important chemicals and bio‐based polymers like poly(butylene succinate) (PBS) (Kim et al. 2023).

Bio‐based SA production has been extensively explored using various metabolically engineered microorganisms, including Actinobacillus succinogenes (Der Werf et al. 1997), Corynebacterium glutamicum (Inui et al. 2004), Escherichia coli (Lee et al. 2005), Mannheimia succiniciproducens (Lee et al. 2002), Basfia succiniciproducens (which has a nearly identical genome sequence to M. succiniciproducens; Kuhnert et al. 2010), Saccharomyces cerevisiae (Agren et al. 2013), and Yarrowia lipolytica (Cui et al. 2023). Among these, M. succiniciproducens, a natural SA producer, is distinguished by its high SA yield and productivity under anaerobic conditions (Ahn et al. 2020; Kim et al. 2023). Leveraging its potential, extensive efforts have been made to enhance its metabolic efficiency and SA production capabilities through systems metabolic engineering and bioprocess engineering (Ahn et al. 2018; Ahn et al. 2020; Choi et al. 2016; Kim et al. 2017; Kim et al. 2024; Lee et al. 2016).

PBS, a key industrial polymer derived from SA and 1,4‐butanediol, has gained significant attention for its bio‐based sustainable manufacturing method and biodegradability. The demand for bio‐based plastics like PBS has surged due to growing environmental concerns about plastic pollution (Choi et al. 2023). PBS is highly valued for its excellent mechanical properties, thermal stability, and biodegradability, making it a promising alternative to conventional plastics, particularly in packaging, agriculture, and biomedical applications (Xu and Guo 2010a). PBS is typically processed through conventional methods such as extrusion, injection molding, and film blowing (Gigli et al. 2016; Xu and Guo 2010b). To enhance its properties, chemically modified PBS materials have been developed to improve mechanical strength and thermal resistance (Platnieks et al. 2021). However, conventional PBS production faces critical challenges, including reliance on fossil resources for producing its monomers and high production costs under relatively harsh conditions (Platnieks et al. 2021; Savitha et al. 2022).

To address these challenges, this study focuses on enhancing SA production and developing an eco‐friendly PBS synthesis method under mild reaction conditions (Figure 1). A metabolically engineered M. succiniciproducens strain, PALK (pMS3‐gltA), was constructed to enhance SA production by optimizing carbon flux towards TCA cycle. High‐purity SA was then purified from the fermentation broth and used in a two‐step polymerization process to synthesize PBS. This process employed reusable catalysts, Amberlyst and Candida antarctica lipase B (CALB), enabling polymerization under mild conditions to minimize energy consumption and environmental impact. The material properties and biodegradability of the resulting PBS were analyzed and compared to its petrochemical‐derived counterpart. This study underscores the feasibility of combining metabolic engineering with eco‐friendly polymer synthesis to produce sustainable and biodegradable materials. By addressing key challenges in PBS production, it provides valuable insights into the potential for developing bio‐based polymers as viable alternatives to conventional plastics.

Figure 1.

Figure 1

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Schematic overview of PBS production. SA is produced by the fermentation of Mannheimia succiniciproducens, then separated and purified. This high‐purity SA is subsequently polymerized via a two‐step process to synthesize PBS. Upon disposal, PBS can be biodegraded.

2. Materials and Methods

2.1. Plasmid and Strain Construction

The strains, plasmids, and oligonucleotides used in this study are listed in Supporting Information S1: Table S1. To construct pMS3‐gltA, the gltA gene fragment was prepared by polymerase chain reaction with primers P1‐2 using M. succiniciproducens genomic DNA as template. The gltA gene fragment was then inserted into pMS3 digested with EcoRI and KpnI using Gibson assembly (Gibson et al. 2009). Correct vector construction was verified using DNA sequencing. Transformation of M. succiniciproducens with the plasmid vector constructed in this study was carried out following the procedure previously developed (Kim et al. 2008).

2.2. In Silico Analysis

Flux response analysis was performed to calculate the variations in metabolic fluxes (i.e., succinate production flux and specific growth rate) in response to changes in the metabolic flux of citrate synthase. The analysis was performed using the M. succiniciproducens genome‐scale metabolic model which comprises 686 metabolic reactions and 519 metabolites (Kim et al. 2007). The succinate yield was calculated by dividing the succinate production flux by the glucose uptake rate. The simulations were carried out in Python environment (Python Software Foundation, Delaware, U.S.A.) with Gurobi Optimizer 9.0.1 (Gurobi Optimization Inc., Houston, TX, USA). Reading, writing, and manipulation of the COBRA‐compliant SBML files were implemented using COBRApy 0.17.1 (Ebrahim et al. 2013).

2.3. Fed‐Batch Fermentation and Analytical Procedures

The glycerol stocks (15%, w/v) of engineered M. succiniciproducens strain, which was previously stored in a deep freezer at −80°C, were used to preculture cells in a 50 mL tube equipped with CO2 gas inlet and outlet ports. The precultured cells were transferred to a 500 mL Erlenmeyer flask equipped with CO2 gas inlet and outlet ports. The MH5 medium containing per L: 2.78 g yeast extract, 2.78 g polypeptone, 0.18 g NaCl, 0.02 g CaCl2 · 2H2O, 0.2 g MgCl2 · 6H2O, 8.06 g K2HPO4, and 9.15 g NaHCO3 supplemented with 18.02 g/L (100 mM) glucose and charged with CO2 as a headspace gas was used for cultivation. The tubes or flasks were incubated in a static incubator at 39°C. Antibiotics were added to the following concentrations when needed: ampicillin (50 mg/L) and kanamycin (25 mg/L).

Fed‐batch fermentations were carried out in a 6.6 L Bioflo 3000 reactor (New Brunswick Scientific Co., Edison, NJ, USA) with the working volume of 2.5 L. Fermentation was conducted using a chemically defined medium (CDM) containing per L: 1 g NaCl, 0.02 g CaCl2·2H2O, 2 g (NH4)2SO4, 0.5 g alanine, 0.5 g asparagine, 0.005 g biotin, 0.5 g methionine, 0.005 g Ca‐pantothenate, 0.005 g pyridoxine‐HCl, 0.005 g thiamine, 0.2 g MgCl2·6H2O, 1.5 g K2HPO4, 9.997 g NaHCO3, 0.005 g ascorbic acid, 0.5 g aspartic acid, 0.5 g cysteine, 0.005 g nicotinic acid, 0.5 g proline, 0.5 g serine, and 5 mL trace metal solution. The trace metal solution contained per L: 5 mL HCl, 10 g FeSO4·7H2O, 2.25 g ZnSO4·7H2O, 1 g CuSO4·5H2O, 0.5 g MnSO4·5H2O, 0.23 g Na2B4O7·10H2O, and 0.1 g (NH4)6Mo7O24. The CDM was supplemented with 18.02 g/L (100 mM) glucose and/or 4.60 g/L (50 mM) glycerol. Antibiotics were added at the following concentrations when needed: ampicillin (50 mg/L) and kanamycin (25 mg/L). Fed‐batch fermentation was initiated by inoculating 300 mL of precultured broth, giving the initial optical density of 600 nm (OD600) of 0.2–0.3. Temperature and the agitation speed of four flat blade turbine impellers in the bioreactor were controlled at 39°C and 200 rpm, respectively. The pH of the fermentation broth was controlled at 6.5 by automatic addition of a mixture of 1.57 M ammonia and 6.84 M Mg(OH)2 solution. The bioreactor was continuously sparged with industrial‐grade CO2 at a flow rate of 0.2 vvm (sparged gas volume per working volume per min) by a mass flow controller. Fed‐batch fermentations were performed in a semi‐continuous feeding mode, which the feeding solution was supplied into the bioreactor via a peristaltic pump, to maintain the carbon source concentration at 5–15 g/L for substrate inhibition minimization by changing the feeding rate.

The concentrations of glucose, glycerol, and organic acids including acetic, formic, lactic, SA, and pyruvic acids were measured using high performance liquid chromatography (Agilent Technologies, Palo Alto, CA, USA) equipped with UV/visible‐light and refractive index detectors (Agilent). A MetaCarb 87H column (300 mm by 7.8 mm; Agilent) was eluted isocratically with 5 mM H2SO4 at 60°C at a flow rate of 0.6 mL/min. Cell growth was monitored using an Ultraspec 3000 spectrophotometer (Amersham Biosciences, Uppsala, Sweden).

2.4. Separation and Purification of SA From Fermentation Broth

Separation of SA from fermentation broth was carried out following the SA recovery procedure reported previously (Choi et al. 2016; Lee et al. 2019). Briefly, the fermentation broth collected at the end of fermentation was centrifuged at 5600g for 20 min to remove cell debris and treated with Activated Charcoal Norit (Sigma‐Aldrich) with stirring for 1 h at room temperature to remove the pigments existing in the fermentation broth. The Activated Charcoal Norit was separated from the decolorized fermentation broth using 0.2 μm filter paper (Whatman International Ltd., Kent, UK). The decolorized SA fermentation broth was concentrated under 700 mbar at 90°C using a rotary vacuum evaporator (EYELA, Tokyo, Japan) equipped with a condenser (LAUDA‐Brickmann, NJ, USA), vacuum pump Labport N‐820 (KNF Lab, NJ, USA), vacuum controller NVC‐2300B (EYELA), and water bath KSB‐202 (EYELA) to remove water and other impurities with low boiling points. Next, the pH of SA fermentation broth was adjusted to 2.0 using H2SO4 to obtain fully protonated SA, and the precipitated SA was then lyophilized. To eliminate elemental impurities, SA was redissolved in ethanol, and the SA/ethanol solution was filtered via 0.2 μm filter paper (Whatman International Ltd). Finally, the SA crystals were obtained by lyophilizing the filtered solution. Concentrations of various elements remaining in the purified samples were analyzed with inductively coupled plasma‐MS (ICP‐MS) using Agilent ICP‐MS 7700S (Agilent).

2.5. Synthesis and Analysis of PBS

PBS was synthesized by mixing 296 mmol of SA, 2960 mmol of ethanol (Supelco), and Amberlyst 15 (5 wt% of total monomer, Supelco) in a reactor at 78°C with vigorous stirring for 30 h. The reaction mixture was then centrifuged at 5600g for 10 min to remove the catalyst. The supernatant, diethyl succinate, was then used for condensation reaction with 1,4‐butanediol. 28.75 mmol of diethyl succinate and 28.75 mmol of 1,4‐butanediol (Sigma‐Aldrich) were mixed with CALB (10 wt% of total monomer, Sigma‐Aldrich) in a reactor at 90°C with vigorous stirring for 72 h. The product mixture was subsequently dissolved in chloroform and filtered to remove CALB. Polymer was extracted using a solvent extraction method, with chloroform as the solvent and methanol used to precipitate the polymer from the concentrated chloroform solution.

The structure, molecular weight, thermal properties of the polymer were determined by nuclear magnetic resonance (NMR) spectroscopy, gel permeation chromatography (GPC), differential scanning calorimetry (DSC), and thermogravimetric analysis (TGA), respectively. For NMR analysis, a Bruker AVANCE III 600 spectrometer (Bruker, Rheinstetten, Germany) equipped with a BBI probe was used with CDCl3 as the solvent and tetramethylsilane as the internal standard for chemical shift. For GPC, at 40°C, we used a Waters Alliance 2695 Separation Module (Waters) equipped with a Waters 2414 RI detector and two PL Gel columns (Mixed C, 5 μm particles, 30 cm, ID 7.5 mm; Polymer Laboratories, Amherst, MA) with chloroform as an eluent and polystyrene as a standard polymer. For DSC, a DSC 204 F1 Phoenix (Netzsch, Selb, Germany) was used with nitrogen as a protective gas. The T m and ΔH f values were determined from the second heating ramp. Approximately, 5 mg of DSC samples were heated at 150°C for 5 min and were reheated from −70°C to 150°C at a rate of 10°C/min. The position of the maximum peak and the area under the melting curve were used to determine T m and ΔH f, respectively. For TGA, a thermogravimetric analyzer Perseus TG209 F1 Libra (Netzsch, Selb, Germany) was used. Approximately, 10 mg of samples were heated from 30°C to 800°C at a rate of 10°C/min under the nitrogen atmosphere. T d5 of PBS were derived from the TGA curves when 5% of the initial weight was loss. T dmax of PBS were derived from the TGA curves with the maximum degradation rate. The crystal structures of PBS were characterized using a high‐resolution X‐ray diffractometer SMARTLAB XRD (Rigaku, Tokyo, Japan) with the Cu Kα1 (λ = 0.15406 nm) radiation source operated at 45 kV and 200 mA. The X‐ray diffraction (XRD) spectra were collected at 2θ = 5–90° with a scan rate of 5°/min.

2.6. Biodegradation of Synthesized Bio‐PBS and Chem‐PBS

Biodegradation of bio‐PBS and chem‐PBS were conducted by flask cultivation of Pseudomonas fluorescens (KCTC 42821) using each PBS as the sole carbon source. The culture of the bacterium was prepared in lysogeny broth (10 g/L tryptone, 5 g/L yeast extract, and 10 g/L NaCl) using a rotary shaker at 30°C and 200 rpm. Next, a 0.5 mL of seed culture was inoculated into a 50 mL of mineral salt medium [3.25 g/L (NH4)2SO4, 1 g/L KH2PO, 0.2 g/L NaH2PO4, 0.5 g/L MgSO4·7H2O, 0.005 g/L thiamine, and 5 mL/L of trace metal solution containing 10 g/L FeSO4·7H2O, 2.25 g/L ZnSO4·7H2O, 1 g/L CuSO4·5H2O, 0.5 g/L MnSO4·5H2O, 0.23 g/L Na2B4O7·10H2O, 2 g/L CaCl2·2H2O, and 0.1 g/L (NH4)6Mo7O24] in a baffled flask with or without 5 g/L of bio‐PBS or chem‐PBS. The flask culture was conducted under the aerobic condition for 2 weeks in a rotary shaker at 30°C and 200 rpm. Cell growth was monitored at an optical density of 600 nm using an Ultrospec 3000 spectrophotometer (Pharmacia Biotech, Uppsala, Sweden). The weight of the degraded PBS sample was measured after filtration to separate PBS from culture broth, washed in methanol to remove possible debris and oven dried at 70°C. The carbon content of the culture broth was measured before and after 2 weeks of culture using the total organic carbon (TOC) analyzer vario TOC cube (Elementar, Hanau, Germany).

3. Results and Discussion

3.1. Bio‐Based SA Production Using Engineered M. succiniciproducens Strain

M. succiniciproducens naturally produces SA through the strong reductive branch of TCA cycle. The M. succiniciproducens PALK strain, which was previously developed by knocking out the pta, ackA, and ldhA genes encoding phosphate acetyltransferase, acetate kinase, and lactate dehydrogenase, respectively, to eliminate byproduct formation such as acetic acid and lactic acid, was selected as the base strain (Choi et al. 2016; Figure 2a).

Figure 2.

Figure 2

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Metabolic engineering of Mannheimia succiniciproducens for enhanced SA production. (a) Metabolic pathway of the M. succiniciproducens PALK (pMS3‐gltA) strain. (b) Flux response analysis result of gltA overexpression on M. succiniciproducens genome‐scale metabolic model using glucose as a carbon source. Fed‐batch fermentation profile of the M. succiniciproducens PALK (pMS3‐gltA) strain using (c) glucose and (d) glucose and glycerol as dual carbon sources. Symbols: black square, glucose; white square, cell growth; white circle, pyruvate; red circle, SA; white diamond, acetate; cross, glycerol. Abbreviations: ackA, acetate kinase; fdh, formate dehydrogenase; frdABCD, fumarate reductase; fumC, fumarase; gltA, citrate synthase; ldhA, lactate dehydrogenase; mdh, malate dehydrogenase; MQ, menaquinone; MQH2, menaquinol; pckA, phosphoenolpyruvate carboxykinase; pfl, pyruvate formate lyase; ppc, phosphoenolpyruvate carboxylase; pta, phosphate acetyltransferase.

To further enhance SA production, the gltA gene encoding citrate synthase was selected for overexpression. Citrate synthase is a key enzyme that catalyzes the entry of carbon flux into the TCA cycle by converting acetyl‐CoA and oxaloacetate into citrate. Its strategic importance arises from its rate‐limiting role in gram‐negative bacteria, where its activity is inhibited under high NADH levels (Huang et al. 2018; Li et al. 2016; Underwood et al. 2002; Vemuri et al. 2006; Vuoristo et al. 2015). This inhibition often leads to the accumulation of upstream intermediates, such as acetyl‐CoA and pyruvate, reducing carbon flux toward SA production. Overexpressing gltA alleviates this metabolic bottleneck, facilitating greater carbon flux through the TCA cycle and thereby enhancing SA production, as demonstrated in previous studies (Li et al. 2016; Mutyala et al. 2023; Zhu et al. 2013). In M. succiniciproducens, we hypothesized that gltA overexpression increases carbon flux into the TCA cycle, thereby potentially enhancing oxidative flux toward SA production and increasing biomass formation. These dual effects can synergistically contribute to the overall improvement in SA production. Consistent with this hypothesis, in silico simulations using the genome‐scale metabolic model of M. succiniciproducens predicted that gltA overexpression can increase biomass flux without compromising succinate production flux when glucose was used as a carbon source (Figure 2b).

To validate this hypothesis, the gltA gene was overexpressed in PALK strain, resulting in PALK (pMS3‐gltA) strain (Figure 2a). Anaerobic fed‐batch fermentation of PALK (pMS3‐gltA) strain produced 100.82 g/L of SA with the yield and productivity of 1.30 mol/mol and 4.00 g/L/h, respectively, using glucose as a carbon source (Figure 2c and Table 1). In contrast, fed‐batch fermentation of the control PALK (pMS3) strain, which harbors an empty vector, produced 74.56 g/L of SA with the yield and productivity of 1.11 mol/mol and 3.03 g/L/h, respectively, using glucose as a carbon source (Ahn et al. 2018). Additionally, PALK (pMS3‐gltA) strain demonstrated improved cell growth (maximum OD600 of 12.2; Figure 2c and Table 1) compared to PALK (pMS3) strain (maximum OD600 of 10.8; Ahn et al. 2018). The enhanced performance of PALK (pMS3‐gltA) strain was further confirmed by an increased initial glucose uptake rate (13.39 mmol/gDCW/h), higher SA production rate (18.00 mmol/gDCW/h), and a shorter doubling time (0.84 h) compared to the control strain (8.76 mmol/gDCW/h, 10.41 mmol/gDCW/h, and 1.16 h, respectively; Table 1). These results confirm that gltA overexpression positively impacts both SA production and cell growth, resulting in higher overall SA production performance compared to previously reported M. succiniciproducens strains (Table 2).

Table 1.

Fed‐batch fermentation results of the engineered Mannheimia succiniciproducens strains.

Strain Fermentation conditiona Maximum OD600 Final SA (g/L) Overall productivity (g/L/h) Yieldb (mol/mol) Glucose uptake rate (mmol/gDCW/h) SA production rate (mmol/gDCW/h) Doubling time (h) Source
PALK (pMS3) Fed‐batch, GLC 10.8 74.56 3.03 1.11 8.76 10.41 1.16 Ahn et al. (2018)
PALK (pMS3) Fed‐batch, GLC + GOL 8.54 89.23 3.16 1.09 12.06 20.86 1.08 Figure S1a
PALK (pMS3‐gltA) Fed‐batch, GLC 12.2 100.82 4.00 1.30 13.39 18.00 0.84 Figure 2c
PALK (pMS3‐gltA) Fed‐batch, GLC + GOL 10.3 110.16 4.37 1.38 14.70 23.63 0.95 Figure 2d
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a

GLC, glucose; GOL, glycerol.

b

As two kinds of carbon source were used in this study, the yield was calculated based on glucose equivalent (mol SA per mol glucose equivalent) for comparison.

Table 2.

Succinic acid production by metabolically engineered strains.

Strain Description Titer (g/L) Yield (mol/mol) Productivity (g/L/h) Carbon source Cultivation mode Source
Actinobacillus succinogenes FZ53 Mutant of wild type 130Z strain 105.8 1.22 1.34 Glucose Anaerobic batch Guettler et al. (1996)
Corynebacterium glutamicum S071/pGEX4‐NCgl0275 ΔldhA, Δpta‐ackA, ΔactA, Δpck, ΔpoxB, ΔptsG, +pyc P458S , +ppc, +pckG, +NCg10275 152.2 1.67 1.11 Glucose Dual‐phase fed‐batch Chung et al. (2017)
Escherichia coli AFP111/pTrc99A‐pyc ΔpflB, ΔptsG, ΔldhA, +pyc 99.2 1.68 1.31 Glucose Dual‐phase fed‐batch Vemuri et al. (2002)
Saccharomyces cerevisiae SUC‐297 Δadh1, Δadh2, Δgpd1, +pckA, +gsh1, +cys3, +glr1, +mdh3, +pyc2p, +fumR, +frdm1, +mae1 43.0 — 0.45 Glucose Dual‐phase fed‐batch Van De Graaf et al. (2015)
Yarrowia lipolytica PGC202 Δsdh5, Δach1, +pck, +scs2 110.7 0.41 0.80 Glycerol

Aerobic fed‐batch

 Cui et al. (2017)
M. succiniciproducens PALK (pMS3) ΔldhA, Δpta‐ackA 74.56 1.11 3.03 Glucose Anaerobic fed‐batch Ahn et al. (2018)
M. succiniciproducens PALK (pMS3‐mgtB) ΔldhA, Δpta‐ackA, +mgtB 93.89 1.27 3.44 Glucose Anaerobic fed‐batch Kim et al. (2024)
M. succiniciproducens PALK (pMS3‐gltA) ΔldhA, Δpta‐ackA, +gltA 100.82 1.30 4.00 Glucose Anaerobic fed‐batch This study
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Based on our previous studies, the utilization of glucose and glycerol as dual carbon sources promoted enhanced SA production, as the conversion of glycerol (C3) to phosphoenolpyruvate (C3) generates twice as much reducing equivalents per six‐carbon equivalent mole compared with glucose (C6) (Ahn et al. 2020; Choi et al. 2016). Fed‐batch fermentation of PALK (pMS3‐gltA) strain using glucose and glycerol as dual carbon sources produced 110.16 g/L of SA with the yield and productivity of 1.38 mol/mol glucose equivalent (mol SA per mol glucose equivalent) and 4.37 g/L/h, respectively (Figure 2d and Table 1). These values represent a significant improvement compared to the control PALK (pMS3) strain under the same conditions (Supporting Information S1: Figure S1a and Table 1). In addition, the observed increase in SA production through gltA overexpression was consistent with the in silico simulation using glucose and glycerol as dual carbon sources (Supporting Information S1: Figure S2). To ensure consistent and reliable comparisons between strains, fed‐batch fermentations of PALK (pMS3) and PALK (pMS3‐gltA) strains were conducted simultaneously under identical conditions, using the same media formulation and operator. The results of these fermentations are presented in Figure 2 and Supporting Information S1: Figure S1,S3–S5.

3.2. Synthesis of PBS Using Purified Bio‐Based SA From Fermentation Broth

Following the strain development for enhanced SA production, purified bio‐based SA was used as a monomer for synthesizing PBS. High‐purity monomers, typically requiring purities above 99.8%, are essential for successful polymerization as impurities can inhibit chain growth and reduce the molecular weight of the resulting polymer (Szwarc 1958; Vardon et al. 2016). Using our previously established SA recovery protocol (Choi et al. 2016; Lee et al. 2020), we achieved a SA purity of 99.91%, as confirmed by ICP‐MS analysis (Supporting Information S1: Table S2).

PBS is traditionally synthesized through the direct polycondensation of SA and 1,4‐butanediol (Barletta et al. 2022; Xu and Guo 2010b). However, this method often results in lower molecular weight polymers due to the incomplete removal of water, a byproduct of the condensation reaction, which shifts the reaction equilibrium away from polymer formation and limits chain elongation (Ferreira et al. 2015; Platnieks et al. 2021). To counteract this, high temperatures and prolonged reaction times are typically required, which increase energy consumption and risk thermal degradation of the polymer (Ferreira et al. 2015; Platnieks et al. 2021).

To address these limitations, we employed a two‐step synthesis strategy (Supporting Information S1: Figure S6). In the first step, SA was esterified with ethanol to produce diethyl succinate, a more reactive intermediate that enables efficient polymerization. A 1:10 molar ratio of SA to ethanol was used to drive the reaction toward completion by shifting the equilibrium in favor of diethyl succinate formation. The excess ethanol also facilitates effective water removal during the reaction, overcoming the equilibrium bottleneck associated with direct condensation (Orjuela et al. 2012). Amberlyst, an ion‐exchange resin catalyst, was selected for its stability, reusability, and ease of separation, making it highly suitable for environmentally friendly processes.

In the second step, equimolar amounts of diethyl succinate and 1,4‐butanediol were polymerized using CALB as a catalyst. Enzymatic polymerization offers several advantages over conventional methods: it operates at a mild temperature (90°C), minimizing thermal degradation; it eliminates the need for toxic metal‐based catalysts, such as titanium butoxide (Ti(OBu)4); and it provides superior control over the polymer structure, ensuring high reproducibility and reduced byproduct formation (Liu et al. 2020). These features make enzymatic polymerization both energy‐efficient and environmentally friendly.

For comparison, chem‐PBS was synthesized using petrochemical‐derived SA under identical reaction conditions. Although bio‐PBS and chem‐PBS share the same chemical structure, minor variations in monomer purity, residual metal ions, or intermediate reaction conditions can influence key polymer properties such as molecular weight, crystallinity, and thermal behavior. Therefore, chem‐PBS served as a baseline reference to validate the quality of bio‐PBS.

3.3. Material Properties of PBS

The 1H and 13C NMR analyses of both bio‐PBS and chem‐PBS showed identical chemical shifts (Figure 3), suggesting that the two types of PBS share the same chemical structures. The M n of bio‐PBS and chem‐PBS were 2697, 4897 g/mol, respectively (Table 3). The polydispersity index (M w/M n) of bio‐PBS and chem‐PBS were 2.41 and 1.92, respectively (Table 3). Although the M n of bio‐PBS was lower, its polydispersity index indicates a uniform molecular weight distribution comparable to that of chem‐PBS. A previous study using CALB for PBS synthesis under mild conditions has reported M n values ranging from 3000 to 5000 g/mol (Azim et al. 2006). The relatively lower M n observed in this study can be attributed to factors such as slower reaction kinetics at mild polymerization temperatures and the intrinsic activity and stability of CALB during prolonged reactions (Liu et al. 2020; Platnieks et al. 2021). Mild temperatures, while reducing the risk of thermal degradation, can limit the enzyme's catalytic efficiency, thereby slowing polymerization rates. Additionally, extended reaction times can cause enzyme instability, further hindering polymer chain elongation. Despite these limitations, the consistent polydispersity of bio‐PBS demonstrates the potential of enzymatic polymerization to produce PBS with acceptable material properties under sustainable conditions.

Figure 3.

Figure 3

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Synthesis of bio‐ and chem‐PBS. 1H NMR spectra of (a) chem‐PBS (top) and bio‐PBS (bottom) and 13C NMR spectra of (b) chem‐PBS (top) and bio‐PBS (bottom) using CDCl3 as the NMR solvent. The peaks in the 1H and 13C NMR spectra were numbered to indicate the corresponding hydrogen and carbon atom, respectively.

Table 3.

Properties of synthesized PBS.

Polymer M n (g/mol) M w/M n T m (°C) ∆H f (J/g) T d5 (°C) T dmax (°C)
Bio‐PBS 2697 2.41 107.1 87.5 312.3 396.6
Chem‐PBS 4897 1.92 110.7 93.0 338.7 403.6
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Thermal properties of bio‐PBS and chem‐PBS were identified by thermogravimetric analysis and differential scanning calorimetry (Figure 4a,b and Table 3). The T d5 was 312.3°C for bio‐PBS and 338.7°C for chem‐PBS. The T dmax was observed at 396.6°C for bio‐PBS and 403.6°C for chem‐PBS. Moreover, the T m was 107.1°C for bio‐PBS and 110.7°C for chem‐PBS. It should be noted that thermal properties (as indicated by T d5, T dmax, T m) are strongly influenced by the molecular weight of polymers due to the greater number of polymer chains contributing to the polymer's overall stability (Toncheva et al. 2011). Moreover, slightly lower Tm and Td values can be advantageous in terms of melt processability, allowing energy‐efficient processing at lower temperatures while maintaining sufficient thermal stability for practical use.

Figure 4.

Figure 4

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Material properties of bio‐ and chem‐PBS. The thermal properties (T m, ΔH f, T d5, T dmax) of bio‐PBS (red) and chem‐PBS (blue) were analyzed using (a) TGA and (b) DSC analyses. The crystal structures of (c) bio‐PBS (top) and chem‐PBS (bottom) were determined by XRD analyses. The properties of bio‐PBS and chem‐PBS analyzed in this study are summarized in Table 3.

The diffraction peaks of bio‐PBS and chem‐PBS crystal structures from power XRD analyses appeared at approximately 2θ = 19.5 and 22.5°, corresponding to the (020) and (110) planes of the α‐crystalline phase of PBS (Chaiwutthinan et al. 2015; Hu et al. 2017; Figure 4c). The close similarity in peak positions confirms that both bio‐PBS and chem‐PBS maintain a well‐defined crystalline structure, supporting their mechanical durability and thermal processability. Therefore, the two‐step polymerization process was effective in synthesizing PBS with adequate molecular weight distribution, thermal stability, and crstallinity, confirming its feasibility for real‐life applications.

3.4. Biodegradability of Synthesized PBS

Most commercially available plastics are derived from petrochemicals, contributing significantly to greenhouse gas emissions throughout their life cycle (Jambunathan and Zhang 2016). In 2019, plastics accounted for 1.8 billion tons of CO2 equivalent emissions, representing 3.4% of global emissions (Choi et al. 2023). These environmental concerns have fueled interest in bio‐based and biodegradable alternatives, such as PBS. While PBS shows promise for applications in packaging and agriculture, its adoption has been limited by relatively slow degradation rates and the scarcity of PBS‐degrading microorganisms in natural environments (Abe et al. 2010; Boonmee et al. 2016; Platnieks et al. 2021). Despite these challenges, preliminary studies have reported the microbial degradation of PBS by bacteria and fungi (Ishii et al. 2008; Kitamoto et al. 2011; Maeda et al. 2005; Savitha et al. 2022; Uchida et al. 2002). This process involves enzymatic hydrolysis mediated by enzymes such as depolymerases, esterases, and lipases, which cleave the ester linkages within PBS (Lee et al. 2008; Savitha et al. 2022; Tserki et al. 2006).

In this study, Pseudomonas flurorescens was selected to assess the biodegradation of PBS. The genus Pseudomonas is widely distributed in natural environments and includes bacteria capable of degrading aliphatic polyesters (Lee and Kim 2010). Moreover, PBS degradation by Pseudomonas sp. has been previously reported (Savitha et al. 2022; Taniguchi et al. 2002). The biodegradation of bio‐ and chem‐PBS was assessed by cultivating P. fluorescens in a mineral salt medium supplemented with 5 g/L of either PBS type. Control experiments without PBS supplementation were conducted to verify the dependence of bacterial growth on PBS availability.

The growth profiles of P. fluorescens in the presence of bio‐ or chem‐PBS were comparable, with negligible growth observed in the absence of PBS (Figure 5a). Notably, 93.3% of the bio‐PBS and 87.8% of the chem‐PBS were degraded after 2 weeks (Figure 5b), outperforming previous reports that observed only 50% PBS degradation within the same time frame (Asheeba et al. 2010). Control experiments in which PBS was incubated without bacteria showed no significant weight loss due to abiotic degradation or polymer erosion, confirming that weight loss was attributable to microbial degradation (Figure 5b). All biodegradability tests were performed in triplicate.

Figure 5.

Figure 5

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Biodegradability of bio‐ and chem‐PBS. The biodegradability of synthesized PBS was investigated by culturing Pseudomonas fluorescens for 14 days in the mineral salt medium using bio‐PBS or chem‐PBS as the carbon source. (a) Growth profiles of P. fluorescens when supplied with no PBS, bio‐PBS, and chem‐PBS. (b) Weight of the PBSs. The degraded amount (%) represents the ratio of the weight of degraded PBS to the weight of PBS before degradation. The rate of biodegradation was determined by dividing the weight of degraded PBS to the total amount of cultivation time (14 days). Statistical significance was determined using an unpaired two‐tailed Student's t test: p < 0.0005 for both bio‐PBS and chem‐PBS when comparing samples with and without P. fluorescens. (c) TOC values of the supernatants of various culture media were analyzed before and after degradation. All experiments were performed in triplicate. Error bars represent standard deviations.

The higher biodegradation rate of bio‐PBS compared to chem‐PBS may be attributed to differences in polymer microstructure, which could arise from impurities or trace elements associated with the starting materials, such as bio‐based SA or petrochemical‐derived SA. These differences may influence polymer crystallinity or enzyme accessibility, thereby affecting microbial attachment and degradation efficiency. Similar effects have been observed in other bio‐based polymers, where variations in material composition or processing conditions significantly impact degradation rates (Savitha et al. 2022; Lee et al. 2008).

As bacteria‐mediated degradation proceeded, bio‐ and chem‐PBS were hydrolyzed into water‐soluble small molecules, as evidenced by increased total organic carbon (TOC) values in the culture supernatant after PBS degradation (Figure 5c). The TOC values corresponded with the extent of bacterial growth in the medium, further corroborating the role of microbial activity in PBS degradation (Figure 5a).

These results confirm that the synthesized bio‐PBS is biodegradable and achieves higher degradation rates compared to conventional PBS. The enhanced biodegradability of bio‐PBS, combined with its sustainable production process, makes it a promising candidate for applications in packaging and agricultural films, particularly for short‐lived products.

4. Conclusions

In this study, an integrated approach was developed for the sustainable production of PBS using bio‐based SA from metabolically engineered M. succiniciproducens. Fed‐batch fermentation of the engineered PALK (pMS3‐gltA) strain using glucose and glycerol as dual carbon sources achieved a high titer of 110.16 g/L, with a yield and productivity of 1.38 mol/mol and 4.37 g/L/h, respectively, demonstrating significant improvements over previously reported strains. The high‐purity bio‐based SA (99.91%) was then used to synthesize PBS through a two‐step polymerization under mild conditions employing reusable catalysts, thereby eliminating the need for toxic metal catalysts and reducing energy demand.

Building on these findings, future research should focus on further enhancing the sustainability and performance of bio‐PBS. First, improving the catalytic activity and stability of CALB could increase the molecular weight of PBS, thereby broadening its industrial applications. Second, optimizing the crystallinity and surface characteristics of PBS could improve its enzymatic hydrolysis and biodegradability under diverse environmental conditions. Third, investigating the potential effects of microplastics generated during PBS biodegradation would be valuable for a comprehensive assessment of its environmental safety. In addition, incorporating life cycle assessment and technoeconomic analysis will be essential to rigorously evaluate the environmental and economic feasibility of bio‐based PBS production. Finally, the sustainable polymerization strategy demonstrated in this study could be extended to other bio‐based monomers, thereby expanding the portfolio of biodegradable plastics.

In conclusion, this study integrates advances in metabolic engineering, bio‐based monomer production, and environmentally friendly polymerization processes to develop a sustainable pathway for PBS synthesis. The strategies reported here provide a robust framework for producing bio‐based and biodegradable polymers, contributing to global efforts toward sustainable materials and reduced plastic waste. By addressing both production and end‐of‐life challenges, this approach represents a significant step forward in creating eco‐friendly materials for a circular economy.

Author Contributions

S.Y.L. conceived the project. J.Y.K., J.A.L., and S.Y.L. designed the experiments. J.Y.K., J.A.L., G.B.K., and Y.L. conducted the experiments and analyzed the data. J.Y.K. and S.Y.L. wrote the manuscript. All authors read and approved the final manuscript.

Conflicts of Interest

Authors declare that they have competing financial interests as the M. succiniciproducens strains described in this paper are covered by patents registered including KR1007803240000, AU20077277593, BE2054502, BRPI07138393, CA2659246, CN200780036066.4, DE2054502, DK2054502, EP2054502, ES2054502, FI2054502, FR2054502, GB2054502, IDP000039187, IN289769, IT2054502, JP5421103, MXa2009001004, MY162581A, NZ575237, PH12009500147, PT2054502, RU2415942, SE2054502, US9217138, VN159559, ZA200901254, US9428774, and NL2054502, and also by other patents in application.

Supporting information

Kim etal SI clean.

BIT-122-3393-s001.pdf (249.7KB, pdf)

Acknowledgments

This study is supported by the Development of next‐generation biorefinery platform technologies for leading bio‐based chemicals industry project (2022M3J5A1056072), Development of platform technologies of microbial cell factories for the next‐generation biorefineries project (2022M3J5A1056117) from National Research Foundation supported by the Korean Ministry of Science and ICT.

Kim, J. Y. , Lee J. A., Kim G. B., Lee Y., and Lee S. Y.. 2025. “Succinic Acid Production by Engineered Mannheimia succiniciproducens and Its Use in Chemoenzymatic Poly(Butylene Succinate) Synthesis.” Biotechnology and Bioengineering 122: 3393–3405. 10.1002/bit.70072.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Kim etal SI clean.

BIT-122-3393-s001.pdf (249.7KB, pdf)

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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