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. 2026 May 14;11(20):29422–29432. doi: 10.1021/acsomega.5c10104

Biosynthesis of High Molecular Weight Poly(3-hydroxybutyrate) by Wild-Type Cupriavidus necator H16 Using Sugar Acids as the Carbon Source

Jia Yi Cheah 1, Kousuke Takashima 1, Masatake Kanai 1, Daisuke Ishii 1,*
PMCID: PMC13216913  PMID: 42222778

Abstract

This study investigated whether pectin-derived sugar acids can serve as effective carbon sources for poly­(3-hydroxybutyrate) [P­(3HB)] production in Cupriavidus necator H16. We attempted to utilize sugar acids, including glucuronic, galacturonic, and gluconic acids, as the carbon sources for the biosynthesis of polyhydroxyalkanoate (PHA) by wild-type Cupriavidus necator H16. Among the three types of sugar acids tested, only gluconic acid was used as the carbon source for PHA production. When cultivated on gluconic acid at a controlled pH, C. necator produced significantly higher biomass, increased P­(3HB) accumulation, and elevated polymer molecular weight compared with fructose-grown cultures. The resulting P­(3HB) exhibited enhanced mechanical strength and crystallinity, demonstrating the material advantages of utilizing gluconic acid. These findings demonstrate the potential of gluconic acid as a viable substrate for the production of high-quality P­(3HB).

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Introduction

Synthetic plastics are an indispensable part of both domestic and industrial economies, with demand increasing annually. The majority of plastics currently produced are of a fossil-fuel-based variety, including polypropylene (PP) and polyethylene (PE). , From 1950 to 2018, approximately 6.3 billion tons of plastic were produced, with only a minor proportion being recycled or incinerated, and the remainder landfilled. The disposal of synthetic plastics and their slow degradation have led to significant environmental issues and public health concerns. The release of synthetic plastic into landfills has been identified as one of the major sources of exposure to microplastics, which are known to have adverse effects on human health, including DNA damage and organ dysfunction. , Consequently, there is a growing interest in developing alternative materials, such as biodegradable plastics, to address these environmental and public health concerns.

Polyhydroxyalkanoates (PHAs) are regarded as a promising substitute due to their biodegradable properties and favorable characteristics, including high durability, UV resistance, hydrophobicity, and stability in a range of environments. PHAs are hydroxylalkanoic acids that form esters with other molecules, with the most well-known PHA being polyhydroxybutyrate (PHB). Poly­(3-hydroxybutyrate) (P­(3HB)) is a short-chain-length (scl-PHA) polymer synthesized by various microorganisms and was the first PHA to be identified in the bacterium Bacillus megaterium. The weight-average molecular weight (M w) of PHAs synthesized by unmodified strains ranges from 0.1 × 106 to 2.01 × 106, with PHAs exhibiting a M w exceeding 3 million classified as ultrahigh-molecular-weight PHA (UHMW-PHA). These PHAs are known to have exceptional mechanical properties and are less affected by the reduction in molecular weight during the extraction or heating process, as they can maintain a high molecular weight even after degradation. UHMW-P­(3HB) shows tensile strength and Young’s modulus significantly higher than P­(3HB) and could be used for high-strength fibers and films. − Although the molecular weight of PHAs synthesized by unmodified strains is generally low, there have been reports of bacterial strains capable of synthesizing high molecular weight PHAs.

At present, a variety of wild-type and recombinant bacteria are employed for large-scale PHA production, with Cupriavidus necator proving to be a particularly favorable native producer of PHAs. C. necator is a Gram-negative bacterium with the capacity to utilize a range of carbon sources, including sugar acids, amino acids, fatty acids, alcohols, and aromatic compounds, for heterotrophic growth. It is also capable of accumulating intracellular P­(3HB) granules at a concentration of up to 90 wt % under nutrient-limiting conditions. , The C. necator H16 strain is the most representative and widely studied wild type. It is metabolically versatile and exhibits heterotrophic growth with carbon sources such as fructose and gluconate.

Gluconic acid and fructose represent two of the most studied carbon sources for C. necator H16. Gluconic acid is a nontoxic, noncorrosive, mild organic acid that can be found in plants, honey, and wine. It can be produced through mild enzymatic or microbial oxidation of glucose. While fructose remains one of the most widely used carbon sources for laboratory and industrial C. necator, gluconate supports faster growth than alternative substrates in wild-type C. necator H16, which grows more rapidly on gluconate than on fructose.

Although PHAs offer numerous benefits over petroleum-based synthetic plastics, their cost of production can reach up to five times higher than that of synthetic plastics, posing a considerable barrier to their implementation on an industrial scale. One potential strategy to mitigate this cost challenge is to utilize less expensive substrates, such as agro-industrial residuals. A plethora of inexpensive alternative carbon sources for C. necator, including sugarcane molasses, glycerol, waste cooking oil, and waste animal fats, have been evaluated. − Pectin, a heteropolysaccharide and one of the most abundant components of plant cell walls, is particularly attractive because its hydrolysis yields sugar acids such as galacturonic and glucuronic acids, which could serve as alternative carbon sources. The potential of pectin and its hydrolyzates as carbon sources for C. necator was previously evaluated, with pectin identified as a suitable substrate for supporting cell growth in this organism. While previous work has shown that pectin hydrolyzates can support the growth of C. necator, it remains unknown whether pectin-derived sugar acids can sustain PHA synthesis, what their relative suitability is compared to conventional sugars, and whether they influence the polymer’s physical properties such as molecular weight. Given that the superior mechanical properties of high molecular weight P­(3HB) make it preferable as a material compared to P­(3HB) of lower molecular weight, it is therefore desirable to produce high molecular weight P­(3HB) with inexpensive carbon sources.

Although many studies have evaluated mixtures of sugars and other substances for PHA production, the production efficiency of simple sugars has seldom been evaluated. Therefore, this study aims to assess the potential of pectin-derived uronic acids and gluconic acid as carbon sources for PHA production in C. necator. Furthermore, we have examined the physical characteristics of PHAs produced with gluconic acid in comparison to the conventional P­(3HB), which was produced using fructose and commercial P­(3HB).

Materials and Methods

Bacterial Strains and Cultivation Conditions

The bacterial strain Cupriavidus necator H16 was employed in all experiments conducted. The reagents for the preparation of culture media were used as received. Initially, C. necator H16 was cultivated at 30 °C for 16 h on a reciprocating shaker at 160 rpm under aerobic conditions in a test tube containing 5 mL of nutrient-rich (NR) medium comprising 10 g L–1 bonito extract, 2 g L–1 yeast extract, and 10 g L–1 tryptone. Subsequently, the medium was inoculated into 500 mL Sakaguchi flasks sealed with breathable porous caps containing 200 mL of NR medium with 10 g L–1 of d-fructose (Wako special grade, FUJIFILM Wako Pure Chemical Corporation, Tokyo, Japan), d-gluconic acid (50% aqueous solution, FUJIFILM Wako Pure Chemical Corporation), d-glucuronic acid (Biochemistry grade, FUJIFILM Wako Pure Chemical Corporation), or d-galacturonic acid (97.0%, monohydrate form, Sigma-Aldrich Co., Milwaukee, U.S.) added as a carbon source. The inoculated volume was 1 mL. The inoculum was prepared by transferring a single colony from an NR agar plate that had been incubated at 30 °C for 72 h into 5 mL of NR. The pH of the culture medium with the added carbon source was adjusted to 7.00 using either 1 M NaOH or 1 M HCl. Culture medium containing 100 mM Tris-HCl (pH 8.5) was used without the adjustment of pH. The culture medium was measured prior to and after cultivation. Following inoculation, the medium was cultivated at 30 °C for 48 h on a reciprocating shaker at 130 rpm.

Biomass Recovery

The cells were recovered by centrifugation at 24 °C for 15 min at 6300 × g, including one washing step with distilled water. The pellet was then freeze-dried for 4 days, after which the dry mass weight was measured. The supernatants obtained from centrifugation were filtered with a 0.45 μm PTFE membrane filter (Minisart Syringe Filter, Poly­(ether sulfone) (PES), Pore Size 0.45 μm, Sartorius, Göttingen, Germany) and used for pH measurement.

PHA Extraction

The lyophilized cell dry mass was suspended in 100 mL of chloroform (reagent grade, FUJIFILM Wako Pure Chemicals, Japan) per 1 g of cell mass at room temperature for 24 h with vigorous stirring, after which the extracted PHA solution was gravitationally filtered with filter paper (No. 1, Advantec Co. Ltd., Tokyo, Japan) and precipitated in 50 mL of methanol per 1 g of cell mass. The precipitated PHA was recovered by paper filtration (Hydrophilic PTFE Membranes, 47 mm diameter, pore size 0.50 μm, Advantec Co. Ltd., Japan) and then subjected to a 24-h drying process at 40 °C in a vacuum oven.

PHA Quantification by Gas Chromatography (GC)

The methyl esterification of PHA was performed in accordance with the following procedure. Approximately 20 mg of lyophilized cells were suspended in 2 mL of chloroform, and then an equivalent volume of methanol containing 15% sulfuric acid was added and subjected to incubation at 100 °C for 140 min. Subsequently, the mixture was cooled to room temperature, followed by the addition of 1 mL of distilled water and vigorous agitation using a vortex mixer. Following a period of rest at room temperature, the lower organic layer was collected. The internal standard, 0.1% methyl octanoate dissolved in chloroform, was then added, and the mixture was filtered using a 0.2 μm PTFE filter (Minisart RC4, Sartorius, Göttingen, Germany). Quantification of 3-hydroxybutyric acid was carried out using a gas chromatography system (Nexis GC-2030, Shimadzu Corporation, Kyoto, Japan). Analysis conditions were as follows: injection volume, 1 mL; split ratio, 40; carrier gas, helium (100 kPa, total flow rate of 54.5 mL/min); column used, InertCap 1 (0.25 mm i.d. × 30 m, film thickness of 0.40 μm, GL Science Co., Ltd., Tokyo, Japan); oven temperature, 50 °C at the injection and elevated to 80 °C at 5 min (6 K/min) and then to 280 °C at 16 min (18.4 K/min), at which temperature the column was kept for 5 min; detector, FID.

Estimation of PHA content, P, from the peak areas of the analyte and the internal standard was performed by using the following equation:

P(wt%)=K×Am×S×100

K: a constant calculated from 3HB samples with known concentrations.

A: peak area of the analyte.

m: dry cell weight of the analyte.

S: peak area of the internal standard.

In the present study, a K value of 7.2 was used for the estimation.

Determination of Molecular Weight by Gel Permeation Chromatography (GPC)

The extracted P­(3HB) was dissolved in chloroform by heating at 80 °C for an hour, after which the concentration was adjusted to 2 mg/mL. The sample was then filtered using a 0.2 μm PTFE filter (DISMIC-13HP, Advantec Co. Ltd., Japan), and the molecular weight was measured using a gel permeation chromatography system (LC-20AD, Shimadzu Corporation, Kyoto, Japan). Analysis conditions were as follows: eluent, chloroform (HPLC grade, Kanto Chemicals Co., Saitama, Japan); flow rate, 0.8 mL/min; column used, KF-806 M (7.8 mm i.d. × 300 mm, Shodex Inc., Japan); column temperature, 50 °C; injection volume, 100 μL; detector, refractive index (RID-20, Shimadzu Corporation). Molecular weight (MW) distribution and the resultant average MWs (number-average molecular weight (M n), weight-average molecular weight (M w), and polydispersity index (PDI)) were estimated by calibrating the elution curves using monodisperse polystyrene (PS) standards. The molecular weights of the standard PSs used were 6.55 × 106, 3.02 × 106, 2.33 × 106 (SH-75, Shodex Inc., Japan), 1.6 × 106, 9.0 × 105, 2.0 × 105, 9.0 × 104, 4.0 × 103 (Pressure Chemicals Co., Pittsburgh, U.S.), and 5.8 × 102 (Agilent Technologies, Santa Clara, U.S.). Standard PSs obtained from different sources were used to cover a broad and finer range calibration curve with commercially available PS standards.

Preparation of P­(3HB) Solvent-Cast Films

0.5 g of P­(3HB) was dissolved in 10 mL of chloroform at 80 °C for 1 h. It was then poured into a 9 cm silicon Petri dish, dried at room temperature overnight, and then subjected to a 24-h drying process at 40 °C in a vacuum oven. The commercial P­(3HB) used in this study was purchased from Sigma-Aldrich Co., Milwaukee, U.S.

Tensile Test of P­(3HB) Films

Tensile tests were performed on an MCT 2150 universal tensile testing machine (A&D Co. Ltd., Tokyo, Japan) equipped with a 500 N load cell. To perform the tensile test, the solvent-cast film of P­(3HB) was cut into small pieces (ca. 3 × 20 mm2), placed between PTFE films, and then melted at 200 °C under a load of 1 MPa for 15 s by using a hydraulic thermal pressing device (IMC-180C, Imoto Machinery Co. Ltd., Kyoto, Japan). After the melting process, the molten P­(3HB) pressed between the PTFE films was instantly coagulated by soaking it in an iced water bath to prevent the crystallization of P­(3HB) during cooling. The coagulated P­(3HB) film was stretched in iced water to prevent the crystallization of P­(3HB) during stretching by using a unidirectional stretching device (IMC-B309, Imoto Machinery Co. Ltd., Japan) with a stretch ratio of 6. Films were stretched to form a uniform and oriented crystal structure. The stretched film, chucked by the stretching device, was then annealed at 100 °C for 1 h in an atmospheric oven. After being cooled down to ambient temperature, the stretched-annealed P­(3HB) film was then cut into a rectangular shape (ca. 2 × 30 mm2) and tested with an initial chuck length of 20 mm and at a crosshead speed of 10 mm/min. The tensile tests were repeated in triplicate, and the elongation at break (in %) and maximum strength (in MPa) were estimated by averaging the values obtained from each stress–strain curve.

Differential Scanning Calorimetry

Thermal properties of the P­(3HB) films were investigated by differential scanning calorimetry (DSC), performed on Thermo Plus EVO2 DSCvesta (Rigaku Corporation, Japan). Stretched-annealed P­(3HB) films were cut into small pieces with a weight of ca. 2 mg and sealed in an aluminum pan. An empty aluminum pan was used as a reference. The first heating cycle was carried out from 30 to 200 °C. The samples were then cooled to −100 °C and reheated to 200 °C. All the cooling and heating cycles were carried out at a rate of 20 °C/min under an inert nitrogen atmosphere. The lamella thickness of the P­(3HB) crystal was calculated according to the Gibbs–Thomson eq :

L=2γeΔHm0Tm0Tm0−Tm 1

where L is the lamella thickness (nm), T m is the melting temperature of the lamellae, γ e is the surface energy, γ e = 46.89 mN/m, ΔHm0 is the enthalpy of fusion of the lamellae, and Tm0 is the melting point of the crystal with infinite lamella thickness (Tm0=200C°) . , The density of P­(3HB) is taken as ρ = 1.2 g/cm3.

Wide-Angle X-ray Diffraction Analysis

The crystalline structure of the samples was studied using a Miniflex 600 (Rigaku Corporation, Tokyo, Japan), an X-ray diffractometer equipped with a Cu–Kα X-ray source (λ = 0.1542 nm), a Bragg–Brentano 2θ–θ goniometer, and a HyPix-400 MF semiconductor two-dimensional detector. The samples were scanned in the range of 3–60° with a step size of 0.02°. After data acquisition, the two-dimensional diffraction profile was transformed into a one-dimensional 2θ–I profile. Deconvolution of the 2θ–I profile was performed using data processing software (2DP, Rigaku Corporation, Japan). The degree of crystallinity was estimated from the deconvoluted profile as the ratio of the crystalline peak area to the total area of the profile.

Statistical Analysis

All experimental data were described as mean ± standard deviation. Each experiment was performed in triplicate. Statistical analysis of the results was performed by conventional methods, using Microsoft Excel. The statistical significance of the results was determined using Student’s t-test (significance level: p ≤ 0.05).

Results

Cell Growth and PHA Production

C. necator H16 has been demonstrated to be an effective platform for PHA production. It is known to synthesize and accumulate P­(3HB) using fructose and gluconic acid as carbon sources. Nevertheless, the effects of different carbon sources on the production and physical properties of P­(3HB) remain unclear.

In this study, we quantified intracellular P­(3HB) produced and its average molecular weight using gluconic acid, glucuronic acid, and galacturonic acid as carbon sources to investigate the potential of using uronic acids in P­(3HB) production with C. necator H16. The results are summarized in Table . P­(3HB) in cells cultivated in this study was compared to commercial P­(3HB) using GC and showed identical peaks; therefore, we concluded that the PHA produced was P­(3HB) (as shown in Supporting Information). In order to evaluate the effects of the addition of acidic sugars on the pH of the culture medium, the pH of the medium was measured prior to culturing. The addition of sugar acids resulted in a sharp decrease in culture medium pH; hence, growth was only observed at low sugar acid concentrations (as shown in Supporting Information). To prevent the acidification of the culture medium at high sugar acid concentrations, Tris–HCl buffer was added to the culture medium. Although phosphate-buffered saline (PBS) was also considered, its buffering capacity was insufficient, and high concentrations appeared to inhibit cell growth (as shown in Supporting Information). Therefore, Tris–HCl buffer was chosen as a more favorable alternative. The pH of the medium with fructose, a neutral sugar commonly employed in the cultivation of C. necator H16, was identical to that of the medium without fructose, at 7.82. The pH of the culture medium with the addition of gluconic acid, glucuronic acid, and galacturonic acid was found to be 7.12, 5.22, and 5.90, respectively. The fructose-added culture medium exhibited a biomass yield of 3.4 ± 1.5 g/L, accompanied by a PHA content of 24.2 ± 18.8%. The biomass and PHA content in the medium with gluconic acid added were found to be 2.6 ± 0.1 g/L and 14.8 ± 2.22%, respectively. The biomass in the medium with glucuronic acid and galacturonic acid added was 1.4 ± 0.0 g/L and 1.3 ± 0.0 g/L, respectively. The accumulation of PHA was not observed.

1. Biosynthesis Conditions and Quantitative Information on the Biosynthesized P­(3HB) by Wild-Type C. necator H16.

  Carbon Source Sugar Concentration (g L–1) Initial pH Final pH CDW (g L–1) PHA Content (wt %) M n (×105 g mol–1) M w (×105 g mol–1) PDI
Cells were cultivated in NR medium with 100 mM Tris–HCl buffer. Initial pH was not adjusted Blank 0 7.82 ± 0.02 8.09 ± 0.02 1.4 ± 0.0 0.7 ± 0.2 8.2 25 3.1
Fructose 10 7.82 ± 0.06 7.25 ± 0.59 3.4 ± 1.5 24.2 ± 18.8 5.3 ± 2.7 12 ± 2.1 2.7 ± 1.4
Gluconic acid 10 7.12 ± 0.02 7.90 ± 0.04 2.6 ± 0.1 14.8 ± 2.2 6.0 ± 1.4 21 ± 2.1 3.5 ± 0.4
Glucuronic acid 10 5.22 ± 0.05 7.49 ± 0.02 1.4 ± 0.0 0.0 ± 0.0 – – –
Galacturonic acid 10 5.90 ± 0.02 7.57 ± 0.02 1.3 ± 0.0 0.0 ± 0.0 – – –
Cells were cultivated in NR medium with 100 mM Tris–HCl buffer. Initial pH was adjusted to 7.0 Blank 0 7.00 7.86 ± 0.17 1.5 ± 0.1 0.0 ± 0.0 – – –
Fructose 10 7.00 7.37 ± 0.21 3.4 ± 2.3 14.7 ± 0.9 3.0 ± 1.2 7.8 ± 3.2 2.8 ± 1.1
Gluconic acid 10 7.00 7.87 ± 0.09 5.5 ± 0.4 52.6 ± 8.1 5.4 ± 0.7 20 ± 1.5 3.6 ± 0.2
Glucuronic acid 10 7.00 7.87 ± 0.14 1.7±0.4 0.0 ± 0.0 – – –
Galacturonic acid 10 7.00 7.86 ± 0.16 1.6±0.1 0.0 ± 0.0 – – –
Cells were cultivated in NR medium. Initial pH was adjusted to 7.00 Blank 0 7.00 8.22 ± 0.26 1.8 ± 0.2 0.7 ± 1.2 – – –
Fructose 10 7.00 7.69 ± 0.11 3.6 ± 0.1 24.2 ± 10.7 4.1 ± 2.0 8.0 ± 5.2 1.9 ± 0.3
Gluconic acid 10 7.00 8.22 ± 0.08 6.6 ± 0.5 53.3 ± 7.2 5.7 ± 0.6 16 ± 2.0 2.7 ± 0.4
Glucuronic acid 10 7.00 8.25 ± 0.24 1.8 ± 0.2 0.0 ± 0.0 – – –
Galacturonic acid 10 7.00 8.23 ± 0.21 1.6 ± 0.1 0.1 ± 0.2 – – –
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a

Cell Dry Weight.

b

Number-Averaged Molecular Weight.

c

Weight-Averaged Molecular Weight.

d

Polydispersity Index.

e

Not estimated because of insufficient PHA yield for GPC measurements. Data are shown as mean ± SD, n = 3.

To evaluate the effects of the addition of Tris on C. necator H16, Tris-containing culture medium was adjusted to a pH of 7.0 and cultured. The biomass amount and PHA content were equivalent to the results of the respective pH without being adjusted for the blank, fructose, glucuronic acid, and galacturonic acid addition, and the weight-average molecular weight of the PHA obtained from the culture in the fructose-added culture medium was also similar to that of the pH without being adjusted. The results also suggest that the utilization efficiency of glucuronic acid and galacturonic acid by C. necator H16 is low regardless of pH. On the other hand, the biomass of the gluconic acid-added culture medium adjusted to pH 7.00 was 5.5 ± 0.4 g/L, and the PHA content was 52.6 ± 8.1%. This indicates that the biomass increased by approximately 2-fold and the PHA content increased by approximately 3-fold compared to the unadjusted pH gluconic acid medium. These results suggest that the P­(3HB) production of C. necator H16 when utilizing gluconic acid increased greatly due to the adjustment of the pH of the medium.

To eliminate the effect of acidification of the culture medium by the addition of acidic sugars without the addition of Tris, the pH of all types of culture medium was adjusted to 7.00 and cultured alternatively. The biomass amount and PHA content were equivalent to the results of the respective Tris-including cultures for the blank, fructose, glucuronic acid, and galacturonic acid additions. These results indicated that the addition of Tris in the culture medium has little effect on the utilization of fructose and the accumulation of PHA in C. necator H16. The results also showed that the utilization efficiency of glucuronic acid and galacturonic acid by C. necator H16 is low regardless of pH and the addition of Tris. Nonetheless, the biomass of the gluconic acid-added culture medium adjusted to pH 7.0 and the PHA content were the highest at 6.6 ± 0.5 g/L and 53.3 ± 7.2%, respectively. This indicates that the utilization of gluconic acid for growth and P­(3HB) production in C. necator is improved when the medium pH is 7.0 at the start of culturing.

Molecular Weight Distribution of PHA

PHAs biosynthesized by C. necator H16 using gluconic acid as the carbon source displayed a much higher weight-averaged molecular weight than those biosynthesized using fructose. The weight-averaged molecular weight of the PHA obtained from the culture in the fructose-added culture medium was almost identical to that of the Tris-including cultures. The addition of gluconic acid as a carbon source in C. necator H16 showed a high weight-averaged molecular weight (≥1.6 × 106 g mol–1). The results are 1.5- to 2-fold higher than the fructose-added cultures. To further investigate the differences between the weight-averaged molecular weight of C. necator H16 biosynthesized PHAs with fructose and gluconic acid as carbon sources, we examined the molecular weight distribution curve from GPC. Figure shows the molecular weight distribution curve of PHAs biosynthesized by C. necator H16 with fructose or gluconic acid as the carbon source. While all curves showed one peak in the fructose-added cultures, two peaks were observed in the gluconic acid-added cultures. The molecular weight distribution curve of PHAs recovered in the gluconic acid-added cultures showed a second peak shifting toward higher molecular weight. This could indicate that the polymerization of PHA shifted to a higher rate during a certain point in cultivation.

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Molecular weight distribution, as estimated by GPC (calibrated by PS), of P­(3HB) biosynthesized by wild-type C. necator H16.

Physical Properties of PHA

Tensile tests were performed on stretched-annealed P­(3HB)­s biosynthesized using fructose and gluconic acid as carbon sources (Table and Figure a). Both films were stretched to 6 times their original length. The stretched-annealed P­(3HB) films have been shown to exhibit improved mechanical properties, especially in tensile strength, due to the highly oriented crystalline structure and the formation of β-crystals during the drawing process. P­(3HB) biosynthesized with gluconic acid exhibited a tensile strength of 40 ± 2.8 MPa, significantly higher (p < 0.0029) than the tensile strength exhibited by P­(3HB) biosynthesized with fructose (22 ± 2.3 MPa). Commercial P­(3HB) exhibited a tensile strength of 25 ± 6.2 MPa that was similar to P­(3HB) biosynthesized with fructose. While P­(3HB) biosynthesized with gluconic acid exhibited higher tensile strength, both fructose and gluconic acid P­(3HB) films displayed similar elongation at break at 45 ± 21.4% and 36 ± 20.2%, respectively, and commercial P­(3HB) film exhibited the lowest elongation at break (19 ± 5.8%). The brittleness of gluconic acid biosynthesized and commercial P­(3HB) films is explained by the relatively high degrees of crystallinity (59.4% and 54.3% for gluconic acid and commercial P­(3HB), as compared to 43.0% for fructose P­(3HB)). On the other hand, the gluconic acid P­(3HB) exhibited the highest strain energy (0.03 ± 0.01 MJ/m3) among the three types of P­(3HB) films, showing the highest tenacity.

2. Physical Properties of Stretched Annealed P­(3HB) Biosynthesized by Wild-Type C. necator H16.

  P(3HB) Gluconic Acid P(3HB) Fructose P(3HB) Commercial
Lamellae thickness (nm) 11.6 8.72 9.66
15.4    
Degree of crystallinity (%) 59.4 43.0 54.3
ΔHm  (J/g) 94.6 82.6 78.9
Tm  (°C) 173.0 178.3 179.5
Strain energy (J) 0.03 ± 0.01 0.01 ± 0.01 0.009 ± 0.003
Elongation at break (%) 36 ± 20.2 45 ± 21.4 19 ± 5.8
Tensile strength (MPa) 40 ± 2.8 22 ± 2.3 25 ± 6.2
Drawing ratio 6 6 6
M w (×105) 20 ± 1.5 7.8 ± 3.2 1.8 ± 0.2
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(a) Strain–stress curves. (b) DSC curves of stretched annealed P­(3HB) biosynthesized by wild-type C. necator H16.

The thickness of each P­(3HB) lamella was estimated with data from DSC (Table and Figure b). Gluconic acid biosynthesized P­(3HB) showed two melting peaks indicating the existence of two lamellae of different thicknesses. Overall, gluconic acid biosynthesized P­(3HB) had thick crystal lamellae (15.5, 11.6 nm) compared to fructose biosynthesized (8.72 nm) and commercial P­(3HB) film (9.66 nm). Gluconic acid biosynthesized P­(3HB) had both thick and thinner lamellae; while the thinner lamella was stretched and formed β-crystals increasing its strength, the thicker lamella was not, which could account for its brittleness. Commercial P­(3HB) formed thick lamellae; therefore, it is brittle. In the case of fructose biosynthesized P­(3HB), it is likely that intermediate and uniform lamellae were formed; therefore, β-crystals were not observed, while the low crystallinity contributes to its flexibility as the amorphous region could be stretched.

Crystalline Structure of P­(3HB)

To further investigate whether the increased tensile strength was due to the formation of β-crystals, XRD analysis was performed on P­(3HB)­s biosynthesized with fructose and gluconic acid as carbon sources (Figure ). While alpha crystals were observed in both films, a slight peak belonging to the β-crystals was observed at 19° in the P­(3HB) film biosynthesized with gluconic acid. These results indicate that the formation of β-crystals contributed greatly to the improved tensile strength of the gluconic acid biosynthesized P­(3HB) film. Furthermore, the results suggest that P­(3HB) film biosynthesized using gluconic acid with a higher molecular weight than that of fructose and commercial P­(3HB) is more likely to form β-crystals even when drawn at the same ratio. Therefore, when drawn and annealed, P­(3HB) film with higher molecular weight exhibited greater mechanical properties.

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XRD chromatograms of P­(3HB)­s biosynthesized by wild-type C. necator H16, assigned by Wang and Tashiro, 2016.

Discussion

Glucuronic acid and galacturonic acid are pectin-derived uronic acids, and the results from this study indicate that the addition of glucuronic acid or galacturonic acid as the carbon source likely does not contribute to an increase in biomass or PHA accumulation. Genomic analysis of C. necator H16 reported that its natural substrate range of C. necator H16 is reported to be fructose, gluconate, and organic acids, which confirms the absence of annotated metabolic pathways or transporters for glucuronic and galacturonic acids. , Therefore, Cupriavidus necator H16 lacks the capacity to metabolize glucuronic or galacturonic acids because its genome does not encode key uronic acid catabolism genes. , This is consistent with empirical screenings of related strains, such as Cupriavidus taiwanensis and Cupriavidus necator DSM 545, which also fail to utilize galacturonic acid. , Interestingly, even within the closely related Burkholderia genus, reports of uronic acid utilization remain rare, and while certain strains like Burkholderia cepacia exhibit pectinolytic activity, the specific mechanisms and metabolic fluxes have not been elucidated. Considering that there are only a limited number of species that can utilize glucuronic acid and galacturonic acid in Burkholderiaceae, it is understandable that C. necator H16 utilizes neither glucuronic acid nor galacturonic acid.

The PHA accumulation in C. necator H16 commonly occurs under a nutrient-limiting environment, indicating that a nitrogen-absent environment would be more favorable. The addition of Tris adversely increases the ratio of nitrogen present in the culture medium; subsequently, a decrease in PHA production could be expected. However, the addition of Tris under our experimental conditions showed no significant influence on the PHA accumulation in C. necator H16. Therefore, we believe that it is important to explore the potential effects of Tris–HCl on the metabolic flux of C. necator H16 in future work.

C. necator H16 synthesizes and accumulates P­(3HB) efficiently using fructose and gluconic acid as carbon sources. When fructose is used as the carbon source, the initial adjustment and the following shift in the pH of the culture medium have little effect on the utilization of fructose and the accumulation of PHA in C. necator H16. However, in the case of gluconic acid, adjustment of the medium pH led to a significant increase in PHA accumulation, indicating that C. necator H16 utilizes gluconic acid more efficiently at a neutral pH. In addition, the use of gluconic acid as the carbon source produced PHAs with a higher molecular weight than PHAs produced with fructose. Furthermore, multiple studies have reported differences between molecular weight and the carbon source of choice. The use of simple sugars such as fructose led to PHA (P­(3HB)) with a molecular weight of 1.3 × 106 g mol–1. Another study showed that the use of plant oil as a carbon source resulted in PHA (PHBV) with a molecular weight of 3.1 × 106 g mol–1 at maximum. It can be seen that C. necator H16 can produce high molecular weight PHA in the case of fatty acids. In the present case, fructose and gluconic acid are both hexoses which contain the same number of carbons per molecule. In C. necator H16, both fructose and gluconic acid act as metabolic intermediates through the Entner–Doudoroff (ED) pathway. Considering that both are metabolized through the ED pathway and contain the same number of carbons per molecule, differences in the molecular weight of PHAs biosynthesized by C. necator H16 are unlikely due to the differences in carbon sources. Thus, the differences in molecular weight could be attributable to the fluctuations in medium pH during cultivation. The pH of the culture media shifted from 7.00 to 8.22 ± 0.08 in gluconic acid-added ones, while from 7.00 to 7.69 ± 0.11 in fructose-added ones, depending on the initial adjustment of pH prior to cultivation or the addition of Tris. These results indicate that the addition of gluconic acid shifted the culture toward more basic conditions. This could imply that the polymerization of PHA occurs at a more efficient rate under basic conditions; hence, PHA produced with gluconic acid as the carbon source exhibited a higher average molecular weight.

The molecular weight distribution of PHAs produced with fructose and gluconic acid as carbon sources also showed different characteristics. PHAs produced with gluconic acid had bimodal molecular weight distribution curves which were not observed when fructose was used as the carbon source. As we speculated, this could possibly be caused by the shift in culture medium pH toward basic conditions, contributing to the production of high molecular weight PHAs synthesized by C. necator H16. Although the data suggest that PHA polymerization may increase during a specific phase of cultivation, the mechanistic basis for this shift cannot be conclusively determined from the current data set. Therefore, detailed mechanistic interpretation remains a limitation of the present work and should be addressed in future studies. Previous studies showed that Escherichia coli harboring C. necator H16 genes produces PHAs with higher molecular weight under slightly acidic conditions. However, there was also a report showing that the molecular weight of PHAs produced was not affected by the pH of the culture medium directly and instead could influence other factors such as the expression level of PHA synthase. This could indicate that the shift of the culture medium’s pH toward basic conditions influenced the PHA synthase expression in the case of C. necator H16. The molecular weight of PHAs synthesized is known to be affected by factors such as the concentration of PHA synthase and its catalytic activity, as well as the degradation of PHA during biosynthesis. , Therefore, we investigated the expression of phaC, the PHA synthase gene in C. necator H16, to elucidate the difference in molecular weight when gluconic acid is used. The expression of PHA synthase was relatively lower throughout cultivation when gluconic acid was used compared to fructose (Figure S2). A previous study has shown that the molecular weight of P­(3HB) is negatively correlated to the specific activity of PhaC. Hence, the reason gluconic acid yielded P­(3HB) of higher molecular weight could be due to the low expression of PHA synthase during cultivation. Though gluconic acid yielding P­(3HB) with a higher molecular weight than fructose may also be attributed to differences in intracellular monomer supply or PHA synthase regulation during gluconate metabolism, which could influence chain-elongation kinetics and chain-transfer frequency, elucidating the precise mechanism will require targeted metabolic and enzymatic analyses, which represent an important direction for future work. While past reports have shown that the lag phase under heterotrophic growth of C. necator can be shortened depending on the choice of carbon source (acetate) and its concentration. Therefore, we believe that it is important to investigate the growth kinetics as well as the changes in the composition of the culture medium throughout the culturing process, and the metabolome and gene expression of C. necator H16 in future work.

P­(3HB) film with higher molecular weight displayed better tensile strength and the formation of β-crystals could be observed, whereas P­(3HB) film with lower molecular weight only formed α-crystals. The formation of β-crystals occurs during the stretching process of P­(3HB) film where the amorphous tie chains transform into an extended all-trans conformation (Figure ). Thus, the formation of β-crystals is highly dependent on the amount of tie chains present in the amorphous region. The formation of β-crystals could only be observed in gluconic acid biosynthesized P­(3HB) film which indicates that there are more tie chains present in high molecular weight P­(3HB). Commercial P­(3HB) possesses the lowest molecular weight; thus, it exhibited comparatively low tensile strength and elongation at break due to the lack of tie chains. In fructose biosynthesized P­(3HB), there are less tie molecular chains, and consequently, better elongation could be observed without the beta structure. As a result, an increase in tensile strength was not observed. However, in the case of gluconic acid biosynthesized P­(3HB), an increase in tensile strength was observed attributed to the formation of β structure, while the elongation was slightly lower than that of fructose, as the transition of amorphous tie molecular chains into β structure enhanced the rigidity. These results also concur with past research where higher molecular weight PHA exhibits better mechanical properties even at the same drawing ratio. Only gluconic acid biosynthesized P­(3HB) film formed β-crystals which indicated that higher molecular weight PHA is more favorable for the formation of β-crystals.

4.

4

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Crystalline structure of P­(3HB)­s biosynthesized by wild-type C. necator H16.

The utilization of gluconic acid, when sourced from biomass hydrolyzates or industrial side streams, offers strong feasibility and cost advantages as it can be generated at low cost through glucose oxidation or derived from agro-industrial residues, potentially lowering feedstock expenses and improving overall process sustainability while producing higher-quality P­(3HB) compared to conventional sugars such as fructose.

Conclusions

This study demonstrates that among the sugar acids examined, only gluconic acid served as an effective carbon source for both growth and P­(3HB) biosynthesis in Cupriavidus necator H16. In contrast, glucuronic acid and galacturonic acid did not support biomass formation or PHA accumulation, likely due to the absence of uronic-acid catabolic genes in the metabolic pathways of C. necator H16. When gluconic acid was supplied at an initial medium pH of 7.00, C. necator exhibited enhanced biomass accumulation and increased intracellular P­(3HB) contentmore than 3-fold compared to unadjusted conditionsindicating that neutral pH strongly facilitates PHA accumulation. More importantly, PHAs synthesized from gluconate displayed significantly higher weight-average molecular weight (M w ≥ 1.6 × 106 g mol–1) than PHAs synthesized from fructose, along with a characteristic bimodal molecular-weight distribution suggestive of pH-dependent shifts in polymerization kinetics or PHA synthase activity during cultivation. The elevated molecular weight correlated directly with improved mechanical performance of stretched-annealed P­(3HB) films, including higher tensile strength and β-crystal formation relative to fructose-derived or commercial P­(3HB).

Collectively, these findings establish gluconic acid as a promising, non-GMO-compatible substrate for generating high-molecular-weight P­(3HB) using wild-type C. necator H16. Because gluconic acid can be produced readily from low-value, glucose-rich biomass streams, its use also holds potential for cost-effective, scalable PHA bioproduction. The genetic modification-free PHA production can also contribute to the production of biodegradable plastic materials that can be utilized in applications where the materials originating from genetically modified microorganisms must be avoided. Further studies integrating metabolic profiling, gene-expression analysis, and controlled bioreactor operation will be essential to elucidate the mechanisms through which gluconate metabolism and medium alkalization influence PHA polymerization and to optimize this strategy for industrial manufacturing of high-performance biodegradable polymers.

Supplementary Material

ao5c10104_si_001.pdf (183.2KB, pdf)

Acknowledgments

J.Y.C. acknowledges the financial support from Ajinomoto Scholarship Foundation, Japan. The present study was also supported by a Grant in Aid for Basic Research (B) (General) from Japan Society for the Promotion of Science, Japan (No. 25K02146).

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.5c10104.

  • Results of P­(3HB) production in NR medium without initial pH adjustment or with 1 M phosphate-buffered saline (PBS), a typical gas chromatogram as raw data of P­(3HB) quantification, and quantitative PCR for the evaluation of phaC expression, and a gene encoding the polymerization enzyme of P­(3HB) in vivo (PDF)

J.Y.C. performed the microbial production and characterization of PHA, data analysis, and manuscript drafting. K.T. supported the microbial production and characterization of PHA. M.K. advised the microbial production and characterization of PHA and revised the manuscript. D.I. performed the conceptualization of the study, advised the microbial production and characterization of PHA, and revised the manuscript.

The authors declare no competing financial interest.

References

  1. Mehnaz S., Javaid A.. Microbes and Plastic Waste Management. Environ. Sustainability. 2020;3:337–339. doi: 10.1007/s42398-020-00149-3. [DOI] [Google Scholar]
  2. Narancic T., O’Connor K. E.. Plastic Waste as a Global Challenge: Are Biodegradable Plastics the Answer to the Plastic Waste Problem? Microbiology. 2019;165:129–137. doi: 10.1099/mic.0.000749. [DOI] [PubMed] [Google Scholar]
  3. Alabi O. A., Ologbonjaye K. I., Awosolu O., Alalade O. E.. Public and Environmental Health Effects of Plastic Wastes Disposal: A Review. J. Toxicol. Risk Assess. 2019;5(2):1510021. doi: 10.23937/2572-4061.1510021. [DOI] [Google Scholar]
  4. Narancic T., Cerrone F., Beagan N., O’Connor K. E.. Recent Advances in Bioplastics: Application and Biodegradation. Polymers. 2020;12:920. doi: 10.3390/polym12040920. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Börner T., Zinn M.. Key Challenges in the Advancement and Industrialization of Biobased and Biodegradable Plastics: A Value Chain Overarching Perspective. Front. Bioeng. Biotechnol. 2024;12:1406278. doi: 10.3389/fbioe.2024.1406278. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Li Y., Tao L., Wang Q., Wang F., Li G., Song M.. Potential Health Impact of Microplastics: A Review of Environmental Distribution, Human Exposure, and Toxic Effects. Environ. Health. 2023;1:249–257. doi: 10.1021/envhealth.3c00052. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Koller, M. ; Mukherjee, A. ; Obruca, S. ; Zinn, M. . Polyhydroxyalkanoates (PHA): Microbial Synthesis of Natural Polyesters. In In Microbial Production of High-Value Products; Microbiology Monographs, Rehm, B. H. A. ; Wibowo, D. , Eds.; Springer International Publishing: Cham, 2022; Vol. 37, pp. 185–236. [Google Scholar]
  8. Sharma V., Sehgal R., Gupta R.. Polyhydroxyalkanoate (PHA): Properties and Modifications. Polymer. 2021;212:123161. doi: 10.1016/j.polymer.2020.123161. [DOI] [Google Scholar]
  9. Saravanan K., Umesh M., Kathirvel P.. Microbial Polyhydroxyalkanoates (PHAs): A Review on Biosynthesis, Properties, Fermentation Strategies and Its Prospective Applications for Sustainable Future. J. Polym. Environ. 2022;30:4903–4935. doi: 10.1007/s10924-022-02562-7. [DOI] [Google Scholar]
  10. Choi S. Y., Rhie M. N., Kim H. T., Joo J. C., Cho I. J., Son J., Jo S. Y., Sohn Y. J., Baritugo K.-A., Pyo J., Lee Y., Lee S. Y., Park S. J.. Metabolic Engineering for the Synthesis of Polyesters: A 100-Year Journey from Polyhydroxyalkanoates to Non-Natural Microbial Polyesters. Metab. Eng. 2020;58:47–81. doi: 10.1016/j.ymben.2019.05.009. [DOI] [PubMed] [Google Scholar]
  11. Ino K., Sato S.. Ultrahigh-Molecular-Weight Polyhydroxyalkanoates: Microbial Production and Mechanical Properties. Oleoscience. 2021;21:517–523. doi: 10.5650/oleoscience.21.517. [DOI] [Google Scholar]
  12. Kusaka S., Iwata T., Doi Y.. Microbial Synthesis and Physical Properties of Ultra-High-Molecular-Weight Poly­[(R)-3-hydroxybutyrate] J. Macromol. Sci., Part A. 1998;35(2):319–335. doi: 10.1080/10601329808001980. [DOI] [Google Scholar]
  13. Tsuge T.. Fundamental Factors Determining the Molecular Weight of Polyhydroxyalkanoate during Biosynthesis. Polym. J. 2016;48:1051–1057. doi: 10.1038/pj.2016.78. [DOI] [Google Scholar]
  14. Higuchi-Takeuchi M., Morisaki K., Toyooka K., Numata K.. Synthesis of High-Molecular-Weight Polyhydroxyalkanoates by Marine Photosynthetic Purple Bacteria. PLoS One. 2016;11:e0160981. doi: 10.1371/journal.pone.0160981. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Dalton B., Bhagabati P., De Micco J., Padamati R. B., O’Connor K.. A Review on Biological Synthesis of the Biodegradable Polymers Polyhydroxyalkanoates and the Development of Multiple Applications. Catalysts. 2022;12:319. doi: 10.3390/catal12030319. [DOI] [Google Scholar]
  16. Bellini S., Tommasi T., Fino D.. Poly­(3-Hydroxybutyrate) Biosynthesis by Cupriavidus Necator: A Review on Waste Substrates Utilization for a Circular Economy Approach. Biores. Technol. Rep. 2022;17:100985. doi: 10.1016/j.biteb.2022.100985. [DOI] [Google Scholar]
  17. Sakamoto M., Kimura Y., Ishii D., Nakaoki T.. Biosynthesis of Poly­(3-Hydroxyalkanoate) from Amino Acids in Medium with Nitrogen, Phosphate, and Magnesium, or Some Combination of These Nutrients. J. Polym. Environ. 2014;22:488–493. doi: 10.1007/s10924-014-0656-9. [DOI] [Google Scholar]
  18. Morlino M. S., Serna García R., Savio F., Zampieri G., Morosinotto T., Treu L., Campanaro S.. Cupriavidus Necator as a Platform for Polyhydroxyalkanoate Production: An Overview of Strains, Metabolism, and Modeling Approaches. Biotechnol. Adv. 2023;69:108264. doi: 10.1016/j.biotechadv.2023.108264. [DOI] [PubMed] [Google Scholar]
  19. Shcherbakova-Sandu M. P., Meshcheryako E. P., Gulevich S. A., Kushwaha A. K., Kumar R., Sonwane A. K., Samal S., Kurzina I. A.. Use of Glucose Obtained from Biomass Waste for the Synthesis of Gluconic and Glucaric Acids: Their Production, Application, and Future Prospects. Molecules. 2025;30(14):3012. doi: 10.3390/molecules30143012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. González-Villanueva M., Galaiya H., Staniland P., Staniland J., Savill I., Wong T. S., Tee K. L.. Adaptive Laboratory Evolution of Cupriavidus necator H16 for Carbon Co-Utilization with Glycerol. Int. J. Mol. Sci. 2019;20:5737. doi: 10.3390/ijms20225737. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Guimarães T. C., Araújo E. S., Hernández-Macedo M. L., López J. A.. Polyhydroxyalkanoates: Biosynthesis from Alternative Carbon Sources and Analytic Methods: A Short Review. J. Polym. Environ. 2022;30:2669–2684. doi: 10.1007/s10924-022-02403-7. [DOI] [Google Scholar]
  22. Kamilah H., Al-Gheethi A., Yang T. A., Sudesh K.. The Use of Palm Oil-Based Waste Cooking Oil to Enhance the Production of Polyhydroxybutyrate [P­(3HB)] by Cupriavidus necator H16 Strain. Arab. J. Sci. Eng. 2018;43:3453–3463. doi: 10.1007/s13369-018-3118-1. [DOI] [Google Scholar]
  23. Mothes G., Schnorpfeil C., Ackermann J.-.. Production of PHB from Crude Glycerol. Eng. In Life Sci. 2007;7(5):475–479. doi: 10.1002/elsc.200620210. [DOI] [Google Scholar]
  24. Zhang L., Jiang Z., Tsui T.-H., Loh K.-C., Dai Y., Tong Y. W.. A Review on Enhancing Cupriavidus Necator Fermentation for Poly­(3-Hydroxybutyrate) (PHB) Production From Low-Cost Carbon Sources. Front. Bioeng. Biotechnol. 2022;10:946085. doi: 10.3389/fbioe.2022.946085. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Scholz P., Chapman K. D., Ischebeck T., Guzha A.. Analysis of Pectin-derived Monosaccharides from Arabidopsis Using GC–MS. Bio-Protocol. 2023;13(16):e4746. doi: 10.21769/BioProtoc.4746. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Locatelli G., Da Silva G. D., Finkler L., Finkler C. L. L.. Acid Hydrolysis of Pectin for Cell Growth of Cupriavidus Necator . Biotechnology. 2011;11:29–36. doi: 10.3923/biotech.2012.29.36. [DOI] [Google Scholar]
  27. Zubairi S. I., Bismarck A., Mantalaris A.. The Effect of Surface Heterogeneity on Wettability of Porous Three Dimendional (3-D) Scaffolds of Poly­(3-Hydroxybutyric Acid) (PHB) and Poly­(3-Hydroxybutyric-co-3-Hydroxyvaleric Acid) (PHBV) J. Teknol. 2015;75:305–312. doi: 10.11113/jt.v75.3960. [DOI] [Google Scholar]
  28. Righetti M. C., Lorenzo M. L. D.. Nonlinear Determination of the Equilibrium Melting Temperature from Initial Nonreorganized Crystals of Poly­(3-Hydroxybutyrate) Polym. Eng. Sci. 2012;52:2383–2390. doi: 10.1002/pen.23199. [DOI] [Google Scholar]
  29. Perret E., Reifler F. A., Gooneie A., Chen K., Selli F., Hufenus R.. Structural Response of Melt-Spun Poly­(3-Hydroxybutyrate) Fibers to Stress and Temperature. Polymer. 2020;197:122503. doi: 10.1016/j.polymer.2020.122503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Kabe T., Tanaka T., Marubayashi H., Hikima T., Takata M., Iwata T.. Investigating Thermal Properties of and Melting-Induced Structural Changes in Cold-Drawn P­(3HB) Films with α- and β-Structures Using Real-Time X-Ray Measurements and High-Speed DSC. Polymer. 2016;93:181–188. doi: 10.1016/j.polymer.2016.04.028. [DOI] [Google Scholar]
  31. Pohlmann A., Fricke W. F., Reinecke F., Kusian B., Liesegang H., Cramm R., Eitinger T., Ewering C., Pötter M., Schwartz E., Strittmatter A., Voß I., Gottschalk G., Steinbüchel A., Friedrich B., Bowien B.. Genome Sequence of the Bioplastic-Producing “Knallgas” Bacterium Ralstonia Eutropha H16. Nat. Biotechnol. 2006;24:1257–1262. doi: 10.1038/nbt1244. [DOI] [PubMed] [Google Scholar]
  32. Cramm R.. Genomic View of Energy Metabolism in Ralstonia Eutropha H16. J. Mol. Microbiol. Biotechnol. 2008;16:38–52. doi: 10.1159/000142893. [DOI] [PubMed] [Google Scholar]
  33. Kutralam-Muniasamy G., Perez-Guevara F.. Genome characteristics dictate poly-R-(3)-hydroxyalkanoate production in Cupriavidus necator H16. World J. Microbiol. Biotechnol. 2018;34(6):79. doi: 10.1007/s11274-018-2460-5. [DOI] [PubMed] [Google Scholar]
  34. Locatelli G. O., Finkler L., Finkler C. L. L.. Comparison of Acid and Enzymatic Hydrolysis of Pectin, as Inexpensive Source to Cell Growth of Cupriavidus Necator . An. Acad. Bras. Ciênc. 2019;91(2):e20180333. doi: 10.1590/0001-3765201920180333. [DOI] [PubMed] [Google Scholar]
  35. Wei Y.-H., Chen W.-C., Huang C.-K., Wu H.-S., Sun Y.-M., Lo C.-W., Janarthanan O.-M.. Screening and Evaluation of Polyhydroxybutyrate-Producing Strains from Indigenous Isolate Cupriavidus Taiwanensis Strains. Int. J. Mol. Sci. 2011;12:252–265. doi: 10.3390/ijms12010252. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Shoily S. S., Fatema K., Dina R. B., Biswas A., Haque P., Rahman M. M., Uddin M., Sajib Z. A.. The Pectinolytic Activity of Burkholderia Cepacia and Its Application in the Bioscouring of Cotton Knit Fabric. J. Gen. Eng. Biotechnol. 2023;21:136. doi: 10.1186/s43141-023-00596-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Arikawa H., Sato S., Fujiki T., Matsumoto K.. A Study on the Relation between Poly­(3-Hydroxybutyrate) Depolymerases or Oligomer Hydrolases and Molecular Weight of Polyhydroxyalkanoates Accumulating in Cupriavidus Necator H16. J. Biotechnol. 2016;227:94–102. doi: 10.1016/j.jbiotec.2016.04.004. [DOI] [PubMed] [Google Scholar]
  38. Lee W.-H., Loo C.-Y., Nomura C. T., Sudesh K.. Biosynthesis of Polyhydroxyalkanoate Copolymers from Mixtures of Plant Oils and 3-Hydroxyvalerate Precursors. Biores. Technol. 2008;99:6844–6851. doi: 10.1016/j.biortech.2008.01.051. [DOI] [PubMed] [Google Scholar]
  39. Bocanegra J. K., Da Cruz Pradella J. G., Da Silva L. F., Taciro M. K., Gomez J. G. C.. Influence of pH on the Molecular Weight of Poly-3-Hydroxybutyric Acid (P3HB) Produced by Recombinant Escherichia Coli . Appl. Biochem. Biotechnol. 2013;170:1336–1347. doi: 10.1007/s12010-013-0257-4. [DOI] [PubMed] [Google Scholar]
  40. Sim S. J., Snell K. D., Hogan S. A., Stubbe J., Rha C., Sinskey A. J.. PHA Synthase Activity Controls the Molecular Weight and Polydispersity of Polyhydroxybutyrate in Vivo . Nat. Biotechnol. 1997;15:63–67. doi: 10.1038/nbt0197-63. [DOI] [PubMed] [Google Scholar]
  41. Hiroe A., Tsuge K., Nomura C. T., Itaya M., Tsuge T.. Rearrangement of Gene Order in the phaCAB Operon Leads to Effective Production of Ultrahigh-Molecular-Weight Poly­[(R)-3-Hydroxybutyrate] in Genetically Engineered Escherichia Coli . Appl. Environ. Microbiol. 2012;78:3177–3184. doi: 10.1128/AEM.07715-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Amer A., Kim Y.. Minimizing the Lag Phase of Cupriavidus Necator Growth under Autotrophic, Heterotrophic, and Mixotrophic Conditions. Appl. Environ. Microbiol. 2023;89(2):e02007–22. doi: 10.1128/aem.02007-22. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Wang H., Tashiro K.. Reinvestigation of Crystal Structure and Intermolecular Interactions of Biodegradable Poly­(3-Hydroxybutyrate) α-Form and the Prediction of Its Mechanical Property. Macromolecules. 2016;49(2):581–594. doi: 10.1021/acs.macromol.5b02310. [DOI] [Google Scholar]

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