Microbial production of polyhydroxyalkanoates and lactate-based biopolymers from C1 resources: current advances and trends

Abstract

With the advancement of industrialization, the environmental burden caused by non-biodegradable petroleum-based plastics has become increasingly severe. Biodegradable plastics have attracted considerable attention due to their inherent degradability and environmental compatibility, with significant potential for applications across food, agriculture, industry and medicine. However, the high production cost of biodegradable plastics remains a critical barrier to their large-scale commercialization. And the synthesis of bioplastics using cost-effective C1 resources has emerged as a promising solution. This review highlights recent progress in the microbial production of polyhydroxyalkanoates (PHAs) and lactate-based biopolymers derived from C1 substrates. Key advancements in microbial strain and process innovations are addressed, and characterization techniques and biodegradation mechanisms are summarized. And current challenges and future prospects for enhancing the sustainable production of PHAs and lactate-based biopolymers from C1 resources are discussed. This comprehensive overview offers viable strategies for promoting environmentally responsible alternatives to petroleum-based plastics and advancing global efforts toward carbon neutrality.

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Yong Wang

Dr Yong Wang received his Ph.D. in Chemical Engineering and Technology from Beijing University of Chemical Technology (BUCT). He currently works as an associate professor at the College of Food Science and Biology, Hebei University of Science and Technology. His research interests focuses on green biomanufacturing of non-food bio-based materials derived from C1 resources, integrated biomass utilization through innovative biorefining technologies, and performance enhancement of industrial microbial fermentation systems.

Jiaxin Liang

Jiaxin Liang is currently a Master's student at the College of Food Science and Biology, Hebei University of Science and Technology. Her research specializes in the construction of polylactic acid metabolic pathways and enzyme engineering for polylactic acid production in Chlorella pyrenoidosa. Her work integrates metabolic engineering and synthetic biology approaches to optimize PLA biosynthesis efficiency, aiming to advance environmentally sustainable bioplastic production.

Dejing Kong

Dr Dejing Kong received his Ph.D. in Genetics from University of Chinese Academy of Sciences (UCAS). He is currently a faculty member in the College of Food Science and Biology at Hebei University of Science and Technology. His research mainly focuses on synthetic biology and the development of gene editing tools (such as CRISPR-Cas) to precisely modify the genetic material of microorganisms (including bacteria, yeasts, and microalgae).

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Green foundation

1. Intensified human activities have substantially elevated exhaust gas emissions. The resulting C1 compounds can serve as sustainable and low-cost feedstocks for biosynthesis of PHAs and lactate-based biopolymers. And this bioconversion approach offers an efficient strategy for environmental sustainability.

2. This review systematically outlines recent progress in the microbial production of PHAs and lactate-based biopolymers derived from C1 substrates. Recent advancements in microbial strains and process innovations are addressed, and characterization techniques and biodegradation mechanisms are summarized.

3. Exploring new microbial resources, regulatory mechanisms and smarter fermentation processes will drive enhancements in bioproduction efficiency of PHAs and lactate-based biopolymers for economically viable bioproduction. And combinatorial characterization methods and deeper understanding of biodegradation mechanisms will further support the environmentally responsible application of PHAs and lactate-based biopolymers.

1. Introduction

Since the invention of the semi-synthetic plastic Parkesine as a substitute for ivory,¹ the global demand for plastics has increased steadily. By mid-2017, approximately 8.3 billion tons of plastics had been produced worldwide, resulting in around 6.3 billion tons of plastic waste, of which 79% accumulated on land or in natural environments, 12% was incinerated, and only 9% was recycled.² During the COVID-19 pandemic in 2019, heightened awareness of personal protection significantly increased the use of single-use plastic products.³ Particularly in February 2020, China's daily production of disposable masks surged to 116 million units.⁴ Weathered plastic particles from personal protective equipment, along with microplastics and nanoplastics (NPs), can adsorb hazardous chemicals and pathogenic microorganisms, thereby threatening ecosystems, biota, occupational safety, and human health.⁵ In response to the surge in plastic production over the past decade, the United Nations has adopted various measures, including the United Nations Environment Assembly resolution on marine debris and microplastics, and the Sustainable Development Goals.⁶ And plastic pollution is now considered a global crisis, necessitating coordinated international efforts to support sustainable economic transitions.⁷

Biopolymers represent a distinctive class of polymers regarded as environmentally friendly due to their biodegradability and biocompatibility. These materials are typically derived from renewable resources, such as forestry and agricultural biomass.^8,9 Polyhydroxyalkanoates (PHAs) and lactate-based biopolymers exhibit biodegradability and biocompatibility and are widely applied in products such as disposable tableware and films. PHAs constitute a class of bioplastics synthesized by microorganisms and are widely recognized as promising alternatives to conventional synthetic plastics (polyethylene or polypropylene). PHAs exhibit desirable properties, including mechanical strength, thermoplastic processability, biodegradability, and biocompatibility.¹⁰ And bacterial monocultures have achieved PHAs accumulation levels of up to 90% of DCW.¹¹ Based on polymer chain length, PHAs are categorized into short-chain-length PHAs (scl-PHAs, C3 to C5) and medium-chain-length PHAs (mcl-PHAs, C6 to C14).¹² Compared with scl-PHAs, mcl-PHAs are more elastic, amorphous, and adhesive in nature.¹³ From a compositional standpoint, microbial PHAs mainly include polyhydroxybutyrate (PHB), poly(3-hydroxyvalerate) (PHV), and their copolymer poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV). The physical properties of PHAs vary with composition, as demonstrated by changes in PHBV characteristics when the molar ratio (mol%) of 3-hydroxybutyrate (3HB) to 3-hydroxyvalerate (3HV) is altered.¹⁴ In addition, a copolyester of 3-hydroxybutyrate and 3-hydroxyhexanoate poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) (PHBHHx), is notable for its superior flexibility and excellent marine biodegradability.¹⁵ And the biocompatibility and lack of genotoxicity of poly(3-hydroxybutyrate-co-4-hydroxybutyrate) (P(3HB-co-4HB)) make it a highly suitable material for absorbable medical applications.¹⁶ Lactate-based biopolymers comprise poly(lactic acid) (PLA) homopolymers and lactic acid (LA)-containing copolymers.¹⁷ Lactate-based biopolymer is a thermoplastic biopolymer that is both biodegradable and renewable, encompassing various polymers synthesized from LA, such as poly(D-lactic acid) (PDLA), poly(D,L-lactic acid) (PDLLA) and poly(L-lactic acid) (PLLA).¹⁸ LA-containing copolymers typically include poly(lactate-co-3-hydroxybutyrate) [P(LA-co-3HB)], poly(lactic acid-co-3-hydroxybutyrate-co-3-hydroxypropionate) [P(LA-co-3HB-co-3HP)],¹⁹ and poly(lactic acid-co-3-hydroxybutyrate-co-3-hydroxyvalerate) [P(LA-co-3HB-co-3HV)].^20,21 Lactate-based biopolymers production offers several advantages, as its monomer, LA, can be obtained via fermentation of renewable feedstocks, including rice bran, sweet sorghum juice, sweet sorghum bagasse, corn stover, cassava bagasse, and rice straw.^22–28 Currently, lactate-based biopolymers are applied in a wide range of areas, including 3D printing filaments, fibers, films, cups, food containers, packaging, textiles, surgical sutures, and tissue scaffolds.¹⁸ Furthermore, lactate-based biopolymers oligomers have been shown to function as antibacterial agents for biosafety and environmental protection.²⁹ High-molecular-weight lactate-based biopolymers (≥100 000 Da) are suitable for producing fibers, nonwovens, and durable structural materials.¹⁸ Despite advantages, production cost of biodegradable plastics remains relatively high compared with conventional plastics, which significantly restricts their broader application.³⁰ Therefore, there is an urgent need for new, sustainable, and low-cost substrates, particularly those derived from C1 carbon sources, which are byproducts of industrial and anthropogenic activities.

Since the onset of the Industrial Revolution, extensive human activities have significantly increased the emission of exhaust gases (e.g. CO₂, CH₄ and CO) into the atmosphere.^31–33 Although physical and chemical approaches can be used to reduce such emissions, these methods are typically energy-intensive and economically inefficient. An alternative and more sustainable strategy involves the bioconversion of C1 gases into biodegradable biopolymers via microbial systems.³⁴ Formic acid and methanol can be generated from CO and CO₂ through electrochemical conversion or inorganic catalysis.^35–41 And the storage of carbon intermediates in the form of formic acid or methanol is advantageous for stability and microbial utilization in synthesizing biopolymers.⁴² Therefore, the C1 resources discussed in this review include CO₂, CH₄, CO, synthetic gas, methanol, and formic acid. The production of biopolymers is closely linked to sustainable development, and the choice of carbon source is pivotal. Compared to traditional sugars and vegetable oils, which rely on agricultural production and are susceptible to multiple constraints, C1 resources break through the agricultural dependency and ecological limitations of conventional carbon sources, demonstrating significant sustainability and long-term cost advantages. The production costs of PHAs and lactate-based polymers using traditional carbon sources are 4–9 times higher than those of conventional plastics,⁴³ representing one of the major bottlenecks in their production. Although C1 resource conversion technologies are still in the developmental stage (e.g. low efficiency in microbial metabolic pathways), their core advantage lies in the ability to replace high-cost traditional organic carbon sources with nearly zero-cost C1 substrates.⁴⁴ Moreover, it offers a more environmentally friendly pathway for biopolymer production, particularly against the backdrop of increasing global attention to climate change and resource sustainability, where the importance of C1 resources is becoming ever more prominent. With continuous advancements in biocatalytic technologies, the application potential of C1 resources in the biopolymer industry will be further unlocked. Therefore, advancing C1 bioconversion technologies is not only a technical challenge in developing new synthetic pathways for biopolymers but also holds strategic significance for building a future green, low-carbon, and more resilient bio-based manufacturing industrial system.

Despite challenges such as low efficiency in natural metabolic pathways and insufficient process intensification, the bioconversion of waste-derived C1 compounds into PHAs and lactate-based polymers has been achieved through genetic modification of microbial strains, optimization of culture conditions, and innovation in bioconversion technologies. Research on microbial platforms for PHB synthesis from CO₂ contributed insights into carbon recycling.⁴⁵ And evaluations of PHAs biosynthesis by wild-type and metabolically engineered strains (including microalgae and cyanobacteria) using C1 substrates enhanced environmental sustainability.^46,47 The development of cyanobacteria and Pichia pastoris (P. pastoris) cell factories for LA-based polymers (or its precursor) production from CO₂ and methanol reflect part of the recent advancements.^48,49 Previous studies highlighted the great potential of C1 resources bioconversion for PHAs and lactate-based biopolymers. However, the C1 bioconversions, degradation mechanisms, and characterization approaches for PHAs and lactate-based polymers have not yet been elaborated to a relatively comprehensive extent. This study is dedicated to comprehensively dissecting the complete microbial synthesis chain, from the utilization of initial substrates and the synthesis of intermediate metabolites to the formation and degradation of final products. It supplements the functional roles and mechanistic actions of key genes and their encoded enzymes within specific metabolic pathways (e.g. regulatory mechanisms in carbon metabolic flux allocation). Furthermore, it systematically evaluates various characterization methods employed to depict this complex process, including their advantages, limitations, and respective applicable scopes. Through this in-depth and integrative analysis, the study aims to provide a solid theoretical foundation and practical reference for advancing the green, efficient, and sustainable development of the biodegradable plastics industry (Fig. 1).

Fig. 1 Microbial production of polyhydroxyalkanoates and lactate-based biopolymers.

2. Microbial cell factories for PHAs and lactate-based polymers biosynthesis from C1 resources

2.1. Microorganisms for producing PHAs based on C1 substrates

2.1.1. Microalgae. Great potential has been demonstrated for microalgae in the field of bioenergy production.^50,51 Unlike heterotrophic microorganisms, microalgae can thrive on waste substrates such as municipal and industrial wastewater and simultaneously accumulate valuable compounds, including pigments, lipids, and proteins, while improving environmental quality.^52,53 And microalgae are favored for simple cultivation requirement, rapid growth rate, and ease of harvesting.⁵⁴ PHAs have been reported to be biosynthesized from CO₂ by microalgae, which include prokaryotic microalgae (e.g. Nostocales, Synechococcus, and Oscillatoriaceae) and eukaryotic microalgae (e.g. Naviculales). As indicated in Table 1, photosynthetic PHAs production offers a promising route for carbon capture, with significant potential for cost reduction.⁴⁸ With advances in genetic engineering, microalgae are emerging as sustainable platforms that integrate microbial biofactory capabilities with photosynthetic efficiency.⁵⁵

Table 1 PHAs production by microalgae utilizing CO₂

Strain	Genetic modification	Culture conditions	PHAs production				Ref.
			Type	Content (wt%)	Productivity	Titer (mg L^−1)
a Wild-type strain. b Not reported.
T. distorta TISTR 8985	W^a	14 days of phototrophic growth followed by 6 days of nitrogen-deficient dark incubation	PHB	N^b	2.20 mg g^−1 d^−1	N	58
N. muscorum TISTR 8871					0.80 mg g^−1 d^−1
C. fritschii TISTR 854					0.30 mg g^−1 d^−1
A. spiroides TISTR 8075			PHBV		0.50 mg g^−1 d^−1
C. scytonemicola TISTR 8095	W	50 μmol photon per m^2 per s light at 28 °C normal growth condition for 30 d and specific nutrient deprivation condition for 14 d	PHB	25.40	8.10 mg L^−1 d^−1	356.50	59
O. okeni TISTR 8549	W	75 μmol m^−2 s^−1 continuous white light at 32 °C for 28 d	PHBV (5.5 mol%HV)	14.00	3.68 mg L^−1 d^−1	103.00	60
	W			∼10.00	N	N
Synechocystis sp. PCC 6803	The native phaAB gene was overexpressed into Synechocystis chromosome	40 μE m^−2 s^−1 continuous illumination at 160 rpm and 30 °C	PHB	26.00	N	N	63
Synechocystis sp. PCC 6803	XfpK overexpression in a double Pta and Ach deletion background	18 h white light sinusoidally varying between 10 and 260 μE m^−2 s^−1, 6 h dark and 28 °C	PHB	∼12.00	7.30 mg L^−1 d^−1	232.00	66
Synechocystis sp. PCC 6803	ΔpirC-REphaAB	Light/dark rhythm (12 h each) illumination of 50 µE and 28 °C at 125 rpm	PHB	63.00	N	N	64
S. elongatus UTEX 2973	Heterologous expression phaCAB genes derived from Cupriavidus necator H16	200 μE m^−2 s^−1 continuous fluorescent light and 38 °C at 130 rpm	PHB	16.70	75.2 mg L^−1 d^−1	420.00	62
P. tricornutum	Heterologous expression of genes phaA, phaB and phaC ptimized for codons (from Ralstonia europha)	50 µmol photons per m^2 per s and 21 °C	PHB	1.70	3.49 mg L^−1 d^−1	27.90	65
Synechocystis sp. PCC 6714	UV light Random mutation	60 µmol photons per m^2 per s and 300 rpm	PHB	37.00	36.90 mg L^−1 d^−1	N	69

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Wild-type strains capable of producing PHAs from CO₂ as the sole carbon source have been extensively screened. As a type of prokaryotic microalgae, cyanobacteria grow autotrophically and store carbon in the form of PHB.^56,57 A study assessed 20 evolutionarily distinct cyanobacteria, identifying Anabaena spiroides TISTR 8075 (PHBV producer) and Tolypothrix distorta TISTR 8985, Nostoc muscorum TISTR 8871, and Chlorogloea fritschii TISTR 854 (PHB producers), though accumulation amount was below 5% DCW.⁵⁸ And further research identified Synechocystis sp. IFA-3 and Chlorogloeopsis sp. Heinrichs-4 as promising PHB producers.⁵⁶ In addition, Calothrix scytonemicola TISTR 8095 and Oscillatoria okeni TISTR 8549 were also reported as efficient producers for PHB (25.4% DCW) and PHBV (14% DCW), respectively.^59,60

Genetic engineering further improves microbial PHAs production.^34,61,62 The metabolic pathways for microbial synthesis of PHAs are illustrated in Fig. 2. Engineering modification in the expression of relative enzymes is the most direct way for PHAs biosynthesis regulation. By enhancing or heterologously expressing the pha gene family (including phaA, phaB, or phaCAB), a PHB metabolic pathway can be constructed. When combined with specific metabolic regulation strategies, such as nitrogen starvation, substrate supplementation, or gene knockout, this approach can significantly increase the titer of PHB and the carbon conversion rate (Fig. 2A). Overexpression of phaAB in Synechocystis sp. PCC 6803 under nitrogen deprivation increased PHB content to 26% DCW, reaching 35% DCW with acetate supplementation.⁶³ Parallelly, deletion of sll0944 (pirC) in Synechocystis sp. PCC 6803 and expression of phaA and phaB from Cupriavidus necator (C. necator) under the strong psbA2 promoter resulted in a high PHB yield of 63% (DCW).⁶⁴ Furthermore, Synechococcus elongatus (S. elongatus) UTEX 2973 engineered to express phaCAB from C. necator H16 achieved 420 mg L⁻¹ PHB (16.7% DCW) and a specific productivity of 75.2 mg L⁻¹ day⁻¹, paving the way for industrial flue gasutilization in bioplastic biosynthesis.⁶² (Fig. 2A). It is worth to mention that, regulation of the metabolic balance is a powerful approach for PHAs overproduction. A heterologous PHB pathway was contructed in eukaryotic microalgae Phaeodactylum tricornutum (P. tricornutum) with metabolic balance regulation, and a maximum of 2.3% DCW content was achieved under ambient CO₂.⁶⁵

Fig. 2 The representative metabolic pathways for biosynthesis of PHAs (A) and lactate-based polymers (B). Dashed arrows represent multi-step enzymatic transformations.

Engineering of the acetyl-CoA pool improves PHB content and productivity in Synechocystis sp. PCC6803, and 12% DCW content with a productivity of 7.3 mg L⁻¹ day⁻¹ was achieved.⁶⁶ By deleting the genes for phosphotransacetylase (pta) and acetyl-CoA hydrolase (ach), the conversion of acetyl-CoA to acetate was reduced, leading to an accumulation of acetyl-CoA. Concurrently, the introduction of the heterologous gene for phosphoketolase (xfpk) from Bifidobacterium breve, which catalyzes the cleavage of sugar phosphates to produce acetyl phosphate, further increased the levels of acetyl-CoA. The synergistic effect of these modifications ultimately resulted in a significant enhancement of PHB yield (Fig. 2A). Furthermore, an ATP-hydrolysis-based driving force module was constructed in S. elongatus PCC 7942 by introducing an enzyme capable of catalyzing ATP hydrolysis and coupling it with the condensation reaction of acetyl-CoA (such as NphT7).⁶⁷ This enabled the irreversible conversion of acetyl-CoA to acetoacetyl-CoA, significantly enhancing the rate and efficiency of carbon flux toward 3HB, resulting in a cumulative 3HB titer of 1.2 g L⁻¹ (Fig. 2A). The genetic modification of microalgae for efficient PHB synthesis primarily relies on the synergistic driving of carbon metabolic flux through the strengthening of synthetic pathways, the blockage of competing pathways, and the reconstruction of metabolic networks. When combined with energy metabolism, these strategies can significantly increase product accumulation and promote industrial production.

Mutagenesis provides an effective alternative strategy to enhance PHAs bioproduction.⁶⁸ A mutant strain, MT_a24, derived from Synechocystis sp. PCC 6714, was obtained through random mutagenesis induced by ultraviolet (UV) radiation. Under nitrogen- and phosphorus-limited conditions, a PHB content of 37% DCW was achieved using CO₂ as the sole carbon source, and the productivity was more than 2.5-fold higher than that of the wild-type strain.⁶⁹ And random mutagenesis could be used to help identify target genes for genetic engineering in cyanobacteria.

2.1.2. Pseudomonadota-Alphaproteobacteria. Alpha-proteobacteria can utilize a broad range of organic and inorganic substances as energy and carbon sources.^70,71 Certain strains capable of utilizing C1 resources for PHAs production predominantly belong to the Rhodospirillales and Hyphomicrobiales orders (Table 2).

Table 2 Key bacterial strains involved in PHAs production using C1 substrates

Strain	Genetic modification	Substrate (carbon source)	PHAs production				Ref.
			Type (mol%)	Content (wt%)	Productivity (mg L^−1 d^−1)	Titer (g L^−1)
a Wild-type strain. b Not reported.
R. rubrum	W^a	Synthetic syngas (CO : CO2 : H2 : N2 = 5 : 1 : 5 : 9)	PHB	30.00	324	1.60	73
	W	Synthetic syngas (CO : N2 = 3 : 17)	PHB	8.00	N^b	0.036	74
	Heterologous expression of phaG, PP_0763, and phaC1(from P. putida KT2440) in R. rubrum ΔphaC1 ΔphaC2 phaZ2C176A(pBBR1MCS-2) under strong promoter PcooF	Synthetic syngas (CO : CO2 : H2 : N2 = 4 : 1 : 4 : 1)	P(3HD-co-3HO)	7.10	N	N	75
M. trichosporium OB3b	Heterologous expression of CoA-dependent succinate semialdehyde and 4HB-CoA transferase (from P. gingivalis) in strain M. trichosporium OB3b driven by promoter Ptac.	40% (v/v) methane	P(3HB-co-4HB) (1.33% 4HB)	18.85	N	N	84
	Heterologous expression of CoA-dependent succinate semialdehyde dehydrogenase and 4HB-CoA transferase gene from C. kluyveri in strain M. trichosporium OB3b driven by promoter Ptac.		P(3HB-co-4HB) (2.03% 4HB)	10.74
	Heterologous expression of CoA-dependent succinate semialdehyde dehydrogenase and 4HB-CoA transferase gene from C. kluyveri in strain M. trichosporium OB3b driven by promoter Ptac, and overexpression of icd gene from M. trichosporium OB3b driven by promoter Ptac.		P(3HB-co-4HB) (3.08% 4HB)	7.10
M. trichosporium OB3b	W	Synthetic syngas (CH4 : CH3OH = 3 : 7)	PHB	52.50	N	0.049	77
M. hirsuta	W	Synthetic syngas (CH4 : CO2 : H2S = 140 : 59 : 1)	PHB	45.00	N	N	79
M. hirsuta CSC1	W	Synthetic syngas (CH4 : CO2 = 1 : 2)	PHB	45.00	N	N	80
		Synthetic syngas (CH4 : CO2 : O2 = 3 : 1 : 6)		3.70
M. hirsuta	W	CH4	PHB	34.60	1400	N	81
M. hirsuta	W	Synthetic syngas (CH4 : O2 = 2.1 : 1)	PHB	14.50	60	N	83
M. extorquens AM1	Endogenous gene phaC and phaA was deleted and the pathway genes bktB, phaJ1, and phaC2, were heterologously expressed, and expression level of bktB was improved by untranslated region (UTR) engineering	Formate, propionate	PHBV (8.9% 3HV)	20.6	N	N	85
M. parvus OBBP	W	Methane	PHB	50.00	N	N	78
		Methane and valerate	PHBV (22.00% 3HV)	54.00
C. necator	MF01ΔB1/pBBP-ccrMeJ4a-emd	Synthetic syngas (H2 : O2 : CO2 = 8 : 1 : 1)	PHBHHx (52.30% 3HB, 47.70%HHx)	61.70	N	∼6.57	15
	MF01/pBPP-ccrMeJ4a-emd		PHBHHx (96.70% 3HB, 3.30%HHx)	85.80	N	∼7.31
C. necator H16	W	Synthetic syngas (H2 : O2 : CO2 : N2 = 3.6 : 7.6 : 12.3 : 76.5)	PHB	70.00	∼45	0.27	90
	Introduction of phaC1 from Pseudomonas sp. 61-3 and overexpression of phaAB and bktB		P(3HB-co-3HV-co-3H4MV)(97.60% 3HB, 1.20% 3HV, 1.2% 3H 4MV)	57.00	∼13	∼0.08
C. necator H16	W	97% air and 3% CO2	PHBV (∼35.16% 3HB, ∼64.84% 3HV)	3.84	N	0.08	93
R. eutropha	Introduction of cyanobacterial RuBisCO from Synechococcus sp. PCC 7002	Synthetic syngas (H2 : CO2 : O2 = 78 : 11 : 11)	PHB	34.00	N	N	94
R. eutropha H16 (ATCC17699)	W	Synthetic syngas (H2 : O2 : CO2 : N2 = 3.8 : 7.3 : 13.0 : 75.9)	PHB	89.00	∼490	2.94	91
R. eutropha	pBBR1MCS-3-PL::coxMSLDEFGoc	30% Synthetic syngas (CO : H2 : CO2 : N2 = 4 : 4 : 1 : 1)	PHB	49.70	∼62	1.30	95
R. eutropha	W	Synthetic syngas (CO : H2 : CO2 : N2 = 2 : 2 : 2 : 4)	PHB	42.00	∼4732	14.20	92
C. coskatii	Construction of PHB-producing recombinant by introducing thlA, hbd, crt, phaJ and phaEC from Clostridium species	Synthetic syngas (CO : H2 : CO2 : N2 = 4 : 4 : 1 : 1)	PHB	1.12	N	N	98

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CO can serve as the sole carbon and energy source for Rhodospirillum rubrum (R. rubrum).⁷² Synthetic gas containing 40 v/v% CO was used alongside acetate under carbon- and phosphorus-limited conditions for PHB production by R. rubrum (ATCC 11170), achieving a maximum product content of 30 w/w% DCW.⁷³ And CO-based acclimation promotes cell growth and PHB production in R. rubrum (ATCC 11170).⁷⁴ In addition, strain engineering was performed through genetic modification, enabling the direct biotransformation of synthesis gas for the production of poly(3-hydroxydecanoate-co-3-hydroxyoctanoate) (P(3HD-co-3HO)), a copolymer of 3-hydroxydecanoic acid and 3-hydroxyoctanoic acid. Genes from Pseudomonas putida KT2440 (P. putida KT2440) (phaG PP_0763, and phaC1) were introducd into a PHA-negative R. rubrum mutant, regulated by the CO-inducible promoter PcooF from R. rubrum (ATCC 11170). The phaG, PP_0763, and phaC1 genes encode a 3-hydroxyacyl-ACP thioesterase, a medium-chain-length fatty acid CoA ligase, and a medium-chain-length PHA synthase, respectively. This enzymatic system efficiently transforms carbon sources from synthesis gas into precursor monomers for poly(3HD-co-3HO) and catalyzes their polymerization (Fig. 2A). And the recombinant mutant strain accumulated P(3HD-co-3HO) up to 7.1% of DCW when using synthetic gas exclusively.⁷⁵

The primary genera responsible for PHAs biosynthesis based on methane and formate include Type II methanotrophs (e.g. Methylosinus and Methylocystis) and Type II methylotroph (e.g. Methylorubrum), which are classified under the Hyphomicrobiales order within Alphaproteobacteria.⁷⁶ The carbon source composition plays a critical role in determining the fermentation performance of Methylosinus trichosporium (M. trichosporium).⁷⁷ For Methylocystis, studies have shown the first evidence from a pure culture for the methane trophic synthesis of PHBV, and methane mass transfer was further improved in P3HB production by Methylocystis parvus OBBP (M. parvus OBBP).⁷⁸ And nitrogen starvation has been validated as a strategy for enhancing PHAs production of Methylocystis.^79–82 Furthermore, as a key intermediate, formate can bridge the conversion of synthetic gas or CO to biodegradable plastics.⁸³ The efficient bioconversion of methane and formate to desired products relies critically on the engineering of microbial cell factories (Fig. 2A). In M. trichosporium OB3b, the biosynthetic pathway for 4-hydroxybutyric acid (4-HB) was engineered by overexpressing genes encoding phosphoenolpyruvate carboxylase, isocitrate dehydrogenase, and 2-oxoglutarate dehydrogenase, which enabled the production of 10.5 mg L⁻¹ 4-HB. Coupled with the native poly(3-hydroxybutyrate) (P(3HB)) pathway, the M. trichosporium OB3b produced P(3HB-co-4HB) from methane, with a 4-HB molar fraction of 3.08% and total PHAs content of 7.01% DCW.⁸⁴ For CO utilization, a segmented bioconversion process with formate as an intermediate product is readily understandable. In the study of Hwang et al.,⁸³Acetobacter woodii (A. woodii) was employed to convert CO into formate. And engineered Methylorubrum extorquens (M. extorquens) AM1 then utilized this formate for PHB production. To enhance biomass production through improved formate uptake and assimilation, the formate transporter homolog and formate-tetrahydrofolate ligase encoded by the ftfL gene were individually overexpressed. The DCW of the overexpressing strain MZ02 reached 3.51 g L⁻¹ at pH 6.6, representing a 30% increase compared to the wild type. And PHB production increased to 1.37 g L⁻¹, reflecting an 87% improvement over the wild type. When the gene encoding the enzyme complex (phaCAB) in the PHB synthesis pathway was co-expressed with ftfL, the overexpression of ftfL improved the utilization efficiency of formate, providing sufficient substrate for PHB synthesis. Additionally, the overexpression of phaCAB directly enhanced the capacity for PHB synthesis. As a result, the PHB titer increased to 1.57 g L⁻¹, which is 2.15 times greater than that of the wild type, and the PHB content in the cells rose by 46.6% (Fig. 2A).⁷⁹ In addition, engineered M. extorquens was developed for the production of PHBV copolymers from formate.⁸⁵ To enhance the incorporation of 3HV, bktB was overexpressed through untranslated region engineering, and the endogenous phaA gene was deleted. The phaA gene encodes acetyl-CoA acetyltransferase, which competes with the β-ketothiolase encoded by the bktB gene for the utilization of acetyl-CoA. Deleting the phaA gene eliminates this competitive bottleneck, while the overexpression of bktB significantly enhances the incorporation of propionyl-CoA, thereby facilitating the targeted synthesis of PHBV copolymers enriched in 3HV (Fig. 2A). The engineered strain produced PHBV with 8.9% 3HV from formate alone. When supplemented with propionate and butyrate, PHBVs containing up to 70.6% 3HV were obtained. These findings demonstrate that a synergistic metabolic engineering strategy, which involves enhancing the expression of key enzymes, optimizing substrate utilization efficiency, and alleviating metabolic competition bottlenecks, significantly increases the titer of PHAs produced by methane-oxidizing bacteria and methylotrophic bacteria, as well as the regulatory capacity of copolymer monomers. Consequently, this provides an efficient pathway for the production of high-performance biopolymers from C1 substrates.

2.1.3. Pseudomonadota-Betaproteobacteria. Beta-proteobacteria are capable of growth and reproduction using a wide range of carbon sources. Strains from the order Burkholderiales are primarily responsible for utilizing C1 compounds to synthesize PHAs (Table 2).

C. necator possesses an intrinsic pathway for PHB biosynthesis, storing the polymer in granules under nutrient-limited conditions, with titers exceeding 70% of total biomass.⁸⁶ The C. necator H16 strain is a facultative Knallgas bacterium capable of using H₂ as an energy source.¹⁵ It is genetically manageable and features a high-throughput carbon fixation system, converting most input CO₂ into biomass.⁸⁷ And C. necator preferentially consumes valeric acid during growth phases, but switches to acetic acid consumption during PHA production.⁸⁸

Researchers have also explored the use of various carbon sources to assess the PHAs production potential of C. necator.⁸⁹ And the results indicated that the type of carbon source influences not only PHAs composition and molecular weight but also the surface microstructure and porosity of the obtained polymer films. The use of low-hydrogen, non-combustible gas mixtures (H₂ : O₂ : CO₂ : N₂ = 3.6 : 7.6 : 12.3 : 76.5) for PHAs biosynthesis by Ralstonia eutropha H16 (ATCC 17699) (R. eutropha) provided a strategy for eliminating the risk of explosion.^90,91 In nitrogen-limited medium, this strain accumulated up to 70% P(3HB) by weight. Moreover, when cultured autotrophically under nitrogen-limiting conditions, the recombinant strain R. eutropha 1F2 produced a 3HB-based copolymer containing 1.2 mol% 3HV and 1.2 mol% 3H4MV, achieving a total polymer content of 57% DCW. Of particular note, a cell-bridging strategy was employed to bridge the nanoscaled cellulose particles with an enzyme complex (including carbon monoxide dehydrogenase, carbon monoxide binding unit and carbonic anhydrase) and R. eutropha. This approach enabled the efficient conversion of syngas (H₂ : CO₂ : CO : N₂ = 2 : 2 : 2 : 4) to accessible carbon source, and provided the production of 14.2 g L⁻¹ PHB, 33.8 g L⁻¹ DCW, and 42% PHAs content, with a productivity of 0.197 g L⁻¹ h⁻¹.^46,92 And CO₂ and volatile fatty acids (VFA) from anaerobic digestion process were also used by C. necator H16 for PHAs production, reaching a product content of 3.84 wt%.⁹³ Thus, Betaproteobacteria, particularly C. necator and R. eutropha, are the predominant strains for the biosynthesis of PHAs. By optimizing carbon sources, cultivation conditions, and the structure of biocatalytic media, the synthesis performance of PHAs can be significantly enhanced.

To further unlock the strain's potential for achieving higher titers, a broader range of PHA types, and more efficient substrate conversion, genetic modification of key metabolic pathways is essential. Through genetic modification, C. necator H16 was engineered to achieve PHAs contents of 20–60% DCW. And the combination of different thioesterases (TEs) with PHA synthases (phaCs) (e.g. Native C. necator C4 phaC_Cn; Pseudomonas aeruginosa PAO1 (P. aeruginosa PAO1) phaC2_Pa; P. aeruginosa PAO1 phaC1_Pa; and a Pseudomonas spp 61-3 phaC1_Ps) enabled selective manipulation of PHAs composition, reaching direct copolymer synthesis from CO₂ (Fig. 2A).⁸⁷ By introducing heterologous RuBisCO from the cyanobacterial Calvin–Benson–Bassham (CBB) cycle and coupling it with the endogenous GroES/EL and RbcX systems, carbon conversion efficiency was enhanced. Concurrently, the expression of membrane-bound hydrogenase (MBH) and soluble hydrogenase (SH) was regulated to optimize the oxidation of H₂ for NADH generation, thereby supplying the essential reducing power for the CBB cycle, which collectively resulted in a 93.4% increase in the OD600 of R. eutropha H16 within 72 hours (Fig. 2A).⁹⁴ Additionally, R. eutropha H16 was engineered to express the Cox subcluster from the carbon monoxide-oxidizing Oligotropha carboxidovorans OM5. The coxOc gene, controlled by the CO₂-induced promoter PL, enabled R. eutropha to oxidize CO into CO₂ for carbon assimilation. Cultivation in synthetic gas containing CO, H₂, and CO₂ improved cell growth and increased P(3HB) synthesis by over 20%.⁹⁵ Moreover, precise genetic engineering was conducted on C. necator H16. The MF01 strain was constructed by replacing phaA with the medium-chain-specific β-ketothiolase gene bktB and phaC with the broad-substrate-specificity mutant phaC_NSDG. Subsequently, the phaB1 gene was knocked out to obtain the MF01ΔB1 strain. By weakening the (R)-acetoacetyl-CoA reduction pathway, the flux towards crotonyl-CoA was directionally enhanced, providing a metabolic basis for the synthesis of 3-hydroxyhexanoate (3HHx) monomers. Meanwhile, the substrate specificity of enzymes was exploited to precisely regulate the abundance of the 3HHx component. And the 3HHx content in PHBHHx reached 47.7 mol% in strain MF01ΔB1/pBBP-ccr_MeJ4a-emd (Fig. 2A).¹⁵

2.1.4. Pseudomonadota-Gammaproteobacteria. Although gammaproteobacteria have relatively limited direct applications in bioplastic production, their metabolic and biosynthetic versatility presents opportunities for future development. Genetic modification is the key to unlocking the production potential of polyhydroxyalkanoates (PHAs) in γ-proteobacteria, and strains capable of utilizing C1 resources to synthesize PHAs are primarily found in the orders Enterobacterales and Pseudomonadales. The (R)-3-hydroxybutyric acid biosynthesis pathway was successfully constructed in Escherichia coli (E. coli). This was achieved by introducing the phaA (β-ketothiolase) and phaB (acetoacetyl-CoA reductase) genes from R. eutropha, along with the pct (propionyl-CoA transferase) gene from Clostridium beijerinckii 8052 (Fig. 2A).⁹⁶ The heterologously expressed propionyl-CoA transferase (PCT) efficiently catalyzes the conversion of acetate derived from syngas into 3HB by facilitating the generation of acetyl-CoA. The engineered strain produced 1.02 g L⁻¹ of 3HB, achieving a yield of 0.26 g g⁻¹. Furthermore, P. putida KT2440 was engineered for enhanced production of medium-chain-length polyhydroxyalkanoates (mcl-PHAs). This involved the deletion of the phaZ (inhibiting PHA degradation) and the overexpression of the acs (enhancing acetyl-CoA supply), along with phaG, phaC1 (driving de novo synthesis and polymerization), and phaJ4 (directing the conversion of fatty acid β-oxidation products into mcl-PHA precursors). And the modified strain provided a PHAs titer of 556.2 mg L⁻¹ within 24 h (Fig. 2A).⁹⁷

2.1.5. Bacillota. Bacillota typically possess thicker cell walls, which enhance their survival under environmental pressures such as osmotic stress. However, studies about the natural synthesis of PHAs within Bacillota from C1 resources remains limited.⁹⁸ Using H₂-CO₂ as substrates, the thermophilic bacterium Kyrpidia spormannii EA-1 (K. spormannii EA-1) provided 26.8 μg cm⁻² PHAs titer with a productivity of 96 mg d⁻¹ m⁻² on a cathode.⁹⁹ Concurrently, the genetic engineering of Bacillota strains to efficiently utilize C1 substrates presents a key strategy for overcoming the metabolic limitations of natural strains and achieving industrial-scale production of PHAs. Anaerobic PHB production was achieved using recombinant clostridial acetogens supplied with synthetic gas (CO : H₂ : CO₂ : N₂ = 4 : 4 : 1 : 1) as the sole carbon and energy source. A new synthetic pathway containing thlA (thiolase A), ctfA/B (CoA-transferase A/B) and bdhA (3-hydroxybutyrate dehydrogenase) was constructed in Clostridium coskatii (C. coskatii) [p83_tcb] for 3-HB production. PHB production was further achieved using a recombinant C. coskatii [p83_PHB_Scaceti] expressing a novel synthetic PHB pathway containing the genes thlA (thiolase A), hbd (3-hydroxybutyryl-CoA dehydrogenase), crt (crotonase), phaJ ((R)-enoyl-CoA hydratase), and phaEC (PHA synthase) (Fig. 2A). And the engineered C. coskatii [p83_PHB_Scaceti] accumulated 1.12% (DCW) PHB (Table 2).⁹⁸

2.1.6. Yeast. Yeast presents a promising platform for large-scale industrial PHAs production.¹⁰⁰ Extensive genomic toolkits and a variety of yeast strains are available for synthetic biology applications, including the model organism brewing yeast and non-model species such as Yarrowia lipolytica (Y. lipolytica), Pichia stipitis, P. pastoris, Kluyveromyces marxianus, Candida utilis, and Rhodotorula glutinis.¹⁰¹

For PHAs production by wild-type strain, biological waste substrates (banana peel and chicken feather hydrolysates) were used for P(3HB-co-3HV) production by the marine-derived Pichia kudriavzevii VITNN02.¹⁰² Optimization using a three-level Box-Behnken design involving six variables enhanced PHAs content from 40% to 79.68%, with a final PHAs concentration of 23.53 g L⁻¹. With the gene-editing technologies such as CRISPR/Cas9,¹⁰³ PHAs biosynthetic pathways compatible with yeast metabolism can be introduced and expressed in different yeast hosts. Although native Saccharomyces cerevisiae (S. cerevisiae) cannot synthesize PHAs, its aerobic oxidative metabolism supports the formation of metabolic precursors required for PHAs biosynthesis, thereby enabling expression of biological PHAs pathways in S. cerevisiae.¹⁰⁴ Research has shown that by introducing the phaA, phaB1, and phaC1 genes into S. cerevisiae, polymers with physicochemical properties comparable to those produced by natural or engineered PHAs-producing bacteria can be synthesized, leading to PHAs accumulation of up to 11% DCW (Fig. 2A).¹⁰⁵ And Arxula adeninivorans was engineered by overexpressing three key PHA pathway enzymes (β-ketothiolase, acetyl-CoA reductase, and PHA synthase) along with a phasin gene.¹⁰⁶ The optimized strain exhibited stable accumulation of PHB and PHBV, reaching up to 52.1% of DCW as poly(hydroxybutyrate-co-hydroxyvalerate) (PHBV), with a titer of 10.8 g L⁻¹.

However, limited studies have explored yeast-based PHAs production from C1 substrates. Current research has successfully achieved the precise modification of metabolic pathways in yeast through genetic engineering, reconstructing the central metabolic network to ensure that yeast can efficiently utilize C1 substrates. And this modification is aimed at providing sufficient supplies of key precursors essential for the biosynthesis of PHAs, such as acetyl-CoA, free fatty acid (FFA), and malate. In P. pastoris, the deletion of two fatty acyl-CoA synthetase genes (FAA1 and FAA2) enhanced free (FFA) production. As acetyl-CoA is the precursor for fatty acid biosynthesis, overexpression of the citrate lyase gene (MmACL) from Mus musculus, which catalyzes the cleavage of citrate into acetyl-CoA and oxaloacetate, increased FFA levels by 23%.¹⁰⁷Ogataea polymorpha (O. polymorpha) was engineered with the MAE1, MDH, and PYC genes from Schizosaccharomyces pombe for creating the PMT-strain, and the T-strain was obtained with only MAE1. PMT- and T-strain utilized methanol as a carbon source and produced substantial amounts of malic acid after 72 h: 4.1 g L⁻¹ and 3.4 g L⁻¹, respectively (Fig. 2A).¹⁰⁸ Malic acid has also been used as a co-substrate for PHAs synthesis in bacterial systems. And a P(3HB) concentration of 2.80 g L⁻¹ was provided by Burkholderia sacchari using a mixture of glucose and malic acid.¹⁰⁹ As previously reported, a hybrid electrobiosystem was developed for the integration of spatially separated CO₂ electrolysis and yeast fermentation.¹¹⁰ This system efficiently converted CO₂ into acetate via electrolysis, which was subsequently utilized by yeast to produce glucose, achieving an average glucose titer of 1.81 g L⁻¹. In S. cerevisiae, a glucose leakage phenotype (LY031) was constructed by deleting all known hexokinase genes (GLK1, HXK1, and HXK2). To demonstrate the system's adaptability for methanol-to-glucose conversion, a similar glucose leakage phenotype was engineered in P. pastoris by eliminating genes associated with glucose utilization, including HXK1, HXK2, GLK1, and the hexokinase isoenzyme 2 gene (HXK iso2) (Fig. 2A). The resulting P. pastoris mutant strain was capable of glucose production, reaching approximately 0.5 g L⁻¹ after 96 h in shaker flasks. Furthermore, to enhance glucose yield via hydrolysis of glucose-1-phosphate, the yihX gene encoding haloacid dehalogenase-like phosphatase 4 from E. coli was expressed in P. pastoris, resulting in a glucose yield of approximately 1.08 g L⁻¹ and a volumetric productivity of 11.25 mg L⁻¹ h⁻¹. The efficient conversion of CO₂ into glucose advances research progress on the biosynthesis of PHAs using C1 resources. By integrating key gene overexpression, competitive pathway knockout, and a coupled CO₂ electrolysis-fermentation system, it is possible to successfully reconstruct the metabolic network of yeast. This enables the efficient utilization of C1 resources to produce key precursors such as acetyl-CoA and malate, thereby laying the foundation for subsequent PHAs biosynthesis (Fig. 2A).

2.2. Microorganisms for producing lactate-based polymers from C1 resources

Lactate-based material is a type of biodegradable and thermoplastic bio-based polyester.⁶⁸ In recent years, biosynthesis has accounted for approximately 90% of global LA production.¹⁸ Bioprocess-based production of lactate monomers offers a significantly lower environmental impact by utilizing renewable feedstocks instead of petrochemical sources.^111,112 Nonetheless, most lactate monomers are still derived from starch-based fermentation, which imposes pressure on land use and agricultural resources.¹¹³ The conversion of methanol to LA has been achieved in engineered yeasts (e.g. P. pastoris and O. polymorpha) through systematic metabolic engineering and adaptive evolution.^49,114 It is worth to mention that engineering lactate dehydrogenase cofactor preference and mitochondrial localization in yeasts provide new insights for promoting LA biosynthesis. These studies provide promising prospects for the biosynthesis of lactate-based polymers from C1 resources.

Lactate-based polymers are commonly synthesized via polycondensation, ring-opening polymerization and copolymerization.^115,116 However, due to the complexity of production process, lactate-based polymers is not economically viable.¹¹⁷ The microbial one-step production of lactate-based polymers has received intensive research attention, and the degradation profile, mechanical strength, hydrophilicity, and lipophilicity of the lactate-based polymers can be adjusted. Within this framework, the profound engineering of host strains using gene-editing technology represents a central methodology for efficient substrate conversion toward target polymers. Previous studies utilizing common carbon sources such as glucose and xylose provide a valuable technical foundation for the biosynthesis of lactic acid-based copolymers from C1 substrates. Microbial production of lactate-based polymers is summarized in Table 3. One-step microbial fermentation of lactate-based polymers involves constructing unnatural metabolic pathways based on the PHA biosynthetic pathway. Typically, LA is converted to lactyl-CoA via Pct, butyryl-CoA transferase (Bct), or isocaprenoyl-CoA : 2HIC CoA-transferase (HadA). Lactyl-CoA is then polymerized into lactate-based polymers using engineered PHAs synthase (Fig. 2B).¹¹⁸ Based on unnatural metabolic pathway construction, logical selection and chromosomal gene targets deletion further minimized the by-product formation, thereby increasing PLA accumulation in S. cerevisiae.¹⁰⁵ And the biosynthesis pathways of D-lactate oligomers and PDLA were constructed in engineered E. coli and Y. lipolytica.^119,120 In addition, directed enzyme evolution enhanced the substrate specificity of both PCT and PHA synthase for LA and lactoyl-CoA, thereby promoting PLA biosynthesis in E. coli.¹²¹ A commonly studied lactide-based block copolymer is P(LA-co-3HB), which is primarily synthesized via synthetic biology and systematic metabolic engineering. In terms of transparency and biodegradability, P(LA-co-3HB) integrates the advantageous properties of both PLA and P(3HB).¹²² For the first time, a one-step microbial metabolic pathway for synthesizing the representative LA copolymer, P(LA-co-3HB), was successfully developed in E. coli.¹²³ In synthesizing P(LA-co-3HB) using glucose in S. cerevisiae, the endogenous DLD1 gene was deleted to reduce the accumulation of L-LA, and the Pct gene was introduced to convert D-LA into the polymerizable D-lactyl-CoA. Concurrently, a modified PHA synthase was incorporated into the constructed PHB synthesis pathway to form the copolymer, ultimately resulting in an accumulation of 3.6% DCW of P(LA-co-3HB).¹⁰⁵ Additionally, Wu et al. engineered the E. coli strains WJ03-02 and WJ03 by replacing the promoter of Pct540Cp with ldhA promoter and knocking out the gene for the glucose-specific PTS enzyme IIBC component (ptsG) to alleviate the repression of carbon catabolites, enabling the utilization of corn stover hydrolysate as a feedstock for the production of P(3HB-co-LA) (Fig. 2B). And the engineered WJ03-02 strain produced P(3HB-co-LA) containing 7.1 mol% LA.¹²⁴ Furthermore, P(LA-co-3HB) has demonstrated potential as an effective solution to various environmental problems associated with plastic use.¹¹⁵ In addition, E. coli BW25113 was engineered by introducing a mutant PHA synthase (F392S) and deleting the pyruvate formate-lyase activating gene (pflA). By blocking the conversion pathway from pyruvate to acetyl-CoA, the intracellular accumulation of LA-CoA is enhanced, ultimately increasing the proportion of LA in the copolymer. Furthermore, the introduction of exogenous key enzyme genes, such as PHA synthase, facilitates the reconstruction of carbon metabolic flux to promote the efficient biosynthesis of lactate-based copolymers (Fig. 2B). And 62 wt% P(LA-co-3HB) was synthesized, with 45 mol% LA content, using 20 g L⁻¹ glucose.²⁰ Biosynthesis of other lactate-based polymers (e.g. P(LA-co-3HB-co-3HV) and P(LA-co-3HB-co-3HP)) currently focuses on traditional carbon source substrates such as glucose, xylose, sucrose and glycerol. For example, using recombinant E. coli as the core chassis, the structural similarity between PLA and PHAs has been leveraged to obtain a lactate polymerase (LPE) with lactyl-CoA (LA-CoA) polymerization activity through the directed modification of PHA synthase. Additionally, the heterologous expression of key enzymes such as PCT and specific enoyl-CoA hydratase (PhaJ4) has been employed to construct a complete pathway for monomer synthesis and polymerization (Fig. 2B).¹¹⁶ Furthermore, using glucose, xylose, sucrose, or glycerol as substrates, glycolysis generates pyruvate, which is subsequently converted into LA by lactate dehydrogenase (ldhA) and further transformed into LA-CoA. Concurrently, the acetyl-CoA intermediate is catalyzed by phaA-β-ketothiolase (PhaA) and phaB-acetoacetyl-CoA reductase (PhaB) to produce 3HB-CoA. In the ternary copolyester, the third monomer precursor can also be supplemented through the directed metabolism of pentanoic acid (3HV-CoA) or glycerol (3-hydroxypropionyl-CoA (3HP-CoA)). Ultimately, the lactate polymerase (LPE) catalyzes the intracellular copolymerization of the various monomer CoA thioesters, resulting in the formation of P(LA-co-3HB), P(LA-co-3HB-co-3HV) or P(LA-co-3HB-co-3HP) copolymers (Fig. 2B).^19,20,116 Although these systems utilize conventional carbon sources, they provide a significant foundation for the sustainable production of bioplastics with the potential for carbon-negative emissions.

Table 3 Microbial production of lactate-based polymers

Strain	Genetic modification	Substrate (carbon source)	Lactate-based polymer production				Ref.
			Type (mol%)	Content (wt%)	Productivity (mg L^−1 d^−1)	Titer (g L^−1)
a Not reported.
S. cerevisiae	D-Lactate dehydrogenase gene (DLD1) was deleted	Glucose	P(LA-co-3HB)	3.6	N^a	N	105
Y. lipolytica	A propionyl-CoA transferase (PCT) converting lactic acid into lactyl-CoA, and an evolved polyhydroxyalkanoic acid (PHA) synthase polymerizing lactyl-CoA, was introduced into the engineered strain	Glucose, lactic acid	PDLA	26.2	N	0.16	120
S. elongatus PCC7942	D-Lactic dehydrogenase (LDH), PCT, and PHA synthase, was introduced into S. elongatus PCC7942	CO2	PLA	1.5	N	0.11	8
E. coli	E. coli express a D-specific lactate-polymerizing enzyme.	Glucose	PDLA	N	N	8.3	119
E. coli JW0885	ΔpflA mutant	Glucose, hemicellulose	P(60 mol% LA-co-3HB)	N	N	7.3	115
E. coli BW25113	Engineered E. coli BW25113 (with mutated PHA synthase, F392S) along with the pyruvate formate lyase activating enzyme gene (pflA)	Glucose	P(45 mol% LA-co-3 HB)	62	N	N	20

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Metabolic pathways for microbial synthesis of lactate-based polymers from C1 resources are illustrated in Fig. 2B. Few studies on direct biosynthesis of PLA from C1 resources have been reported. The systematic remodeling of the metabolic network in host strains can enable the de novo production of lactate-based polymers from C1 substrates. A heterologous pathway consisting of engineered D-lactate dehydrogenase, propionate CoA transferase, and polyhydroxyalkanoate synthase was introduced into S. elongatus PCC7942.⁴⁸ By optimizing the promoters for the expression of key genes and constructing an acetyl-CoA pool, PLA production was increased approximately 19-fold to 15.0 mg g⁻¹ DCW compared to the wild-type strain. In addition, the high density culture (HDC) has been validated as an effective supplementary approach for increasing PLA titer with a 270-fold increase. As ideal phototrophic autotrophic microbial chassis, cyanobacteria can directly utilize CO₂ to produce PLA through carbon-negative synthetic biology pathways. The core mechanism involves the combination of metabolic engineering and synthetic biology to reconstruct the carbon metabolic flux and PLA synthesis pathway within cyanobacteria (Fig. 2B).⁴⁸ Therefore, to achieve efficient biosynthesis of lactic acid-based biopolymers from C1 substrates, current research primarily focuses on: reconstructing the central carbon metabolic flux (e.g. knocking out competitive pathway genes such as ptsG and pflA) to enhance the supply of key precursors (e.g.D-lactic acid and LA-CoA); introducing and optimizing exogenous key enzymes (e.g. PHA synthase and PCT) to improve the strain's ability to polymerize monomers; and deleting endogenous competitive genes (e.g. DLD1) to minimize substrate diversion, thereby directing carbon flux towards the synthesis pathway of the target polymer.

3. Process innovations for PHAs and lactate-based biopolymers

Production efficiency and cost-effectiveness of PHAs and lactic acid-based polymers as sustainable materials are critical for their industrial applications. Microorganisms that metabolize sugars or oils, such as Rhodococcus species, frequently accumulate PHB to levels as high as 80% of DCW.⁴³ And the lactate-based biopolymers from glucose can reached up to 81.7% DCW in engineered E. coli JX041.²⁰ In contrast, microorganisms that utilize C1 substrates typically show limited PHAs and lactate-based biopolymers production.¹²⁵Oscillatoria okeni TISTR 8549 produced PHBV using CO₂ with a titer of only 14% DCW.⁶⁰ Similarly, S. elongatus PCC 7942 produced only 23 mg g⁻¹ DCW PLA using CO₂.⁴⁸ To overcome this limitation, bioprocess engineering can be employed for improving the PHAs and lactate-based biopolymers production from C1 substrates, as shown in Fig. 3 and Table 4.

Fig. 3 Main fermentation processes for PHAs and lactate-based polymers. (A) Single-stage or dual-stage bioprocess; (B) Bio-electrochemical system; (C) Mixed fermentation; (D) Enzyme-assisted catalytic system.

Table 4 Fermentation processes for PHAs and lactate-based polymers

Fermentation process	Strain	Substrate (carbon source)	PHAs and lactate-based polymers production				Ref.
			Product type	Content (wt%)	Productivity	Titer (g L^−1)
a Not reported.
Single-stage or dual-stage bioprocess	E. coli JLX10	Glucose	PLA	11	N^a	N	118
	S. meliloti	Lactic acid	PLA	3.2	N	N	130
	S. elongatus PCC7942	CO2	PLA	1.5	N	0.11	48
	A. woodie, M. extorquens AM1	CO	PHB	6.5	N	0.097	83
	C. autoethanogenum DSM 10061	CO : CO2 : H2/N = 3 : 1 : 2 : 4	PHA	24	N	N	136
	A. woodie, R. eutropha H16	CO2	PHB	N	N	0.5	137
	C. necator DSM 545	H2 : O2 : CO2 = 84 : 2.8 : 13.2	PHBV	N	N	N	133
	S. ovata, C. basilensis	CO2	PHB	N	300.96 mg L^−1 d^−1	N	135
Bio-electrochemical system	K. spormannii EA-1	CO2	PHA	N	117 mg m^−2 d^−1	0.027	99
	C. necator	CO2	PHB	34	N	0.056	150
Mixed fermentation	Methanotrophic community	CH4	PHA	14	N	N	155
	II methanotrophs	CH4	PHB	48.3	N	N	156
	Microalgae and purple phototrophic bacteria	Domestic wastewater, sugar molasses	PHBV	26.1	520 mg L^−1 d^−1	N	157
Enzyme-assisted catalytic system	R. eutropha	CO	PHB	N	N	N	159
	R. eutropha	CO2	PHB	42	N	14.2	92
	E. coli	HbCoA	PHB	N	N	N	160

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3.1. Single-stage or dual-stage bioprocess

The design and selection of production processes for PHAs and lactic acid-based polymers are crucial for achieving efficient and sustainable bioconversion. Different bioprocesses can be adapted to accommodate various substrate characteristics and target product requirements, while single-stage and dual-stage bioprocesses have emerged as prominent strategies, exhibiting distinct characteristics and broad applicability.

Previous studies by Liu et al.¹²⁶ and Wang et al.^127–129 have adopted single-stage or dual-stage fermentations for the biosynthesis of L-LA and hyaluronic acid. However, the significant breakthrough lies in the development of a direct one-step fermentation strategy for the production of PLA and lactic acid-containing copolymers using metabolically engineered microorganisms (Fig. 3A).¹¹⁸ For the single-stage bioprocess, expression of a broad-host-range plasmid (pTAM) carrying PctCp and PhaC1_Ps6–19 resulted in 3.2% DCW PLA and 42% DCW P(3HB-co-LA-co-3HHx-co-3HO) in Sinorhizobium meliloti and P. putida, respectively.¹³⁰ In addition, a single-stage bioprocess was used for PLA biosynthesis, which achieved the direct conversion of CO₂ to PLA in engineered S. elongatus PCC7942.⁴⁸ The single-stage bioprocessing eliminates the need for separation and purification of intermediate products inherent in multi-stage biological processes, thereby streamlining the production workflow. However, the extensive genetic modifications imposed on engineered microorganisms for multifunctionalization in single-stage systems may potentially elevate intracellular metabolic burden, which could adversely affect product biosynthesis efficiency while concomitantly increasing byproduct formation.

For utilization of CO which is toxic to most living organisms, Acetobacterium woodii (A. woodii)-driven conversion of CO to formate has attracted special attention, as it is independent of external reducing agents (e.g. H₂). A dual-stage whole-cell biocatalytic process using industrial waste was developed by Hwang et al.,⁸³ and CO was bioconverted to formate with high conversion yield and specificity (∼100%) in the first stage. In the second stage, PHB production from formate using engineered Methylbacterium extorquens AM1 was achieved via the overexpression of ftfL and phaCAB, reaching a PHB yield of 2.24%. While strong growth is essential for a productive cell factory, excessive diversion of resources toward biomass accumulation can compromise the biosynthesis of target products. By decoupling the cell proliferation phase from the product synthesis phase, the dual-stage bioprocess allows for the independent optimization of cultivation conditions at each stage (Fig. 3A). This decoupling facilitates the redirection or enhancement of metabolic flux towards product synthesis, while simultaneously ensuring adequate flux for essential growth.⁴⁶ As reported by Salem et al.,¹³¹ PHB accumulation was triggered by nitrogen limitation using a gas mixture (CH₄ : O₂ = 1 : 1), and cultivation proceeded in dual-stage: initial biomass generation with nitrogen, followed by a nitrogen-depleted phase to activate the serine pathway for PHB accumulation. Dual-stage bioprocess significantly enhanced PHB accumulation, achieving 59.4% DCW, which is substantially higher than the 37.8% DCW obtained in single-stage process. This demonstrates the significant advantages of the dual-stage bioprocess in the metabolic regulation of microbial synthesis. Notably, the benefits of this approach extend beyond fermentation systems utilizing CO and methane; it also exhibits considerable application potential in phototrophic autotrophic systems that utilize CO₂ as the sole carbon source. When Synechocystis sp. CCALA192 was cultivated in a 200 L tubular photobioreactor using CO₂ as the sole carbon source in dual-stage bioprocesses, the biomass concentration reached 1.0 g L⁻¹ and the average PHB content was 12.5% DCW.¹³² Parallelly, A dual-stage process was initiated with glucose serving as the substrate for large-scale biomass generation in the first stage.¹³³ And the second stage utilized CO₂ and valeric acid as carbon sources to produce PHBV copolymer. To gain a deeper understanding and control of this process, a physics-based kinetic model describing the production of 3HB and 3HV monomers was developed. And this calibrated and validated model accurately predicts the proportion of 3HV monomers in the polymer with a deviation of only 2%. It also enables the precise design and production of PHBV copolymers with predetermined compositions, offering a practical framework for the synthesis of tailored bioplastics. Furthermore, experimental cycles and personnel input could potentially be reduced by approximately 70–80%.¹³⁴ And the similar process routes were also intensively reported.^135–137 Integrating artificial intelligence (AI) and machine learning (ML) into bioprocess development could further accelerate fermentation optimization and predictive modeling construction, facilitating more efficient PHAs or lactate-based biopolymers production at scale. For example, the chemical diversity introduces difficulties in determining the optimal application-specific chemical compositions of biopolymers. Melting temperature (T_m) prediction has been used to demonstrate the promise of ML-based techniques for establishing efficient structure–property mappings in PHAs-based chemical space. For PHAs homo- and copolymer chemistries, a manually curated data set of T_m values was developed by experimental measurement. And descriptors based on topology, shape, and charge/polarity of specific motifs forming the polymer backbone were then used to numerically represent the polymers. Combined with glass transition temperature (T_g) prediction model and an evolutionary algorithm-based search strategy, the ML models were used to address polymer design with multiobjective optimization.¹³⁸ And existing data sets play a significant role in the training, validation, and evaluation of ML models. The comprehensive database includes ChemSpider, the Materials Project, Material Hub Springer, PubChem, MatWEB, NIST, PoLyInfo, PolyIE and TPSX.¹³⁹ Thus, the decoupling of microbial growth and product accumulation, sequential utilization of substrates, and the construction of AI-assisted mathematical models can significantly enhance the biosynthesis of PHAs and lactate-based polymers, with two-stage processes demonstrating greater potential for industrial production compared to single-stage processes.

3.2. Bio-electrochemical system

With the emergence of green electricity generation from sources such as wind, solar, and tidal energy, their pivotal role in optimizing energy structures and achieving deep decarbonization has become increasingly evident.^140–143 Bio-electrochemical system has successfully reached electrochemical reduction of CO₂ into intermediates (e.g. CO, formate, methanol and methane), which can be further bioconverted to PHAs and lactate-based biopolymers.^144–147 And bioelectrochemical systems have achieved negative carbon emissions, demonstrating significant environmental benefits.^148,149

In microbial electrochemical synthesis systems (Fig. 3B), commonly used microorganisms include C. necator and K. spormannii. And the key enzyme systems involved in electrochemical conversion involves [NiFe]-hydrogenases and Calvin cycle-related enzymes, which collectively facilitate CO₂ reduction and fixation. The production of PHAs and lactic acid-based polymers in these systems primarily focuses on enhancing Faraday efficiency, which can be improved by optimizing electrode materials and structures, as well as electrode spacing. K. spormannii EA-1 was used for PHAs production directly on the cathode. The cathode structure was optimized, and a reduced anode-cathode distance improved PHA productivity, providing a 3-fold enhancement in PHA production (117 mg day⁻¹ m⁻²).⁹⁹ In the study by Stöckl et al., PHB was produced from CO₂ using a tin-based gas diffusion electrode in combination with C. necator. In this system, formate was generated and utilized as an intermediate during the electroreduction of CO₂ in a physiological electrolytic buffer.¹⁵⁰ The overall Faraday efficiency of the complete pathway (from CO₂ capture to PHB production) reached 4%, with a PHB concentration of 56 mg L⁻¹. A more recent study reported the development of a defect-rich bismuth catalyst (P–Bi) through plasma bombardment followed by an in situ electrochemical reduction scheme. Introducing numerous defects on the surface of P–Bi catalyst promoted the faraday efficiency for formate production. Using this electrogenerated formate as a carbon source in a variable-rate feeding batch fermentation, the PHB concentration reached 99.6 mg L⁻¹, with a corresponding PHB content of 2.16%.¹⁵¹ Although bioelectrochemical systems present a highly promising alternative for the production of PHAs and lactic acid-based polymers by utilizing low-cost CO₂, thereby achieving carbon-negative emissions and significant environmental benefits, it is important to note that the scale-up of microbial electrochemical synthesis systems necessitates attention to issues related to system structural optimization, energy consumption reduction, and faradaic efficiency enhancement.¹⁵²

3.3. Mixed fermentation

Mixed fermentation, an alternative to pure culture, simplifies biopolymer production by eliminating the rigorous sterilization required in pure culture methods.^153,154 By selecting complementary microbial partners, synthetic co-cultures can be constructed with clearly defined metabolic roles, thereby enabling or enhancing the biosynthesis of specific biopolymers (Fig. 3C).^23,150,151

The microbial consortia used for mixed fermentation to produce PHAs or lactate-based biopolymers primarily consist of methanotrophic and phototrophic mixed communities. In the research of Luangthongkam et al., a methanotrophic community was enriched in a semi-continuous reactor and supplemented with valeric acid to increase the 3HV fraction. This strategy achieved up to 65 mol% 3HV and improved PHAs titer (14.1% DCW).¹⁵⁵ Methanotrophic communities achieved stable long-term methane consumption in semi-continuous reactors under non-sterile conditions, exhibiting a consistent exponential growth pattern while maintaining robust growth and metabolic capacity under unsterilized environment. A novel setup for methanotrophic enrichment and PHB production was reported, achieving maximum PHB accumulation of 59.4% and 54.3% in ammonium and nitrite salt media, respectively.¹³¹ Parallelly, methane-trophic and heterotrophic microbes were selectively enriched from sewage sludge to form stable mixed cultures. Using methane as the sole carbon source, these cultures accumulated PHB contents ranging from 43.2% to 45.9%.¹⁵⁶ Notably, using waste CH₄ as feedstock in mixed fermentation reduces PHB production costs by approximately 30–35% and enhances adaptability to complex environmental conditions; even at low methane concentrations (25%), PHB content can reach 37.0%.¹⁵⁶ Additionally, photosynthetic mixed cultures of microalgae and purple phototrophic bacteria have also been reported for PHAs biosynthesis.¹⁵⁷ However, the dynamics of microbial communities in mixed fermentation systems are complex and variable. Research utilizing 16S rRNA gene sequencing has analyzed the microbial composition under different cultivation conditions, revealing that varying inorganic salt additions (e.g. CuCl₂·2H₂O) can significantly alter community composition, leading to fluctuations in product titers.^156,158 Thus, while mixed fermentation offers notable advantages in operational simplification, cost reduction, and environmental adaptability, its drawbacks include the difficulty in regulating complex microbial dynamics, and ensuring stability in production processes.

3.4. Enzyme-assisted catalytic system

In the biosynthesis of PHAs and lactate-based polymers, conventional whole-cell catalysis suffers from high substrate transfer resistance and low conversion efficiency, which limits the product accumulation. To promote biosynthesis, microbial strains are increasingly being used as carriers for enzyme immobilization through surface display techniques, which provide biological activity and are closely linked to metabolic engineering and synthetic biology (Fig. 3D).¹⁵⁹

A composite CO conversion system was developed and displayed on the surface of R. eutropha. This system assembled carbon monoxide dehydrogenase (CooS) and carbon monoxide-binding protein (CooA) on the scaffolding protein (mCbpA) through dockerin-cohesin interactions. The CooA facilitated substrate delivery to cell-associated CooS, enabling efficient conversion of CO to CO₂, which was then metabolized by R. eutropha via its chemoautotrophic pathways to synthesize PHB.¹⁵⁹ This system achieved a 3.1-fold increase in CO conversion efficiency; however, the long-term continuous operation and recyclability of the enzyme-assisted catalytic system require further validation. As a more comprehensive way, a cell-bridging enzyme complex strategy was proposed to convert synthetic gas into PHB.⁹² A multi-enzyme complex of carbon monoxide dehydrogenase (CooS), CO-binding protein (CooA), and carbonic anhydrase (CA), anchored by minimized scaffoldin (mCbpA), was assembled onto nanoscale cellulose particles. The assembly was further anchored to R. eutropha through the specific binding of the BspA domain in the mCbpA-BspA (MB) fusion protein. The enzyme complex (CooS, CooA, CA) converts CO in syngas to CO₂, which is then transformed into HCO₃⁻. This approach offers the advantage of converting the low-solubility gaseous substrate into a high-solubility carbonate, thereby reducing mass transfer resistance to the microbial cells and enhancing substrate delivery efficiency. After 72 h fermentation using synthetic gas (H₂ : CO₂ : CO : N₂ = 2 : 2 : 2 : 4), the PHB concentration reached 14.2 g L⁻¹. This work expands the substrate range of the enzyme-assisted catalytic system, significantly enhancing the accumulation of PHB produced by R. eutropha from syngas, with a titer increase of 4.5 times (14.2 g L⁻¹). Additionally, immobilizing enzymes on nanocellulose particles improves enzyme stability; for instance, immobilized CA retains 77.7% activity after storage at 4 °C for 5 days, compared to only 52.3% for free enzymes. This provides a viable direction for improving enzyme stability in display technology on cell surfaces. Furthermore, transitioning from traditional intracellular metabolic engineering to a more modular and controllable synthetic biology approach allows for precise metabolic reprogramming of host cells, enabling them to function not only as production factories for key catalytic enzymes but also as customizable living catalytic interfaces. By relocating synthetic reactions to the extracellular space, interference with intracellular metabolic pathways is minimized, representing a significant innovation in bioplastics synthesis. For example, expressing the fusion protein lipoprotein-outer membrane protein A-phaC (lpp-ompA-phaC) in E. coli enables the display of PHA synthase on the outer membrane, facilitating extracellular PHB synthesis.¹⁶⁰ Thus, while the enzyme-assisted catalytic system holds great potential for increasing biopolymer titers and substrate conversion efficiency, the long-term stability of surface enzyme complexes remain unverified, and large-scale production applications have not been realized yet.

4. Characterization of PHAs and lactate-based biopolymers

Precise characterization of PHAs and lactic acid-based polymers primarily focuses on core aspects including complex component separation, chiral structure identification, molecular weight distribution determination, and degradation product analysis.¹⁶¹ The compositional profile and residual by-products of PHAs and lactate-based biopolymers are critical indicators for predicting their biodegradability and physical properties.¹⁶² Currently, in addition to chromatography, which remains the most widely used technique, a range of other methods including spectroscopic, physicochemical and thermal analyses are employed (Fig. 4).¹⁶³ And these techniques are essential for assessing polymer structure, thermal behavior, and microbial degradation profiles.

Fig. 4 Characterization methods for PHAs and lactate-based biopolymers.

4.1. Chromatographic analysis

The precise characterization of biopolymers relies heavily on chromatographic techniques for separation and quantification, notably high-performance liquid chromatography (HPLC), gel permeation chromatography (GPC), gas chromatography (GC), and gas chromatography-mass spectrometry (GC-MS) (Fig. 4).¹⁶⁴ And the selection of appropriate analytical strategies is vital for achieving accurate and reliable characterization.

HPLC enables the separation and quantification of biopolymer components.¹⁵ PHBV monomers are not directly detectable by conventional HPLC. Therefore, PHBV monomers were converted into detectable small molecule acids: crotonic acid (from 3HB hydrolysis) and 2-pentenoic acid (from 3HV hydrolysis). Quantitative analysis of crotonic acid and 2-pentenoic acid was performed using HPLC coupled with photodiode-array detection and an Inertsil 100A ODS-3 C18 reverse-phase column (5 μm, 250 × 4.6 mm, MZ-Analysentechnik GmbH, Mainz, Germany), with UV detection at 210 nm. And the detection limit achieved 0.01 g L⁻¹.¹⁶⁵ Additionally, the LA content in P(LA-co-3HB) was evaluated using HPLC to monitor structural changes in complex materials during degradation.¹⁶⁶ And HPLC coupled with a Chiralpak IG column (5 μm, 2.0 mm × 250 mm, Daicel, Osaka, Japan) facilitated the analysis of enantiomeric ratios of LA in lactate-based biopolymers.^167,168 Although HPLC analysis may exhibit strong dependence on derivatization pretreatment,¹⁶¹ its high sensitivity, accuracy, and robust quantitative capacity have driven its extensive adoption in industrial applications GPC, also known as size exclusion chromatography (SEC), measures molecular weight distributions and enables the determination of key parameters for biopolymers, including the number-average molecular weight (M_n), weight-average molecular weight (M_w), peak molecular weight (M_p), and polydispersity index (PDI).¹⁶⁹ In GPC analysis of P(3HB-co-4HB), reproducible results were obtained within specific ranges for number-average molecular weight (M_n, 1 × 10⁴–1 × 10⁶) and polydispersity index (PDI, 0.2–3.9).¹⁷⁰ Furthermore, GPC confirmed that hydrothermal treatment of PLA microplastics (PLA-MPs) at 180 °C led to reduced polymerization degrees (e.g., Mn, M_w, M_p) and increased PDI due to hydrolytic chain scission.¹⁶⁹ Additionally, GPC characterization revealed that the molecular weights of PHAs varied with extraction methods. Mondal et al.¹⁷¹ demonstrated that chloroform extraction coupled with ultrasonication caused a slight decrease in weight-average molecular weight (M_w) of PHAs (from 280.8 kDa to 275.0 kDa), indicating that ultrasonication primarily physically disrupts cell walls to release PHAs without significantly altering their chemical structures. In PHAs or lactate-based polymer production, GPC enables rapid and reliable determination of polymerization degree and polydispersity, facilitating consistent monitoring of molecular weight uniformity.

GC commonly determines biopolymer chemical composition after pretreatments like transmethylation and solvent extraction.^172,173 And GC detection of P(3HB-co-3HV-co-3HHx) copolymers enables precise quantification of 3HV monomers within the range of 2–60 mol%.¹⁷⁴ In contrast, thermally assisted hydrolysis and methylation-gas chromatography (THM-GC) offers a rapid and accurate approach for determining the composition of biopolymers. Its advantage lies in integrating the depolymerization and derivatization processes, which solves the problems of low volatility and thermal instability of biopolymers in traditional GC analysis. By reducing side reactions and coking caused by high-temperature pyrolysis, the accuracy of the analysis is improved. In a comparative study, THM-GC efficiently determined the PHB composition produced by C. necator, yielding a strong linear correlation (r² = 0.9972) with conventional GC results. This confirms THM-GC as a reliable and practical technique for analyzing microbial PHAs.¹⁷⁵ In addition, GC-MS is widely used for identifying chemical structures and degradation products of biopolymers, enabling trace analysis.¹⁷⁶ However, the complex pretreatment and derivatization steps of GC-MS hinder rapid online detection and necessitate a highly comprehensive database for comparative analysis. GC-MS analysis of PHB extracted from Bacillus paramycoides MCCC 1A04098 identified 2,4-ditert-butylphenol as a major component. In contrast, Azotobacter salinestris NBRC 102611 and Brevundimonas naejangsanensis BIO-TAS2-2 primarily produced the isopropyl ester of 2-butenoic acid.¹⁷⁷ And GC-MS was also employed to confirm P(3HB) accumulation in Ralstonia solanacearum,¹⁷⁸ and to validate the presence of PHAs copolymers in P. aeruginosa EO1.¹⁷⁹ Generally, GC/THM-GC is a fundamental separation method suitable for the quantification of known components in biopolymers and is a commonly used technique in industrial applications. GC-MS combines the excellent separation ability of GC with the powerful identification ability of MS, demonstrating advantages in high sensitivity and specificity for the identification of unknown substances and the analysis of complex sample components.

4.2. Spectroscopic analysis

Spectroscopic techniques, including Nuclear Magnetic Resonance (NMR), Fourier Transform Infrared Spectroscopy (FTIR), X-ray Diffraction (XRD), and X-ray Fluorescence (XRF), are widely applied for the characterization of PHAs and lactate-based biopolymers (Fig. 4).

Nuclear Magnetic Resonance (NMR) spectroscopy is extensively used to quantify moisture, fat, oil content and identify biopolymers composition. In addition, NMR can achieve quantitative analysis of monomer components in different biopolymers based on internal or external standard methods.¹⁷¹ In the detection of P(3HB-co-3HV) by NMR, the detection limits for 3HB and 3HV reached a monomer mole fraction of 1 mol%.¹⁸⁰ Online low-field NMR analysis was performed using a Spinsolve 43 Carbon Ultra instrument with a flow-assembly based on PTFE tubing to enable real-time monitoring of PHAs extraction. The results obtained were highly correlated with both offline gas chromatography (GC) and high-field NMR (correlation coefficient >94%), confirming the suitability of low-field NMR as a practical analytical technology for optimizing downstream PHAs processing.¹⁸¹ Moreover, proton NMR (¹H NMR) was performed on PHAs synthesized by Saccharophagus degradans from glucose and seaweed. Characteristic chemical shifts at 1.27 ppm, 2.33–2.58 ppm, and 5.3 ppm were attributed to methyl (CH₃), methylene (CH₂), and methine (CH) protons, respectively. And ¹H NMR analysis confirmed that the carbon source had no significant effect on the polymer composition.¹⁸² Similarly, PHB production by Bacillus cereus SH-02 was validated using ¹H NMR, ¹³C NMR, and GC-MS.¹⁸³ The chemical structure of PHBV containing 18.0 mol% 3HV in Haloferax mediterranei was identified via¹H NMR.¹⁸⁴ And combined ¹H NMR, ¹³C NMR, and 2D COSY NMR analyses confirmed the structure of the product from engineered S. elongatus as a PLA homopolymer.⁴⁸ These results demonstrated that NMR is suitable for precise chemical structure identification of biopolymers (e.g., PHAs and lactate-based biopolymers). However, the high cost of NMR equipment and the intricate sample detection procedures limit its application for rapid, routine quality control in industrial production.¹⁷¹

Fourier Transform Infrared Spectroscopy (FTIR) analyzes molecular vibrations and identifies chemical bonds, facilitating both screening and quantification of intracellular PHAs.¹⁸⁵ And FTIR offers several advantages, including improved signal-to-noise ratio, faster analysis speed, and simultaneous full-wavelength detection. FTIR analysis of P(3HB) and P(3HB-co-3HV) was performed using the KBr pellet method with a scanning range of 4000–400 cm⁻¹, requiring minimal sample preparation.¹⁶⁵ However, this approach only enabled qualitative identification of characteristic ester functional groups and peak variations induced by comonomer incorporation.¹⁶¹ In a study of Hagagy et al.,¹⁸⁶ FTIR identified PHAs produced by two Halolamina species, with characteristic carbonyl group (C=O) absorption peaks at 1628.98 and 1629.28 cm⁻¹, confirming PHB presence. Additionally, FTIR spectra of P(3HB) and three fillers (peat, clay, and birch wood flour) were distinguished. And crystalline phase bands appeared at 1228 cm⁻¹, while amorphous phase bands shifted to 1182 cm⁻¹.¹⁸⁷ In another study, FTIR analysis was conducted on PHB extracted during a 28 h biocalorimetric co-culture experiment. A strong ester C=O peak at 1723 cm⁻¹ indicated a crystalline structure, while a C–O stretch appeared at 1231 cm⁻¹ and a broad OH absorption at 3425 cm⁻¹.¹⁸⁸ The functional group specificity and operational convenience of FTIR align with industrial requirements for rapid, cost-effective, and straightforward analysis. And FTIR has become an essential qualitative analytical technique for biopolymers synthesis, characterization, and application, proving particularly valuable for monitoring biosynthesis and analyzing multi-component material systems.

X-ray Diffraction (XRD) can analyze the crystalline structure of biopolymers, thereby providing detailed information on purity, crystallinity, and morphology.¹⁶¹ The XRD analysis of P(3HB) and P(3HB-co-3HV) was performed at the BL45XU beamline (λ = 0.09 nm) at SPring-8 (Harima, Japan), achieving sensitivity to detect lattice parameter shifts and crystallinity variations induced by 3HV content changes across a 3HV content gradient of 5.3–16.0 mol%.¹⁸⁹ In addition, XRD analysis showed that PHAs produced from glucose by S. degradans exhibited higher crystallinity than that produced from seaweed, likely due to reduced microcrystalline size in the latter.¹⁸² And XRD combined with FTIR confirmed good compatibility between PLA and PPC in polymer blends, attributed to structural similarity and hydrogen bonding.¹⁹⁰ XRD offers a crucial theoretical basis for material performance optimization and composite compatibility design by analyzing crystal characteristics.¹⁹¹ However, due to the weak diffraction signals of polymer crystals, XRD requires slow scanning to ensure the signal-to-noise ratio. Additionally, XRD data processing is intricate, requiring cross-validation with complementary characterization data.¹⁸⁹ X-ray Fluorescence (XRF) is a preferred technique for non-destructive analysis of trace elements in polymers, which include various inorganic additives, such as plasticizers, stabilizers, antioxidants, pigments, and catalytic agents, with minimal sample preparation.¹⁶² Using an energy-dispersive X-ray fluorescence (EDXRF) model (FXL 950, Niton, USA) equipped with a 50 kV silver X-ray tube, the limits of detection (LOD) for Mn, Co, Zn, and As in seafood products were determined to be 5 μg g⁻¹, 2 μg g⁻¹, 1 μg g⁻¹, and 1 μg g⁻¹, respectively.¹⁹² Parallelly, energy-dispersive X-ray fluorescence (EDXRF) was used to analyze 33 types of single-use plastics (SUPs) collected in Mexico. Concentrations of metal elements, including Cu (1898 μg g⁻¹), Cr (1586 μg g⁻¹), Mo (95 μg g⁻¹), Zn (1492 μg g⁻¹), Fe (1900 μg g⁻¹), and Pb (7528 μg g⁻¹), were found to exceed international safety standards.¹⁹³ This highlights the capability of XRF for rapid, simultaneous elemental analysis of solid samples like bioplastics, making it suitable for industrial production.

4.3. Physicochemical analysis

Physicochemical analysis are commonly used to characterize the microstructure and mechanical properties of biopolymers (Fig. 4). Electron microscopy, including scanning electron microscopy (SEM) and transmission electron microscopy (TEM), provides detailed morphology of surface and internal structure.¹⁹⁴ Scanning electron microscopy (SEM) offers high sensitivity and can resolve the surface morphology of nanometer-scale biopolymers without the need for complex sample preparation processes.^161,195 SEM was used to observe surface degradation of P(3HB), P(3HB-co-3HV).¹⁹⁴ And untreated PLA samples exhibited smooth and uniform surfaces, whereas samples treated with Aspergillus flavus CCUG 28296 showed signs of surface deterioration.¹⁹⁶ In addition, SEM micrographs of Trichoderma and Fusarium oxysporum-treated PLA samples after 14 d revealed roughened surfaces, the presence of surface cracks and pores, and fungal hyphae with spore formation. These features were absent in untreated samples, confirming fungal-mediated biodegradation.¹⁹⁷ SEM was also used to evaluate the morphological changes in 28% P(LA-co-3HB), 69% P(LA-co-3HB), and P(3HB) during a 10-week river water degradation study. All polymers developed surface holes post-immersion, indicating biodegradation.¹⁶⁶ SEM directly confirms microbial/enzymatic erosion and degradation of polymeric materials by detecting surface pores, corrosion, and cracks. Transmission electron microscopy (TEM) was applied to examine ultrathin cross-sections of samples, enabling direct visualization of intracellular polymer accumulation.^161,194 TEM characterization of P(3HB) and P(3HB-co-3HV) precisely detects nanostructural features and defects, resolving microcracks (≥2 nm), amorphous region boundaries (5–10 nm), and microfibril bundles (50–100 nm), while effectively visualizing subtle interactions at cell-polymer interfaces.¹⁹⁸ And TEM enabled the characterization of internally formed PLA/PHAs, revealing details such as particle aggregation, a smooth circular morphology, and consistent particle size.^199,200 TEM not only precisely characterizes the microstructure of biopolymers, but also directly resolves internal ultrastructural details in biological samples, highlighting its advantages in analyzing complex systems at nano- to subcellular scales. However, SEM and TEM analyses of PHAs and lactate-based biopolymers require stringent conditions, such as high-vacuum environments, making them unsuitable for volatile, fragile, or vacuum-sensitive samples. And the qualitative morphological information obtained must be combined with complementary analytical techniques to definitively determine the specific composition of biopolymers.¹⁶¹

In addition to electron microscopy, mechanical properties of polymers can be assessed using an extensometer, which measures tensile strength and viscoelastic properties.²⁰¹ Another widely used physical technique is the analytical balance method, which quantifies polymer degradation through mass loss, which is direct, practical, and broadly accessible. Gravimetric analysis typically employs high-precision analytical balances, achieving sensitivity levels of approximately 0.01–0.1 mg.¹⁶² As indicated by research, weight loss of P(3HB), 28% P(LA-co-3HB), and 69% P(LA-co-3HB) samples in river water was monitored over 10 weeks.¹⁶⁶ However, mass loss measurements alone may not reliably predict complete biodegradation.¹⁶¹ Therefore, additional analytical methods such as respirometry, FTIR, and NMR should besupplemented.²⁰² The combination of weight loss and respirometric techniques was demonstrated to compare the degradation of P(HB-co-HV) and poly(butylene succinate-co-adipate) (PBSA). And weight loss at 450 d reached 5.5% and 8.0%, respectively.²⁰³ In addition, respirometric analysis showed corresponding CO₂ emissions, emphasizing that weight loss does not always indicate microbial mineralization of polymer material. The respirometric method encompasses indirect techniques used to assess the respiratory metabolic activity of microorganisms.¹⁶² Detection of oxygen sensor-based respirometry can be reliably conducted with a minimum liquid foods microbial concentration of 4 CFU mL⁻¹.²⁰⁴ The application of respirometry combined with a modified Gompertz growth model to fit dynamic oxygen uptake rates enables determination of biosynthesis/degradation rates for PHAs and lactate-based biopolymers.²⁰⁵ The indirect nature of respirometry in assessing microbial activity determines its status as a supplementary tool for evaluating biopolymer synthesis and degradation processes.

4.4. Thermal analysis

Thermal analysis is a valuable technique for evaluating the physicochemical properties of polymers, offering the advantage of requiring minimal sample pretreatment. Common thermal analysis techniques include differential scanning calorimetry (DSC), differential thermal analysis (DTA), and thermogravimetric analysis (TGA) (Fig. 4).²⁰⁶

DSC/DTA involves the application of a controlled heating or cooling rate to a sample while monitoring enthalpic/temperature changes associated with thermal transitions such as melting, crystallization, glass transitions, and decomposition.^207,208 A Diamond-DSC instrument (PerkinElmer) was employed to assess the influence of curcumin on the thermal behavior of PLA/PPC blends, which possesses low crystallinity for high transparency.¹⁹⁰ DSC/DTA analysis of P(3HB) or P(3HB-co-3HV) demonstrates sensitivity to phase transition temperature differences (0.5 °C) and thermal effect variations (1 J g⁻¹).^209,210 Linking DSC/DTA findings to the end-use performance of PHAs and lactate-based biopolymers provides critical guidance for understanding their processing behavior, mechanical properties, thermal stability, and potential applications.^211,212 TGA provides information on the thermal stability of polymer blends/biopolymers as a function of composition, revealing the dynamic characteristics of the degradation process.^196,213 And TGA exhibits high sensitivity among rapid detection techniques with a limit of detection (LOD) of 0.041 mg for polystyrene microplastics in water.²¹⁴ TGA performed on a TA Q500 device (TA Instruments, USA) demonstrated that PLA/PPC blends containing curcumin remained thermally stable during melt extrusion and compression molding, indicating material durability.¹⁹⁰ In addition, TGA analysis revealed a downward shift in maximum weight loss temperature with broadened weight loss peaks, indicating PLA degradation through reduced molecular weight, diminished thermal stability, and increased material heterogeneity.¹⁹⁶ Despite certain limitations in practical applications of thermal analysis techniques, such as their inability to provide chemical information on the degradation of biopolymers, their integration with other methods is imperative for comprehensive polymer analysis.

5. Biodegradation of PHAs and lactate-based biopolymers

PHAs and lactic acid-based biopolymers demonstrate significant potential across multiple fields due to their biodegradability. In biomedical applications, PHAs show promise in tissue engineering and drug delivery systems. High-crystallinity P(3HB) emerges as an optimal candidate for rigid scaffolds, while elastic poly(4-hydroxybutyrate) (P(4HB)) serves as ideal suture material for in vivo degradation without secondary surgery.^215,216 In addition, P(4HB) fiber membranes act as artificial dermal layers, enhancing full-thickness skin defect healing with excellent biocompatibility, gas permeability, and anti-contraction properties.²¹⁷ And PHAs also enable eco-friendly production of compostable tableware and food packaging, reducing plastic waste.²¹⁸ PHBV, notable for high mechanical strength, is utilized in daily disposables (cosmetics, packaging, containers, bags, hygiene products) and industrial components (printed circuit boards, automotive panels, bicycle helmets).^219,220 Moreover, lactic acid-based polymers exhibit agricultural applications through biodegradable mulch films for cotton cultivation, effectively suppressing weeds. These films degrade post-harvest, minimizing retrieval costs and pollution compared to conventional mulch, while enabling nutrient release. And studies reported complete degradation of PHA/PLA blended mulch within 176 d.^10,221 When compounded with nanoparticles (e.g. TiO₂), lactic acid-based biopolymers can be applied in advanced coatings, sensors, and electronic devices.²²² However, PHAs and lactic acid-based polymers exhibit variable degradation rates and extents under different oxygen conditions, categorizing their biodegradation into aerobic and anaerobic processes. The overall biodegradation pathways of PHAs and lactate-based biopolymers are illustrated in Fig. 5.

Fig. 5 Aerobic (A) and anaerobic (B) biodegradation pathways of PHAs and lactate-based biopolymers.

5.1. Aerobic biodegradation of PHAs and lactate-based biopolymers

5.1.1. Aerobic biodegradation mechanism of PHAs and lactate-based biopolymers. Aerobic biodegradation process of PHAs and lactate-based biopolymers generally involves four stages: surface biodegradation, biological fragmentation, assimilation, and final mineralization.^223–226 Under aerobic conditions, the biodegradation of different PHAs and lactic acid-based polymers exhibits marked variations in enzymatic involvement, intermediate products, and metabolic pathways, reflecting the influence of polymer structural characteristics on degradation (Fig. 5A). PHB is depolymerized by PHB depolymerase into β-hydroxybutyrate, which is subsequently converted to acetyl-CoA via the β-oxidation pathway and finally fully oxidized to CO₂ and H₂O through the TCA cycle. In contrast, PHBV is catalyzed by PHA depolymerase to yield D-3-hydroxybutyrate, which is then transformed into acetoacetate by D-3-hydroxybutyrate dehydrogenase, further generating acetyl-CoA before entering the TCA cycle for metabolism. PHA depolymerases are diverse and have been isolated and identified from various microorganisms, including fungi (e.g. Penicillium funiculosum) and several bacteria (Paucimonas lemoignei, Bacillus thuringiensis, Alcaligenes faecalis, Caldimonas manganoxidans and Ralstonia pickettii T1).^227–230 The degradation mechanism of PLA differs significantly from that of PHAs, primarily involving esterases or proteases that catalyze the production of LA. LA is converted to pyruvate by lactate dehydrogenase, which is then transformed into acetyl-CoA by pyruvate dehydrogenase, ultimately entering the TCA cycle. For P(LA-co-3HB), the action of phaZs depolymerases generates β-hydroxybutyrate and LA. β-Hydroxybutyrate is converted to acetyl-CoA via β-oxidation, while LA is transformed into acetyl-CoA through lactate dehydrogenase and pyruvate dehydrogenase, with both pathways leading to the TCA cycle for oxidative degradation. Specific proteases and esterases play crucial roles in the degradation of lactate-based biopolymers, with reported sources including bacteria (Amycolatopsis sp. strain K104-1, Actinomadura keratinilytica T16-1) and fungi (Arthrobotrys oligospora),^231,232 while esterases have been identified from P. aeruginosa strain S3.²³¹ Based on these degradation pathways, all routes ultimately converge at acetyl-CoA, a central metabolic intermediate, which then enters the TCA cycle for complete oxidation. This reveals the common steps in the aerobic biodegradation of PHAs and lactic acid-based biopolymers.

Achieving efficient degradation and recycling of PHAs and lactic acid-based biopolymers is a core step in establishing a green closed-loop lifecycle for bio-based plastics. Protein engineering offers the potential to enhance the performance of biodegradation enzymes for bioplastics and traditional petrochemical plastics. For instance, to develop an efficient PET-degrading enzyme, the keratinase ThcCut1 from the aerobic bacterium Thermobifida cellulosilytica was engineered through rational design. Mutations at the substrate-binding site (G63A/F210I/D205C/E254C/Q93G) resulted in a 1.165-fold increase in enzyme activity and an over 20 °C rise in T_m, enabling the enzyme to efficiently degrade 96.2% of discarded PET bottle fragments within 96 h at 70 °C.²³³ This improvement is attributed to the enlarged substrate-binding pocket, facilitating polymer entry and binding to the enzyme's active site, thereby enhancing enzyme activity. Additionally, the formation of disulfide bonds increased the rigidity of the protein structure, improving thermal stability. Additionally, in the aerobic Pseudomonas putida KT2440, the activity of the PHA depolymerase (PhaZKT) is critically dependent on a lid region. Mutations in a region distant from the lid, such as S184F, enhanced esterase activity towards small substrates but reduced depolymerization activity. Molecular dynamics revealed lid rigidity hindering polymer binding, while G286R mutation boosted depolymerase activity and long-chain substrate affinity. The lid structure is crucial for enzyme function and difficult to directly modify, and engineering mutations at distal sites can optimize the enzyme's catalytic performance.²³⁴ Similarly, site-saturation mutagenesis of a PLA hydrolase from A. keratinilytica yielded mutant PAM (S101F, S103L, T106I) with 4.5-fold higher activity, achieving 90% PLA degradation at pH 7.5 and 45 °C within 24 h versus 20% for wild-type.²³⁵ Activity enhancement stemmed from optimized hydrophobic interactions (via phenylalanine/leucine substitutions) and steric complementarity stabilizing transition states. Rational design and site-directed mutagenesis emerge as key strategies to improve enzymatic activity and stability, advancing efficient degradation and recycling of PHAs and lactate-based biopolymers.

5.1.2. Aerobic biodegradation conditions of PHAs and lactate-based biopolymers. Aerobic degradation of PHAs and lactate-based biopolymers is often considered more environmentally favorable.²³⁶ And leveraging the enzymatic activity of various microorganisms under controlled aerobic conditions facilitates the degradation process.⁸ Aerobic degradation can take place in soil, aquatic environments, and composting systems. Microbial degradation of PHAs and lactate-based biopolymers under different environmental conditions is summarized in Table 5.

Table 5 Microbial degradation of PHAs and lactate-based biopolymers under different environmental conditions

Microorganisms	Degradation substrates	Enzymes involved in the degradation	Degradation environment	Ref.
a Not reported. b Aerobic biodegradation. c Anaerobic biodegradation.
M. chlamydosporia, P. pedernalense, Gliocladiopsis sp., M. hiemalis, P. lilacinum	P(3HB)	PHA depolymerase	Red soil^b	240
P. chrysogenum, T. purpureogenus, T. funiculosus	P(3HB)	PHA depolymerase	Soil^b
B. pumilus, Paraburkholderia sp., Pseudomonas sp., Rhodococcus sp., S. rhizophila, S. prunicolor, and V. paradoxus	P(3HB)	P(3HB)-depolymerase	Soil^b	241
Gemmobacter sp., Caulobacter sp., C. Planktophila	P(LA-co-3HB)	N^a	River water^b	166
Alcanivorax sp. 24	PHB and PHBV	Esterase ALC24_4107	Seawater^b	248
A. plasticoclasticus MED1	PHBV	depolymerase ALSP_2657	Marine^b	246
Shewanella sp. JKCM-AJ-6,1α	P(3HB) and (R)-3HB	Poly(3-hydroxybutyrate) depolymerase (EC 3.1.1.75)	Sea water^b	247
Marinobacter sp. NK-1	P(3HB) and (S)-3HB	Poly(3-hydroxybutyrate) depolymerase (EC 3.1.1.75)
Alcanivorax sp.24	PHB	Depolymerase ALC24_4107
DTU014, Clostridiales, Methanobacterium	PLA	Esterase/protease	Anaerobic digestion^c	265
T. xylanilyticum	PLA	N	Anaerobic digestion^c	266
N	PHBHHx	N	Anaerobic digestion^c	268
N	P(3HB-co-4HB)	N	Anaerobic digestion^c	258

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The degradation rate of PHAs and lactate-based biopolymers in soil is heavily dependent on both the soil environment and the local microbial community.²³⁷ The degradation of lactate-based biopolymers is more effective at higher soil pH levels, as most degrading enzymes for lactate-based polymers exhibit optimal activity at alkaline or neutral pH.²³¹ However, It is worth to mention that PHAs films exhibited only 47% degradation in nearly neutral soil (pH 6.63, Dam Bai, Vietnam), whereas 98% degradation occurred in slightly acidic soil (pH 5.48, Hoa Lac, Vietnam)²³⁸ Additionally, soil moisture influences microbial activity and biopolymers hydrolysis. The optimal soil moisture content for aerobic degradation for PHAs and lactate-based biopolymers has been identified as 50–60%.²³⁹ Moreover, the composition of microbial communities is critical to the biodegradation of PHAs and lactate-based biopolymers. A comparison of P(3HB) degradation in red ferralitic soil under tropical conditions (Kerala, India) and chernozem soil under conditions of a sharply continental climate (Eastern Siberia, Russia) was conducted. And the results indicated that the abundance of microorganisms in red ferralitic soil was notably lower than in chernozem soil, and correspondingly, the degradation rate of P(3HB) films was slower in red ferralitic soil than in chernozem soil.²⁴⁰ This highlights the role of microbial community composition in determining biodegradation efficiency under different climatic conditions. Interestingly, microbial communities may evolve the capability to degrade bioplastics even in extreme environments. A study analyzed the microbial community structure in low-temperature soils of the Siberian subarctic region, focusing on the ability of the soil microbial community to metabolize degradable P(3HB), and the results indicated substantial biodegradation activity even under cold environmental conditions.²⁴¹ It should be pointed out that the composition of biopolymers also affects their degradation. The study of van der Zee et al.²⁴² demonstrated that blending PLA with polycaprolactone (PCL) significantly enhances its biodegradability in soil. After a 12 months soil incubation, the CO₂ conversion rate of PLA/PCL mixtures reached 39%.

Plastic pollution in marine environments has become a critical concern, with 70–80% of plastic waste from terrestrial sources eventually infiltrating marine ecosystems.²⁴³ The ability of PHAs and lactate-based biopolymers to undergo biodegradation in marine conditions has been confirmed.²⁴⁴ Furthermore, lactate-based biopolymers have demonstrated high degradation capability in subtropical marine environments, achieving complete decomposition within 1 to 3 mo. And the degradation efficiency of PLA in marine environments was found to significantly exceed that in open-air conditions, with mass loss values reaching up to 50%.²⁴⁵ Many marine bacteria possess specialized enzymatic systems for degrading PHAs and lactate-based biopolymers. For PHAs, Alteromonas plasticoclasticus MED1 employs tandem depolymerases (ALSP_2657 and ALSP_2202/2201) to degrade PHBV, while strains such as Shewanella sp. and Marinobacter sp. produce substrate- or monomer-induced depolymerases. Additionally, Alcanivorax sp.24 secretes depolymerase ALC24-4107 capable of hydrolyzing PHB. These findings collectively demonstrate marine microbes have evolved diverse enzymatic repertoires to utilize bioplastics as carbon sources.^246–248 For lactate-based polymers, marine genera including Symbiobacterium, Sphingobacterium, Comamonas, Pseudomonas, and Alcaligenes degrade PLA via esterases and lipases,²⁴⁹ while Agarifytica secretes PHA depolymerases for P(LA-co-3HB) degradation.²⁵⁰ Beyond marine systems, biodegradation of PHAs and lactate-based polymers in other aquatic environments exhibits ecological significance. Kadoya et al.¹⁶⁶ reported 20% weight loss of P(LA-co-3HB) (28–69 mol% LA) in river water after 10 weeks, with enriched bacteria (Gemmobacter sp., Caulobacter sp., Candidatus Planktophila, etc.) harboring PHB depolymerase homologs. This highlights how environmental conditions and microbial consortia influence degradation efficiency across aquatic systems.

Composting provides an efficient approach for converting bioplastic waste into bioresources.²⁵¹ Compared to mechanical recycling or incineration, it offers sustainable organic recycling with economic viability.²⁵² Industrial composting accelerates degradation of PHAs/lactate-based biopolymers through controlled temperature, humidity, oxygenation, and pH.^253–255 Thermophilic conditions (50–60 °C) selectively enrich thermophilic polymer-degrading microbiota, enhancing enzymatic performances.²³⁹ For instance, PHBV (26% HV) showed 59% mass loss over 186 d in composting (40–63 °C).²⁵⁶ And successful implementation in German industrial facilities yields marketable compost from biodegraded plastics,²⁵³ demonstrating composting's capacity to optimize decomposition of biopolymers through parameter modulation.

5.2. Anaerobic biodegradation of PHAs and lactate-based biopolymers

5.2.1. Anaerobic biodegradation mechanism of PHAs and lactate-based biopolymers. The anaerobic biodegradation of PHAs and lactate-based biopolymers proceeds through four stages in oxygen-free environments: hydrolysis, acidogenesis, acetogenesis, and methanogenesis, ultimately generating biogas (CH₄/CO₂ mixture).²⁵⁷ PHAs are depolymerized by PHA depolymerases/esterases/lipases into short-chain monomers (e.g. 3HB, 3HV), which enter β-oxidation pathways to form acetyl-CoA. Subsequent conversion to acetate occurs via phosphotransacetylase and acetate kinase, with acetoclastic methanogens (Methanosaeta) further transforming acetate into CH₄ and CO₂.²⁵⁸ This process relies on synergistic microbial consortia.²⁵⁹ In addition, PLA is hydrolyzed by esterases/proteases to lactate, converted by acidifying bacteria to volatile fatty acids (e.g. acetate, propionate and butyrate), which methanogens ultimately metabolize to CH₄ and CO₂.^260,261 Understanding these mechanisms is critical for optimizing anaerobic digestion of biopolymers (Fig. 5B).

Notably, computationally guided rational design strategies have been successfully applied to modify PLA-degrading enzymes, enhancing their performance under high-temperature anaerobic digestion conditions. The PLA-degrading esterase RPA1511 exhibited significantly improved thermal stability through the introduction of five key mutations (S153L, I245V, R276D, F151L, and A267G), resulting in an increase in T_m by 8.57 °C and an 11.5-fold enhancement in relative enzyme activity. The increase in the T_m of the degrading esterase facilitates a reduction in the rigidity of the polymer during degradation, improving enzyme–substrate contact. Additionally, a further V202W mutation allows the mutant enzyme to form hydrogen bonds with the substrate, enhancing enzyme–substrate binding. Free energy calculations indicate that the binding free energy of the mutant-substrate complex is reduced, contributing to increased enzyme activity. At 65 °C, the mutant enzyme (S153L/I245V/R276D/S128Y/V202W) converted PLA powder (molecular weight M_w of 10 000–18 000 Da) into LA monomers within 72 h, and the conversion rate reached 85.38%, which was 3.3 times higher than that of the wild-type enzyme.²⁶² The dual enhancement of enzyme activity and thermal stability in PLA-degrading enzymes presents a promising approach for efficient anaerobic degradation of PLA.

5.2.2. Anaerobic biodegradation conditions of PHAs and lactate-based biopolymers. Biodegradable plastics are primarily treated via aerobic composting (emitting CO₂) or landfilling (generating non-collectable CH₄), both contributing to greenhouse gas release.²⁶³ In contrast, anaerobic digestion in closed reactors converts waste into biogas (renewable energy) and digestate (fertilizer), offering controlled, eco-friendly, and resource-efficient processing with shorter treatment durations. The collectable methane produced can be utilized for power generation, heating, or upgraded to biomethane.^263,264 Temperature critically governs anaerobic degradation of PHAs and lactate-based biopolymers. Lactate-based polymers show negligible degradation (4.4%) under mesophilic (37 °C) conditions but achieve 93.0% degradation under thermophilic (55 °C) conditions. Thermophilic consortia (e.g. DTU014, Clostridiales, Methanobacterium) drive PLA degradation, with DTU014 likely mediating propionate breakdown and Methanobacterium participating in methanogenesis.²⁶⁵ Thermophilic microbes encode heat-resistant esterases/hydrolases that cleave PLA ester bonds efficiently. In addition, studies corroborate temperature-dependent degradation: PLA-based coffee capsules degraded 24% at 38 °C vs. 58% at 58 °C over 100 d,²⁶⁶ while lactate-based cups degraded 90% in 40 d at 58 °C versus 66% over 280 d at 37 °C.²⁶⁷ Notably, injection-molded PLA pots exhibited minimal degradation (13%) despite 60 d at 58 °C, attributed to enhanced crystallinity impeding microbial and moisture penetration.²⁶⁴ It is worth to mention that, PHAs demonstrate superior anaerobic degradability at moderate temperatures. PHAs achieved 93% degradation within 10 d at 38 °C,²⁶⁸ while P(3HB-co-4HB) films degraded 70% at 37 °C.²⁵⁸ These results confirmed that PHAs-based materials are more readily decomposed under anaerobic conditions compared to lactate-based biopolymers. Notably, material morphology significantly influences degradation rates. For instance, PHBHHx particles degraded by 54.6%, while films achieved 77.1% degradation within 85 d at 38 °C.²⁶⁸ The larger specific surface area of films (10 µm to 50 µm) compared to particles (125 µm to 1 cm) increases the contact area between microorganisms and the material, facilitating microbial attack and degradation. Thus, compared to traditional aerobic treatment, anaerobic digestion offers a more environmentally friendly and resource-efficient approach for processing biodegradable plastics. This is particularly evident under high-temperature conditions, which can significantly enhance the degradation efficiency of lactate-based polymers, with material morphology also playing a crucial role in influencing degradation rates.

6. Challenges and prospects

Significant progress has been achieved over the past decade in the biosynthesis of bioplastics and their monomers from C1 substrates. However, achieving high titer, substrate conversion rate, and productivity of PHAs and lactate-based biopolymers derived from C1 resources remains a major prerequisite for economically viable bioproduction. In parallel, characterization methods and understanding of dynamic change mechanisms for these biopolymers must be further developed to support their environmentally responsible application. Key challenges and prospective directions are outlined below:

6.1. Exploring new microbial resources and elucidating novel biosynthesis regulatory mechanisms

Various microorganisms, particularly microalgae and bacteria, have been extensively studied for PHAs and LA-based biopolymers production. However, direct biosynthesis of PHAs from C1 substrates by yeast has been rarely reported, and the production of LA-based polymers via similar pathways remains largely unexplored. The microbial resource pool for PHAs and LA-based biopolymers biosynthesis from C1 substrates remains underdeveloped, and comprehensive insights into the associated metabolic networks are still lacking. Further studies are required to screen novel microbial strains, especially extremophilic species and non-conventional microorganisms, with potential for efficient conversion of C1 resources into PHAs and LA-based biopolymers. And integrated multi-omics analyses, combined biochemical characterization, and computational modeling can systematically resolve intracellular carbon flux allocation, precursor supply, biopolymer synthesis, and critical metabolic regulatory nodes, thereby enabling precise target libraries for iterative optimization of engineered microbial strains. In addition, one of the key barriers to commercialization is the high production cost of PHAs and LA-based biopolymers. It is essential to improve microbial fermentation performances through targeted engineering of rate-limiting enzymes, metabolic network optimization to redirect intracellular carbon flux, balanced regulation of reducing power and energy metabolism, and strain adaptation, thereby maximizing carbon source conversion into target products.

6.2. Establishment of innovative fermentation processes

Multiple fermentation strategies have been investigated to enhance the production of PHAs and lactate-based polymers from C1 substrates. These include single-stage or dual-stage bioprocess, bio-electrochemical system, mixed-culture fermentation, and enzyme-assisted catalytic system. These approaches aim to improve mass transfer of gas-phase substrates, expand the adaptability to diverse C1 feedstocks, and enhance the stability of the biotransformation process. And bioelectrochemical system represents an emerging field, with a focus on optimizing electrode material design and enhancing electron transfer efficiency as key areas for future research.

To address the bottleneck of low control accuracy and dynamic regulation challenges in industrial fermentation of bio-based polymers caused by complex microbial metabolic networks and high multivariable coupling, a hybrid intelligent model combining data-driven ML with mechanism-based metabolic regulatory frameworks can be developed. This integration enhances predictive capability and generalization performance while maintaining interpretability. An emerging frontier in this area is the application of artificial intelligence (AI) in fermentation process control. To address the challenge of dynamic nutrient influences on PHAs synthesis in continuous bioreactors, a hybrid model integrating metabolic mechanism-based frameworks with AI-driven neural network analysis was developed. This model combines microbial growth kinetics and substrate uptake mechanisms while leveraging AI for nonlinear relationship learning, enabling successful simulation and prediction of bioreactor performance under steady-state and dual-nutrient-limited conditions. Furthermore, the AI-based predictive analysis demonstrated exceptional accuracy (R² = 0.918), significantly outperforming traditional purely mechanism-based models.²⁶⁹ Parallelly, a genetic algorithm (GA)-optimized artificial neural network (ANN) model has been applied to predict PHAs concentration in C. necator, based on mixture-process design data.²⁷⁰ The GA-ANN model achieved an R² value of 0.935, markedly surpassing a conventional polynomial model (R² = 0.301), demonstrating AI's efficacy in process prediction and establishing a foundation for intelligent, precise regulation of PHAs and lactate-based polymers production. Future integration of such models with bioreactor systems promises to maximize production efficiency across operational cycles.

6.3. Utilizing combinatorial characterization methods and clarifying dynamic change mechanisms

Integrated characterization techniques (e.g., chromatographic, spectroscopic, physicochemical, and thermal analyses) are essential for elucidating the biodegradation mechanisms and material properties of PHAs and lactate-based biopolymers, providing critical insights into the development of novel degradative enzymes. And identifying key mutation sites in high-efficiency PHAs- and PLA-degrading enzymes is pivotal for developing enzymes with enhanced degradation efficacy. Structure-guided rational engineering has demonstrated substantial potential in constructing hyperthermostable PLA hydrolases. Strategic selection of mutation sites in PLA-degrading enzymes can be achieved through the integration of structural biology and computational simulations. Specific approaches include: resolving three-dimensional enzyme structures and constructing enzyme–substrate transition-state models to precisely identify substrate-binding residues via molecular dynamics simulations; and validating critical sites through sequence conservation analysis.²³⁵

Real-time and in situ monitoring has emerged as a research frontier. To address the limitations of traditional offline detection and delayed analysis in PHAs and lactate-based polymer synthesis/extraction processes, real-time monitoring of poly(hydroxybutyrate-co-hydroxyhexanoate) (P(HB-co-HHx)) extraction via single-scan low-field NMR spectroscopy has recently been demonstrated.¹⁸¹ This approach reveals extraction kinetics through dynamic signal tracking and enables precise endpoint determination to avoid overprocessing or incomplete extraction.

Furthermore, the production and degradation processes of biopolymers are hindered by unclear dynamic mechanisms, leading to imprecise process control, inconsistent product structures, and unquantifiable degradation profiles. By analyzing characteristic peaks associated with monomers and functional groups via in situ Raman spectroscopy, advanced dynamic process control can be achieved during biopolymer manufacturing and degradation.²⁷¹ Online in situ feedback mechanisms not only enhance the predictability and stability of PHAs and PLA-based polymers production/degradation but also provide robust analytical tools for optimizing molecular structures and material properties. These advancements lay the foundation for establishing a comprehensive “synthesis-usage-degradation” evaluation platform, potentially reducing enzyme-dependent degradation costs and supporting sustainable lifecycle management of recyclable bio-based materials.

7. Conclusions

The extensive pollution caused by traditional fossil-based plastics poses a significant threat to the environment, human health, and sustainable development. The utilization of C1 substrates offers a promising avenue for reducing the production cost of bioplastics while supporting global carbon neutrality goals. This review has comprehensively summarized recent advances in the microbial synthesis of PHAs and lactate-based biopolymers from C1 resources, including progress in microbial cell factories, fermentation strategies, characterization techniques, and biodegradation mechanisms. Future research should prioritize the improvement of titer, yield, and productivity in biopolymer production from C1 feedstocks. This can be achieved through the exploration of novel microbial strains, deeper insights into regulatory pathways, and the establishment of innovative fermentation processes. Furthermore, the development of integrated characterization methods and a better understanding of efficient biodegradation mechanisms are essential for enhancing the environmental viability and practical application of these bioplastics.

Author contributions

Yong Wang: conceptualization, funding acquisition, writing – original draft, supervision. Jiaxin Liang: data curation, writing – original draft. Guangye Hu: data curation, writing – original draft. Yumeng Zhen: writing – review&editing. Xu Zhang: visualization. Di Cai: writing – review&editing. Bin Wang: project administration. Jiazheng Sun: visualization. Dejing Kong: funding acquisition, supervision.

Conflicts of interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Abbreviations

AI	Artificial intelligence
ACE	Acetate
ACE-CoA	Acetate-CoA
ANN	Artificial neural network
DCW	Dry cell weight
DSC	Differential scanning calorimetry
DTA	Differential thermal analysis
FTIR	Fourier transform infrared spectroscopy
G3P	Glyceraldehyde 3-phosphate
GA	Genetic algorithm
GC	Gas chromatography
GC-MS	Gas chromatography–mass spectrometry
GPC	Gel permeation chromatography
HPLC	High-performance liquid chromatography
MBH	Membrane-bound hydrogenase
MES	Microbial electrosynthesis systems
MET	Secondary microbial electrochemical technologies
NMR	Nuclear magnetic resonance
OAA	Oxaloacetate
1,3-PGA	1,3-Bisphosphoglycerate
PCT	Propionyl CoA transferase
PDLA	Poly(D-lactic acid)
PDLLA	Poly(D,L-lactic acid)
PEP	Phosphoenolpyruvate
PGA	Phosphoglyceric acid
P(3HB)	Poly(3-hydroxybutyrate)
PHAMCL	Medium-chain-length PHAs
PHAs	Polyhydroxyalkanoates
PHB	Polyhydroxybutyrate
P(HB-co-HHx)	Poly(hydroxybutyrate-co-hydroxyhexanoate)
PHBHHx	Poly(3-hydroxybutyrate-co-hydroxyhexanoate)
PHBV	Poly(3-hydroxybutyrate-co-3-hydroxyvalerate)
PHV	Poly(3-hydroxyvalerate)
PLLA	Poly(L-lactic acid)
R-3-HB-CoA	3-Hydroxybutyryl-CoA
RuBisCO	Ribulose-1,5-bisphosphate carboxylase/oxygenase
SEM	Scanning electron microscopy
SH	Soluble hydrogenase
TCA	The tricarboxylic acid cycle
TEM	Transmission electron microscopy
TGA	Thermogravimetric analysis
THM-GC	Thermally assisted hydrolysis and methylation-gas chromatography
VFA	Volatile fatty acids
XRD	X-ray diffraction
XRF	X-ray fluorescence

Data availability

No primary research results, software or code have been included and no new data were generated or analysed as part of this review.

Acknowledgements

We would like to acknowledge the establishment of the Fermentation Technology Innovation Center of Hebei Province. This work was supported by the Scientific and Technological Project of Shijiazhuang (grant no. 241790927A), the National Natural Science Foundation of China (grant no. 21908042), the Technology Project of Hebei Education Department (grant no. ZD2021049), the S&T Program of Hebei (grant no. 21372902D), and then Open Funding Project of the Fermentation Technology Innovation Center of Hebei Province.

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