Cephalosporin C acylases (CCAs) catalyze the hydrolysis of cephalosporin C to 7-aminocephalosporanic acid, a key intermediate for semisynthetic cephalosporin antibiotics. The functional secretion of heterologous CCAs in Escherichia coli is often constrained by signal peptide efficiency. To enhance the production of the engineered CCA mutant A14 from Bosea sp. OK403, we performed systematic signal peptide screening and identified the native SP AsPGA as most effective. Following codon optimization to generate SP AsPGA∗ , targeted mutagenesis of its N-, H-, and C-regions produced the superior H9 mutant (C16A). This variant increased extracellular A14 expression and activity by 6.26-fold and 2.0-fold, respectively. Molecular dynamics simulations indicated the C16A substitution stabilizes the signal peptide conformation and facilitates SecA translocon interaction. This work establishes that systematic mining and engineering of signal peptides, particularly through mutations that enhance structural stability, is a powerful strategy for optimizing the secretory production of complex enzymes in recombinant systems. Strategy to enhance CPC acylase secretion in Escherichia coli via signal peptide mining, screening, and optimization, followed by structural analysis and molecular dynamics simulations of the resulting mutants.

Bacillus subtilis is widely employed for lignocellulose degradation. However, wild-type strains typically exhibit low and incomplete cellulase activities. This study aimed to engineer recombinant B. subtilis strains harboring complete, high-efficiency cellulase systems using CRISPR/Cas9-mediated genome editing. After optimization of signal peptides, transcriptional terminators, and chromosomal integration sites, two endoglucanase expression cassettes were integrated into B. subtilis 168, yielding a strain with a 16.29-fold increase in endoglucanase activity relative to the parental strain. In parallel, a bifunctional cellulase was successfully expressed and optimized, achieving exoglucanase (549.77 U/mL) and β-glucosidase (349.26 U/mL) activities. Moreover, the strain BSKI3Cel, containing three optimized expression cassettes in its genome, exhibited high endoglucanase (129.59 U/mL), exoglucanase (596.75 U/mL), and β-glucosidase (447.42 U/mL) activities. Subsequent transformation of BSKI3Cel with plasmid pJOE2006Bf, carrying genes encoding a composite cellulase system, yielded BSK3P2C, which achieved peak cellulase activities of 533.16, 2959.83, and 2829.61 U/mL on day 8. As a proof of concept, fermentation of wheat straw using BSK3P2C was found to significantly reduce hemicellulose (16.70 %), neutral detergent fiber (7.46 %) and acid detergent fiber (9.93 %) contents. Microscopic analyses confirmed extensive lignocellulose degradation. Overall, this study establishes a high-performance B. subtilis platform with complete cellulase systems for efficient cellulosic biomass conversion.

Antibiotic biosynthesis in actinomycetes is controlled by various transcription factors, with TetR family regulators (TFRs) serving as important modulators of this process. We previously discovered a TFR, SACE_1906, which hinders erythromycin yield in Saccharopolyspora erythraea. However, the precise mechanism by which SACE_1906 regulates erythromycin biosynthesis remains elusive. Herein, we confirmed that SACE_1906 directly inhibits the transcription of its own and neighboring alcohol dehydrogenase gene SACE_1905, as well as all genes within the erythromycin biosynthetic cluster. It was found that two identical conserved sites (5′-CACCGGTCGGTATA-3′) in the intergenic spacer between SACE_1905 and SACE_1906 (SACE_1905-1906-int) are necessary for SACE_1906 binding. Deletion of SACE_1906 resulted in decreased propionyl-CoA content but increased the intracellular supply of methylmalonyl-CoA. We further proved that SACE_1906 restricts methylmalonyl-CoA production from propionyl-CoA to influence intracellular levels of these two erythromycin biosynthetic precursors through the direct repression of several propionyl-CoA carboxylase genes. Moreover, acetaldehyde has been characterized as an effector molecule of SACE_1906 that weakens its binding affinity for SACE_1905-1906-int. This study reveals a novel mechanism by which SACE_1906 orchestrates erythromycin biosynthesis in response to acetaldehyde, expanding our understanding of small-molecule-mediated regulation for antibiotic biosynthesis in actinomycetes.

Prodigiosin, a bioactive tripyrrole pigment, exhibits a broad spectrum of biological activities-including antimicrobial, anticancer, and antimalarial properties-thereby holding significant promise for use in pharmaceutical applications and industrial biotechnology. In this study, three isolates of Serratia marcescens were recovered from the cabbage rhizosphere. Genomic analysis revealed a highly conserved prodigiosin biosynthetic gene cluster embedded within the chromosomes of all three isolates. Though prodigiosin production was detected in these three S. marcescens isolates, the relatively low yield severely limits the feasibility of its large-scale production. To address this issue, we employed a stepwise strategy involving heterologous expression, promoter engineering, genome-wide transposon mutagenesis, and optimization of fermentation media with the aim to achieve high-level prodigiosin production. The introduction of an engineered prodigiosin gene cluster into a tailored Pseudomonas putida KT2440 chassis strain yielded a maximum prodigiosin titer of 665 mg/L in shake-flask cultures, significantly outperforming production levels of the native S. marcescens isolates. When cultured in a small-scale stirred-tank bioreactor, the engineered strain further elevated the prodigiosin yield to 1161 mg/L. Our study presents a robust platform for prodigiosin overproduction, which can be adapted to improve the titers of other prodiginine family compounds.

Escherichia coli (E. coli) has long served as a versatile workhorse for recombinant protein production. As synthetic biology expands the demand for coordinated expression of multiple genes, co-expression systems in E. coli have evolved from basic dual-gene constructs to programmable, polygenic expression platforms. This review critically examines the major strategies enabling multigene co-expression in E. coli, including internal ribosome entry sites (IRES), 2A self-cleaving peptides, dual-promoter cassettes, multicistronic operons, and multi-plasmid configurations. We highlight the mechanistic principles, design trade-offs, and regulatory bottlenecks associated with each approach, such as translational imbalance, inclusion body formation, and plasmid compatibility. Real-world applications in metabolic engineering, complex protein assembly, and biomanufacturing are analyzed to demonstrate the functional advantages of these systems. Finally, we explore emerging programmable toolkits that integrate modular architecture, expression modeling, and AI-assisted design, paving the way for next-generation synthetic expression control in microbial chassis. This review offers a comprehensive and strategic roadmap for researchers engineering multi-gene systems in E. coli and beyond.

Precise regulation of protein abundance is essential for understanding dynamic cellular processes and for advancing therapeutic development. However, existing approaches lack the spatiotemporal resolution required to these cellular processes. Recent advances in optogenetics have enabled the design of optogenetic targeted protein degradation systems (Opto-TPD) allowing reversible and non-invasive control of protein stability with high spatiotemporal precision. In this review, we systematically summarize the design principles of Opto-TPD tools, including those based on light-oxygen-voltage (LOV)-domain conformational systems, light-inducible dimerization systems, and light-controlled degradation tool expression systems. We further highlight their applications in probing protein function, modulating signaling pathways, and therapeutic translations. By comparing the mechanistic features, performance, and limitations of each platform, we aim to provide a comprehensive resource for guiding future tool optimization. Altogether, these Opto-TPD tools represent a powerful and versatile complement to existing protein manipulation technologies, expanding the toolbox for precise control of protein homeostasis in living systems.