New biotechnology最新文献

Polyhydroxyalkanoates (PHAs) are biodegradable polymers produced by various microorganisms as intracellular carbon and energy reserves. Their potential to replace petroleum-based plastics has made them central to sustainable materials research. However, their large-scale commercialization is hindered by high production costs, prompting efforts to improve yield and identify low-cost, non-food carbon sources. This review examines PHA biosynthesis optimization via advances in microbial strain-substrate selection ensuring economic feasibility. Focus is placed on three key bacterial genera, Cupriavidus, Pseudomonas, and Escherichia coli analyzing their metabolic flexibility, suitable substrate range and PHA content (g product/ g substrate). Escherichia coli species and Cupriavidus necator demonstrate high polymer contents from largely simple sugars and fatty-acid substrates respectively, while Pseudomonas species offer a broad substrate adaptability, particularly waste streams. The environmental impact assessment of some Cupriavidus necator and Pseudomonas strains highlight low carbon footprint from waste lipids compared to PHA production from glucose or bottle-grade PET, however these results are contingent to specific scenarios without a reproducible and quantifiable GWP values across these genera. Integrating microbial engineering of agro-industrial and organic wastes in adherence to regulatory frameworks around waste valorization and optimization, genus-specific LCA bioprocessing approach offers a path toward economically viable and environmentally sustainable PHA production.

Microbial communities play a key role in biogeochemical transformations in a wide range of ecosystems, but they also hold significant potential to enhance the bioproduction of desired chemicals. Although designing synthetic microbial consortia has generated a lot of interest, a more in-depth understanding of the interactions between strains is required, particularly when strains are engineered to cross-feed, but are not isolated from related environments. Challenges include enhancing stability, productivity and controllability. Here, we used a synthetic microbial co-culture consisting of engineered strains of the photosynthetic cyanobacterium Synechococcus elongatus PCC 7942 cscB/SPS and nitrogen-fixing bacterium Azotobacter vinelandii AV3. Each relies on the other for conversion of atmospheric carbon (CO₂) and nitrogen (N₂) into organic forms, i.e. sucrose and ammonia, respectively, resources which can be shared. As both strains have such contrasting growth dynamics in co-culture compared to monoculture, we applied a label-free quantitative proteomics approach to characterise metabolism in both strains. The proteomes of both shifted when in co-culture to reflect adaptive restructuring of carbon and nitrogen metabolism, although A. vinelandii appeared to transition to a more stressed state, inducing proteins linked to polymer biosynthesis. An analysis of the co-culture over 16 days led to phenotypic changes, including cell structure alterations in A. vinelandii AV3 over time, with the proteome suggesting cell envelope remodelling and potentially encystment. These findings suggest that physiological control of parameters, such as oxygen and nutrient availability, may enable cultivation of more stable co-cultures.

The unicellular red alga Cyanidioschyzon merolae is a valuable model organism for studying pre-mRNA splicing, stress adaptation, and biotechnological applications. However, the limited availability of selectable markers has constrained its potential in genetic engineering. In this study, we evaluated the sul1 gene, which encodes a sulfadiazine-resistant variant of dihydropteroate synthase, as a new selectable marker (SUL) for C. merolae transformation. SUL has previously been used for this purpose in plants and green algae. We analyzed the sensitivity of C. merolae to sulfadiazine and determined the concentration that effectively inhibited cell growth. To test the effectiveness of SUL as a selectable marker, we designed a transformation construct containing SUL directed to the algal mitochondria through a native targeting peptide, along with mVenus to visualize transformation. We integrated the construct into a neutral genomic locus via homologous recombination. Fluorescence microscopy confirmed stable mVenus expression, and sulfadiazine selection successfully enriched transformed cells. As a demonstration of the utility of this marker, we rescued the large-cell phenotype of a cell division cycle-like kinase 2 (CmClk2) mutant by replacing the CAT-marked kinase domain deletion with the SUL-marked native kinase domain, thereby restoring CmClk2 function and recycling the CAT marker. The deletion phenotype provides evidence for a conserved cell-cycle regulatory role for CmClk2 in C. merolae. Beyond establishing SUL as an effective selectable marker, this highlights how SUL facilitates functional genetic studies of essential cellular regulators.

Protein biopharmaceuticals play a key role in providing effective, targeted, and personalized therapies for diverse diseases, while also preventing and mitigating a broad range of infections. N-glycosylation is a key post-translational modification influencing the biological activity of many protein-based therapeutics, yet structure-function relationships of N-glycans remain poorly understood due to challenges in producing homogeneous glycoforms. Current go-to production hosts, mammalian and yeast cells, often yield heterogeneous glycan profiles and require extensive genetic manipulation. Alternative production hosts such as the plant Nicotiana benthamiana, provide more homogeneous glycosylation and flexibility through transient expression, but are limited in the generation of certain glycoforms. In vitro glycoengineering can overcome these limitations but is time consuming and requires expensive resources. In this study, we show that by combining in planta and in vitro glycoengineering strategies, we can quickly produce a wide range of homogeneous glycoforms of pharmaceutical proteins with high mannose, paucimannose, hybrid and complex N-glycan structures. Using N. benthamiana as a transient expression host, we produced two pharmaceutical glycoproteins - the monoclonal antibody rituximab and the helminth vaccine candidate OoASP-1 - and modified them in vitro using Escherichia coli produced glycoenzymes. The combination of these two glycoengineering systems minimizes the amount of time and resources required, while maintaining high glycan homogeneity. This scalable, flexible, and cost-effective platform opens the door to glycan structure-function relationship studies and can support rational design of next-generation biopharmaceuticals.

Prenylated plant phenolics are a large group of secondary metabolites known for their bioactivity that is beneficial for plant and human alike, for example as antimicrobial agents against pathogenic microbes. However, the limited availability of prenylated phenolics, especially O-prenylated phenolics, and the complexity of the plant metabolite mixtures hinder bioactivity studies and further application. To explore approaches for more efficient production of prenylated phenolics, we produced and characterized a novel prenyltransferase (ScPT_MY) from the edible mushroom Sparassis crispa. ScPT_MY belongs to the dimethylallyl tryptophan synthase (DMATS) family and was found to primarily catalyze O-prenylation of structurally diverse phenolics. The best substrates included l-tryptophan, l-tyrosine, stilbenes, and isoflavonoids. The ScPT_MY reactions predominantly yielded a single mono-O-prenylated product with an exception of the isoflavonoids, for which more products were obtained, including di-prenylated ones. Notably, we demonstrated the potential of this O-prenylating DMATS to enhance bioactivity by showing that enzymatic O4'-prenylation conferred antimicrobial activity to resveratrol, a compound with otherwise very poor antimicrobial activity. Finally, our phylogenetic analysis suggested the possibility of combining evolutionary relationships with structural insights to predict substrate scope and regioselectivity, while also revealing seven largely unexplored fungal DMATS clades that may harbor novel functions for biotechnological applications.

The BacMam platform is a scalable and efficient gene delivery system for mammalian cells, enabling the production of recombinant proteins or highly complex structures like bionanoparticles. This study represents the first comprehensive investigation to evaluate the performance of the BacMam platform in a common producer cell line HEK293-6E, on stable transgene expression mediated by the REMBAC (rapid efficient manifold baculovirus transduction) cassette. In this study, six transient high expression cassettes were adopted for stable gene expression and evaluated in HEK293-6E and Vero cells. The constructs included either the wild-type viral CMV promoter, a methylation-resistant mutant of the CMV promoter, or the mammalian EF-1α promoter. To further enhance expression, endogenous mammalian introns or viral long terminal repeats (LTRs) were included, along with β-globin insulators placed at both the 5' and 3' ends of the cassette. Additionally, the constructs were equipped with two antibiotic resistance genes, Hygromycin and G418, and a full-length Woodchuck Hepatitis Virus Posttranscriptional Regulatory Element (WPRE), known to improve mRNA stability and enhance protein expression by modulating RNA structure [1].The results indicated that transduction of HEK293-6E cells achieved higher efficiency, resulted in more clones with a stably integrated expression cassette, and stronger protein expression than transduction of Vero cells. The inclusion of β-globin insulators significantly enhanced gene expression in HEK293-6E cells, while their effect in Vero cells was less pronounced. These findings highlight the importance of cell line selection and genetic design in optimizing recombinant protein production.

Euglena gracilis, a Generally Recognized as Safe (GRAS) microorganism, is of growing biotechnological interest due to its ability to accumulate diverse cellular metabolites under heterotrophic conditions. This study evaluated canola meal protein hydrolysate (CMPH) as a protein-derived organic nitrogen source to partially substitute conventional complex supplements (yeast extract and tryptone) in heterotrophic E. gracilis cultivation. CMPH was produced by enzymatic hydrolysis of canola meal using trypsin (3000 mL^-1, pH 8.0, 37 °C, 24 h). The hydrolysate contained 74.28 ± 5.92 % protein (micro-Kjeldahl), and X-ray fluorescence analysis indicated phosphorus (6.8 %), calcium (16.7 %), potassium (1.7 %), and magnesium (1.2 %) as major mineral components. A Box-Behnken design coupled with response surface methodology (BBD-RSM) was applied to optimize growth conditions in CMPH-supplemented medium. Cultures inoculated at 1.0 × 10⁵ cells mL^-1 were incubated at 25 °C in the dark for 168 h. RSM identified pH 6.8, 1 g L^-1 CMPH, and 120 h as statistically optimal conditions within the tested design space, yielding 9.8 × 10⁵ cells mL^-1 in the absence of added glucose, with a specific growth rate of 0.69 ± 0.23 d^-1. Higher cell densities were obtained at increased CMPH concentrations and upon glucose supplementation, reaching levels comparable to standard Hutner's medium. These results indicate that the RSM-derived optimum reflects efficient nitrogen utilization rather than maximal biomass accumulation. Overall, CMPH demonstrates potential as a functional organic nitrogen supplement for heterotrophic E. gracilis cultivation, supporting the valorization of canola meal as a protein-rich agricultural by-product.