ACS Synthetic Biology最新文献

Triterpenoids possess a range of biological activities and are extensively utilized in the pharmaceutical, food, cosmetic, and chemical industries. Traditionally, they are acquired through chemical synthesis and plant extraction. However, these methods have drawbacks, including high energy consumption, environmental pollution, and being time-consuming. Recently, the de novo synthesis of triterpenoids in microbial cell factories has been achieved. This represents a promising and environmentally friendly alternative to traditional supply methods. Saccharomyces cerevisiae, known for its robustness, safety, and ample precursor supply, stands out as an ideal candidate for triterpenoid biosynthesis. However, challenges persist in industrial production and economic feasibility of triterpenoid biosynthesis. Consequently, metabolic engineering approaches have been applied to improve the triterpenoid yield, leading to substantial progress. This review explores triterpenoids biosynthesis mechanisms in S. cerevisiae and strategies for efficient production. Finally, the review also discusses current challenges and proposes potential solutions, offering insights for future engineering.

Industrial biotechnology uses Design–Build–Test–Learn (DBTL) cycles to accelerate the development of microbial cell factories, required for the transition to a biobased economy. To use them effectively, appropriate connections between the phases of the cycle are crucial. Using p-coumaric acid (pCA) production in Saccharomyces cerevisiae as a case study, we propose the use of one-pot library generation, random screening, targeted sequencing, and machine learning (ML) as links during DBTL cycles. We showed that the robustness and flexibility of the ML models strongly enable pathway optimization and propose feature importance and Shapley additive explanation values as a guide to expand the design space of original libraries. This approach allowed a 68% increased production of pCA within two DBTL cycles, leading to a 0.52 g/L titer and a 0.03 g/g yield on glucose.

The presence of a hydration layer in humid and underwater environments challenges adhesive–substrate interactions and prevents effective bonding, which has become a significant obstacle to the development of adhesives in the industrial and biomedical fields. In this study, ultrahigh-molecular-weight (UHMW) silk-elastin-like proteins (SELP) with 3,4-dihydroxyphenylalanine (DOPA) converted from tyrosine residues by tyrosinase exhibited excellent adhesive properties on different interfaces, such as glass, aluminum, wood, polypropylene sheets, and pigskin, under both dry and wet conditions. Additionally, by incorporating trace amounts of cross-linking agents like Fe³⁺, NaIO₄, and tris(hydroxymethyl) phosphine (THP), the mussel-inspired adhesives maintained a stable and excellent adhesion, broadening the conditions of application. Notably, the UHMW SELP adhesive exhibited remarkable underwater adhesion properties with a shear strength of 0.83 ± 0.17 MPa on glass. It also demonstrated good adhesion to biological tissues including the kidney, liver, heart, and lungs. In vitro cytocompatibility testing using L929 cells showed minimal toxicity, highlighting its potential application in the biomedical field. The sustainable, cytocompatible, cost-effective, and highly efficient adhesive provides valuable insights for the design and development of a new protein-based underwater adhesive for medical application.

Directed evolution is often limited by the throughput of accurate screening methods. Here we demonstrate the feasibility of utilizing a singular transcription factor (TF)-system that can be refactored in two ways (both as an activator and repressor). Specifically, we showcase the use of previously evolved 5-halo- or 6-halo-tryptophan-specific TF biosensors suitable for the detection of a halogenated tryptophan molecule in vivo. We subsequently validate the biosensor’s utility for two halogenase-specific halo-tryptophan accumulation screens. First, we isolated 5-tryptophan-halogenase, XsHal, from a mixed pool of halogenases with 100% efficiency. Thereafter, we generated a targeted library of the catalytic residue of 6-tryptophan halogenase, Th-Hal, and isolated functioning halogenases with 100% efficiency. Lastly, we refactor the TF circuit to respond to the depletion of halogenated tryptophan and prototype a high-throughput biosensor-directed evolution scheme to screen for downstream enzyme variants capable of promiscuously converting halogenated tryptophan. Altogether, this work takes a significant step toward the rapid and higher throughput screening of halogenases and halo-tryptophan converting enzymes to further reinforce efforts to enable high-level bioproduction of halogenated chemicals.

As the use of engineered cell therapies expands from pioneering efforts in cancer immunotherapy to other applications, an attractive but less explored approach is the use of engineered red blood cells (RBCs). Compared to other cells, RBCs have a very long circulation time and reside in the blood compartment, so they could be ideally suited for applications as sentinel cells that enable in situ sensing and diagnostics. However, we largely lack tools for converting RBCs into biosensors. A unique challenge is that RBCs remodel their membranes during maturation, shedding many membrane components, suggesting that an RBC-specific approach may be needed. Toward addressing this need, here we develop a biosensing architecture built on RBC membrane proteins that are retained through erythropoiesis. This biosensor employs a mechanism in which extracellular ligand binding is transduced into intracellular reconstitution of a split output protein (including either a fluorophore or an enzyme). By comparatively evaluating a range of biosensor architectures, linker types, scaffold choices, and output signals, we identify biosensor designs and design features that confer substantial ligand-induced signal in vitro. Finally, we demonstrate that erythroid precursor cells engineered with our RBC-protein biosensors function in vivo. This study establishes a foundation for developing RBC-based biosensors that could ultimately address unmet needs including noninvasive monitoring of physiological signals for a range of diagnostic applications.

In this study, we proposed a biological approach to efficiently produce pseudouridine (Ψ) from glucose and uracil in vivo using engineered Escherichia coli. By screening host strains and core enzymes, E. coli MG1655 overexpressing Ψ monophosphate (ΨMP) glycosidase and ΨMP phosphatase was obtained, which displayed the highest Ψ concentration. Then, optimization of the RBS sequences, enhancement of ribose 5-phosphate supply in the cells, and overexpression of the membrane transport protein UraA were investigated. Finally, fed-batch fermentation of Ψ in a 5 L fermentor can reach 27.5 g/L with a yield of 89.2 mol % toward uracil and 25.6 mol % toward glucose within 48 h, both of which are the highest to date. In addition, the Ψ product with a high purity of 99.8% can be purified from the fermentation broth after crystallization. This work provides an efficient and environmentally friendly protocol for allowing for the possibility of Ψ bioproduction on an industrial scale.

The de novo construction of a living organism is a compelling vision. Despite the astonishing technologies developed to modify living cells, building a functioning cell “from scratch” has yet to be accomplished. The pursuit of this goal alone has─and will─yield scientific insights affecting fields as diverse as cell biology, biotechnology, medicine, and astrobiology. Multiple approaches have aimed to create biochemical systems manifesting common characteristics of life, such as compartmentalization, metabolism, and replication and the derived features, evolution, responsiveness to stimuli, and directed movement. Significant achievements in synthesizing each of these criteria have been made, individually and in limited combinations. Here, we review these efforts, distinguish different approaches, and highlight bottlenecks in the current research. We look ahead at what work remains to be accomplished and propose a “roadmap” with key milestones to achieve the vision of building cells from molecular parts.

Ruminant livestock produce around 24% of global anthropogenic methane emissions. Methanogenesis in the animal rumen is significantly inhibited by bromoform, which is abundant in seaweeds of the genus Asparagopsis. This has prompted the development of livestock feed additives based on Asparagopsis to mitigate methane emissions, although this approach alone is unlikely to satisfy global demand. Here we engineer a non-native biosynthesis pathway to produce bromoform in vivo with yeast as an alternative biological source that may enable sustainable, scalable production of bromoform by fermentation. β-dicarbonyl compounds with low pKa values were identified as essential substrates for bromoform production and enabled bromoform synthesis in engineered Saccharomyces cerevisiae expressing a vanadate-dependent haloperoxidase gene. In addition to providing a potential route to the sustainable biological production of bromoform at scale, this work advances the development of novel microbial biosynthetic pathways for halogenation.

Conventional biological experiments often focus on in vitro assays because of the inherent limitations when handling multiple variables in vivo, including labor-intensive and time-consuming procedures. Often only a subset of samples demonstrating significant efficacy in the in vitro assays can be evaluated in vivo. Nonetheless, because of the low correlation between the in vitro and in vivo tests, evaluation of the variables under examination in vivo and not solely in vitro is critical. An emerging approach to achieve high-throughput in vivo tests involves using a barcode system consisting of various nucleotide combinations. Unique barcodes for each variant enable the simultaneous testing of multiple entities, eliminating the need for separate individual tests. Subsequently, to identify crucial parameters, samples were collected and analyzed using barcode sequencing. This review explores the development of barcode design and its applications, including the evaluation of nucleic acid delivery systems and the optimization of gene expression in vivo.

The important roles that protein glycosylation plays in modulating the activities and efficacies of protein therapeutics have motivated the development of synthetic glycosylation systems in living bacteria and in vitro. A key challenge is the lack of glycosyltransferases that can efficiently and site-specifically glycosylate desired target proteins without the need to alter primary amino acid sequences at the acceptor site. Here, we report an efficient and systematic method to screen a library of glycosyltransferases capable of modifying comprehensive sets of acceptor peptide sequences in parallel. This approach is enabled by cell-free protein synthesis and mass spectrometry of self-assembled monolayers and is used to engineer a recently discovered prokaryotic N-glycosyltransferase (NGT). We screened 26 pools of site-saturated NGT libraries to identify relevant residues that determine polypeptide specificity and then characterized 122 NGT mutants, using 1052 unique peptides and 52,894 unique reaction conditions. We define a panel of 14 NGTs that can modify 93% of all sequences within the canonical X_–1-N-X₊₁-S/T eukaryotic glycosylation sequences as well as another panel for many noncanonical sequences (with 10 of 17 non-S/T amino acids at the X₊₂ position). We then successfully applied our panel of NGTs to increase the efficiency of glycosylation for three protein therapeutics. Our work promises to significantly expand the substrates amenable to in vitro and bacterial glycoengineering.