ACS Synthetic Biology最新文献

2-amino-deoxyadenosine (dZ) occurs naturally in certain bacteriophage genomes, where it replaces deoxyadenosine, forming three hydrogen bonds with thymidine. This noncanonical deoxyribonucleoside underlies the unique biophysical properties of dZ-DNA. Its corresponding ribonucleoside has been introduced to RNA to form Z-modified RNA, with promising applications in vaccine production and biomedicine. Unlike dZ-DNA, Z-modified RNA has only been synthesized in vitro, which required the addition of chemically synthesized 2-amino-adenosine triphosphate (ZTP) as a precursor. Here, we describe enzyme engineering studies on dZMP-succinate-synthetase (PurZ), a key enzyme in the bacteriophage dZ-DNA biosynthetic pathway that natively catalyzes the conversion of deoxyguanosine monophosphate (dGMP) to dZMP-succinate. Through site-saturation mutagenesis, we generated mutants with altered substrate specificity, capable of catalyzing the conversion of GMP to ZMP-succinate. We further demonstrated that these mutants, in combination with bacterial adenylosuccinate lyase, guanylate kinase and nucleoside diphosphate kinase, efficiently convert GMP to ZTP, marking a critical step in developing a biosynthetic pathway for Z-modified RNA, and enabling enzymatic synthesis of ZTP on a semipreparative scale. Our work provides the basis for further research on the impacts of Z-modified RNA in living organisms, and supports the cost-effective production of Z-modified RNA vaccines and therapeutics.

In synthetic biology, DNA assembly is a routine process where increasing demands for standardization, high-throughput capacity, and error-free execution are driving the development of accessible, automated solutions. Here, we present Slowpoke, a user-friendly and flexible workflow for Golden Gate-based cloning designed for the popular entry-cost, open-source liquid-handling platforms Opentrons OT-2 and Flex. Slowpoke automates the key steps of the DNA assembly process, including cloning, Escherichia coli transformation, plating, and colony PCR, requiring user intervention primarily for colony picking and plate transfers. To further simplify the usage, we developed a free graphical user interface (GUI), available at https://slowpoke.streamlit.app/, which enables rapid protocol generation through simple file uploads. We validated the workflow using two Golden Gate-based toolkits, the MoClo Yeast Toolkit (YTK), and SubtiToolKit (STK). High assembly efficiencies were achieved across platforms for basic transcript unit constructions: 17/17 positive colonies with YTK on OT-2, 11/12 on Flex, and 8/13 with STK on OT-2. High-throughput assemblies were also performed with six parts in Flex using YTK-compatible parts, and 55 out of 57 combinations resulted in correct constructs. These results confirm the robustness and adaptability of the workflow across toolkit complexity and automation platforms. The Slowpoke suite, including code scripts and templates, is freely available at https://github.com/Tom-Ellis-Lab/Slowpoke, offering an accessible and modular solution for automating Golden Gate cloning in synthetic biology laboratories.

Regulating gene expression with precision is essential for cellular engineering and biosensing applications, where rapid, programmable, and sensitive control is desired. Current approaches to regulatory circuit design often rely on control at a single regulatory level, primarily the transcriptional level, thereby limiting the capability of fine-tuning the regulatory dynamics in response to complex stimuli. To address this challenge, we developed four novel RNA-protein hybrid type-1 incoherent feed-forward loop (I1-FFL) circuits in Escherichia coli that integrate transcriptional and translational regulators to achieve multilevel control of gene expression. These hybrid circuits leverage the modularity and rapid dynamics of RNA-based activators alongside the versatile inhibition capabilities of the protein-based repressors to endow tunable pulse dynamics through engineered delays that act as transient repressor decoys. By repurposing synthetic RNA regulators at multiple regulatory levels together with aptamers and RNA-binding proteins, we demonstrate previously unexplored circuits with tunable dynamics. Complementary simulation results highlighted the importance of the engineered delays in achieving tunable pulse dynamics in these circuits. Integrating modeling insights with experimental validation, we demonstrated the flexibility of designing the RNA-protein hybrid I1-FFL circuits, as well as the tunability of their dynamics, highlighting their suitability for applications in environmental monitoring, metabolic engineering, and other engineered biological systems where precise temporal control and adaptable gene regulation are desired.

The research on synthetic methylotrophic bacteria for one-carbon (C1) feedstock assimilation has garnered substantial interest and is regarded as the forefront of biomanufacturing advancements. Nevertheless, the effective utilization of C1 feedstocks faces challenges due to inadequate tolerance toward C1 compounds. This study elucidates that the buildup of formaldehyde causes severe DNA-protein cross-linking (DPC), and thus hampers growth and methanol assimilation in Escherichia coli. To tackle this issue, we exploited a metalloproteinase, SpWss1, from Schizosaccharomyces pombe. By fine-overexpressing SpWss1 in the E. coli genome, we were able to alleviate DPC damage and enhance formaldehyde tolerance. Remarkably, the engineered strain displayed a 10-fold increase in the amount of methanol assimilated (142 mM) compared to that of the control strain lacking SpWss1 (14 mM). Moreover, through iterative substrate feeding of methanol and xylose in shake-flask experiments, the genetically modified strain exhibited improved consumption levels, reaching up to 309 mM (∼10 g/L), making it one of the highest methanol-consuming strains among all E. coli strains without adaptive evolution. Additionally, the modified strain significantly enhanced the sustainable production of valuable products, such as triacetic acid lactone and fatty acids, from methanol. Overall, our findings underscore the significant scientific and biotechnological importance of addressing DPC to optimize C1 assimilation, providing valuable insights for sustainable chemistry, engineering, and industrial biotechnology applications.

Dopamine deficiency resulting from nigrostriatal dopaminergic neuronal damage manifests as extrapyramidal motor symptoms of Parkinson's disease (PD). Oral tablet dosing of levodopa, administered 3-4 times a day, remains the standard of care due to its tolerability and effectiveness; however, it is prone to deleterious side effects, including off-periods and levodopa-induced dyskinesia after long-term use. Herein, using synthetic biology approaches, we developed and systematically evaluated the feasibility of a probiotic-based live-biotherapeutic system to continuously deliver L-DOPA stably, thereby relieving motor symptoms. Our data demonstrate that our engineered plasmid-based L-DOPA-expressing Escherichia coli Nissle 1917 probiotic strain (EcN²_LDOPA-P3) efficiently produced up to 12,000 ng/mL L-DOPA in vitro. In mouse model systems, EcN²_LDOPA-P3 readily colonized for up to 48 h, achieved steady-state plasma L-DOPA concentrations, and increased brain L-DOPA and dopamine levels by 1- to 2-fold. Lastly, EcN²_LDOPA-P3 significantly diminished motor and nonmotor behavioral deficits in a mouse model of PD compared to traditional chemical L-DOPA therapy. These findings support the therapeutic feasibility of a noninvasive, orally administered bioengineered bacterial therapy for the chronic delivery of L-DOPA, which may address limitations associated with current treatment alternatives.

Antibiotic resistance is escalating, highlighting the urgent need for novel antimicrobial strategies. Defensin-like antimicrobial peptides (AMPs) are considered ideal candidates due to their broad-spectrum activity and engineerable potential; however, their limited antimicrobial efficacy and complex chemical synthesis constrain practical applications. In this study, we aimed to enhance the antimicrobial properties of defensin-like AMPs through rational design, directed evolution, and structural fusion strategies. The engineered variant XC1 demonstrated significantly improved antimicrobial activity against a broad range of pathogens, including methicillin-resistant Staphylococcus aureus, while maintaining broad-spectrum efficacy. Comprehensive evaluation of toxicity and stability showed that XC1 exhibited good functional stability in serum, low hemolysis, and low cytotoxicity, indicating excellent therapeutic potential. In addition, high-level secretory expression of defensin-derived AMPs and their engineered variants was achieved using Pichia pastoris GS115, demonstrating strong biosynthetic capability. Together, these results provide a viable strategy for enhancing the antimicrobial activity and scalable biosynthesis of defensin-like AMPs.

Mixotrophy offers a promising strategy for biosynthesis by simultaneously utilizing organic carbon and CO₂; however, mixotrophic microorganisms are rarely isolated outside of photoautotrophic microalgae. In this study, the chemoautotroph Cupriavidus necator H16 was found to preferentially consume fructose when coexisting with CO₂ and H₂, switching to utilization of only CO₂ and H₂ after fructose depletion. Transcriptomic analysis revealed significant differences in genes involved in energy metabolism, electron generation, and the respiratory chain. The molecular mechanism underlying the inability of C. necator H16 to simultaneously utilize carbohydrates and CO₂ was identified as the suppression of hydrogenase expression in the presence of fructose. By inducing regulator hoxA to activate hydrogenase expression, an engineered C. necator strain capable of mixotrophic growth was developed. This engineered strain can simultaneously utilize fructose, CO₂, and H₂, maintain optimal growth, and approach carbon-neutral cultivation. This work provides insights for the mixotrophic cultivation of C. necator and serves as a reference for developing mixotrophic microorganisms in future studies.

β-Elemene, a sesquiterpene with anticancer activity, faces limited microbial production due to low yields and inefficient enzyme coordination. This study established a modular covalent enzyme cascade in Escherichia coli for high-efficiency β-elemene biosynthesis. Systematic screening identified the SnoopTag/SnoopCatcher-mediated covalent assembly of farnesyl diphosphate synthase and germacrene A synthase as the optimal configuration (strain FS07). Subsequent enzyme stoichiometry modulation or linker engineering did not surpass FS07's performance, indicating a near-optimal design. Fermentation optimization elevated the β-elemene titer to 7.21 g/L in FS07. Fed-batch fermentation in a 1.3 L bioreactor subsequently increased the final titer to 31.21 g/L, representing a 4-fold improvement over scaffold-free controls and achieving the highest reported yield in E. coli to date. This work provided a robust enzymatic scaffolding strategy for high-level terpenoid production.

Genetic manipulation of core gut probiotics remains challenging due to endogenous cellular barriers and a scarcity of efficient molecular tools, limiting progress in live biotherapeutic development. Here, we characterized the native type II-C CRISPR-Cas system in Bifidobacterium longum subsp. longum GNB (B. longum GNB). Through integrated bioinformatic analysis and high-throughput protospacer adjacent motif (PAM) screening, we identified a novel 5'-NNRMAT-3' (where R = A/G, M = A/C) motif recognized by its compact Cas9 nuclease (BLCas9). The stringent PAM dependency of BLCas9 was unequivocally confirmed by in vitro cleavage assays. Leveraging this endogenous mechanism, we developed a dual-plasmid editing platform for robust and multiplex genome engineering in the probiotic strain Escherichia coli Nissle 1917 (EcN). Application of this system notably enhanced extracellular γ-aminobutyric acid (GABA) production in EcN through targeted metabolic engineering. Our work provides the first molecular dissection of a type II-C system in Bifidobacterium longum and establishes a generalizable framework for the discovery and application of compact programmable nucleases, suggesting a viable strategy for modulating host physiology via the gut-brain axis.

Inducible translocation to subcellular compartments is a common strategy for protein switches that control a variety of cell behaviors. However, existing switches achieve translocation through induced dimerization, requiring constitutive anchoring of one component into the target compartment and optimization of relative expression levels between the two components. We present a simpler, single-component strategy called Avidity-assisted targeting (Aviatar). Aviatar achieves translocation with only a single protein by converting low-affinity monomers into high-avidity assemblies through inducible clustering. We demonstrated the Aviatar concept and its generality using optogenetic clustering to drive translocation to the plasma membrane, endosomes, golgi, endoplasmic reticulum, and microtubules using binding domains for lipids or endogenous proteins that were specific to those compartments. Aviatar recruitment regulated actin polymerization at the cell periphery and revealed compartment-specific signaling of receptor tyrosine kinase fusions associated with cancer. Finally, GFP-targeting Aviatar probes allowed inducible localization to any GFP-tagged target, including endogenously tagged stress granule proteins. Aviatar is a straightforward platform that can be rapidly adapted to a broad array of targets without the need for their prior modification or disruption.