ACS Synthetic Biology

Combinatorial DNA design and assembly is an efficient and pragmatic way to obtain high-performing metabolic pathway designs quickly. However, implementation may require organism-specific technical barriers to be overcome. Firstly, suitable expression control parts such as promoters and ribosome-binding sites (RBSs) which provide a suitable range of expression levels need to be identified or developed. Secondly, these need to be assembled into pathway-encoding combinatorial libraries of sufficient size, quality and diversity. For organisms with transformation frequencies too low to allow direct transformation of library assembly reactions, such as many Clostridium spp., assembly and amplification is typically carried out using Escherichia coli. However, if constructs are deleterious (or burdensome) to E. coli, which is often the case when using Clostridium genetic parts, poor libraries may be obtained. Here we develop a new approach called integration-coupled activation of promoterless sequences (ICAPS) to overcome this barrier and therefore enable combinatorial assembly in Clostridium. Libraries were designed and assembled as promoterless synthetic operons, preventing expression during DNA assembly, and expression was only activated later, when constructs were integrated into the host genome downstream of a promoter. Variation of expression levels was achieved using a range of context-resistant RBS sequences. This approach was used to produce a Clostridium acetobutylicum library with combinatorial expression variants of an introduced hexanol pathway. This proof of concept provides a generally-applicable approach to implement combinatorial metabolic pathway-encoding libraries in Clostridium spp., circumventing the excessive strength of Clostridium expression control parts in E. coli, and is applicable to other organisms.

Cell-free expression systems offer a method for rapid prototyping of DNA circuits and functional protein synthesis. While crude extracts remain a black box with many components carrying out unknown reactions, PURE contains only the required transcription and translation components for protein production. All proteins and small molecules are at known concentrations, enabling detailed modeling for reliable computational predictions. However, there is little to no experimental data supporting the expression of target proteins for PURE-based models. In this work, we generalized the PURE detailed translation model for proteins with arbitrary amino acid compositions and lengths. We then built a chemical reaction network for transcription in PURE, validating the transcription models using DNA expression for the malachite-green aptamer (MGapt) to measure mRNA production. Lastly, we coupled the transcription and the generalized translation models to create a PURE protein synthesis model built purely of mass-action reactions. We used the combined model to capture the kinetics of MGapt and deGFP expressed from plasmids at varying concentrations.

There is growing interest in engineering animal and plant microbiomes to deliver double-stranded RNA (dsRNA) for RNA interference (RNAi) applications. We developed a genetically encoded biosensor that uses bimolecular fluorescence complementation to monitor dsRNA levels within bacterial cells to accelerate the symbiont-mediated RNAi design-build-test cycle. We validated performance of the sensor in Escherichia coli and demonstrated enhanced dsRNA accumulation in engineered strains of the aphid symbiont Serratia symbiotica.

Operons are gene clusters controlled by a single promoter that enable coordinated translation from a single messenger RNA. Here we describe an expansion of the CyanoGate MoClo toolkit to assemble synthetic operons. The versatile CyanOperon system includes two Level 0 acceptor vectors for building interchangeable promoter-ribosome bind site (RBS) combinations and 15 Level 1 acceptor vectors for the hierarchical assembly and expression of up to six genes within a single operon. The system also allows for operon assembly into a self-replicating vector or for chromosomal integration by homologous recombination. To showcase CyanOperon, we assembled the violacein biosynthesis pathway as an operon and demonstrated violacein production in Escherichia coli. We then constructed a 20-part RBS library to examine how spacer length between the Shine-Dalgarno sequence and start codon affects translation in E. coli and the model cyanobacterium Synechocystis sp. PCC 6803. Lastly, we compared the expression of up to three operonic fluorescent markers following chromosomal integration or from a self-replicating vector in E. coli and Synechocystis sp. PCC 6803. The CyanOperon system is publicly available and can be readily integrated with other MoClo systems to accelerate the development of standardized operon assemblies. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=81 SRC="FIGDIR/small/707249v1_ufig1.gif" ALT="Figure 1"> View larger version (11K): org.highwire.dtl.DTLVardef@29557borg.highwire.dtl.DTLVardef@1ab1d18org.highwire.dtl.DTLVardef@1033249org.highwire.dtl.DTLVardef@da6db5_HPS_FORMAT_FIGEXP M_FIG C_FIG

Despite transformative advances in DNA synthesis, sequencing, and automation that have accelerated recombinant DNA workflows, molecular cloning hosts have scarcely evolved past the Escherichia coli strains adopted out of convenience in the 1970s. We present NBx CyClone - an engineered strain of Vibrio natriegens - as a next-generation host for molecular cloning. This non-pathogenic marine bacterium combines broad plasmid and genetic tool compatibility, a versatile metabolism, and the fastest known doubling time of any free-living organism. By shortening growth-dependent steps, this host offers a practical route to faster, more efficient recombinant DNA workflows across research and industry.

In vitro reconstitution of protein systems - e.g., metabolic pathways, genetic circuits or biosensors - often requires optimization to enhance their activity. Combinatorial DNA libraries that simultaneously target multiple genes allow for a holistic optimization strategy by studying the interplay between the systems components, which may reveal DNA variants that would be hidden when testing each element in isolation. Here, we screen large populations of synthetic vesicles that express combinatorial DNA variants of a DNA self-replicator or a phospholipid synthesis pathway. We simultaneously vary the strengths of multiple RBSs or synonymously mutate the first codons of multiple genes to explore the effects of the protein translation rates directly on the functionality of the two core synthetic cell modules. We isolated high performers through DNA self-selection or functional screening by fluorescence-activated cell sorting. Long-read sequencing of the fittest variants informed on the optimal RBS strengths and base substitutions in the first codons and indicated which genes were most impactful in regulating the functionality of the protein systems. Single-mutation data were used to predict the fitness of combinatorial variants, which was compared with the experimental fitness observed. The theoretical fitness of combinatorial variants was extremely predictive for the two-gene library of the DNA replicator but less for the larger pathway library. Altogether, our approach exemplifies how combinatorial testing can be expanded from single proteins to multiprotein systems, which can in the future be extended to the evolutionary engineering of even larger genetic and metabolic networks, and eventually an entire artificial cell. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=68 SRC="FIGDIR/small/707944v1_ufig1.gif" ALT="Figure 1"> View larger version (28K): org.highwire.dtl.DTLVardef@1ac1d47org.highwire.dtl.DTLVardef@b62339org.highwire.dtl.DTLVardef@1c2a7ddorg.highwire.dtl.DTLVardef@9abedf_HPS_FORMAT_FIGEXP M_FIG C_FIG

Reporter cell assays, such as those used to detect estrogenic chemicals, can detect target chemicals at low concentrations and can be used to analyze chemical mixtures without a priori knowledge of the mixture components. However, the outputs of these assays are affected by biological variability, which complicates their interpretation. Here, we describe and demonstrate a workflow that is useful for determining potential sources of biological variability and optimizing the performance of cell-based assays. The workflow involves developing an appropriate mathematical model for a transcriptional activation assay, calibrating it with experimental data, and conducting sensitivity analysis to characterize individual components of the genetic circuit based on their effect on the reporter signal output. This workflow was tested using an estrogen receptor transcriptional activation assay. For this circuit, our analysis predicts that controlling estrogen response element number, promoter strength, and reporter signal degradation rates minimizes reporter output variability. We show that careful model development, calibration, and analysis can offer biologically relevant insights to minimize the variability of cell-based assays and improve genetic circuits for increased sensitivity and dynamic range.

BackgroundMicrocins are small antibacterial proteins secreted by gram-negative bacteria. The activities of new microcins discovered using bioinformatic searches need to be validated and characterized to understand how they mediate competition in microbiomes and to evaluate their potential as new therapeutics for combating antibiotic resistance. Engineered plasmids containing the type I secretion system associated with Escherichia coli Microcin V (MccV) can secrete heterologous proteins, including other class II microcins, and this system functions in other bacterial hosts. However, existing microcin secretion constructs are not designed for easily swapping components -- such as origins of replication, resistance genes, promoters, and signal peptides -- that may need to be changed for compatibility with other chassis. ResultsWe refactored the E. coli MccV type I secretion system into genetic parts compatible with a modular Golden Gate assembly scheme and used these parts to construct two-plasmid microcin secretion systems. In our design, one plasmid encodes the type I secretion system proteins, and the other encodes a signal peptide fused to the cargo protein that will be secreted. We tested two versions of a system with inducible promoters separately controlling expression of the components on each plasmid. One used plasmids that replicate in E. coli and its close relatives. The other used broad-host-range plasmids. When induced to secrete MccV, both versions produced similar zones of inhibition against a susceptible strain of E. coli. Next, we identified putative class II microcins in genomes of bacteria from plant-associated genera (Pantoea, Erwinia, and Xanthomonas) using an existing bioinformatics pipeline. We screened 23 of these putative microcins for E. coli self-inhibition. Seven exhibited some inhibition, mostly later in growth curves, but none had effects that were comparable in strength to MccV. ConclusionsThe genetic parts we created can be assembled in various combinations into tailored systems for secreting small proteins from diverse bacterial chassis. These systems can be used to further characterize the targets of novel microcins and to secrete these or other small proteins for various applications. For example, beneficial bacteria used in crop protection could be engineered to secrete microcins that kill or inhibit plant pathogens to increase their efficacy.

Microbes have the potential to manufacture plastics from sustainable feedstocks while enabling novel material properties and functions that are not easily accessible through conventional chemical synthesis. Realising this potential requires a comprehensive genetic and process engineering framework that spans chassis and bioprocess optimisation, polymer property control, and downstream functionalisation. Here we develop such a platform in Cupriavidus necator, with a focus on high-value polyhydroxyalkanoate (PHA) nanoparticles. To this end we first optimise the transformation protocol for the organism. Next, we create a library of PhaC synthase variants from C. necator, Aeromonas caviae and Brevundimonas sp. in a {Delta}phaC background, demonstrating that they allow customisation of the material properties of produced PHA particles. Our results combine data from Flow cytometry, Transmission Electron Microscopy (TEM), Fourier Transform InfraRed Spectroscopy (FTIR), and Differential Scanning Calorimetry (DSC) to show that it is possible to generate materials ranging from highly crystalline PHAs to softer P(3HB-co-3HHx) copolymers and that an A. caviae PhaC variant can double the yield of large PHA granules. To improve bioprocess sustainability, we coupled C. necator with B. subtilis in sucrose-fed co-cultures, using tetracycline tolerance differences and inoculation ratios to enhance PHA production from inexpensive, sugar-rich feedstocks. Finally, we add function to the produced PHA nanoparticles by using the molecular protein-fusion technology SpyTag-SpyCatcher, showing it is possible to efficiently capture SpyCatcher-GFP on PHA granules as a proof of concept for PHAs use as a customisable bio-based nanoparticle. Together, our work offers an innovation to produce bio-PHA nanoparticles in a customisable way, with potential applications in sustainable biomanufacturing, biosensing, drug delivery and future bioremediation technologies.

Legitimacy screening, the process of verifying the identity and purpose of customers ordering synthetic nucleic acids, is a primary safeguard against the misuse of synthetic biology. However, the associated costs discourage the adoption of screening practices. To evaluate whether AI tools can facilitate this process, we tested five large language models on five verification tasks using customer profiles of life sciences researchers from around the world. We compared AI performance against an expert human baseline on flag accuracy, source quality, source fidelity, and cost. The best-performing model, Gemini 2.5 Pro aided by four bibliographic and sanctions APIs, achieved comparable flag accuracy to the human baseline (90% and 89%, respectively). Gemini 2.5 Pro outperformed the human baseline on source quality and fidelity, at roughly one-tenth of the cost ($1.18 vs. $14.04 per customer). For information-gathering tasks, which excluded the human review step, costs averaged $0.23 per customer, around 50 times cheaper than human screening. These results support piloting AI-assisted legitimacy screening at providers of synthetic nucleic acids and other dual-use biotechnology products, with AI systems handling information gathering and human reviewers retaining authority over order fulfillment decisions.

Efficient genome writing in mammalian cells requires robust methods for integrating large DNA payloads. The previously described method mammalian Switching Antibiotic resistance markers Progressively for Integration (mSwAP-In) enables iterative, biallelic genome rewriting in mammalian stem cells with DNA payloads exceeding 100 kb. However, the lack of standardized vectors and certain technical constraints have limited its broader adoption. Here we present an improved plasmid toolkit designed to streamline the implementation of mSwAP-In. The toolkit includes two core vectors. pLP-TK (pCTC174) is a landing-pad plasmid compatible with Golden Gate assembly of genomic homology arms and supports both mSwAP-In and the recombinase-mediated cassette exchange method Big-IN. mSwAP-In MC2v2 (pKBA135) is a versatile Big DNA assembly and delivery vector that supports Gibson-based assembly and incorporates positive, negative, and fluorescent selection markers, as well as a backbone counterselection cassette to minimize unwanted plasmid integration. The vector architecture also enables propagation in yeast and bacterial hosts, inducible plasmid copy-number amplification in standard E. coli strains, and CRISPR/Cas9-mediated payload release through preinstalled guide RNA target sites. We further characterize the FCU1/5-FC counterselection system in mouse embryonic stem cells and define conditions that minimize its bystander toxicity. Finally, we provide a set of Cas9-gRNA expression plasmids optimized for common mSwAP-In applications. Together, these reagents constitute a standardized and experimentally validated toolkit that simplifies large-scale genome writing using mSwAP-In.

Course-based Undergraduate Research Experiences (CUREs) have emerged as a transformative approach to science education, expanding access to authentic research opportunities beyond the traditional undergraduate research assistant (URA) training. By embedding research into a curriculum, CUREs engage a broad and diverse population of students in a classroom environment that emphasizes experimental design, data analysis, and scientific communication. However, this has been difficult to develop for fields such as plant synthetic biology due to the long timescales of plant transformation. One avenue around this problem is to utilize a recent innovation that enables high throughput and rapid screening of gRNA efficacy by leveraging viral-based delivery of guide RNAs (gRNAs). In this work, we develop and validate a CURE with undergraduate students at Colorado State University (CSU). Students worked in teams to design and test efficacy of gRNAs targeting a Cas9-based transcriptional repressor to different regions of the promoters of the three GIBBERELLIN INSENSITIVE 1 genes (GID1a, GID1b, and GID1c) in Arabidopsis thaliana. Over the semester, students generated and analyzed gene expression data to understand the efficiency of twelve new gRNAs. We further validated CURE student-identified gRNAs with an undergraduate research assistant (URA) that assessed target gene expression and phenotypic outcomes in stable transgenic lines expressing SynTF constructs with the strongest gRNAs from the class. We further describe the curriculum structure to facilitate adoption at other institutions and present student-generated datasets demonstrating the utility of ViN-based screening for identifying effective SynTF gRNAs for plant functional genomics and engineering. Graphical Abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=111 SRC="FIGDIR/small/715601v1_ufig1.gif" ALT="Figure 1"> View larger version (35K): org.highwire.dtl.DTLVardef@13869f5org.highwire.dtl.DTLVardef@b469feorg.highwire.dtl.DTLVardef@9aa51borg.highwire.dtl.DTLVardef@cdc129_HPS_FORMAT_FIGEXP M_FIG C_FIG

Photosynthetic cyanobacteria are promising platforms for sustainable chemical production, as they can convert light and CO2 into valuable compounds. Achieving this often requires engineering cyanobacteria with non-native enzymes with strong promoters to maximize enzyme accumulation. However, despite extensive engineering efforts, the extent to which heterologous proteins misfold and undergo degradation in cyanobacteria remains unknown. Here, we systematically investigate the fate of recombinant proteins in Synechocystis sp. PCC 6803 by quantifying metabolic enzyme degradation. To do this, we developed a quantitative approach that combines split-GFP protein reporting with inducible CRISPRi knockdown of Clp protease system, enabling detection of proteins that would otherwise be degraded. Applying this method to 103 heterologous proteins previously used in cyanobacterial metabolic engineering studies, we find that nearly half undergo significant degradation, with some losing over 95% of their potential expression. Furthermore, we demonstrate that replacing enzymes with homologs is often a more effective strategy to address expression issues than optimizing genetic elements. These findings provide the first quantitative overview of heterologous protein expression in cyanobacteria and identify enzymes that are poorly expressed and suboptimal for their respective pathways, information usable to increase production titers in photosynthetic cell factories.

Cells have the potential to utilize biological pathways to synthesize semiconductor nanomaterials, such as CdS quantum dots. As in chemical reaction schemes, biogenic synthesis requires control of the concentration and redox state of starting materials during the nucleation and growth of nanoparticles. Biological pathways regulate these key processes of particle synthesis, and manipulation of such pathways enables biological control of multiple aspects of nanoparticle synthesis. Here, strains of Escherichia coli were engineered to biosynthesize cadmium sulfide (CdS) quantum dots through the coordinated action of three pathways controlling sulfide generation, cadmium uptake, and nanoparticle nucleation. When exposed to low, micromolar concentrations of external cadmium, strains combining all three pathways produced CdS quantum dots. The synthesis of nanoparticles, nanoparticle yield, and nanoparticle size depended on the combination of pathways found in each strain. Cells lacking all three pathways produced no detectable nanomaterials, cells with specific combinations of one or two pathways produced small particles in the range of 1.95 to 7.9 nm, and cells with all three pathways produced the largest particles with average diameters of 11.78 nm. These results demonstrate that cells can be engineered to control multiple aspects of biogenic nanoparticle synthesis and that these pathways act together to tune the biosynthesis of semiconductor nanomaterials within cells. ImportanceMicrobes synthesize materials, including metallic and semiconductor nanomaterials. This capability stems from the natural ability of microbes to interact with and precisely manipulate metal atoms. Here, multiple biological pathways were combined within a single strain of Escherichia coli, creating a cell capable of producing CdS nanoparticles. This engineered cell controls multiple steps of particle synthesis, including metal uptake, reduction of starting materials, and binding cadmium and sulfide ions to initiate particle formation. Metal uptake by the cells was improved through the modification of a metal ion transport protein, improving cadmium uptake across the outer membrane and creating higher concentrations of cadmium within the cell. Cells with all three pathways were able to produce CdS nanoparticles, called quantum dots, even when exposed to low concentrations of external cadmium. This biotechnology enables nanomaterial synthesis under environmentally friendly conditions and may improve technologies using bacteria to clean up toxic metals.

Cancer therapeutics are increasingly incorporating engineered receptors due to their ability to detect extracellular ligands and initiate intracellular responses that regulate gene expression. By redesigning these natural signaling systems, synthetic receptors hold great potential for use in novel cell-based therapies. One particularly promising direction is modifying the Notch receptor, a transmembrane protein that naturally mediates ligand-dependent signaling at the cell surface to regulate cell proliferation and differentiation in neurogenesis. Both the intracellular and extracellular domains of Notch can be replaced with alternative domains, creating the family of modified Notch receptors known as synthetic Notch (synNotch). In existing synNotch-activated chimeric antigen receptor (CAR) T-cells, the extracellular domain can be engineered to adjust binding affinity for a specific cancer antigen, enabling precise tuning of therapeutic activity while minimizing off-target effects. To quantify and inform such tuning, we develop differential equations models of synNotch receptor signaling and subsequent gene expression. The mathematical models couple activation dynamics on fast timescales (characteristic of receptor-ligand interactions) and on slow timescales (characteristic of downstream gene expression dynamics). Global Sobol sensitivity analysis of the proposed models highlights parameters that yield the greatest variability in synNotch signal transduction and gene expression, indicating their potential to be engineered for different functions in future cancer therapeutics. For the receptor-ligand interactions in the synNotch model, we find that ligand association and ligand-independent activation are the most sensitive parameters. In the downstream gene expression model, promoter strength and degradation rates of mRNA and gene product are found to be most amenable to engineering.

Zymomonas mobilis is an ethanologenic Alphaproteobacterium with many interesting characteristics for fundamental research and applied microbial engineering. Although genetic engineering has been established for Z. mobilis since the 1980s, a rich set of inducible transcriptional regulators is still unavailable. In this work, seven different chemically inducible promoters have been systematically tested for their functionality in Z. mobilis. In particular, for the first time, NahR-PsalTTC, VanRAM-PvanCC, CinRAM-Pcin and LuxR-PluxB have been characterized in Z. mobilis, alongside the commonly used regulator-promoter pairs TetR-Ptet and LacI-PlacT7A1_O3O4, and the less commonly used XylS-Pm. All promoters investigated in this work are compatible with the Golden Gate modular cloning framework Zymo-Parts. Characterization was carried out with a shuttle vector backbone based on pZMO7, which has so far been rarely used for applications in Z. mobilis but seems to be completely stable without selection and generates high and uniform levels of expression. From the experimental results presented, it can be concluded that VanRAM-PvanCC and CinRAM-Pcin are particularly promising for broad use in the Z. mobilis community. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=126 SRC="FIGDIR/small/712268v1_ufig1.gif" ALT="Figure 1"> View larger version (39K): org.highwire.dtl.DTLVardef@16579e6org.highwire.dtl.DTLVardef@1262533org.highwire.dtl.DTLVardef@15456a2org.highwire.dtl.DTLVardef@3af98_HPS_FORMAT_FIGEXP M_FIG C_FIG

The thermoacidophilic red alga Cyanidioschyzon merolae represents one of the simplest photosynthetic eukaryotes and an ancient divergent group in the primary endosymbiotic Viridiplantae. Because of its [~]16 Mbp genome, containing few introns, and capacity for transgene integration by homologous recombination, it is an emerging chassis for synthetic biology. However, genomic integration sites and scalable transformation methods have not been established to systematically investigate the effect of genome position on transgene expression. Here, we combined bioinformatic genome analysis, liquid-handling robotics, and assays of heterologous protein and metabolite production to establish a reproducible framework for nuclear genome engineering in C. merolae. We mapped and annotated 40 intergenic loci as candidate neutral sites across 16 out of 20 chromosomes and could validate 38 of them through robotic-assisted transformation. Reporter gene expression analysis revealed highly uniform expression at all integration sites across broad populations of transformants, indicating surprising minimal positional effects and transcriptional neutrality. The functional equivalence of these genomic landing pads was determined by expression of a heterologous isoprene synthase, and coupling algal photobioreactors to headspace analysis to quantify isoprene production driven by transgene expression from different integration sites. Single copy transgene integrants, regardless of genome position, exhibited comparable reporter signals and consequent isoprene production. Together, these results provide the first experimentally validated set of neutral integration sites in C. merolae and establish a high-throughput transformation protocol for its genetic engineering in the context of synthetic genome biology.

Cell-free expression systems (CFES) are increasingly used alongside conventional biotechnological approaches to accelerate early-stage prototyping and are particularly valuable in point-of-use settings. However, their broader adoption remains limited by time- and cost-intensive preparation, as well as stringent cryogenic storage requirements. To address this, several studies have explored lyophilization with protective additives to generate stable, solid-state CFES. These approaches had to balance the protection gained with a loss of activity due to the additives. In this study, we present a CFES that contains a tardigrade-derived Cytosolic-Abundant Heat-Soluble (CAHS) protein to protect the biosynthetic machinery in lysates from damages during drying. We show that the CAHS protein, without any other additives, preserves protein synthesis activity during low-cost room temperature desiccation, while unprotected lysates are affected in mRNA synthesis kinetics and translation yields. The diversity of tardigrade-derived protective proteins is a treasure trove for cell-free synthetic biology, in particular for making CFES more accessible and portable. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=85 SRC="FIGDIR/small/715078v1_ufig1.gif" ALT="Figure 1"> View larger version (27K): org.highwire.dtl.DTLVardef@8ecc2eorg.highwire.dtl.DTLVardef@ff0432org.highwire.dtl.DTLVardef@6c940eorg.highwire.dtl.DTLVardef@6c5390_HPS_FORMAT_FIGEXP M_FIG C_FIG

Selective encapsulation of target enzymes is an increasingly well studied field with a host of potential applications for biotechnology. Natively, many bacteria utilize bacterial microcompartments (BMCs) for enzyme encapsulation to enhance catalysis. BMCs are protein shells that enable selective localization of targeted metabolic enzymes and may improve catalytic rates by co-localizing pathway enzymes and/or serve to sequester toxic or volatile intermediates. The microcompartment shell of Haliangium ochraceum (HO) is a notable BMC chassis because of its modularity and versatility; it is easily expressed and assembled outside its native host and can accept a wide array of cargo. Recently, it was demonstrated that assembly of HO BMC shells can be easily achieved in vitro. Following up on our previous work on in vivo assembly of HO-BMCs with triose phosphate isomerase (TPI) as model enzyme cargo, here we have demonstrated the advantages of in vitro assembly (IVA) for targeted enzyme encapsulation. We achieved variable loading of BMC shells with targeted amounts of TPI and demonstrated enhanced thermal stability of encapsulated TPI versus free TPI up to 62{degrees}C.

Technologies developed over the past decade have made Saccharomyces cerevisiae a promising platform for producing various natural products. Balancing multi-enzyme expression, while maintaining robust microbial growth, remains a limiting factor for engineering long biosynthetic pathways in yeast. Here, we improved the transcriptional capacity of our previously developed isopropyl {beta}-D-1-thiogalactopyranoside (IPTG)-inducible synthetic transcription factors (synTFs) derived from the plant JUB1 DNA-binding domain. To this end, at cysteine positions within surface-exposed loop regions of a JUB1-derived DNA-binding scaffold, we introduced a short peptide to enhance loop flexibility while providing local stability and orientation. The generated synTFs, so-called JUB1-X synTFs, varying in strength, have been successfully used to improve the synthesis of 3-phosphoadenosine 5-phosphosulfate (PAPS), a universal sulfate donor necessary for the synthesis of bioactive molecules, including therapeutic glycosaminoglycans and sulfolipids. Using only this engineered yeast strain in simple batch culture, PAPS accumulation of 21.4 {+/-} 5.8 mg g-1 cdw was achieved after only 5 hours of inducing the expression of JUB1-X synTFs. Beyond PAPS production, the design principle demonstrated here provides a generalizable strategy to fine-tune other plant-derived synTFs, expanding the regulatory capabilities of existing synTF collections. Together, this work offers a modular, scalable approach to constructing high-performance gene circuits and supports the development of yeast cell factories for complex metabolic and synthetic biology applications.