ACS Synthetic Biology

Bacillus subtilis is an important chassis for biotechnology, but its use in multiplex genome engineering is limited by low natural transformation efficiency. Here, we compared inducible promoter systems for synthetic activation of the competence regulator ComK and evaluated their effects on the comG operon competence reporter and transformation efficiency. Xylose- and mannitol-inducible systems outperformed IPTG-based constructs and shifted 96-99% of cells into a reporter-positive competent state. However, reporter activation alone did not predict transformation potential. Optimization of culture density and induction timing increased transformant yield 45-fold relative to the initial protocol and 2800-fold relative to the conventional Spizizen method. Disruption of native competence regulatory genes did not improve performance and often reduced transformation output, highlighting the importance of endogenous regulatory circuitry. Using the optimized strain and protocol, we achieved co-transformation frequencies of 11-18% and constructed multiplex spore-display libraries containing fluorescent protein fusions integrated at multiple loci. Screening identified strong dual-display combinations and showed that cargo loading depends on anchor protein, integration locus, and genetic background. SscA fusions supported the highest display capacity and promoted synergistic co-display. Together, these results show improvements in natural transformation-based genome engineering in B. subtilis and provide insight into the construction of multifunctional engineered spores.

Engineered microbial communities hold significant biotechnological potential because their collective metabolism can produce functions beyond those achievable by individual strains. However, multicellular synthetic gene circuits require orthogonal communication systems that enable precise, programmable signaling between cells. Quorum sensing (QS), where cells both produce and detect small diffusible signal molecules, offers a natural framework for such intercellular communication. However, the construction of complex multicellular circuits for applications such as biobased production is currently hampered by the limited number of orthogonal QS channels available in yeast. Here, we expand the QS toolkit in Saccharomyces cerevisiae by characterizing four LuxR-type biosensors based on EsaR, LasR, TraR and RpaR, alongside the previously established LuxR biosensor. We functionally expressed acyl-CoA-dependent HSL synthases in yeast, producing a diverse range of aliphatic and aromatic HSL signals. LuxR and RpaR, were compatible with in vivo ligand production and established as orthogonal QS signaling pair with synthases MesI and RpaI, respectively. Co-culture experiments demonstrated QS-dependent intercellular signaling, with 3.9-fold and 6.4-fold induction relative to monocultures. Together, these results establish a modular and extensible platform for orthogonal intercellular communication in yeast, enabling the construction of multicellular synthetic gene circuits.

Improving the reconstituted translation system is a key requirement for bottom-up synthetic biology. Here, we developed a two-step in vitro evolutionary method that can be used for improving translational proteins. In this method, two distinct conditions were sequentially applied while maintaining genotype-phenotype linkage in water-in-oil droplets. Using this method, we performed in vitro evolution of four translation factors, IleRS, PheRS, EF-G, and EF-Tu, and identified mutations that modestly enhanced translation activity in in vitro expression assays. One of the EF-G mutations (P610S) increased activity per protein approximately 2-fold for the recombinant protein purified from E. coli. This selection method is useful for improving translational proteins for bottom-up synthetic biology.

Temporal gradient sensing is a fundamental capability observed across diverse natural biological systems, contributing to the coordination of their functions. Harnessing this ability is also of significant interest in synthetic biology, particularly for sensing and control applications. In this work, we focus on a biomolecular topology that exemplifies a broader class of signal-differentiating architectures, while introducing a structural variant of it. We examine their behavior under both nominal and non-ideal conditions, accounting for stochastic noise arising from different sources. Our investigation includes scenarios where these topologies operate independently, as well as when embedded within minimal regulatory architectures based on negative as well as positive feedback. We analyze the stability of the resulting macroscopic dynamics--a prerequisite for practical deployment--and quantify stochastic fluctuations in system output, providing comparisons with the corresponding input/unregulated process. Importantly, our results demonstrate that signal differentiation can be effectively implemented in a biomolecular setting without incurring deleterious noise amplification--a major concern in the utilization of derivative action across disciplines.

Wickerhamomyces ciferrii is a non-model diploid yeast that naturally produces tetraacetyl phytosphingosine (TAPS), a sphingoid base used in cosmetic and dermatological applications. However, its strong preference for non-homologous end joining (NHEJ) over homologous recombination (HR) limits conventional genome editing, while disruption of LIG4, a core NHEJ gene, compromises cellular fitness. Here, we repurposed native NHEJ activity to develop a homology-independent multicopy genome integration platform for W. ciferrii. The platform combines three optimized donor-design features: telomeric end-shielding with two tandem copies of an 11 bp repeat to improve linear donor persistence, a defective URA5 auxotrophic marker to enrich multicopy integrants, and 5'-phosphorylated donor termini to enhance transformant recovery and integration output. These features were consolidated into the platform vector pTdmVU5. As a metabolic engineering demonstration, multicopy integration of LCB1 and LCB2, encoding the two subunits of serine palmitoyltransferase, increased TAPS titer by 2.7-fold. This work converts the native NHEJ bias of W. ciferrii from a barrier to precise genome editing into a practical tool for pathway amplification and establishes a framework for engineering NHEJ-dominant non-model yeasts.

Cyanobacterial natural products are a rich source of bioactive compounds, yet their heterologous production remains challenging. This study investigates the feasibility of expressing the lyngbyatoxin A (LTXA) biosynthetic gene cluster in a fungal host. The lyngbyatoxin biosynthetic genes (ltxA, ltxB, ltxC) were individually cloned and expressed in Aspergillus oryzae NSAR1 under the control of an inducible promoter. Metabolite production was assessed using LC- MS, and transcriptional analysis was performed by RT-PCR. Codon-optimized constructs and precursor feeding experiments were employed to evaluate pathway functionality. No production of LTXA or pathway intermediates was detected upon co-expression of ltxA-C despite confirmed transcription of ltxB and ltxC. RT-PCR analysis revealed truncation of the ltxA transcript, suggesting incompatibility with fungal transcriptional or splicing machinery. In contrast, expression of a codon-optimized ltxC enabled biotransformation of indolactam V to LTXA in A. oryzae, confirming functional expression of the prenyltransferase. These results highlight transcriptional limitations as a key barrier to heterologous expression of cyanobacterial NRPS pathways in fungal hosts, while demonstrating that downstream tailoring enzymes can remain functional. This work provides insights for future engineering of fungal platforms for cyanobacterial natural product biosynthesis.

The genus Mycobacterium is increasingly recognized as a major clinical concern due to diseases such as tuberculosis, along with the emergence of antimicrobial-resistant strains, underscoring the urgent need for advanced genetic tools to study mycobacterial biology and pathogenesis. Progress in this area relies heavily on the functional characterization of previously unannotated genes, which necessitates tightly regulated expression systems. Here, we report the development of an improved tetracycline-regulated vector platform, comprising the episomal pM(R)T2 and integrative pMI(R)T2 series, which builds upon the previously described pMT vector system. The T2 vector series incorporates a fine-tuned TetRO system for enhanced transcriptional control. The pMT2 vectors function as tetracycline-inducible systems, whereas the pMRT2 variants utilize a reverse tetracycline repressor (RevTetR) to enable tetracycline-repressible gene regulation. Additionally, the integrative variant, pMI(R)T2 switches the oriM element with the integrase and attP sites derived from mycobacteriophage L5, facilitating stable genomic integration and controlled expression of concentration-sensitive genes, including toxins. To expand the selection flexibility, the pAN(R)Tet series replaces the kanamycin resistance cassette with a hygromycin resistance cassette. Functional validation of gene regulation in M. smegmatis and M. bovis BCG shows that both TetR and RevTetR systems provide reliable inducible and repressible controls, respectively, upon anhydrotetracycline addition. Taken together, these vectors constitute a versatile, tightly regulated genetic toolkit with significant potential to accelerate research and therapeutic development in mycobacterial systems.

Budding yeast Saccharomyces cerevisiae is a workhorse chassis for producing added food and agricultural compounds. However, building multi-enzymatic pathways for these chemicals often requires iterative genomic integration, underscoring the need for efficient, rapid genome-editing tools that can reliably target transcriptionally active chromosomal regions. In this study, to accelerate strain construction, we established a genome-editing toolkit to rapidly engineer eight loci, highly expressed hot-spots, but nonessential genomic sites suitable for stable pathway assembly. Our approach integrates three key design features: (i) selectable markers to enable rapid screening of edited cells, (ii) extended homology arms that leverage the yeast homology-directed repair machinery for robust genomic integration, and (iii) co-delivery of Cas9 and guide RNAs to promote efficient double-stranded DNA breaks at specific integration sites. The sequence independence of FASTOP relies on the release of integration cassettes from integrative vectors, mediated by restriction digestion at two flanking multiple-cutting sites in the integration module to minimize the risk of introducing sequence errors during PCR amplification of the integration cassettes. Following the introduction of a fluorescent reporter cassette, we observed high integration efficiencies across the target sites. We then integrated the biosynthetic pathway of plant-derived flavonoid naringenin into the hot-spots of the yeast genome using the FASTOP toolkit. Our results demonstrated that upon expressing the five essential genes in simple shake flask culture, naringenin production reached 505.7 mg/L, representing a significant (69-fold) increase over previously reported titers for comparable minimal heterologous pathways in S. cerevisiae. Together, the FATSOP toolkit provides a user-friendly platform for reliably modifying hot-spot loci to rapidly construct multi-enzymatic metabolic pathways in S. cerevisiae, while achieving high production levels for high-value food-relevant metabolites.

Escherichia coli is a well-established model organism in molecular biology and biotechnology. Despite its long history as a laboratory workhorse, the efficient single-step chromosomal integration of large DNA fragments remains a challenge. Currently known methods are either simple but have limitations on insert size, or flexible but laborious requiring plasmid construction or multi-step procedures. Here, we present PhAGE (Phage-Assisted Genome Engineering), which enables the integration of [~]20 kb DNA fragments into E. coli genome within a single day. PhAGE method uses in vitro packaging of recombinant DNA into bacteriophage capsids, followed by general transduction to introduce pre-assembled DNA with flanking homology arms into recipient cells. This approach allows efficient and landing pad-free integration of large constructs into the target loci. We demonstrate its usefulness through rapid integration of multi-gene operons. PhAGE resolves the long-standing trade-off between simplicity and insert size in E. coli genome engineering, accelerating strain construction across a wide range of applications, from biosynthetic pathway engineering to genome-scale design.

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.

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.

Translation initiation has become an attractive target for engineering orthogonal translation systems, yet the extent to which these systems retain functionality across distinct host backgrounds remains poorly defined. In bacteria, start codon recognition depends on pairing between the initiator tRNA anticodon and a suitable start codon within the appropriate distance from the Shine-Dalgarno sequence. These sequence-specific interactions enable translation initiation to be reprogrammed through anticodon engineering. What is currently missing is an understanding of how anticodon mutants of initiator tRNAs function across different bacterial strains. Here, we systematically evaluated the portability of a library of twelve i-tRNA anticodon mutants paired with their complementary non-canonical start codons. Most i-tRNA-start codon pairs supported detectable translation initiation across multiple strains, demonstrating broad functional portability. However, initiation efficiency, absolute system output, and fitness effects varied substantially between strains. Comparative genomic analyses revealed host-specific gene differences broadly, and endogenous tRNA gene sequence and copy number specifically, was associated with this variability. While most i-tRNA variants were well tolerated, a subset produced strain-dependent growth defects that primarily affected growth rate rather than final culture density. Together, these findings show that translation initiation efficacy of engineered i-tRNAs is partially strain-dependent and that host background must be considered a key design variable when deploying these translation systems. Looking forward, this study provides a framework for host-aware selection of microbial chassis for orthogonal translation applications in synthetic biology. Graphical Abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=100 SRC="FIGDIR/small/719103v1_ufig1.gif" ALT="Figure 1"> View larger version (20K): org.highwire.dtl.DTLVardef@118b02borg.highwire.dtl.DTLVardef@1d5dab0org.highwire.dtl.DTLVardef@1088d0borg.highwire.dtl.DTLVardef@63eb74_HPS_FORMAT_FIGEXP M_FIG C_FIG

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.

Non-model bacteria offer unique metabolic capabilities for sustainable bioproduction, yet their limited genetic accessibility hinders systematic strain development. Here we present conjugation-based serine recombinase-assisted genome engineering (cSAGE), a broad-host-range platform that enables predictable, iterative genomic integration in transformation-resistant bacteria. cSAGE combines conjugative DNA delivery, standardized low-copy vectors, orthogonal recombinases, and modular genetic parts to support rapid pathway assembly and cross-host benchmarking. Using purple nonsulfur bacteria as a testbed, we integrate promoter engineering, multi-payload genome modification, and genome-scale metabolic modeling to empirically evaluate host-dependent pathway performance. Applying this workflow, we identify strain-specific differences in photosynthetic conversion of lignin-derived p-coumarate to the thermoplastic precursor p-vinylphenol. By enabling genome engineering and functional comparison across diverse bacteria using a single plasmid system, cSAGE provides a general framework for non-model strain prototyping and biotransformation discovery.

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 disparity between the production and demand of recombinant proteins (r-proteins) has significantly hindered their commercial viability. Leveraging genomic resources offers substantial promise in enhancing our comprehension of metabolic and regulatory networks, thus facilitating the development of highly productive protein cell factories. However, the considerable gap between high-throughput strategies for monitoring r-protein secretion and genome perturbation in P. pastoris continues to obstruct the systematic linkage of genotype and phenotype, thereby limiting the optimization of production. Here, we developed a novel strategy combining dual-base editor-mediated in-situ genome engineering with nanobody-regulated biosensor-assisted droplet sorting to enhance r-protein secretion (BINDER) in P. pastoris. We successfully employed BINDER to screen recombinant human serum albumin (rHSA) hyper-producers and identified two critical SNVs conferring up to a 1.78-fold improved secretion titer from 113,632 mutants, providing valuable insights into the secretion mechanism. Fed-batch cultivation of the engineered strain resulted in the highest reported rHSA titer, 23.43g/L, in P. pastoris, demonstrating its substantial potential for industrial applications. Given the high transferability of base editors and the novel biosensors independence from the properties of the target protein, the strategy developed here might be expanded to a variety of microbial species and r-proteins.

The SCN1A gene encodes NaV1.1, a voltage-gated sodium channel protein that is necessary for neuronal excitability and whose loss-of-function mutations cause Dravet syndrome, a treatment-resistant childhood onset epilepsy. Gene replacement strategies for this syndrome are challenged by the large size of SCN1A and difficulty achieving stable cellular expression. Lentiviral vectors (LVVs) offer sufficient packaging capacity and genomic integration for defective SCN1A gene replacement. Here, we evaluated LVV-mediated delivery of different engineered SCN1A transgene sequences in human cells. LVV-transduced cells expressed full-length NaV1.1 protein that trafficked to the membrane and produced functional sodium currents. However, SCN1A transgene expression declined over time despite stable vector copy number, indicating post-integration regulatory limitations. Expression efficiency varied by SCN1A transgene sequence, with a codon-optimized variant showing higher expression despite lower LVV copy number. Treatment with sodium butyrate, a histone deacetylase inhibitor, significantly enhanced SCN1A transgene expression and partially rescued expression decay in a sequence-dependent manner. Incorporation of a ubiquitous chromatin opening element (UCOE) upstream of the promoter to maintain expression resulted in a trend of increased expression and increased responsiveness to butyrate. These findings demonstrate that sequence-specific and epigenetic factors may influence expression of large transgenes following lentiviral delivery, highlighting key challenges and design considerations for therapeutic SCN1A transgene expression.