Synthetic and Systems Biotechnology

DNA-based information storage offers a promising alternative to conventional media due to its high density, long-term stability, and low energy requirements. However, its application remains hindered by synthesis costs, limited sequence length and poor scalability. DNA polymerase is a critical enzymatic tool in the DNA storage systems by enabling high-fidelity data writing and targeted sequence amplification. In this study, we engineered chimeric DNA polymerases by fusing the high-fidelity 9{degrees}N DNA polymerase with double-stranded DNA binding proteins derived from thermophilic archaea. These fusions significantly enhanced processivity, thermal stability, and salt tolerance by stabilizing enzyme-template interactions, mimicking sliding clamps while preserving catalytic efficiency. Leveraging these properties, we demonstrated precise file retrieval from a mixed oligonucleotide pool using orthogonal barcode primers. Compared with wild-type 9{degrees}N, the chimeric polymerases, particularly PLS, exhibited reduced substitution error rates and improved read accuracy. We then applied these enzymes to a DNA movable type storage system, where prefabricated DNA modules were assembled into encoding blocks. Using engineered polymerases, these blocks were recombined to enable flexible data rewriting without de novo DNA synthesis. Moreover, a multi-enzyme assembly strategy enabled the construction of kilobase-scale DNA sequences encoding a classical Chinese poem, achieving complete data recovery. All assembled fragments remained stable in E. coli over 100 generations, exhibiting the potential for in vivo storage. Collectively, our findings demonstrated the role of engineered DNA polymerases for DNA-based information storage. Moreover, this system reduced synthesis demands, supported scalable rewriting, and ensured long-term preservation, offering a practical route to sustainable, high-fidelity DNA data storage.

CRISPR-Cas systems have revolutionized genome editing with their precision and versatility, enabling transformative applications in various fields, especially in the treatment of genetic diseases. However, the clinical translation of this technology is hindered by challenges such as off-target effects and uncontrolled nuclease activity. At the same time, it has the possibility of causing biosecurity risks, underscoring the urgent need for reliable regulatory tools. Existing CRISPR inhibitors, primarily anti-CRISPR protein or exogenously synthesized small molecules, are limited by their specificity or bioavailability and long research period, unable to address the diverse CRISPR nucleases used in research and therapy. Based on the phenomena obtained from various in vitro and cell experiments, combining molecular dynamics simulation and bio - layer interferometry (BLI) analysis, here we report a naturally occurring small-molecule {beta}-nicotinamide mononucleotide (NMN), the first known endogenous metabolite with broad-spectrum inhibitory activity against multiple CRISPR-associated proteins (Cas9, Cas12, and Cas13) through various mechanisms. Our findings establish NMN as a dual-purpose tool, which reduces cell damage caused by gene editing and mitigates risks of unintended genetic modifications in research and clinical settings. This discovery further shortens the distance between basic medicine and translational medicine, providing a new approach for developing endogenous regulatory molecules in genome engineering.

DNA nucleases TnpB and IscB were regarded as new antibacterial strategy to combat the drug-resistant bacteria represented by Escherichia coli due to its specificity in targeting DNA and smallest size, but the genome-editing of TnpB/IscB in E. coli remains unclear. This study characterized the genome-editing of TnpB/IscB in E. coli strains. First, the toxicity and cleavage results showed TnpB only worked in E. coli MG1655, while IscB and enIscB could perform in ATCC9637 and BL21(DE3). Next, TnpB-based genome-editing tool was established in MG1655, while IscB/enIscB achieved in ATCC9637/BL21(DE3). The copy number of TnpB/IscB/enIscB were changed to explore the impact of editing efficiency. Moreover, the editing plasmids were successfully cured. Finally, the escaping mechanism of E. coli under editing of TnpB/IscB was revealed. Overall, this study successfully applied TnpB/IscB/enIscB to genome-editing in E. coli, which will broaden genetic manipulation toolbox in E. coli and facilitate the development of new antimicrobial drugs.

The accelerating environmental crisis has intensified the demand for switching from traditional economy to a renewable one with a reduced carbon footprint. Here we reported a hybrid system, coupling chemical process of CO2 hydrogen reduction and biological process for polyhydroxybutyrate (PHB) synthesis, that utilized CO2 as a raw material to produce PHB in vitro. The synthetic pathway of PHB was optimized by screening more efficient methanol oxidases, high activity mutants of glycolaldehyde synthase and coordinating enzyme dosages in the pathway, which achieved the carbon yield of 93.6% for producing PHB from methanol. Finally, by combining with the chemical process from CO2 to methanol, a scaling-up bio-system was performed to convert CO2 into PHB, yielding 5.8 g/L with the productivity of 1.06 g-1L-1h-1. This approach represents a promising carbon-neutral way to produce biodegradable plastics.

Since the demonstration that rapid polyethylene terephthalate (PET) decomposition using enzymes is feasible, a number of efficient depolymerases have been reported with the aim of resolving the plastic pollution issues. However, sporadic studies on enzymes with PET hydrolysis activity hinder the understanding of the distribution of potential PETases hidden in natures repertoire, and subsequently, the identification of potent enzymes. Here, we present the clustering of 1,894 PETase candidates, which include the majority of known PETases, and describe their profiling. An archipelago landscape of 170 lineages shows distribution of 289 representative sequences with features associated with PET-degrading capabilities. A birds-eye view of the landscape identifies three highly promising yet unexplored PETase lineages and two potent PETases, Mipa-P and Kubu-P. The engineered Mipa-PM19 and Kubu-PM12 variants exhibit both high PET hydrolysis activity and thermal stability. In particular, Kubu-PM12 outperformed the engineered benchmarks in terms of PET depolymerization in harsh environments, such as with high substrate load and ethylene glycol as the solvent. Our landscape framework and the identified variants assist in the understanding of how biological processes respond to solid-state and non-natural PET plastics.

Promoters are one of the most critical regulatory elements controlling metabolic pathways. However, in recent years, researchers have simply perfected promoter strength, but ignored the relationship between the internal sequences and promoter strength. In this context, we constructed and characterized a mutant promoter library of Ptrc through dozens of mutation-construction-screening-characterization engineering cycles. After excluding invalid mutation sites, we established a synthetic promoter library, which consisted of 3665 different variants, displaying an intensity range of more than two orders of magnitude. The strongest variant was 1.52-fold stronger than a 1 mM isopropyl-{beta}-D-thiogalactoside driven PT7 promoter. Our synthetic promoter library exhibited superior applicability when expressing different reporters, in both plasmids and the genome. Different machine learning models were built and optimized to explore relationships between the promoter sequences and transcriptional strength. Finally, our XgBoost model exhibited optimal performance, and we utilized this approach to precisely predict the strength of artificially designed promoter sequences. Our work provides a powerful platform that enables the predictable tuning of promoters to achieve the optimal transcriptional strength.

The fusion expression of DNA replication-related proteins with nucleotide deaminase enzymes promotes random mutations in bacterial genomes, thereby increasing genetic diversity among population. Most previous studies have focused on cytosine deaminase, which produces only C[->]T mutations, significantly limiting the variety of mutation types. In this study, we developed a fusion expression system by combining DnaG (RNA primase) with adenine deaminase TadA-8e (DnaG-TadA) in Escherichia coli, which is capable of rapidly introducing A[->]G mutations into the E. coli genome, resulting in a 664-fold increase in terms of mutation rate. Additionally, we engineered a dual-functional TadA variant, TadAD, and then fused it with DnaG. This construct introduced both C[->]T and A[->]G mutations into the E. coli genome, with the mutation rate further increased by 370-fold upon co-expression with an uracil glycosylase inhibitor (DnaG-TadAD-UGI). We applied DnaG-TadA and DnaG-TadAD-UGI systems to the adaptive laboratory evolution for Cd2+ and kanamycin resistance, achieving an 8.0 mM Cd2+ and 200 g/mL kanamycin tolerance within just 17 days and 132 hours, respectively. Compared to conventional evolution methods, the final tolerance levels were increased by 320% and 266%, respectively. Our work offers a novel strategy for random mutagenesis in E. coli and potentially other prokaryotic species. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=98 SRC="FIGDIR/small/610230v1_ufig1.gif" ALT="Figure 1"> View larger version (22K): org.highwire.dtl.DTLVardef@176e445org.highwire.dtl.DTLVardef@848b7corg.highwire.dtl.DTLVardef@1e190b9org.highwire.dtl.DTLVardef@1e47b44_HPS_FORMAT_FIGEXP M_FIG C_FIG For TOC only

Non-ribosomal peptide synthetases (NRPS) are multimodular enzymes that produce complex peptides with diverse biological activities, potentially being used as clinical drugs. However, the pharmaceutical applications of such natural peptides often require further derivatisation and modification of the peptide backbone, mainly performed by chemical synthesis. A sustainable alternative resembles the in vivo engineering of NRPS to change and modify the enzyme properties rationally and, thus, the produced products. The novel NRPS engineering concept, the eXchange Unit Thiolation domain (XUT), allows the efficient modular assembly of different natural NRPS fragments to form hybrid NRPS that produce defined peptides. In this study, we describe a Golden Gate assembly (GGA) method for efficient high-throughput generation of novel and engineered NRPS libraries utilising the XUT concept. This method was applied to generate over 100 novel NRPS with the possibility of changing starter, elongation, and termination modules, respectively. Additionally, we applied this method for targeted modification of the xenoamicin biosynthetic gene cluster (BGC) XabABCD from Xenorhabdus doucetiae, resulting in the generation of 25 novel xenoamicin derivatives. Graphical AbstractA Golden Gate assembly (GGA) method was developed for the efficient assembly of natural and engineered non-ribosomal peptide synthetases (NRPS). This method has enabled the creation of NRPS libraries to generate novel peptides in high-throughput as well as the targeted derivatisation of natural products (NP). O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=179 SRC="FIGDIR/small/650154v1_ufig1.gif" ALT="Figure 1"> View larger version (60K): org.highwire.dtl.DTLVardef@1161805org.highwire.dtl.DTLVardef@1832a43org.highwire.dtl.DTLVardef@4bc6b8org.highwire.dtl.DTLVardef@e3908d_HPS_FORMAT_FIGEXP M_FIG C_FIG

Functional genetic elements are one of the most essential units for synthetic biology. However, both knowledge-driven and data-driven methodology can hardly accomplish the complicated task of genetic elements design efficiently due to the lack of explicit regulatory logics and training samples. Here, we proposed a knowledge-constraint deep learning model named PccGEO to automatically design functional genetic elements with high success rate and efficiency. PccGEO utilized a novel "fill-in-the-flank" strategy with a conditional generative adversarial network structure to optimize the flanking regions of known functional sequences derived from the biological prior knowledge, which can efficiently capture the implicit patterns with a reduced searching space. We applied PccGEO in the design of Escherichia coli promoters, and found that the implicit patterns in flanking regions matter to the properties of promoters such as the expression level. The PccGEO-designed constitutive and inducible promoters showed more than 91.6% chance of success by in vivo validation. We further utilized PccGEO by setting a limited frequency of nucleotide modifications and surprisingly found that the expression level of E. coli sigma 70 promoters could show up to a 159.3-fold increase with only 10-bp nucleotide modifications. The results supported that the implicit patterns are important in the design of functional gene elements and validated the strong capacity of our method in the efficient design of functional genetic elements. Availabilityhttps://github.com/WangLabTHU/PccGEO

Streptomyces has enormous potential to produce novel natural products (NPs) as it harbors a huge reservoir of uncharacterized and silent natural product biosynthetic gene clusters (BGCs). However, the lack of efficient gene cluster engineering strategies has hampered the pace of new drug discovery. Here, we developed an easy-to-use, highly flexible DNA assembly toolkit for gene cluster engineering. The DNA assembly toolkit are compatible with various DNA assembling approaches including Biobrick, Golden Gate, CATCH, yeast homologous recombination-based DNA assembly and homing endonuclease-mediated assembly. This compatibility offers great flexibility in handling multiple genetic parts or refactoring large gene clusters. To demonstrate the utility of this toolkit, we quantified a library of modular regulatory parts, and engineered a gene cluster (act) using characterized promoters that led to increased production. Overall, this work provides a powerful part assembly toolkit that can be used for natural product discovery and optimization in Streptomyces.

CO2 recycle is crucial to the global carbon neutrality. Though cyanobacteria are known to be photoautotrophic cell factories capable of converting CO2 into valuable chemicals, their slower growth rate and lower biomass accumulation compared to those of the heterotrophic organisms significantly restrict their application at commercial scale. The newly discovered marine cyanobacterium, Synechococcus sp. PCC 11901 (hereafter PCC 11901) offers several advantages like rapid growth, high biomass and high salinity tolerance, and could become a new generation of cyanobacterial chassis. To promote its application, in this study we developed genetic toolboxes applicable to PCC11901. First, a cobalamin (VB12)-independent chassis was constructed, allowing for its cheaper cultivation. Second, genome copy numbers and transformation methods were respectively measured and optimized. The 14 neutral sites were identified and characterized within the genome PCC 11901, providing locations for genetic integration of exogenous cassettes. Subsequently, libraries were developed, reaching an expression range of approximately 800 folds for constitutive promoters and an induction fold of up to approximately 400 for inducible promotor, respectively. As a proof of concept of its utilization, we engineered the synthetic pathways of glucosylglycerol (GG) into PCC 11901 using the established toolboxes, yielding 590.41 {+/-} 21.48 mg/L for GG production. Notably, we found the cobalamin-independent PCC 11901 chassis exhibited superior self-sedimentation ability compared to the wild-type chassis. Our work here made it possible to develop the fast-growing PCC 11901 as efficient carbon-neutral cell factory in the future.

Molecular editing of an amino group from -position of amino acids to its {beta}-position is of scientific interest and could be economically appealing. Here we reconstructed an in vitro biotransformation pathway composed of two cascade decarboxylases, i.e., aspartate {beta}-decarboxylase and aspartate -decarboxylase, and implemented molecular editing to change -alanine into {beta}-alanine. In it, we discovered a new reaction of aspartate {beta}-decarboxylase that can fix CO2 directly. This cascade enzymatic pathway enabled an aminomutation reaction with 100% carbon atom economy. This work presented the first CO2-fixing biological reaction catalyzed by the amino acid decarboxylases and demonstrated a new means for the molecular editing of -amino acids.

Gene therapy studies have been of great importance in the elimination of genetic diseases, and the capability of the CRISPR/Cas9 genome editing technique to correct genetic defects has shown great promise. As DNA-based Cas9 nuclease delivery is preferable because of its low cost and higher stability, effective vector-based CRISPR/Cas9 administration is urgently needed. Here, we used the multicellular organism Caenorhabditis elegans to optimize the polymer-mediated DNA delivery system to generate mutants with CRISPR/Cas9. Toward this end, the cationically quaternized polymer of POEGMA-b-P4VP (POEGMA-b-QP4VP) as a carrier of CRISPR/Cas9 components was first synthesized, followed by the formation of plasmid DNA-polymer complex called polyplexes. 1H NMR, Zeta-Sizer, Scanning Electron Microscopy (SEM) analysis, and gel retardation experiments confirmed the polyplexes formation, including pRF4 (Roller) and sgRNA dpy-10, which were then incubated with C. elegans. The polymer-mediated delivery system facilitated the generation of transgenic Roller animals and heritable Dumpy mutants with CRISPR/Cas9. Our study for the first time demonstrated optimized administration of CRISPR/Cas 9 components to C. elegans.

Developing logic gate circuits that are programmable and reliable for specific functions is a central goal in synthetic biology. Traditional synthetic circuits often rely on protein regulators, which face scalability and resource burden limitations. To overcome these challenges, we introduce a novel approach utilizing dual-function and multi-level logic gates based on the CRISPRi system with dCas9 and sgRNA. This method, implemented in an in vitro transcription system, enables the rapid design and validation of complex logic gates. Our dual-function and multi-level design strategies achieve robust functionality in NOR, NAND, AND, and OR gates, allowing different logic outputs by altering only the inputs while reusing all other modules. This showcases significant advancements in efficiency and scalability for synthetic circuit construction. This work reduces development time and simplifies circuit design, paving the way for more efficient synthetic biology applications.

Reliable, efficient, and cheap detection of infectious diseases, especially in the wake of the SARS-CoV-2 pandemic, is of increasing importance. CRISPR/Cas systems have the capabilities to be optimized for this purpose. There is a broad diversity among Cas12a nucleases with immense detection capability, but only a few have been purified and studied biochemically. Here we present the investigation of 22 Cas12a orthologs, with a focus on their cis- and trans-cleavage activity, and thermal stability in combination with non-canonical crRNAs. We noticed that some non-canonical crRNA:Cas12a effector complexes outperformed its corresponding wild-type crRNA:Cas12a complex in trans-cleavage assays. In particular, TsCas12a, ErCas12a, and ArCas12a showed an increase in thermal stability in binary complex compared to its wild-type structure and apo form. Moreover, Cas12a was observed to have the ability to recruit segments of truncated crRNAs providing insights into crRNA:Cas12a catalytic complex with potential for further applications. We also discovered that ErCas12a, BsCas12a, BoCas12a, and TsCas12a possess robust trans-cleavage activity with a shorter PAM sequence requirement. Finally, we applied these effector complexes to discriminately detect SARS-CoV-2 and its B.1.1.7 lineage in clinical nasopharyngeal swabs, saliva samples, and tracheal aspirates. Our findings further expand the toolbox for next-generation CRISPR-based diagnostics.

Cyanophages are considered a promising biological management option for treating cyanobacterial blooms. Broadening the host range of cyanophages and/or shortening the lysis cycle by designing and synthesizing artificial cyanophages are potential strategies to enhance their effectiveness and efficiency. However, the rescue of artificial cyanophage genomes remains unexplored. In this study, we achieved the integration of a full-length cyanophage genome, PP, which originally infects Plectonema boryanum FACHB-240, into the model cyanobacterium Synechococcus elongatus PCC 7942. Since the integration of these large fragments ([~]42 kb) into cyanobacteria depended on conjugation via Escherichia coli, the toxic open reading frames (ORFs) of PP to E. coli were first identified, leading to the identification of toxic ORF6, ORF11, and ORF22. The original PP genome was then rearranged, and the three toxic ORFs were controlled using a tandem induction switch. The full length of the PP genome was integrated into the genome of S. elongatus PCC 7942 via two rounds of homologous recombination. Interestingly, compared to the control strain, the integration of the PP genome decreased photosynthesis and carbon fixation in S. elongatus PCC 7942, exhibiting cyanophage-like behavior. Transcriptomic analysis revealed that 32 of the 41 ORFs of the PP genome were transcribed in S. elongatus PCC 7942, significantly altering the energy metabolism and carbon fixation pathways. These influences were further demonstrated using metabolomics. This study provides a comprehensive approach for the artificial design and integration of cyanophage genomes in cyanobacteria, laying the foundation for their real rescue in the future.

Escherichia coli Nissle 1917 (EcN) is a probiotic microbe that has the potential to be developed as a promising chassis for synthetic biology applications. However, the molecular tools and techniques for utilizing EcN have not been fully explored. To address this opportunity, we systematically expanded the EcN-based toolbox, enabling EcN as a powerful platform for more applications. First, two EcN cryptic plasmids and other compatible plasmids were genetically engineered to enrich the manipulable plasmid toolbox for multiple gene coexpression. Next, we developed two EcN-based enabling technologies, including the conjugation strategy for DNA transfer, and quantification of protein expression capability. Finally, we expanded the EcN-based applications by developing EcN native integrase-mediated genetic engineering capabilities and establishing an in vitro cell-free protein synthesis (CFPS) system. Overall, this study expanded the toolbox for manipulating EcN as a commonly used probiotic chassis, providing several simplified, dependable, and predictable strategies for researchers working in synthetic biology fields. For Table of Contents Use Only O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=133 SRC="FIGDIR/small/543671v1_ufig1.gif" ALT="Figure 1"> View larger version (27K): org.highwire.dtl.DTLVardef@1f8e91forg.highwire.dtl.DTLVardef@90fb70org.highwire.dtl.DTLVardef@6badbforg.highwire.dtl.DTLVardef@15be840_HPS_FORMAT_FIGEXP M_FIG C_FIG

Unspecific peroxygenases (UPOs) are capable of catalyzing the selective oxidation of organic substrates under mild conditions, using hydrogen peroxide (H2O2) as the sole oxidant. This makes them one of the most promising biocatalysts for chemical synthesis. However, the major limitation restricting the application of UPOs to date is their difficulty in heterologous expression. Although more than 4,000 putative UPO enzymes have been recorded in databases, only about 50 of them can currently be heterologously expressed. All UPOs discovered so far originate from eukaryotes (mainly basidiomycetes and some ascomycetes), and they rely on the complex expression systems and post-translational modifications of their native hosts. This further exacerbates the challenges associated with heterologous expression of eukaryotic UPOs. In this work, we developed a deep-learning-based enzyme mining strategy and, for the first time, discovered novel UPO enzymes from bacteria, achieving successful heterologous expression in Escherichia coli. Bacterial UPOs differ greatly from fungal UPOs in sequence similarity, displaying completely distinct evolutionary trajectories. The discovery of bacterial UPOs advances our understanding of the catalytic mechanisms and expression characteristics within the UPO family, breaking the long-held assumption that UPOs can only originate from eukaryotes. Their excellent heterologous expression performance and broad catalytic versatility will further expand the application potential of UPOs.

Multiplex and precise control of the gene expression based on CRISPR/Cas9 is important to metabolic regulation in synthetic biology. However, employing single guide RNAs (sgRNAs) that possess repetitive DNA sequences and exhibit uniform activity could detrimentally affect the editing process, undermining both its stability and regulatory potential. In this study, we developed a deep generative model based on a decoder-only Transformer architecture (sgRNAGen) for the de novo generation of a series of nonrepetitive and diverse sgRNAs with activity. To assess the quality of sgRNAs generated by sgRNAGen, we evaluated their activity by targeting essential genes, with the results indicating that 98% of the generated sgRNAs were active in Bacillus subtilis. The generated sgRNAs were further validated for applications in single-gene editing, large fragment knockouts, and multiplex editing. Notably, the efficiency of knocking out long fragments up to 169.5 kb reached 100%, and targeting multiple sites allowed for the creation of strains with various combinations of mutations in a single editing. Furthermore, we developed a CRISPRi system utilizing the designed sgRNAs to regulate gene expression with desired strength and high precision. SgRNAGen offers a method for devising nonrepetitive and diverse activity sgRNAs, enhancing metabolic control and advancing applications within synthetic biology. TOC O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=112 SRC="FIGDIR/small/596019v1_ufig1.gif" ALT="Figure 1"> View larger version (30K): org.highwire.dtl.DTLVardef@f68faaorg.highwire.dtl.DTLVardef@151f5e2org.highwire.dtl.DTLVardef@1e5cb28org.highwire.dtl.DTLVardef@17cd3f8_HPS_FORMAT_FIGEXP M_FIG C_FIG

Corynebacterium glutamicum is a promising microbial chassis for the industrial production of L-tryptophan, which has exhibited increasing demand due to its diverse applications and high market value. In previous work, we developed an L-tryptophan-overproducing C. glutamicum strain TR26 through multiple rounds of rational metabolic engineering. Here, comparative transcriptome and metabolome analyses were conducted between TR26 and its progenitor strain MB001 to reveal the underlying mechanisms and potential bottlenecks for L-tryptophan production in TR26. Furthermore, by systematically down- and up-regulating differentially expressed genes of interest, two novel genetic targets, glnK and sugR, were identified as being associated with L-tryptophan synthesis. Specifically, the repression of glnK and overexpression of sugR in strain TR26 increased the titer of L-tryptophan by 6.7% and 20.9%, respectively. Gene transcription profiling and intracellular metabolite analysis further suggested that the observed improvements in L-tryptophan synthesis could be attributed to optimized nitrogen transport and metabolism, efficient reallocation of cellular resources and enhanced supply of phosphoenolpyruvate (PEP). This study advances our understanding of the regulation mechanisms governing L-tryptophan synthesis in C. glutamicum and provides valuable insights for further optimization of industrial cell factories.