Accurate delineation of bacterial translation initiation sites (TISs) remains a major challenge, as conventional genome annotation and ribosome profiling (Ribo-seq) often lack the resolution to discriminate closely spaced start codons. To overcome these limitations, we developed TRAINSPOTTER (TRAnslation INitiation SPOTTER), a deformylation-assisted N-terminomics workflow that enables direct, proteome-wide detection of nascent N-termini indicative of active translation initiation. TRAINSPOTTER exploits the universal N-terminal formylation of initiator methionine in bacteria: in vitro enzymatic deformylation by peptide deformylase (PDF) generates a diagnostic hydrophilic shift, allowing selective isolation of previously formylated, initiation-derived peptides by COFRADIC-based chromatography. Optional in vivo PDF inhibition transiently enriches formylated N-termini, primarily enhancing detection sensitivity. Integration of pulse SILAC (Stable Isotope Labeling by Amino acids in Cell Culture) labeling confirmed that deformylation-shifted peptides represent newly synthesized N-termini, validating TRAINSPOTTER’s specificity for nascent translation products. Application to E. coli enabled precise mapping of >1000 TIS-indicative N-termini, including numerous alternative and near-cognate start sites, providing direct proteomic evidence for co-expressed N-terminal proteoforms. The method complements and refines Ribo-seq datasets, offering amino acid-level resolution for otherwise ambiguous initiation events. TRAINSPOTTER thus establishes a robust biochemical framework for proteome-wide identification of TISs and advances the experimental annotation of bacterial proteomes.

Endophytic fungi residing within medicinal plants represent a rich and sustainable source of bioactive secondary metabolites with diverse pharmacological applications. These symbiotic microorganisms establish mutualistic relationships with their host plants, contributing to enhanced stress tolerance, growth promotion, and defense against pathogens. In recent years, endophytic fungi have gained considerable attention as alternative biofactories for the production of valuable compounds such as alkaloids, terpenoids, phenolics, lignans, and polysaccharides. These metabolites exhibit a wide range of biological activities, including antimicrobial, antiviral, anticancer, antioxidant, anti-inflammatory, and antidiabetic effects. Advances in cultivation strategies, such as OSMAC, co-culture, and epigenetic modification, have significantly improved metabolite yield and diversity. Moreover, the integration of genomics, transcriptomics, and metabolomics has revolutionized the understanding of biosynthetic gene clusters and metabolic pathways, enabling the discovery of novel compounds and optimization of production processes. Despite these advances, challenges such as low yield, silent gene clusters, and difficulties in large-scale production remain significant barriers. Nevertheless, continued progress in multi-omics technologies, synthetic biology, and biotechnological tools holds great promise for unlocking the full potential of endophytic fungi. Herein, endophytic fungi represent a powerful and eco-friendly platform for the development of new therapeutic agents and sustainable pharmaceutical applications.

Streptomycetes are prolific producers of bioactive natural products, but many of the biosynthetic gene clusters (BGCs) are silent in the laboratory. Genetic manipulation is important to unlock their full potential. CRISPR–Cas-based genome editing has greatly advanced genetic engineering in Streptomyces. However, several challenges remain, including Cas nuclease toxicity, unintended genomic rearrangements, and elimination of the delivery plasmid. Here, we present a novel genome editing strategy that harnesses cumate-inducible CRISPR interference (CRISPRi) to transiently knockdown essential genes such as divIVA or dnaA as counterselectable marker. This enforces loss of the vector backbone, promotes homologous recombination, and yields markerless mutants by loss of the antibiotic resistance cassette during the final recombination step. We demonstrate the versatility of the ICE system (Inducible CRISPRi targeting an Essential gene) by (i) deleting four BGCs in Streptomyces coelicolor M145, (ii) inserting both a promoter and a large BGC, and (iii) introducing precise single-nucleotide substitutions. Furthermore, deletion of the prodigiosin BGC elicited expression of a poorly expressed BGC for prolinolexin lipopeptides in Streptomyces roseifaciens DSM 106196T. Considering that different essential genes may be targeted, we anticipate that inducible CRISPRi-based counterselection may be adaptable to genome editing strategies in a broad range of microbial systems.

Base editing shows great potential in research and clinical applications. Current iterations of the deaminases used to create precise single-nucleotide changes via base editing exhibit undesirable effects, including off-targeting, off-base editing, and bystander editing. Current deaminases are derived from either larger eukaryotic deaminases, which exhibit high levels of Cas-independent DNA targeting, or from evolved variants of the smaller E. coli TadA protein (ecTadA), which exhibits off-base editing. To overcome the limitations inherent to using a single protein sequence for engineering, we diversified newly identified TadA orthologs by DNA shuffling to yield millions of training sequences for measuring base editor efficiency. We trained generative models on the performance data from the pools of variants and drew on information-theoretic insights to efficiently explore the sequence space to generate diverse and high-performing deaminases. From a single round of diversification, we created a small set of novel and specific cytosine and adenosine deaminases that were markedly distinct in sequence from published base editor deaminases. We found that our model-created deaminases generally outperform those we identified through typical directed evolution. The novel compact deaminases identified here show high on-base activity, comparable to the leading published base editors, and with demonstrably lower off-base activity.

Chemically inducible expression systems enable transgene expression regulation in response to external small molecules. Tetracycline repressor (TetR)-based gene switches work in plants, but antibiotics are neither approved nor advisable for crop use. Here we report engineering of TetR mutants that respond to approved sulfonylurea (SU) herbicides instead of antibiotics. Designed variants show low-nanomolar EC50 values for ethametsulfuron-methyl (Es) or chlorsulfuron and tightly bind the Tet operator sequence, but only in the absence of corresponding SUs. Crystal structures of two repressors in complex with their respective SU ligands reveal extensive interactions explaining their strong binding. The Es repressor-based gene switch is introduced into tobacco, soybean, maize, rice, and Arabidopsis, and robust reporter gene activation is observed upon herbicide application. Addition of a repressor-regulated siRNA targeting the repressor transcript increases the magnitude and spatial distribution of the response following herbicide treatment and results in a partially bistable gene switch. The SU repressors also function well in mammalian cell culture and may enable regulation of additional genes in conjunction with TetR. Practical chemically inducible expression systems for crop plants remain lacking. Here, the authors report the design and evolution of the tetracycline repressor into sulfonylurea herbicide-responsive gene switches and show their application in multiple crops as well as mammalian cell.