Molecular Biology

Nanobodies are recombinant single-domain antibodies (VHHs) derived from the heavy chain–only subset of camelid immunoglobulins that can be reverse-engineered into bivalent antibodies by fusion to immunoglobulin Fc constant regions. Mammalian cells are the system of choice to produce VHH-Fcs to ensure authentic folding and post-translation glycosylation of the expressed VHH-Fcs. In a recent project to find neutralising VHH-Fc binders to the spike proteins of SARS-CoV-2 viruses, we identified a need for rapid expression and purification of multiple VHH-Fc fusions from nanobodies selected by phage display. Here, we present a protocol for the construction of expression vectors by parallel ligase-independent cloning, transient small-scale expression in mammalian cells (4 mL culture volume), screening antigen-binding activity, and midi-scale purification (30 mL culture volume) for downstream activity assays. The workflow is completely transferable between different vector formats, of which three are described herein: Fc fusion dimers, monomeric CD4 fusions, and His-tagged monomers.

While cell hashing enhances single-cell RNA sequencing (scRNA-seq) efficiency and minimizes batch effects, commercial mouse hashtags often fail in FVB/N and several other strains due to antibody-epitope incompatibility. We describe a robust alternative utilizing biotinylated antibody cocktails and streptavidin-conjugated oligos to enable reliable sample multiplexing. This approach was validated in FVB/N lung tissues, yielding high-quality single-cell libraries. Our protocol offers a practical solution for researchers requiring strain-specific or custom-designed multiplexing strategies for single-cell transcriptomics.

While traditional kinetic studies of the cytochrome b₆f complex have frequently relied on measurements within the complex environment of intact leaves or whole-organism systems, such approaches can be limited by overlapping signals and physiological variables. This protocol advances existing frameworks by introducing a streamlined, multi-wavelength spectroscopic approach utilizing a reconstituted in vitro system to elucidate the inter-complex electron transfer kinetics between photosystem I and cytochrome b₆f. Utilizing the JTS-150 pulsed spectrometer, supplied with a Smart Lamp, we monitored the redox transitions of P700⁺ and Cytf by simultaneously measuring the absorbance changes of our isolated complexes system in six different wavelengths (546, 554, 563, 574, 705, and 740 nm). Kinetic analysis was divided into two phases: laser-induced flash kinetics and steady-state actinic induction. We resolved the second-order re-reduction of P700⁺ by plastocyanin, accounting for detector saturation constraints with a 2 ms post-flash delay. Steady-state measurements under actinic light revealed complex Cytf turnover, characterized by a double-exponential decay. Furthermore, dark relaxation kinetics were used to quantify ferredoxin-mediated re-reduction of the cytochrome pool. By allowing the incorporation of specific regulatory and inhibitory factors, this methodology sets the ground for the deconvolution of competing electron pathways. It can therefore be used as a robust framework for assessing the mechanism of regulatory processes on photosynthetic flux.

Plant genome editing is a powerful approach for modifying plant DNA to investigate gene function and to engineer desirable traits. Several genome-editing technologies have been developed, among which CRISPR/Cas systems and transcription activator-like effector nucleases (TALENs) are widely used to introduce targeted double-stranded DNA breaks. While CRISPR/Cas systems are highly efficient for nuclear genome editing, their application to plant organellar genomes remains limited, largely due to difficulties in guide RNA delivery into mitochondria and chloroplasts. Here, we present a detailed and reproducible protocol for constructing TALEN-based binary vectors for targeted genome editing in Arabidopsis thaliana. This protocol describes the assembly of TALE repeat arrays, the generation of nuclear-, mitochondrial-, and plastid-targeted TALEN expression vectors using MultiSite Gateway cloning, and subsequent Agrobacterium-mediated plant transformation and genotyping. The workflow enables the production of nTALENs, mitoTALENs, and ptpTALENs using a unified vector design strategy. In addition, the protocol briefly outlines the construction principles of TALE-based cytidine deaminases (TALECDs) for targeted C-to-T base editing in plant organellar genomes. The protocol provides a flexible and robust framework for plant nuclear and organellar genome editing and can be readily adapted to different target genes and experimental purposes. Its modular design and compatibility with standard molecular cloning techniques make it accessible to laboratories aiming to perform precise genome manipulation in plants.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.

Protein–protein interactions (PPIs) govern nearly all aspects of cellular physiology, yet identifying these interactions under native conditions remains challenging. Here, we present TIE-UP-SIN (targeted interactome experiment for unknown proteins by stable isotope normalization), a robust method for in vivo identification and quantification of PPIs in bacterial systems. The protocol combines metabolic labeling with ¹⁵N isotopes, reversible formaldehyde crosslinking, affinity purification, and quantitative mass spectrometry. TIE-UP-SIN preserves transient or weak interactions during purification and quantifies interaction partners using internal light/heavy peptide ratios, reducing experimental variability. The method employs a triple-sample design to distinguish specific from nonspecific interactors and can be adapted to various bacterial species and affinity tags. Data analysis is streamlined through a user-friendly web application (https://shiny-fungene.biologie.uni-greifswald.de/TIE_UP_SIN_app) that automates statistical analysis, normalization, and visualization, requiring no programming expertise. The entire workflow from cell culture to mass spectrometry data acquisition takes approximately 4–5 days, with data analysis completed in 1–2 days using the web application.

We established a step-by-step approach for generating a single-nucleotide mutation in the promoter region of an immune regulatory gene in human monocyte THP-1 cells by employing a plasmid-based CRISPR-Cas9 system delivered via transfection with a homology-directed repair template DNA (HDR). Key steps include designing a single-guide RNA (sgRNA), cloning it into a CRISPR plasmid encoding the Cas9 protein, transfection of the plasmid constructs along with single-stranded oligonucleotide repair template (ssODNs) into THP-1 cells, followed by selection and validation. This approach provides a precise and relevant model to investigate the role of single polymorphisms in the regulation of inflammatory gene expression in human monocytes. In addition to the rs1024611 single-nucleotide polymorphism (SNP), this CRISPR/Cas9-based strategy is broadly applicable to functional studies of noncoding and coding variants in innate immune genes.

Cyanobacteria have been widely used as model organisms in photobiochemical research and have recently been exploited as hosts in numerous pilot studies to produce valuable biochemicals via genetic and metabolic modifications. Analyzing cellular RNA is a suitable method for studying genetic changes in cells. Several methods have previously been reported for cyanobacterial RNA extraction. However, the majority of these methods rely heavily on phenol and chloroform, which are hazardous. Additionally, these methods are time-consuming and difficult to perform. Using Synechocystis sp. PCC 6803 as a model, this study developed a novel method for extracting total ribonucleic acid (RNA) using standard centrifugation techniques and laboratory chemicals such as citric acid, ethylenediaminetetraacetic acid, sodium dodecyl sulfate, sodium chloride, and tri-sodium citrate dihydrate to extract RNA from cyanobacterial cells. This method does not necessitate the use of hazardous chemicals, especially phenol and chloroform. Furthermore, it is cost-effective since it does not require expensive chemicals. The results of the quantification, purity, and integrity checks show the effectiveness of this method for extracting good-quality RNA. Furthermore, RT-qPCR results demonstrate that the quality of the extracted RNA is suitable for downstream applications.

Human tissue samples represent the gold standard for obtaining clinically relevant and translatable insight into disease processes that in vitro systems cannot fully reproduce. However, patient-derived samples are often limited in size and availability, limiting the number of downstream assays that can be performed. To maximize the use of invaluable human samples, we present a protocol for the tandem extraction of high-quality RNA and protein from the same tissue section. This method has been optimized for 15–30 mg tissue sections, enabling more experimental conditions and technical replicates, while minimizing intrasample variability associated with heterogeneous tissues. This protocol also avoids potentially hazardous solvents present in phenol-chloroform-based methods such as TRIzol, providing a safer and more accessible workflow without compromising biomolecule integrity. This protocol was developed and validated using atherosclerotic plaque tissue from carotid endarterectomy, a very challenging tissue type to work with due to extensive calcification, necrosis, and limited surgical availability. We have also validated this method using mouse aortic tissue and cultured THP-1 cells, demonstrating its versatility across sample input types. As this protocol relies on standard column-based RNA extraction kits and commonly available reagents for protein precipitation and extraction, this methodology is widely accessible and easy to implement as a standard, streamlined workflow.

Transcription factors (TFs) regulate gene expression by binding to cis-regulatory elements in the genome. Understanding transcriptional regulation requires genome-wide characterization of TF occupancy across different chromatin contexts, yet simultaneous assessment of TF binding for multiple factors remains technically challenging. Here, we describe a detailed and reproducible protocol for cFOOT-seq, a cytosine deaminase–based genomic footprinting assay by sequencing, which enables antibody-independent, base-resolution profiling of chromatin accessibility, nucleosome organization, and TF occupancy. In cFOOT-seq, the double-stranded DNA (dsDNA) cytosine deaminase SsdA_tox converts cytosine to uracil in accessible chromatin, whereas TF binding and nucleosome occupancy locally protect DNA from deamination. Using the FootTrack analysis framework, deamination patterns generated by cFOOT-seq are quantitatively analyzed to derive standardized footprint and chromatin organization profiles at base resolution across the genome. Because cFOOT-seq preserves genomic DNA integrity during deamination-based footprinting, it is compatible with ATAC-seq-based chromatin enrichment. ATAC-combined implementations reduce sequencing depth requirements and improve scalability for footprint-focused analyses, supporting applications in low-input and single-cell settings. This protocol provides a practical framework for genome-wide TF footprint profiling and can be readily applied to dissect gene regulatory mechanisms in development, immunity, and disease, including cancer.