Molecular Biology

Telomeres are structures that cap the ends of linear chromosomes and play critical roles in maintaining genome integrity and establishing the replicative lifespan of cells. In stem and cancer cells, telomeres are actively elongated by either telomerase or the alternative lengthening of telomeres (ALT) pathway. This pathway is characterized by several hallmark features, including extrachromosomal C-rich circular DNAs that can be probed to assess ALT activity. These so-called C-circles are the product of ALT-associated DNA damage repair processes and simultaneously serve as potential templates for iterative telomere extension. This bifunctional nature makes C-circles highly sensitive and specific markers of ALT. Here, we describe a C-circle assay, adapted from previous reports, that enables the quantitation of C-circle abundance in mammalian cells subjected to a wide range of experimental perturbations. This protocol combines the Quick C-circle Preparation (QCP) method for DNA isolation with fluorometry-based DNA quantification, rolling circle amplification (RCA), and detection of C-circles using quantitative PCR. Moreover, the inclusion of internal standards with well-characterized telomere maintenance mechanisms (TMMs) allows for the reliable benchmarking of cells with unknown TMM status. Overall, our work builds upon existing protocols to create a generalizable workflow for in vitro C-circle quantitation and ascertainment of TMM identity.

Malaria molecular surveillance has great potential to support national malaria control programs (NMCPs), informing policy for its control and elimination. Here, we present a new three-day workflow for targeted resequencing of markers in 13 resistance-associated genes, histidine rich protein 2 and 3 (hrp2&3), a country (Peru)-specific 28 SNP-barcode for population genetic analysis, and apical membrane antigen 1 (ama1), using Illumina short-read sequencing technology. The assay applies a multiplex PCR approach to amplify all genomic regions of interest in a rapid and easily standardizable procedure and allows simultaneous amplification of a high number of targets at once, therefore having great potential for implementation into routine surveillance practice by NMCPs. The assay can be performed on routinely collected filter paper blood spots and can be easily adapted to different regions to investigate either regional trends or in-country epidemiological changes.

Nucleotide excision repair (NER) removes a wide variety of structurally unrelated lesions from the genome, including UV-induced photolesions such as 6–4 pyrimidine–pyrimidone photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs). NER removes lesions by excising a short stretch of single-stranded DNA containing the damaged DNA, leaving a single-stranded gap that is resynthesized in a process called unscheduled DNA synthesis (UDS). Measuring UDS after UV irradiation in non-dividing cells provides a measure of the overall NER activity, of which approximately 90% is carried out by the global genome repair (GGR) sub pathway. Here, we present a protocol for the microscopy-based analysis and quantification of UDS as a measurement for GGR activity. Following local UV-C irradiation, serum-starved human cells are supplemented with the thymidine analogue 5-ethynyl-2'-deoxyuridine (EdU), which is incorporated into repair patches following NER-dependent dual incision. The incorporated nucleotide analogue is coupled to a fluorophore using Click-iT chemistry, followed by immunodetection of CPD photolesions to simultaneously visualize both signals by fluorescence microscopy. Accompanying this protocol is a custom-built ImageJ plug-in to analyze and quantify UDS signals at sites of CPD-marked local damage. The local UDS assay enables an effective and sensitive fluorescence-based quantification of GGR activity in single cells with application in basic research to better understand the regulatory mechanism in NER, as well as in diagnostics to characterize fibroblasts from individuals with NER-deficiency disorder.

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CRISPR/Cas9 screening has revolutionized functional genomics in biomedical research and is a widely used approach for the identification of genetic dependencies in cancer cells. Here, we present an efficient and versatile protocol for the cloning of guide RNAs (gRNA) into lentiviral vectors, the production of lentiviral supernatants, and the transduction of target cells in a 96-well format. To assess the effect of gene knockouts on cellular fitness, we describe a competition-based cell proliferation assay using flow cytometry, enabling the screening of many genes at the same time in a fast and reproducible manner. This readout can be extended to any parameter that is accessible to flow-based measurements, such as protein expression and stability, differentiation, cell death, and others. In summary, this protocol allows to functionally assess the effect of a set of 50–300 gene knockouts on various cellular parameters within eight weeks.

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Cloning systems like Gateway and Golden Gate/Braid are known because of their efficiency and accuracy. While the main drawback of Gateway is the expensive cost of the enzymes used in its two-step (LR and BP) reaction, Golden Gate requires non-reusable components due to their specific restriction sites. We present the Brick into the Gateway (BiG) protocol as a new cloning strategy, faster and more economic method that combines (i) reusable modules or bricks assembled by the GoldenBraid approach, and (ii) Gateway LR reactions [recombination of attachment sites: attL (L from left) and attR (R from right)] avoiding the BP reaction [recombination of attachment sites: attP (P from phage) and attB (B from bacteria)] usually necessary in the Gateway cloning. The starting point is to perform a PCR reaction to add type IIS restriction sites into DNA fragments generating specific fusion sites. Then, this PCR product is used to design GoldenBraid bricks, including the attL Gateway recombination sites. Using the Golden Gate method, these bricks are assembled to produce an attL1–gene of interest–attL2 fragment, which is integrated into a compatible vector producing a Gateway entry vector. Finally, the fragment containing the target gene is recombined by LR reaction into the Gateway destination vector.

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8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) is considered to be a premutagenic DNA lesion generated by 2'-deoxyguanosine (dG) oxidation due to reactive oxygen species (ROS). In recent years, the 8-oxodG distribution in human, mouse, and yeast genomes has been underlined using various next-generation sequencing (NGS)–based strategies. The present study reports the OxiDIP-Seq protocol, which combines specific 8-oxodG immuno-precipitation of single-stranded DNA with NGS, and the pipeline analysis that allows the genome-wide 8-oxodG distribution in mammalian cells. The development of this OxiDIP-Seq method increases knowledge on the oxidative DNA damage/repair field, providing a high-resolution map of 8-oxodG in human cells.

Directed evolution is a powerful technique for identifying beneficial mutations in defined DNA sequences with the goal of improving desired phenotypes. Recent methodological advances have made the evolution of short DNA sequences quick and easy. However, the evolution of DNA sequences >5kb in length, notably gene clusters, is still a challenge for most existing methods. Since many important microbial phenotypes are encoded by multigene pathways, they are usually improved via adaptive laboratory evolution (ALE), which while straightforward to implement can suffer from off-target and hitchhiker mutations that can adversely affect the fitness of the evolved strain. We have therefore developed a new directed evolution method (Inducible Directed Evolution, IDE) that combines the specificity and throughput of recent continuous directed evolution methods with the ease of ALE. Here, we present detailed methods for operating Inducible Directed Evolution (IDE), which enables long (up to 85kb) DNA sequences to be mutated in a high throughput manner via a simple series of incubation steps. In IDE, an intracellular mutagenesis plasmid (MP) tunably mutagenizes the pathway of interest, located on the phagemid (PM). MP contains a mutagenic operon (danQ926, dam, seqA, emrR, ugi, and cda1) that can be expressed via the addition of a chemical inducer. Expression of the mutagenic operon during a cell cycle represses DNA repair mechanisms such as proofreading, translesion synthesis, mismatch repair, and base excision and selection, which leads to a higher mutation rate. Induction of the P1 lytic cycle results in packaging of the mutagenized phagemid, and the pathway-bearing phage particles infect naïve cells, generating a mutant library that can be screened or selected for improved variants. Successive rounds of IDE enable optimization of complex phenotypes encoded by large pathways (as of this writing up to 36 kb), without requiring inefficient transformation steps. Additionally, IDE avoids off-target genomic mutations and enables decoupling of mutagenesis and screening steps, establishing it as a powerful tool for optimizing complex phenotypes in E. coli.

Graphical abstract:

Figure 1. Overview of Inducible Directed Evolution (IDE).

Pathways of interest are cloned into a P1 phagemid (PM) backbone and transformed into a strain of E. coli containing MP (diversification strain). The mutagenesis plasmid is induced to generate mutations. Phage lysate is produced and used to infect a strain that expresses the phenotype of interest (screening/selection strain). The resulting strain library is screened to identify those with improved properties. Narrowed-down libraries can then go through another IDE cycle by infecting a fresh diversification strain.

R-loops, or RNA:DNA hybrids, are structures that arise co-transcriptionally when a nascent RNA hybridizes back with the template ssDNA, leading to a displaced ssDNA. Because accumulation of R-loops can lead to genomic instability and loss of cellular homeostasis, it is important to determine the genome-wide distribution of R-loops in different physiological conditions. Current R-loop mapping strategies are based on R-loop enrichment—mediated by the S9.6 antibody, such as DRIP-seq, or by the exonuclease RNase H1, such as MapR—or the latest R-loop CUT&Tag, based on an artificial R-loop sensor derived from an RNase H1 sub-domain. Because some of these techniques often require high input material or expensive reagents, we sought to apply MapR, which does not require expensive reagents and has been shown to be compatible with low input samples. Importantly, we demonstrate that incorporation of improved CUT&RUN steps into the MapR protocol yields R-loop-enriched DNA when using low input Drosophila nuclei.

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Workflow for mapping tissue-specific, genome-wide R-loops in Drosophila.

Purify GST-tagged and catalytically inactive RNase H1 tethered MapR enzymes, GST-ΔRH-MNase, and GST-MNase, from transformed E. coli. Perform tissue-specific nuclei immuno-enrichment from UAS-EGFP.KASH-Msp300 Drosophila using magnetic bead–bound green fluorescent protein (GFP) antibody. Incubate isolated nuclei with MapR enzymes and activate MNase DNA cleavage with low salt/high calcium buffers. Purify released, R-loopenriched DNA fragments and generate sequencing-ready libraries. Align MapR data to reference genome and compare R-loop enrichment peaks in genome browser.

Nucleic acids in living organisms are more complex than the simple combinations of the four canonical nucleotides. Recent advances in biomedical research have led to the discovery of numerous naturally occurring nucleotide modifications and enzymes responsible for the synthesis of such modifications. In turn, these enzymes can be leveraged towards toolkits for DNA and RNA manipulation for epigenetic sequencing or other biotechnological applications. Here, we present the protocol to obtain purified 5-hydroxymethylcytosine carbamoyltransferase enzymes and the associated assays to convert 5-hydroxymethylcytosine to 5-carbamoyloxymethylcytosine in vitro. We include detailed assays using DNA, RNA, and single nucleotide/deoxynucleotide as substrates. These assays can be combined with downstream applications for genetic/epigenetic regulatory mechanism studies and next-generation sequencing purposes.

Geobacillus kaustophilus, a thermophilic Gram-positive bacterium, is an attractive host for the development of high-temperature bioprocesses. However, its reluctance against genetic manipulation by standard methodologies hampers its exploitation. Here, we describe a simple methodology in which an artificial DNA segment on the chromosome of Bacillus subtilis can be transferred via pLS20-mediated conjugation resulting in subsequent integration in the genome of G. kaustophilus. Therefore, we have developed a transformation strategy to design an artificial DNA segment on the chromosome of B. subtilis and introduce it into G. kaustophilus. The artificial DNA segment can be freely designed by taking advantage of the plasticity of the B. subtilis genome and combined with the simplicity of pLS20 conjugation transfer. This transformation strategy would adapt to various Gram-positive bacteria other than G. kaustophilus.

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