assembling and annealing nucleic acid duplexes for characterizing enzymatic dna processing with modified oligonucleotides

Assays for DNA Enzymes Using Synthetic Oligonucleotides

All life forms require nucleic acid processing enzymes to carry out fundamental biological processes, including replication, transcription, and DNA repair. Key enzymatic functionalities for these pathways are polymerases, which generate copies of RNA/DNA molecules, ligases which join polynucleotide substrates, nucleases that degrade them, and helicases and topoisomerases, which melt nucleic acid duplexes or change their topology1,2,3,4,5,6,7,8,9,10. In addition, many of these enzymes provide essential molecular tools for applications such as cloning, diagnostics, and high-throughput sequencing11,12,13,14,15.
The functional characteristics, kinetics, and substrate specificities of these enzymes can be determined using labeled DNA/RNA substrates produced by annealing oligonucleotides. Tracking substrates and products has traditionally been achieved by introducing a radioactive label (32P) at either the 5&#39; strand end, which can then be detected by photographic film or with a phosphor imaging system16,17. While radiolabeled substrates offer the benefit of increased experimental sensitivity and do not alter the chemical properties of a nucleotide, the potential health hazards from working with radioisotopes have encouraged the development of non-radioactive nucleic acid labeling to provide a safer alternative for DNA and RNA detection18,19,20. Among these, fluorescence detection, including direct fluorescence detection, time-resolved fluorescence, and energy transfer/fluorescence quenching assays stand out as the most versatile21,22,23,24. The extensive array of fluorophores enables different designs of DNA/RNA substrates featuring unique reporters on each oligonucleotide25. Additionally, the stability of fluorophores, when compared to radioisotopes, allows users to produce and preserve significant quantities of fluorescently labeled DNA substrates19. These fluorophore-labeled substrates can be incubated with the protein of interest, along with different combinations of metal and nucleotide cofactors, to analyze binding and or enzyme activity. Visualization of binding or activity can be observed using various fluorophore dye channels with a gel imaging system. As only the fluorescently labeled oligonucleotides will be visible using this technique, any increase or decrease in the size of the labeled oligonucleotide will be easy to follow. Gels can also be stained afterward, with nucleic acid staining dyes to visualize all DNA bands present on the gel.
Poly-nucleic acid ligases are enzymes that join fragments of DNA/RNA, catalyzing the sealing of breaks by the formation of a phosphodiester bond between 5&#39; phosphorylated DNA termini and the 3&#39; OH of DNA. They can be divided into two groups according to their nucleotide substrate requirement. The highly conserved NAD-dependent ligases are found in all bacteria26 while the structurally diverse ATP-dependent enzymes can be identified through all domains of life8,27. DNA ligases play an important role in Okazaki fragment processing during replication as well as being involved in various DNA repair pathways, such as nucleotide and base excision repair, through the sealing of spontaneous nicks and nicks that are left after repair8,10. Different DNA ligases exhibit varying capacities to join different conformations of DNA breaks, including nicks in a duplex, double-stranded breaks, mismatches, and gaps, as well as RNA and DNA hybrids28,29,30. A diverse range of ligatable substrates can be assembled by annealing oligonucleotides with a 5&#39; phosphate to generate juxtaposed 5&#39; and 3&#39; termini in a nucleic acid duplex31,32,33. The most common method of analysis is resolution by urea PAGE in an end-point assay format; however, recent innovations have included the use of capillary gel electrophoresis, which allows high throughput34, mass-spectrometric profiling35, as well as a homogenous molecular beacon assay, which allows time-resolved monitoring36.
The first step in a ligation reaction is the adenylation of the ligase enzyme by adenosine triphosphate (ATP) or Nicotinamide adenine dinucleotide (NAD), resulting in a covalent enzyme intermediate. The second step in the reaction is adenylation of the nucleic acid substrate on the 5&#39; end of the nick site, which is followed by ligation of the nucleic acid nick strands. Many ligase enzymes that are recombinantly expressed in E. coli are purified in the adenylated form and, therefore, are able to successfully ligate nucleic acids without the addition of a nucleotide cofactor. This makes it difficult to determine what particular type of nucleotide cofactor they require for the ligation of nucleic acids. In addition to describing assays to evaluate DNA ligase activity, a method to reliably determine the cofactor usage by de-adenylating the enzyme using unlabeled substrates is also presented.
Nucleases are a large and diverse group of DNA/RNA modifying enzymes and catalytic RNAs that cleave the phosphodiester bonds between nucleic acids37. Nuclease enzyme functionalities are required in DNA replication, repair, and RNA processing and can be classified by their sugar specificity for DNA, RNA, or both. Endonucleases hydrolyze the phosphodiester bonds within a DNA/RNA strand, while exonucleases hydrolyze DNA/RNA strands one nucleotide at a time from the 3&#39; or 5&#39; end and may do so from either the 3&#39; to 5&#39; or the 5&#39; to 3&#39; end of the DNA38.
While many nuclease proteins are non-specific and may be involved in multiple processes, others are highly specific for a particular sequence or DNA damage6,39,40. Sequence-specific nucleases are used in a wide range of biotechnological applications, such as cloning, mutagenesis, and genome editing. Popular nucleases for these applications are restriction nucleases41, zinc-finger nucleases42, transcriptional activator-like effector nucleases, and most recently, the RNA-guided engineered CRISPR nucleases43. Damage-specific nucleases have recently been identified, such as the EndoMS nuclease, which has specificity for mismatches in the DNA through its mismatch-specific RecB-like nuclease domain5,44. Nuclease activity assays, historically, have been done as discontinuous assays with radiolabeled substrates; however, in addition to their other drawbacks, these do not allow the identification of the site that is cut by a nuclease protein, which is possible when using fluorescently labeled substrates45,46. More recently, continuous nuclease assays have been developed which work by using different DNA dyes that interact with DNA in different states; for example, emitting a higher fluorescent signal when interacting with dsDNA than in its unbound state, or binding specifically to short RNAs47. Other continuous nuclease assays use DNA hairpins with a fluorophore group on the 5&#39; and a quencher on the 3&#39; end so that fluorescence increases as the oligonucleotide is degraded due to a separation of the fluorophore and the quencher48.&#160;While these assays allow one to characterize the kinetics of DNA-degrading proteins, they require previous knowledge of the enzyme&#39;s function and substrate and are also limited to enzymes that change the DNA conformation to cause a difference in dye binding. For this reason, endpoint assays that resolve individual nuclease products are still desirable to provide insight into DNA modifications caused by protein activity.
Here, a detailed procedure is presented for the design of fluorescently labeled DNA/RNA oligonucleotides that can be mixed and matched to generate substrates for testing the activity of novel nuclease, polymerase, and ligase enzymes. The validation of this basic set of oligonucleotide sequences simplifies experimental design and facilitates economical profiling of a wide range of enzymatic functionalities without needing to purchase a large number of bespoke substrates. A detailed procedure is provided for running a standard DNA-processing enzyme assay with these substrates, using the example of DNA ligase activity and modifications for assaying and analyzing nuclease and polymerase enzymes are described. In addition, a modified assay for determining the cofactor specificity of the DNA ligase enzyme with high accuracy is given, and dual-labeled probes are used to evaluate the assembly of multi-component ligations. Finally, modifications to the basic assay format are discussed to allow it to be used to determine protein-DNA interactions with the same substrates by the electrophoretic mobility shift assay (EMSA).

Author Spotlight: Characterizing Novel Enzymes from Extremophiles and Common Pathogens to Understand DNA Repair and Replication

Using Modified Synthetic Oligonucleotides to Assay Nucleic Acid-Metabolizing Enzymes

So my research group, we're really interested in enzymes involved in DNA replication and repair. In particular, we're studying enzymes from extremophilic microorganisms and also from pathogens because we want to understand how these enzymes let those organisms survive environmental challenges. We're interested both in the biological function of the enzymes and also in the biotechnological applications.So we work a lot with DNA and RNA ligases, and we've recently established the role of a minimal type of DNA ligase which seems to be trafficked to the outside of bacterial cell. And we've determined that this plays a role in biofilm formation in the pathogen Neisseria gonorrhoeae. We've also, together with our collaborators from ArcticZymes characterized a novel RNA-to-DNA ligase which is able to attach DNA adapters to either end of a piece of RNA.So our collaborators at ArcticZymes, who've also contributed to this protocol, are characterizing different DNA and RNA-metabolizing enzymes which have potential applications in molecular biology. And so they're using this type of protocol as a first step for understanding the spectrum of activities these enzymes have and benchmarking them relative to what's already available on the market. The protocol offers an easier and safer alternative for assaying nucleic acid metabolizing enzymes compared to the use of radiolabeled substrates.The fluorescently labeled DNA or RNA substrates make our approach more efficient, as we can mix and match the oligonucleotides to make a variety of substrates. So at the moment, we're really focused on understanding the biological function of some novel nucleases that we've discovered in an Antarctic environmental genome. These nucleases have homologs in other extremophilic organisms, and so we're trying to understand what role they play in allowing these microbes to survive.We're also really interested in the activity of some of our novel enzymes on non-natural nucleic acid analogs and their potential uses as part of a molecular biology toolkit with non-natural DNA and RNA substrates.

Assembling and Annealing Nucleic Acid Duplexes for Characterizing Enzymatic DNA Processing with Modified Oligonucleotides

To begin, centrifuge the lyophilized oligonucleotide-containing tube at full speed in a benchtop centrifuge for two to five minutes. Resuspend the oligonucleotides in TE buffer to prepare a master stock of 100 micromolar and gently vortex the tube. Dilute an aliquot of master stock with TE buffer to prepare a 10-micromolar stock and prepare the reaction master mix.After dispensing the mix to the required tubes, anneal the oligonucleotides in a thermocycler at 95 degrees Celsius for five minutes. Let the tube cool to room temperature for 30 minutes to one hour. Then add nucleotide co-factors and other heat-sensitive buffer components to the master mix.If required, store the mix at 20 degrees Celsius for future use. Combine 22.5 microliters of the substrate master mix with 2.5 microliters of the DNA ligase in a PCR tube. Immediately place the reaction tubes in a PCR machine at 25 degrees Celsius for 30 minutes.To quench the reaction, add five microliters of loading dye and incubate at 95 degrees Celsius for five minutes.

assembling-annealing-nucleic-acid-duplexes-for-characterizing

Research

JoVE Journal

Biochemistry

34 vistas. Para comenzar, centrifuga el tubo que contiene oligonucleótidos liofilizados a toda velocidad en una centrífuga de sobremesa durante dos a cinco minutos. Vuelva a suspender los oligonucleótidos en el tampón TE para preparar un material maestro de 100 micromolares y vórtice suavemente el tubo. Diluir una alícuota de material maestro con tampón TE para preparar un material de 10 micromolares y preparar la mezcla maestra de reacción.Después de di...