Использование модифицированных синтетических олигонуклеотидов для анализа ферментов, метаболизирующих нуклеиновые кислоты

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 topology^{1,2,3,4,5,6,7,8,9,10}. In addition, many of these enzymes provide essential molecular tools for applications such as cloning, diagnostics, and high-throughput sequencing^{11,12,13,14,15}.

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' strand end, which can then be detected by photographic film or with a phosphor imaging system^16,17. 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 detection^18,19,20. Among these, fluorescence detection, including direct fluorescence detection, time-resolved fluorescence, and energy transfer/fluorescence quenching assays stand out as the most versatile^21,22,23,24. 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' phosphorylated DNA termini and the 3' 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 life^8,27. 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 repair^8,10. 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 hybrids^28,29,30. A diverse range of ligatable substrates can be assembled by annealing oligonucleotides with a 5' phosphate to generate juxtaposed 5' and 3' termini in a nucleic acid duplex^31,32,33. 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' 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' or 5' end and may do so from either the 3' to 5' or the 5' to 3' 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 damage^6,39,40. 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 domain^5,44. 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 substrates^45,46. 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' and a quencher on the 3' end so that fluorescence increases as the oligonucleotide is degraded due to a separation of the fluorophore and the quencher⁴⁸. While these assays allow one to characterize the kinetics of DNA-degrading proteins, they require previous knowledge of the enzyme'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).