AW is supported by a Rutherford Discovery Fellowship (20-UOW-004). RS is the recipient of a New Zealand Post Antarctic Scholarship. SG and UR acknowledge the Chemical Institute at the University of Troms&#248; - The Norwegian Arctic University for technical support.

1. Design and purchase of oligonucleotides
NOTE: Design single-stranded oligonucleotides to be assembled and annealed into the desired duplexes. One or more of the strands in a duplex must bear a fluorescent moiety for tracking oligonucleotide processing by the enzyme of interest. A basis set of single-stranded sequences that can be assembled for a range of activities is provided in Table 1.
<ol>
	<li>Incorporate the specific modifications needed for the enzyme of interest as described below.
	<ol>
		<li>For DNA ligase substrates (Figure 1): Assemble the simplest substrate from three oligonucleotides: a 5&#39; phosphorylated donor strand (NL2), a 5&#39; FAM-labeled acceptor strand (NL1), and a complement that bridges the two (NL3).
		<ol>
			<li>Ensure strands providing the 5&#39; terminus of the ligatable nick are phosphorylated before assembly of the substrate master mix in step 2. Order this as a modification on NL2 (as given in Table 1) or use enzymatic phosphorylation with T4 polynucleotide kinase after resuspending the oligonucleotides.</li>
			<li>Include 5&#39;-terminal phosphorylation of NL6 and NL8, which comprise the complement of double-stranded breaks depicted in Figure 1A (NL6/NL7 and NL8/NL9) as this most closely resembles the natural substrate produced from a restriction endonuclease. Use a dual-labeled substrate to determine the relative extents of ligation for multi-part assemblies (see step 6).</li>
			<li>Alter the complement strand to produce mismatches (NL10) and gaps (NL11). 
			NOTE: Variations on the simple nicked substrate are depicted in Figure 1A. It is possible to use other sequences to produce an even wider range of mismatches or longer gaps by varying the underlined position.</li>
			<li>Substitute DNA oligonucleotides for RNA oligonucleotides. 
			NOTE: Variations on the simple nicked substrate are depicted in Figure 1B. A wider range of DNA/RNA duplexes can be generated by additional combinations of the basis set given here to generate, for example, double-stranded breaks containing both RNA and DNA. An example of this variation is given in step 6 below, where a dual-label strategy is used.</li>
		</ol>
		</li>
		<li>For DNA polymerase substrates: Assemble the oligonucleotides NL1 and NL3 listed in Table 1 to give a simple primer-extension assay. Investigate additional aspects of polymerase activity by introducing modifications into either the NL1 (primer) or NL3 (template) strands.
		<ol>
			<li>Incorporate damaged base analogs into the NL3 oligonucleotide prior to position 20 to determine the ability to bypass damaged lesions on the template strand.</li>
			<li>Incorporate damaged base analogs into the NL1 oligonucleotide at position 20 to determine the ability to extend a damaged primer.</li>
			<li>Use either RNL1 or RNL3 in the duplex to investigate the extension of an RNA primer or the use of an RNA template.</li>
		</ol>
		</li>
		<li>For nuclease substrates (Figure 2): Assemble oligonucleotides to give a non-exhaustive range of double and single-stranded substrates (Figure 2Ai) as well as a range of flapped and splayed junctions (Figure 2Aii) and damaged substrates (Figure 2B).
		<ol>
			<li>To probe ribonuclease activities, iteratively substitute NL1, NL2, and NL3 with RNL1, RNL2, and RNL3. Use additional RNA versions of HJ5 and HJ6 to expand this set further.</li>
			<li>Use oligonucleotides MD5, MD6, and MD9 that have a centrally placed modification that mimics oxidative damage, an abasic repair intermediate, or a deamination product (Figure 2B). The substrates will detect cleavage of the strand at this position. Label the complement NL3 strand with an orthogonal fluorophore such as TAMRA to detect double-strand cleavage (see step 6).</li>
			<li>Use orthogonal labeling of the complement to detect double-stranded cutting at the mismatched sites on both the probe (NL5 and ND9) and the complement (MD10 and NL10) strands.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Order synthetic oligonucleotides incorporating relevant fluorophores and other modifications from a commercial vendor. 
	NOTE: A 100 nM synthesis scale and HPLC purification subsequent to synthesis are suitable for the assays described.</li>
</ol>
2. Assembling and annealing nucleic acid duplexes
<ol>
	<li>Resuspension and dilution of oligonucleotides
	<ol>
		<li>Before opening, centrifuge the lyophilized oligonucleotides in their 2 mL tubes at full speed in a benchtop centrifuge for 2-5 min to ensure the nucleic acid is on the bottom of the tube.</li>
		<li>Prepare a master stock of 100 &#181;M by resuspending the oligonucleotides in TE buffer (10 mM tris(hydroxymethyl)aminomethane (Tris), 1 mM ethylenediaminetetraacetic acid (EDTA)). Ensure the oligonucleotides are thoroughly resuspended by repeated gentle vortexing and brief centrifugation at full speed.</li>
		<li>Prepare a 10 &#181;M stock by diluting an aliquot of master stock with TE buffer. Dilute the 10 &#181;M stock with ultrapure water (MQ water) to prepare working stocks with concentrations of 0.5 &#181;M, 0.7 &#181;M, or 2.5 &#181;M as per Table 2.</li>
	</ol>
	</li>
	<li>Assembling and annealing the reaction master mixes
	<ol>
		<li>Use working stocks to make up the reaction master mixes using the combinations provided in Table 2 and volumes given in Table 3. For the standard DNA ligase assay and most other assays described here, the final buffer composition is 50 mM Tris pH 8.0, 50 mM NaCl, 10 mM Dithiothreitol (DTT) with 10 mM Mg as the divalent cation.</li>
		<li>Anneal the oligonucleotides in a PCR or microcentrifuge tube by heating at 95 &#176;C for 5 min using a heating block or thermocycler. Allow to cool to room temperature for 30 min (volumes &#60;1 mL) to 1 h (volumes &#62;1 mL). For longer oligonucleotides (&#62;40 nt), carry out slower cooling by using a thermocycler with a down ramp of 95 &#176;C to 25 &#176;C over 45 min, or float the tube containing annealing mixture in a 1 L beaker of boiling water and allow to cool to room temperature overnight.</li>
		<li>Add nucleotide cofactors and other heat-sensitive buffer components to the master mix after cooling to room temperature. Use the final reaction mixture directly for the assay by the addition of enzyme (see step 3 below) or store at -20 &#176;C for future use.</li>
	</ol>
	</li>
</ol>
3. Standard assay setup
<ol>
	<li>Assembly and initiation of the assay reaction
	<ol>
		<li>Combine 22.5 &#181;L of the substrate master mix of interest with 2.5 &#181;L of the DNA ligase or other enzyme of interest in a PCR tube. Run reactions in duplicate or triplicate, especially if the results will be quantitated.</li>
		<li>Include a no-protein control (buffer only) in the assay samples. Include no cofactor controls at this point, if needed. 
		NOTE: Enzymes can be stored at -20 &#176;C in 50% v/v glycerol, allowing them to be pipetted directly from solution. Ensure enzyme solutions with glycerol are well mixed prior to addition, either by pipetting to mix or by gentle vortexing.</li>
	</ol>
	</li>
	<li>Immediately transfer the reactions to a PCR machine at 25 &#176;C and incubate for 30 min. Vary the temperature and duration depending on the optimal conditions for enzyme activity.</li>
	<li>Quench the reactions by adding 5 &#181;L of loading dye (95% formamide, 0.5 M Ethylenediaminetetraacetic acid (EDTA), bromophenol blue) and incubate at 95 &#176;C for 5 min.</li>
</ol>
4. Analysis of assay results
<ol>
	<li>Prepare Tris-Borate-EDTA (TBE)-Urea PAGE gels as described below.
	<ol>
		<li>Prepare a stock of 20% acrylamide, 7 M Urea, and 1x TBE solution. For the oligonucleotide set described here, use Acrylamide/Bis Solution in a 29:1 ratio for optimal resolution.</li>
		<li>For one gel, combine 10 mL of 20% acrylamide and 7 M urea solution with 100 &#181;L of APS (10 %) and 3 &#181;L of Tetramethylethylenediamine (TMED) and cast in a gel caster.</li>
	</ol>
	</li>
	<li>After the gel solidifies, run the samples on the TBE urea gel at 45 - 55 &#176;C.
	<ol>
		<li>Pre-run the gel in 1x TBE buffer for 30 min at 10 mA per gel with external heating.</li>
		<li>Remove excess urea in the wells of the gel by flushing with 1x TBE using a pasture pipette.</li>
		<li>Load 10 &#181;L of each reaction and run at 10 mA for 1.0-1.5 h with external heating.</li>
	</ol>
	</li>
	<li>Visualize the gel on the imager with the correct settings for the chosen fluorophore. For FAM, use a filter set that gives excitation/emission at 495/519 nm, which is stored as a pre-set in most imagers.</li>
	<li>Quantify the band intensity of the product and substrate using image processing software with the imager, or an external program such as ImageJ49,50 and calculate the percentage of the product using the formula 
	<img alt="Equation 1" src="/files/ftp_upload/66930/66930eq01.jpg" style="margin: auto;" /> 
	Where P is the integrated value of the product band, and S is the integrated area of the substrate band. In the case of the DNA ligase reaction example, the product band runs at 40 nucleotides (nt) and the substrate band at 20 nt.</li>
</ol>
5. De-adenylation of the DNA ligase to test cofactor specificity
<ol>
	<li>Preparation of reaction master mixes
	<ol>
		<li>Prepare one set of the master mix containing the FAM-labeled NL1 oligonucleotide as described in Table 4. Prepare a second set containing the NL1 oligonucleotide with no FAM label, as described in Table 4.</li>
		<li>Separately, heat both DNA duplexes to 95 &#176;C for 5 min and cool for 30 min to 1 h at 25 &#176;C. Do not add nucleotide cofactor to either master mix.</li>
	</ol>
	</li>
	<li>Assembly and initiation of de-adenylation reaction
	<ol>
		<li>Prepare a single de-adenylation reaction for each cofactor type/ concentration to be tested by combining 10 &#181;L of the unlabeled master mix with 2.5 &#181;L of ligase enzyme.</li>
		<li>Prepare additional tubes as no-cofactor control and no protein control (2.5 &#181;L of buffer added in place of enzyme).</li>
		<li>Incubate the reactions at a temperature specific to the enzyme&#39;s optimum activity, for 1-2 h. Incubation time can be increased if the enzyme is still adenylated.</li>
	</ol>
	</li>
	<li>Run the ligation reaction with the cofactor.
	<ol>
		<li>Add 10 &#181;L of the labeled master mix and 2.5 &#181;L of the desired nucleotide cofactors (e.g., ATP, NAD, ADP, or GTP) directly to the de-adenylated reaction (0.1-1 mM final concentration).</li>
		<li>Add 2.5 &#181;L of reaction buffer to the no nucleotide cofactor control.</li>
		<li>Incubate the reactions for the same time period and temperature as previously used. Quench and visualize as described in step 4.</li>
	</ol>
	</li>
</ol>
6. Using dual-labeled substrates for splinted ligation or multi-part assembly
<ol>
	<li>Design and purchase an oligonucleotide with a florescent moiety that has a different excitation/ emission spectrum to the fluorophore already used.
	<ol>
		<li>In the set-up described here, use NL2 (TAMRA) oligonucleotide having 5-Carboxytetramethylrhodamine (TAMRA) on the 3&#39; end (Table 1).</li>
	</ol>
	</li>
	<li>Assemble the master mix as described below.
	<ol>
		<li>Combine the components of the reaction described in step 2, including equimolar ratios of all oligonucleotides used in the assembly, as well as buffer and divalent cations.</li>
		<li>Anneal by heating at 95 &#176;C for 5 min and cooling at 25 &#176;C for 30 min - 1 h. Add the cofactor and the enzyme and incubate as described in step 3.</li>
	</ol>
	</li>
	<li>Run and image the samples as described in step 4 using the appropriate channels for the fluorophore pair in the substrate. In the case of FAM and TAMRA, these are the Fluorescein (FITC) and Tetramethyl rhodamine (TRITC) channels present on most imagers.</li>
</ol>
7. Evaluation of DNA binding by Electrophoretic Mobility Shift Assay (EMSA) on native gel
<ol>
	<li>Prepare a 10% native TBE PAGE gel as described below.
	<ol>
		<li>Combine 2.5 mL of 40% acrylamide, 1 mL of 10x TBE, 100 &#181;L of 10% APS, 3 &#181;L of TMED, and 6.5 mL of MQ water and cast in a gel caster.</li>
	</ol>
	</li>
	<li>Assemble the binding reaction as described below.
	<ol>
		<li>Assemble the EMSA substrate according to Table 5 so that EDTA (10 mM) is included and metal ions are omitted.</li>
		<li>Combine 20 &#181;L of the EMSA substrate master mix with 5 &#181;L of the protein in a PCR tube. Include a no protein control sample. Incubate for 30 min at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Analyze by native electrophoresis as described below.
	<ol>
		<li>Add 5 &#181;L of 5x native loading dye (100 mM EDTA, 0.25 % bromophenol blue, 25% v/v glycerol, and MQ water up to 1 mL) to the samples.</li>
		<li>Load on the prepared gel and run at 60 V for 2-3 h with cooling by water circulation until the dye front is a few cm above the end of the gel.</li>
		<li>Visualize and analyze gels as described in step 4.</li>
	</ol>
	</li>
</ol>

Assays for DNA Enzymes Using Synthetic Oligonucleotides

SEG and UR are employees of ArcticZymes Technologies AS which distributes the R2D ligase. AW, ER-S and RS have no competing interests.

Critical steps in the protocol 
Oligonucleotide design and purchase: When purchasing the oligonucleotides for duplex formation, it is essential to consider sequence design. It is recommended to use an oligo analyzer tool to predict properties of the nucleotide sequence, such as GC content, melting temperature, secondary structure, and dimerization potential, before ordering57.
Assembly and annealing of nucleic acid duplexes: When preparing RNA/RNA-...

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).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>30% Acrylamide/Bis Solution (29:1)</td>
			<td>BioRad</td>
			<td>1610156</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine triphosphate (ATP)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium persulfate (APS)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Benchtop centrifuge</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Borate</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bromophenol blue</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol (DTT)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electrophoresis system with circulating water bath</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoresnence imager, e.g. iBright FL1000</td>
			<td>Thermo Fisher Scientific</td>
			<td>A32752</td>
			<td></td>
		</tr>
		<tr>
			<td>Formamide</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gel casting system</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heating block</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Many suppliers</td>
			<td></td>
			<td>Other metal ions may be preferred depending on the protein studied</td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes (1.5 mL)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipettes and tips</td>
			<td>Many suppliers</td>
			<td></td>
			<td>1 mL, 0.2 mL, 0.02 mL, 0.002 mL</td>
		</tr>
		<tr>
			<td>Nicotinamide adenine dinucleotide (NAD+)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Oligonucleotides</td>
			<td>Integrated DNA Technologies</td>
			<td>NA</td>
			<td>Thermo Fisher, Sigma-Aldrich,&#160; Genscript and others also supply these</td>
		</tr>
		<tr>
			<td>pasture pipette</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PCR thermocycler</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PCR tubes</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>RNAse away</td>
			<td>ThermoFisher</td>
			<td>7002PK</td>
			<td>Only needed when working with RNA oligos</td>
		</tr>
		<tr>
			<td>RNase AWAY</td>
			<td>Merck</td>
			<td>83931-250ML</td>
			<td>Surfactant for removal of RNAse contamination on surfaces</td>
		</tr>
		<tr>
			<td>RNAse-free water</td>
			<td>New England Biolabs</td>
			<td>B1500L</td>
			<td>Only needed when working with RNA oligos</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SUPERase IN RNase inhibitor</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM2694</td>
			<td>Broad spectrum RNAse inhibitir (protein-based)</td>
		</tr>
		<tr>
			<td>SYBR Gold</td>
			<td>Thermo Fisher Scientific</td>
			<td>S11494</td>
			<td>This may be used to post-stain gels and visualise unlabelled oligonucleotides</td>
		</tr>
		<tr>
			<td>Tetramethylethylenediamine (TMED)</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tris, or tris(hydroxymethyl)aminomethane</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrapure water (Milli-Q)</td>
			<td>Merck</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>urea</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex</td>
			<td>Many suppliers</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

using modified synthetic oligonucleotides to assay nucleic acid-metabolizing enzymes

The availability of a range of modified synthetic oligonucleotides from commercial vendors has allowed the development of sophisticated assays to characterize diverse properties of nucleic acid metabolizing enzymes that can be run in any standard molecular biology lab. The use of fluorescent labels has made these methods accessible to researchers with standard PAGE electrophoresis equipment and a fluorescent-enabled imager, without using radioactive materials or requiring a lab designed for the storage and preparation of radioactive materials, i.e., a Hot Lab. The optional addition of standard modifications such as phosphorylation can simplify assay setup, while the specific incorporation of modified nucleotides that mimic DNA damages or intermediates can be used to probe specific aspects of enzyme behavior. Here, the design and execution of assays to interrogate several aspects of DNA processing by enzymes using commercially available synthetic oligonucleotides are demonstrated. These include the ability of ligases to join or nucleases to degrade different DNA and RNA hybrid structures, differential cofactor usage by the DNA ligase, and evaluation of the DNA-binding capacity of enzymes. Factors to consider when designing synthetic nucleotide substrates are discussed, and a basic set of oligonucleotides that can be used for a range of nucleic acid ligase, polymerase, and nuclease enzyme assays are provided.

Ligation by DNA ligase 
DNA ligase enzymatic activity will result in an increase in the size of the fluorescently labeled oligonucleotide when visualized on a urea PAGE gel. In the case of the substrates for both DNA- and RNA-ligation listed in Table 2, this corresponds to a doubling in size from 20 nt to 40 nt (Figure 3A). Optimal enzyme activity can be determined by changing conditions such as temperature, protein concentration, or incubation time (<s...

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

Here, a protocol for assaying nucleic acid metabolizing enzymes is presented, using examples of ligase, nuclease, and polymerase enzymes. The assay utilizes fluorescently labeled and unlabeled oligonucleotides that can be combined to form duplexes mimicking RNA and/or DNA damages or pathway intermediates, allowing for the characterization of enzyme behavior.

Using Modified Synthetic Oligonucleotides to Assay Nucleic Acid-Metabolizing Enzymes

The availability of a range of modified synthetic oligonucleotides from commercial vendors has allowed the development of sophisticated assays to ...

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Research

JoVE Journal

Biochemistry

489 Views.  University of Waikato. Here, a protocol for assaying nucleic acid metabolizing enzymes is presented, using examples of ligase, nuclease, and polymerase enzymes. The assay utilizes fluorescently labeled and unlabeled oligonucleotides that can be combined to form duplexes mimicking RNA and/or DNA damages or pathway intermediates, allowing for the characterization of enzyme behavior. 

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

High-throughput Screening of Carbohydrate-degrading Enzymes Using Novel Insoluble Chromogenic Substrate Assay Kits

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper and food industries. A deeper understanding of CAZymes is important from both fundamental biology and industrial standpoints. Vast numbers of CAZymes exist in nature (especially in microorganisms) and hundreds of thousands have been cataloged and described in the carbohydrate active enzyme database (CAZy). However, the rate of discovery of putative enzymes has outstripped our ability to biochemically characterize their activities. One reason for this is that advances in genome and transcriptome sequencing, together with associated bioinformatics tools allow for rapid identification of candidate CAZymes, but technology for determining an enzyme's biochemical characteristics has advanced more slowly. To address this technology gap, a novel high-throughput assay kit based on insoluble chromogenic substrates is described here. Two distinct substrate types were produced: Chromogenic Polymer Hydrogel (CPH) substrates (made from purified polysaccharides and proteins) and Insoluble Chromogenic Biomass (ICB) substrates (made from complex biomass materials). Both CPH and ICB substrates are provided in a 96-well high-throughput assay system. The CPH substrates can be made in four different colors, enabling them to be mixed together and thus increasing assay throughput. The protocol describes a 96-well plate assay and illustrates how this assay can be used for screening the activities of enzymes, enzyme cocktails, and broths.

A high-throughput assay for enzyme screening is described. This multiplexed ready-to-use assay kit comprises of pre-chosen Chromogenic Polymer Hydrogel (CPH) substrates and complex Insoluble Chromogenic Biomass (ICB) substrates. Target enzymes are polysaccharide degrading endo-enzymes and proteases.

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper ...

A Polyaniline-based Sensor of Nucleic Acids

Detection of nucleic acids is at the center of diagnostic technologies used in research and the clinic. Standard approaches used in these technologies rely on enzymatic modification that can introduce bias and artifacts. A critical element of next generation detection platforms will be direct molecular sensing, thereby avoiding a need for amplification or labels. Advanced nanomaterials may provide the suitable chemical modalities to realize label-free sensors. Conjugated polymers are ideal for biological sensing, possessing properties compatible with biomolecules and exhibit high sensitivity to localized environmental changes. In this article, a method is presented for detecting nucleic acids using the electroconductive polymer polyaniline. Simple DNA "probe" oligonucleotides complementary to target nucleic acids are attached electrostatically to the polymer, creating a sensor system that can differentiate single nucleotide differences in target molecules. Outside the specific and unbiased nature of this technology, it is highly cost effective.

Nucleic acids are common analytes for assessing biological systems; however, bias from enzymatic manipulation can cause concern. Here a method is described for label-free detection of nucleic acids using polyaniline. This sensitive, cost-effective sensor technology can distinguish single nucleotide differences between molecules.

Detection of nucleic acids is at the center of diagnostic technologies used in research and the clinic. Standard approaches used in these technologies ...

An Optimized Protocol for Electrophoretic Mobility Shift Assay Using Infrared Fluorescent Dye-labeled Oligonucleotides

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. Radioactivity has been the predominant method of DNA labeling in EMSAs. However, recent advances in fluorescent dyes and scanning methods have prompted the use of fluorescent tagging of DNA as an alternative to radioactivity for the advantages of easy handling, saving time, reducing cost, and improving safety. We have recently used fluorescent EMSA (fEMSA) to successfully address an important biological question. Our fEMSA analysis provides mechanistic insight into the effect of a missense mutation, G73E, in the highly conserved HMG transcription factor SOX-2 on olfactory neuron type diversification. We found that mutant SOX-2G73E protein alters specific DNA binding activity, thereby causing olfactory neuron identity transformation. Here, we present an optimized and cost-effective step-by-step protocol for fEMSA using infrared fluorescent dye-labeled oligonucleotides containing the LIM-4/SOX-2 adjacent target sites and purified SOX-2 proteins (WT and mutant SOX-2G73E proteins) as a biological example.

We describe here an optimized protocol of fluorescent Electrophoretic Mobility Shift Assays (fEMSA) using purified SOX-2 proteins together with infrared fluorescent dye-labeled DNA probes as a case study to tackle an important biological question.

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. ...

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

Localization of SUMO-modified Proteins Using Fluorescent Sumo-trapping Proteins

Here we are presenting a novel method to study the sumoylation of proteins and their sub-cellular localization in mammalian cells and nematode oocytes. This method utilizes a recombinant modified SUMO-trapping protein fragment, kmUTAG, derived from the Ulp1 SUMO protease of the stress-tolerant budding yeast Kluyveromyces marxianus. We have adapted the properties of the kmUTAG for the purpose of studying sumoylation in a variety of model systems without the use of antibodies. For the study of SUMO, KmUTAG has several advantages when compared to antibody-based approaches. This stress-tolerant SUMO-trapping reagent is produced recombinantly, it recognizes native SUMO isoforms from many species, and unlike commercially available antibodies it shows reduced affinity for free, unconjugated SUMO. Representative results shown here include the localization of SUMO conjugates in mammalian tissue culture cells and nematode oocytes.

SUMO is an essential and highly conserved, small ubiquitin-like modifier protein. In this protocol we are describing the use of a stress-tolerant recombinant SUMO-trapping protein (kmUTAG) to visualize native, untagged SUMO conjugates and their localization in a variety of cell types.

Here we are presenting a novel method to study the sumoylation of proteins and their sub-cellular localization in mammalian cells and nematode oocytes. ...

Kinetic Screening of Nuclease Activity using Nucleic Acid Probes

Nucleases are a class of enzymes that break down nucleic acids by catalyzing the hydrolysis of the phosphodiester bonds that link the ribose sugars. Nucleases display a variety of vital physiological roles in prokaryotic and eukaryotic organisms, ranging from maintaining genome stability to providing protection against pathogens. Altered nuclease activity has been associated with several pathological conditions including bacterial infections and cancer. To this end, nuclease activity has shown great potential to be exploited as a specific biomarker. However, a robust and reproducible screening method based on this activity remains highly desirable.
Herein, we introduce a method that enables screening for nuclease activity using nucleic acid probes as substrates, with the scope of differentiating between pathological and healthy conditions. This method offers the possibility of designing new probe libraries, with increasing specificity, in an iterative manner. Thus, multiple rounds of screening are necessary to refine the probes&#39; design with enhanced features, taking advantage of the availability of chemically modified nucleic acids. The considerable potential of the proposed technology lies in its flexibility, high reproducibility, and versatility for the screening of nuclease activity associated with disease conditions. It is expected that this technology will allow the development of promising diagnostic tools with a great potential in the clinic.

Altered nuclease activity has been associated with different human conditions, underlying its potential as a biomarker. The modular and easy to implement screening methodology presented in this paper allows the selection of specific nucleic acid probes for harnessing nuclease activity as a biomarker of disease.

Nucleases are a class of enzymes that break down nucleic acids by catalyzing the hydrolysis of the phosphodiester bonds that link the ribose sugars. ...

Preparation of Sample Support Films in Transmission Electron Microscopy using a Support Floatation Block

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant bottleneck. Macromolecular samples are ideally imaged directly from random orientations in a thin layer of vitreous ice. However, many samples are refractory to this, and protein denaturation at the air-water interface is a common problem. To overcome such issues, support films-including amorphous carbon, graphene, and graphene oxide-can be applied to the grid to provide a surface which samples can populate, reducing the probability of particles experiencing the deleterious effects of the air-water interface. The application of these delicate supports to grids, however, requires careful handling to prevent breakage, airborne contamination, or extensive washing and cleaning steps. A recent report describes the development of an easy-to-use floatation block that facilitates wetted transfer of support films directly to the sample. Use of the block minimizes the number of manual handling steps required, preserving the physical integrity of the support film, and the time over which hydrophobic contamination can accrue, ensuring that a thin film of ice can still be generated. This paper provides step-by-step protocols for the preparation of carbon, graphene, and graphene oxide supports for EM studies.

Sample preparation for cryo-electron microscopy (cryo-EM) is a significant bottleneck in the structure determination workflow of this method. Here, we provide detailed methods for using an easy-to-use, three-dimensionally printed block for the preparation of support films to stabilize samples for transmission EM studies.

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant ...

Making Precise and Accurate Single-Molecule FRET Measurements using the Open-Source smfBox

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes measurements on freely diffusing biomolecules more accessible. This overview includes a step-by-step protocol for using this instrument to make measurements of precise FRET efficiencies in duplex DNA samples, including details of the sample preparation, instrument setup and alignment, data acquisition, and complete analysis routines. The presented approach, which includes how to determine all the correction factors required for accurate FRET-derived distance measurements, builds on a large body of recent collaborative work across the FRET Community, which aims to establish standard protocols and analysis approaches. This protocol, which is easily adaptable to a range of biomolecular systems, adds to the growing efforts in democratising smFRET for the wider scientific community.

This article provides step-by-step instructions for making fully-corrected accurate FRET measurements on individual, freely diffusing biomolecules using the open-source, inexpensive smfBox, from switch on, through alignment and focusing, to data collection and analysis.

The smfBox is a recently developed cost-effective, open-source instrument for single-molecule F&#246;rster Resonance Energy Transfer (smFRET), which makes ...

Detection and Quantification of Calcitonin Gene-Related Peptide (CGRP) in Human Plasma Using a Modified Enzyme-Linked Immunosorbent Assay

Calcitonin gene-related peptide (CGRP) is a vasoactive neuropeptide that plays a putative role in the pathophysiology of migraine headaches and may be a candidate for biomarker status. CGRP is released from neuronal fibers upon activation and induces sterile neurogenic inflammation and arterial vasodilation in the vasculature that receives trigeminal efferent innervation. The presence of CGRP in the peripheral vasculature has spurred investigations to detect and quantify this neuropeptide in human plasma using proteomic assays, such as the enzyme-linked immunosorbent assay (ELISA). However, its half-life of 6.9 min and the variability in technical details of assay protocols, which are often not fully described, have yielded inconsistent CGRP ELISA data in the literature. Here, a modified ELISA protocol for the purification and quantification of CGRP in human plasma is presented. The procedural steps involve sample collection and preparation, extraction using a polar sorbent as a means of purification, additional steps to block non-specific binding, and quantification via ELISA. Further, the protocol has been validated with spike and recovery and linearity of dilution experiments. This validated protocol can theoretically be used to quantify CGRP concentrations in the plasma of individuals not only with migraine, but also with other diseases in which CGRP may play a role.

Detection and Quantification of Calcitonin Gene-Related Peptide CGRP in Human Plasma Using a Modified Enzyme-Linked Immunosorbent Assay

Published data pertaining to calcitonin gene-related peptide (CGRP) concentrations in human plasma are inconsistent. These inconsistencies may be due to the lack of a standardized, validated methodology to quantify this neuropeptide. Here, we describe a validated enzyme-linked immunosorbent assay (ELISA) protocol to purify and quantify CGRP in human plasma.

Calcitonin gene-related peptide (CGRP) is a vasoactive neuropeptide that plays a putative role in the pathophysiology of migraine headaches and may be a ...

A Fibrinolytic Enzyme-Based Method to Study Cerebral Thrombus Cell Components

A Comprehensive Approach to Analyze the Cell Components of Cerebral Blood Clots

Cerebral thrombosis, a blood clot in a cerebral artery or vein, is the most common type of cerebral infarction. The study of the cell components of cerebral blood clots is important for diagnosis, treatment, and prognosis. However, the current approaches to studying the cell components of the clots are mainly based on in situ staining, which is unsuitable for the comprehensive study of the cell components because cells are tightly wrapped in the clots. Previous studies have successfully isolated a fibrinolytic enzyme (sFE) from Sipunculus nudus, which can degrade the cross-linked fibrin directly, releasing the cell components. This study established a comprehensive method based on the sFE to study the cell components of cerebral thrombus. This protocol includes clot dissolving, cell releasing, cell staining, and routine blood examination. According to this method, the cell components could be studied quantitatively and qualitatively. The representative results of experiments using this method are shown.

Author Spotlight: A Novel Method for Comprehensive Cell Component Analysis of Cerebral Blood Clots 

This study describes a fast and effective method for the cell component analysis of cerebral blood clots through clot dissolving, cell staining, and routine blood examination.

Cerebral thrombosis, a blood clot in a cerebral artery or vein, is the most common type of cerebral infarction. The study of the cell components of ...