The authors would like to acknowledge Luiza I. Hernandez (Link&#246;ping University) for her careful revision of the manuscript and valuable advice. This work was supported by The Knut and Alice Wallenberg Foundation and The Swedish Government Strategic Research Area in Materials Science on Advanced Functional Materials at Link&#246;ping University (Faculty Grant SFO-Mat-LiU No. 2009-00971).

1. Oligonucleotide library design and preparation
<ol>
	<li>Library design
	<ol>
		<li>Based on the following criteria design a library of quenched fluorescent oligonucleotide probes between 8 and 12-mer long to avoid secondary structure formation, with equal length for all probes.
		<ol>
			<li>Include at least one DNA and one RNA random sequence containing a combination of adenine (A), guanine (G), cytosine (C) and thymine (T)/uracil (U) (DNA and RNA probes in Table 1). 
			NOTE: The DNA and RNA sequences described in Table 1 have been useful as the initial substrates for classifying an unknown type of nuclease activity, either DNase or RNase. These two sequences are recommended as the starting point for any screening, as they have shown a wide capability to detect nuclease activity profiles in bacteria and tissue samples. If nuclease activity is not observed with these DNA and RNA sequences, the design of additional oligonucleotides is required.</li>
			<li>Include polynucleotide sequences consisting of a single type of nucleotide to determine sequence dependence for a given nuclease activity (DNA-Poly A, DNA-Poly T, DNA-Poly C, DNA-Poly G, RNA-Poly A, RNA-Poly U, RNA-Poly C and RNA-Poly G in Table 1).</li>
			<li>Using the same sequence of nucleotide residues as the initial DNA and RNA sequences, include sequences that contain nucleoside analogs harboring chemical modifications at the 2&#39;-position of the ribose sugar, such as 2&#39;-Fluoro and 2&#39;-O-Methyl, as a stringent step for increasing the selectivity of the nucleases.
			<ol>
				<li>Include fully modified sequences consisting entirely of 2&#39;-Fluoro nucleoside analogs or 2&#39;-O-Methyl nucleoside analogs (All 2&#39;-F and All 2&#39;-OMe, Table 1)</li>
				<li>Include chimeric sequences consisting of unmodified natural purines and 2&#39;-Fluoro pyrimidine nucleoside analogs (RNA Pyr-2&#39;F, Table 1).</li>
				<li>Include chimeric sequences consisting of unmodified natural purines and 2&#39;-O-Methyl pyrimidine nucleoside analogs (RNA Pyr-2&#39;OMe, Table 1).</li>
				<li>Include chimeric sequences consisting of unmodified natural pyrimidines and 2&#39;-Fluoro purine nucleoside analogs (RNA Pur-2&#39;F, Table 1).</li>
				<li>Include chimeric sequences consisting of unmodified natural pyrimidines and 2&#39;-O-Methyl purine nucleoside analogs (RNA Pur-2&#39;OMe, Table 1).</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Oligonucleotide probe preparation and storage 
	NOTE: Oligonucleotides are synthesized by the phosphoramidite method. After synthesis, the oligonucleotides are purified using high performance liquid chromatography (HPLC) and the mass is measured by mass spectrometry.
	<ol>
		<li>Spin down the lyophilized oligonucleotide probes using a centrifuge and dilute them in Tris-EDTA (TE) buffer to prevent nuclease degradation. Determine the dilution volume according to the yield of each probe to render a stock solution with a concentration of 500 pmol/&#956;L.</li>
		<li>Store lyophilized oligonucleotides at 4 &#176;C or room temperature for a short term. For long term storage (months to years), store lyophilized oligonucleotides at -20 &#176;C. Upon resuspension of lyophilized oligonucleotides in TE, store the stock solutions at -20 &#176;C or, preferably, at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Bacterial culture
<ol>
	<li>Take one porous glass bead from the cryogenic storage vial under sterile conditions.</li>
	<li>To isolate individual bacterial colonies (Salmonella and E. coli), use the quadrant method24 by streaking/rolling the bead directly onto a Petri dish containing Tryptic Soy Agar (TSA) medium supplemented with defibrinated sheep blood.</li>
	<li>Incubate the culture at 37 &#176;C for 24 h.</li>
</ol>
3. Supernatant preparation
<ol>
	<li>Transfer a single colony from solid media to 50 mL Tryptic Soy Broth (TSB) and incubate at 37 &#176;C for 24 h at 200 rpm.</li>
	<li>Dilute the culture (1:500) in TSB (sub-culture) and incubate at 37 &#176;C for 24 h at 200 rpm in a shaking incubator. 
	NOTE: It is expected that after 24 h of incubation the bacterial cultures are in stationary phase reaching values higher than 109 CFU/mL.</li>
	<li>After the incubation period, transfer the cultures to capped sterile tubes and centrifuge at 4,500 x g for 30 min.</li>
	<li>Collect the supernatant and use it immediately. Alternatively, store the supernatants at 4 &#176;C or -20 &#176;C.</li>
</ol>
4. Nuclease Activity Assay
<ol>
	<li>Preparation of working solutions
	<ol>
		<li>Prepare a 20 &#956;L working solution for each probe in 1.5 mL nuclease free microcentrifuge tubes by diluting (1:10 ratio) the stock solution (500 pmol/&#956;L) for a final working concentration of 50 pmol/ &#956;L. For that purpose, mix 18 &#956;L of Phosphate Buffer Saline (PBS) containing MgCl2 and CaCl2 with 2 &#956;L of probe stock solution. 
		NOTE: Be aware that in the working solution, the oligonucleotides are more vulnerable to nuclease degradation, therefore, it is recommended to prepare the working solutions just before setting up the reaction.</li>
		<li>Avoid direct light contact during the preparation and keep in the dark (wrapped in foil), when dealing with fluorophores.</li>
	</ol>
	</li>
	<li>Reaction set up
	<ol>
		<li>Pre-warm the fluorometer (e.g. Cytation 1) to 37 &#176;C.</li>
		<li>Prepare a set (one tube/probe) of 1.5 mL nuclease free microcentrifuge tubes for each sample (TSB, E. coli and Salmonella) and label accordingly.</li>
		<li>Carefully add 96 &#956;L of TSB sterile culture media, Salmonella supernatant or E. coli supernatant (from the step 3.4.). Subsequently, add 4 &#956;L/tube of probe working solution accordingly. Perform this step at room temperature. 
		NOTE: 10 probes were used for the first round of the screening and 6 probes for the second round.</li>
		<li>Mix thoroughly by pipetting up and down to obtain a homogenous solution. Avoid introducing air bubbles into the samples while mixing.</li>
		<li>Load 95 &#956;L of each solution (probe + supernatant or culture media) into a separate well of a black bottom, non-treated 96 well plate. Minimize the formation of bubbles in the wells upon loading by dispensing carefully with the tip close to the wall of the well.</li>
		<li>Cover the plate with its lid. Visually inspect the lid and check for pen markings or dust particle accumulation that may introduce measurement artifacts. Replace the lid for a new one if that is the case.</li>
	</ol>
	</li>
	<li>Measurement set up
	<ol>
		<li>Open the acquisition software (Gen5 3.05) by clicking the software shortcut icon (Figure S1A).</li>
		<li>Select Read Now from the task manager window and choose New&#8230; to create the kinetic measurement protocol (Figure S1B).</li>
		<li>Click on Set Temperature in the dialog window titled Procedure and select 37 &#730;C. Confirm and save the settings by pressing OK (Figure S1C).</li>
		<li>Click on Start Kinetics in the dialog window titled Procedure. Then in the pop-up dialog window, titled Kinetic Step, select 2 h in the Run Time input box and 2 min in the Interval input box. Confirm and save the settings by pressing OK (Figure S1D).</li>
		<li>Click on Read in the dialog window titled Procedure. Then in the pop-up dialog window, titled Read Method, select Fluorescence Intensity as a detection method, Endpoint/Kinetic as a read type and Filters as optics type. Confirm and save the settings by pressing OK (Figure S1E).</li>
		<li>In the pop-up dialog window titled Read Step (Kinetic), select Green from the Filter Set. Confirm and save the settings by pressing OK (Figure S2A).</li>
		<li>In the dialog window titled Procedure, select use lid and click on validate to ensure that the created protocol is valid. This is confirmed by a pop-up dialog window (Figure S2B).</li>
		<li>Select protocol in the menu bar and choose procedure (Figure S2C).</li>
		<li>In the dialog window titled Procedure, define the wells to be measured (Figure S2D).</li>
		<li>Enter the name of the experiment in the file name input box (Figure S2E).</li>
		<li>Load the plate with its lid into the plate reader. Ensure the plate is in the right orientation.</li>
		<li>Start the acquisition by clicking read new button in the toolbar (Figure S2F).</li>
	</ol>
	</li>
	<li>Data Analysis
	<ol>
		<li>Click on one of the measured wells in the dialog window titled Plate 1 (Figure S3A).</li>
		<li>Click select wells and include all the measured wells in the well selection dialog window. Confirm and save the settings by pressing OK. (Figure S3B).</li>
		<li>Select data in the dialog window titled Plate 1 to visualize the tabulated results (Figure S3C).</li>
		<li>Export the data to a spread sheet by selecting quick export from the context menu (Figure S3C).</li>
		<li>In the spread sheet, label the data columns accordingly for each sample and probe.</li>
		<li>Generate kinetic graphs, using line with markers style, by plotting relative fluorescence units (RFU) versus time (x-axis: timeline of the reaction and y-axis: RFUs). 
		NOTE: The description of generating a graph is applicable when using spreadsheet programs (e.g., Excel). However, any other programs can be used to generate graphs from the raw data obtained, by plotting RFU versus time.</li>
		<li>Calculate the rate of the enzymatic reaction for each measured interval according to the following formula25: rate= <img alt="Equation 1" src="/files/ftp_upload/60005/60005eq1.jpg" />, where If is RFU at the maximal interval time point and Ii is RFU at the minimal interval time point, and Tf and Ti are the maximal and minimal interval time points, respectively.</li>
		<li>Select the rate with the highest value (Rmax) for each curve. If there is more than one Rmax per sample, select the one that occurs at the earliest measurement time point.</li>
		<li>Calculate the ratio between Rmax and the average value of the time interval at which Rmax occurs: rate coefficient = <img alt="Equation 2" src="/files/ftp_upload/60005/60005eq2.jpg" /></li>
		<li>Calculate the fold difference (FD) between the rate coefficient of Salmonella and E. coli for each probe.</li>
		<li>Consider a fold difference value higher than 3 as significant.</li>
		<li>Consider the significant probes with the highest fold difference values (FD) as candidate probes and as the basis for the library design in the next screening round.</li>
	</ol>
	</li>
</ol>
5. Screening Rounds&#8217; Selection Criteria (Figure 1)
<ol>
	<li>First Round of Screening
	<ol>
		<li>Perform the nuclease activity assay (as described in section 4) using DNA, RNA and polynucleotide probes.</li>
		<li>Evaluate the preference for DNA chemistry or RNA chemistry by comparing the number and performance (FD value) of DNA and RNA candidate probes.</li>
		<li>Select the type of nucleic acid chemistry rendering the greatest number of candidate probes, as illustrated in Figure 1.</li>
	</ol>
	</li>
	<li>Second Round of Screening
	<ol>
		<li>Perform the nuclease activity assay (as described in section 4) using fully modified probes and chimeric probes designed based on the nucleic acid substrate selected in the first screening round.</li>
		<li>Evaluate the preference towards sequences containing 2&#39;-Fluoro and 2&#39;-O-Methyl nucleoside analogs by comparing the results obtained for 2&#39;-Fluoro and 2&#39;-O-Methyl modified candidate probes.</li>
		<li>Select the type of nucleoside analog modification rendering the greatest number of candidate probes, as illustrated in Figure 1.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Alterations of nuclease activity have been associated with a wide variety of disease phenotypes, including different types of cancer and bacterial infections. These alterations are proposed to be the causative agent of a condition14, while in other cases they are the consequence of a detrimental physiological event20 or pathogenic agent16,26. Not surprisingly, attempts to use nucleases and nuclease activity as a dia...

Nucleases are a class of enzymes capable of cleaving the phosphodiester bonds that form the backbone structure of nucleic acid molecules. The vast diversity of nucleases makes their classification rather difficult. However, there are some common criteria used to describe nucleases, such as substrate preference (deoxyribonucleic acid (DNA) or ribonucleic acid (RNA)), cleavage site (endonucleases or exonucleases), or metal ion dependency, among others1. Nucleases are highly conserved catalytic enzymes that have fundamental roles in both, prokaryotic and eukaryotic organisms and have been used, and continue to be used, as gene editing tools2. They are also fundamental actors in DNA maintenance and replication, helping to keep genome stability and participating in proof-reading processes3. In bacteria, for example, nucleases have been identified as important virulence factors, able to promote bacterial survival by reducing the efficacy of the host&#39;s immune system4,5,6,7,8. In mammals, nucleases have been suggested to be implicated in apoptosis9, mitochondrial biogenesis and maintenance10 and mediation of antibacterial and antiviral innate immune responses11. Not surprisingly, nuclease activity alterations, whether enhancement or lack of, have been implicated in a wide array of human diseases. These diseases range from a wide variety of cancers12,13 to cardiac hypertrophy10 or autoimmune diseases14. Therefore, nucleases have become interesting candidates as biomarkers for a heterogeneous group of human conditions. In fact, nucleases have already shown their potential as successful diagnostic tools for the detection of infections caused by specific bacterial agents, such as Staphylococcus aureus or Escherichia coli15,16. In many cancer types, expression of staphylococcal nuclease domain-containing protein 1 (SND1) ribonuclease is indicative of poor prognosis17. In pancreatic cancer patients, elevated ribonuclease I (RNase I) serum levels have been reported18 and proposed to be associated with cancerous cell phenotypes19. In ischemic heart conditions, such as myocardial infarction or unstable angina pectoris, deoxyribonuclease I (DNase I) serum levels have been shown to be a valid diagnostic marker20,21.
It has been hypothesized that the global blueprint of nuclease activity may be different in healthy and disease states. In fact, recent reports have used differences in nuclease activity to distinguish between healthy and cancerous phenotypes22 or to identify pathogenic bacterial infections in a species-specific manner15,23. These findings have opened a new avenue for the use of nucleases as biomarkers of disease. Therefore, there exists a necessity for the development of a comprehensive screening method able to systematically identify disease associated differences in nuclease activity, which may be of key importance in the development of new diagnostic tools.
Herein, we introduce and describe a new in vitro screening approach (Figure 1) to identify sensitive and specific probes capable of discriminating between nuclease activity in healthy and unhealthy, or activity specific to a type of cell or bacteria. Taking advantage of the modularity of nucleic acids, we designed an initial library of quenched fluorescent oligonucleotide probes consisting of a comprehensive set of different sequences and chemistries, both being important parameters for library design. These oligonucleotide probes are flanked by a fluorophore (fluorescein amidite, FAM) and a quencher (tide quencher 2, TQ2) at the 5&#39; and 3&#39; ends respectively (Table 1). By using this fluorescent resonance energy transfer (FRET) based fluorometric assay to measure the kinetics of enzymatic degradation, we were able to identify candidate probes with the potential to discriminate differential patterns of nuclease activity associated with healthy or disease states. We designed an iterative process, in which new libraries are created based on the best candidate probes, that allows the identification of ever more specific candidate probes in subsequent screening steps. Moreover, this approach takes advantage of the catalytic nature of nucleases to increase sensitivity. This is achieved by taking advantage of the activatable nature of the reporter probes and the ability of nucleases to continually process substrate molecules, both representing key advantages over alternative antibody or small molecule-based screening methods.
This approach offers a highly modular, flexible and easy to implement screening tool for the identification of specific nucleic acid probes capable of discriminating between healthy and disease states, and an excellent platform for the development of new diagnostic tools that can be adapted for future clinical applications. As such, this approach was used to identify the nuclease activity derived from Salmonella Typhimurium (herein referred to as Salmonella) for the specific identification of this bacteria. In the following protocol, we report on a method to screen for bacterial nuclease activity using kinetic analysis.

<table><tbody><tr><td>Black bottom, non-treated 96 well plate</td><td>Fisher Scientific</td><td>10000631</td><td></td></tr><tr><td>Cytation1</td><td>BioTek</td><td>CYT1FAV</td><td></td></tr><tr><td>Eppendorf tubes</td><td>Thermofisher</td><td>11926955</td><td></td></tr><tr><td>Escherichia coli</td><td>ATCC</td><td>25922</td><td></td></tr><tr><td>Microbank cryogenic storage vial containing beads</td><td>Pro-Lab Diagnostics</td><td>22-286-155</td><td></td></tr><tr><td>Nucleic acid probes</td><td>Biomers.net</td><td>#</td><td></td></tr><tr><td>Phosphate Buffer Saline containing MgCl&lt;sub&gt;2&lt;/sub&gt; and CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Gibco&amp;trade;</td><td>14040117</td><td></td></tr><tr><td>Salmonella enterica subs. Enterica</td><td>ATCC</td><td>14028</td><td></td></tr><tr><td>Tris-EDTA</td><td>Fisher Scientific</td><td>10647633</td><td></td></tr><tr><td>Tryptone Soya Agar with defibrinated sheep blood</td><td>Thermo Fisher Scientific</td><td>10362223</td><td></td></tr><tr><td>Tryptic Soy Broth</td><td>Sigma Aldrich</td><td>22092</td><td></td></tr></tbody></table>

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.

Figure 1 shows the work flow of this methodology, which is divided into two screening rounds. In the first round of screening, we used 5 DNA probes (DNA, DNA Poly A, DNA Poly T, DNA Poly C and DNA Poly G) and also 5 RNA probes (RNA, RNA Poly A, RNA Poly U RNA Poly C and RNA Poly G). The raw data of this screening round can be found in Supplementary Table 1. In the second round, chemically modified probes were synthesized by replacing the RNA sequence with chemically modified...

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Kinetic Screening of Nuclease Activity using Nucleic Acid Probes

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. ...

kinetic-screening-of-nuclease-activity-using-nucleic-acid-probes

Universidad San Sebastian

Research

JoVE Journal

Biochemistry

8.2K Views.  Linköping University. 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.

Video: Kinetic Screening of Nuclease Activity using Nucleic Acid Probes

Kinetics of Lagging-strand DNA Synthesis In Vitro by the Bacteriophage T7 Replication Proteins

Here we provide protocols for the kinetic examination of lagging-strand DNA synthesis in vitro by the replication proteins of bacteriophage T7. The T7 replisome is one of the simplest replication systems known, composed of only four proteins, which is an attractive feature for biochemical experiments. Special emphasis is placed on the synthesis of ribonucleotide primers by the T7 primase-helicase, which are used by DNA polymerase to initiate DNA synthesis. Because the mechanisms of DNA replication are conserved across evolution, these protocols should be applicable, or useful as a conceptual springboard, to investigators using other model systems. The protocols described here are highly sensitive and an experienced investigator can perform these experiments and obtain data for analysis in about a day. The only specialized piece of equipment required is a rapid-quench flow instrument, but this piece of equipment is relatively common and available from various commercial sources. The major drawbacks of these assays, however, include the use of radioactivity and the relative low throughput.

Kinetics of Lagging-strand DNA Synthesis In Vitro by the Bacteriophage T7 Replication Proteins

We describe sensitive, gel-based discontinuous assays to examine the kinetics of lagging-strand initiation using the replication proteins of bacteriophage T7.

Here we provide protocols for the kinetic examination of lagging-strand DNA synthesis in vitro by the replication proteins of bacteriophage T7. The T7 ...

Genetics

In Vitro Biochemical Assays using Biotin Labels to Study Protein-Nucleic Acid Interactions

Protein-nucleic acid interactions play important roles in biological processes such as transcription, recombination, and RNA metabolism. Experimental methods to study protein-nucleic acid interactions require the use of fluorescent tags, radioactive isotopes, or other labels to detect and analyze specific target molecules. Biotin, a non-radioactive nucleic acid label, is commonly used in electrophoretic mobility shift assays (EMSA) but has not been regularly employed to monitor protein activity during nucleic acid processes. This protocol illustrates the utility of biotin labeling during in vitro enzymatic reactions, demonstrating that this label works well with a range of different biochemical assays. Specifically, in alignment with previous findings using radioisotope 32P-labeled substrates, it is confirmed via biotin-labeled EMSA that MEIOB (a protein specifically involved in the meiotic recombination) is a DNA-binding protein, that MOV10 (an RNA helicase) resolves biotin-labeled RNA duplex structures, and that MEIOB cleaves biotin-labeled single-stranded DNA. This study demonstrates that biotin is capable of substituting 32P in various nucleic acid-related biochemical assays in vitro.&#160;

Presented here are protocols for in vitro biochemical assays using biotin labels that may be widely&#160;applicable for studying protein-nucleic acid interactions.

Protein-nucleic acid interactions play important roles in biological processes such as transcription, recombination, and RNA metabolism. Experimental ...

Nonradioactive Assay to Measure Polynucleotide Phosphorylation of Small Nucleotide Substrates

Polynucleotide kinases (PNKs) are enzymes that catalyze the phosphorylation of the 5&#39; hydroxyl end of DNA and RNA oligonucleotides. The activity of PNKs can be quantified using direct or indirect approaches. Presented here is a direct, in vitro approach to measure PNK activity that relies on a fluorescently-labeled oligonucleotide substrate and polyacrylamide gel electrophoresis. This approach provides resolution of the phosphorylated products while avoiding the use of radiolabeled substrates. The protocol details how to set up the phosphorylation reaction, prepare and run large polyacrylamide gels, and quantify the reaction products. The most technically challenging part of this assay is pouring and running the large polyacrylamide gels; thus, important details to overcome common difficulties are provided. This protocol was optimized for Grc3, a PNK that assembles into an obligate pre-ribosomal RNA processing complex with its binding partner, the Las1 nuclease. However, this protocol can be adapted to measure the activity of other PNK enzymes. Moreover, this assay can also be modified to determine the effects of different components of the reaction, such as the nucleoside triphosphate, metal ions, and oligonucleotides.

This protocol describes a nonradioactive assay to measure kinase activity of polynucleotide kinases (PNKs) on small DNA and RNA substrates.

Polynucleotide kinases (PNKs) are enzymes that catalyze the phosphorylation of the 5&#39; hydroxyl end of DNA and RNA oligonucleotides. The activity of ...

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment screening. NMR-based activity assays are ideally suited to work at the higher concentrations of test compounds required to detect these weaker inhibitors. The dynamic range and chemical shift dispersion in an NMR experiment can easily resolve resonances from substrate, product, and test compounds. This contrasts with spectrophotometric assays, in which read-out interference problems often arise from compounds with overlapping UV-vis absorption profiles. In addition, since they lack reporter enzymes, the single-enzyme NMR assays are not prone to coupled-assay false positives. This attribute makes them useful as orthogonal assays, complementing traditional high throughput screening assays and benchtop triage assays. Detailed protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells. The methods are demonstrated using two nucleoside ribohydrolase enzymes. The use of 1H NMR is shown for the purine-specific enzyme, while 19F NMR is shown for the pyrimidine-specific enzyme. The protocols are generally applicable to any enzyme where substrate and product resonances can be observed and distinguished by NMR spectroscopy. To be the most useful in the context of drug discovery, the final concentration of substrate should be no more than 2-3x its Km value. The choice of NMR experiment depends on the enzyme reaction and substrates available as well as available NMR instrumentation.

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR-based activity assays have been developed to identify and characterize inhibitors of two nucleoside ribohydrolase enzymes. Protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells.

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment ...

Chemistry

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) establish and maintain the DNA methylation pattern. Changes to the normal DNA methylation pattern are linked to cancer development and progression, making DNMTs potential cancer drug targets. Thus, identifying and characterizing novel small molecule inhibitors of these enzymes is of great importance. This paper presents a protocol that can be used to screen for DNA methyltransferase inhibitors. The continuous coupled kinetics assay allows for initial velocities of DNA methylation to be determined in the presence and absence of potential small molecule inhibitors. The assay uses the methyl-sensitive endonuclease Gla I to couple methylation of a hemimethylated DNA substrate to fluorescence generation.
This continuous assay allows for enzyme activity to be monitored in real time. Conducting the assay in small volumes in microtiter plates reduces the cost of reagents. Using this assay, a small example screen was conducted for inhibitors of DNMT1, the most abundant DNMT isozyme in humans. The highly substituted anthraquinone natural product, laccaic acid A, is a potent, DNA-competitive inhibitor of DNMT1. Here, we examine three potential small molecule inhibitors &#8212; anthraquinones or anthraquinone-like molecules with one to three substituents &#8212; at two concentrations to describe the assay protocol. Initial velocities are used to calculate the percent activity observed in the presence of each molecule. One of three compounds examined exhibits concentration-dependent inhibition of DNMT1 activity, indicating that it is a potential inhibitor of DNMT1.

DNA methyltransferases are potential cancer drug targets. Here, a protocol is presented to assess small molecules for DNA methyltransferase inhibition. This assay utilizes an endonuclease to couple DNA methylation to fluorescence generation and allows for enzyme activity to be monitored in real time.&#160;

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) ...

Precision DNAzyme Nanomachines for Discriminative RNA and DNA Analysis

DNAzyme 10-23 - Based Nanomachines for Nucleic Acid Recognition

DNAzyme-based nanomachines (DNM) for the detection of DNA and RNA sequences (analytes) are multifunctional structures made of oligonucleotides. Their functions include tight analyte binding, highly selective analyte recognition, fluorescent signal amplification by multiple catalytic cleavages of a fluorogenic reporter substrate, and fluorogenic substrate attraction for an increase in sensor response. Functional units are attached to a common DNA scaffold for their cooperative action. The RNA-cleaving 10-23 DNMs feature&#160;improved sensitivity in comparison with non-catalytic hybridization probes. The stability of the DNM and the increased chances of substrate recognition are provided&#160;by a double-stranded DNA fragment,&#160;a tile. DNM can differentiate two analytes with a single nucleotide difference in a folded RNA and a double-stranded DNA and detect analytes at concentrations ~1000 times lower than other protein-free hybridization probes. This article presents the concept behind the diagnostic potential of DNA-nanomachine activity and overviews DNM design, assembly, and application&#160;in nucleic acid detection assays.

Author Spotlight: Advancements in DNA Nanosensors &#8211; Addressing Sensitivity and Selectivity Challenges in Molecular Detection

The DNAzyme-based nanomachines can be used for highly selective and sensitive detection of nucleic acids. This article describes a detailed protocol for the design of DNAzyme-based nanomachines with a 10-23 core using free software and their application in the detection of an Epstein-Barr virus fragment as an example.

DNAzyme-based nanomachines (DNM) for the detection of DNA and RNA sequences (analytes) are multifunctional structures made of oligonucleotides. Their ...

Assays for DNA Enzymes Using Synthetic Oligonucleotides

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.

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.

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

A Fluorescence-based Exonuclease Assay to Characterize DmWRNexo, Orthologue of Human Progeroid WRN Exonuclease, and Its Application to Other Nucleases

WRN exonuclease is involved in resolving DNA damage that occurs either during DNA replication or following exposure to endogenous or exogenous genotoxins. It is likely to play a role in preventing accumulation of recombinogenic intermediates that would otherwise accumulate at transiently stalled replication forks, consistent with a hyper-recombinant phenotype of cells lacking WRN. In humans, the exonuclease domain comprises an N-terminal portion of a much larger protein that also possesses helicase activity, together with additional sites important for DNA and protein interaction. By contrast, in Drosophila, the exonuclease activity of WRN (DmWRNexo) is encoded by a distinct genetic locus from the presumptive helicase, allowing biochemical (and genetic) dissection of the role of the exonuclease activity in genome stability mechanisms. Here, we demonstrate a fluorescent method to determine WRN exonuclease activity using purified recombinant DmWRNexo and end-labeled fluorescent oligonucleotides. This system allows greater reproducibility than radioactive assays as the substrate oligonucleotides remain stable for months, and provides a safer and relatively rapid method for detailed analysis of nuclease activity, permitting determination of nuclease polarity, processivity, and substrate preferences.

Exonucleases play critical roles in ensuring genome stability. Loss of WRN exonuclease function results in premature aging. Studying substrates and other requirements of the nuclease in vitro can help elucidate its role in vivo. Here we demonstrate a rapid and reproducible fluorescence-based assay to measure its nuclease activity.

WRN exonuclease is involved in resolving DNA damage that occurs either during DNA replication or following exposure to endogenous or exogenous genotoxins. ...

Biology

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and DNA accessibility by catalyzing ATP-dependent nucleosome remodeling. Biochemically dissecting the contributions of individual subunits of such complexes to the multi-step ATP-dependent chromatin remodeling reaction requires the use of assays that monitor the production of reaction products and measure the formation of reaction intermediates. This JOVE protocol describes assays that allow one to measure the biochemical activities of chromatin remodeling complexes or subcomplexes containing various combinations of subunits. Chromatin remodeling is measured using an ATP-dependent nucleosome sliding assay, which monitors the movement of a nucleosome on a DNA molecule using an electrophoretic mobility shift assay (EMSA)-based method. Nucleosome binding activity is measured by monitoring the formation of remodeling complex-bound mononucleosomes using a similar EMSA-based method, and DNA- or nucleosome-dependent ATPase activity is assayed using thin layer chromatography (TLC) to measure the rate of conversion of ATP to ADP and phosphate in the presence of either DNA or nucleosomes. Using these assays, one can examine the functions of subunits of a chromatin remodeling complex by comparing the activities of the complete complex to those lacking one or more subunits. The human INO80 chromatin remodeling complex is used as an example; however, the methods described here can be adapted to the study of other chromatin remodeling complexes.

Here we describe biochemical assays that can be used to characterize ATP-dependent chromatin remodeling enzymes for their abilities to 1) catalyze ATP-dependent nucleosome sliding, 2) engage with nucleosome substrates, and 3) hydrolyze ATP in a nucleosome- or DNA-dependent manner.

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and ...

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable of measuring SSDC in many single DNA molecules simultaneously. Bead-tethered substrate DNAs, each containing a single copy of the target sequence, are prepared in a microfluidic flow channel. An external magnet applies a weak force to the paramagnetic beads. The integrity of up to 1,000 individual DNAs can be monitored by visualizing the microbeads under darkfield imaging using a wide-field, low magnification objective. Injecting of a restriction endonuclease, NdeI, initiates the cleavage reaction. Video microscopy is used to record the exact moment of each DNA cleavage by observing the frame in which the associated bead moves up and out of the focal plane of the objective. Frame-by-frame bead counting quantifies the reaction, and an exponential fit determines the reaction rate. This method allows collection of quantitative and statistically significant data on single molecule SSDC reactions in a single experiment.

A highly parallel method for measuring the site-specific cleavage of DNA at the single molecule level is described. This protocol demonstrates the technique using the restriction endonuclease NdeI. The method can easily be modified to study any process that results in site-specific DNA cleavage.

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable ...

Kinetic Screening of Nuclease Activity using Nucleic Acid Probes

Summary

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Chapters in this video

Kinetics of Lagging-strand DNA Synthesis In Vitro by the Bacteriophage T7 Replication Proteins

In Vitro Biochemical Assays using Biotin Labels to Study Protein-Nucleic Acid Interactions

Nonradioactive Assay to Measure Polynucleotide Phosphorylation of Small Nucleotide Substrates

NMR-Based Activity Assays for Determining Compound Inhibition, IC₅₀ Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors

DNAzyme 10-23 - Based Nanomachines for Nucleic Acid Recognition

Using Modified Synthetic Oligonucleotides to Assay Nucleic Acid-Metabolizing Enzymes

A Fluorescence-based Exonuclease Assay to Characterize DmWRNexo, Orthologue of Human Progeroid WRN Exonuclease, and Its Application to Other Nucleases

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage