We thank Dr. Julie K. Horton for critical reading of the manuscript and Dr. Rajendra Prasad for helpful suggestions and discussions. Portions of this research were originally published in The Journal of Biological Chemistry, Sassa A et. al., "DNA Sequence Context Effects on the Glycosylase Activity of Human 8-Oxoguanine DNA Glycosylase." J Biol Chem. 287, 36702-36710 (2012) 2. This work was supported, in whole or in part, by National Institutes of Health Research Project Grant Z01-ES050158 in the Intramural Research Program, NIEHS.

1. Preparation of Enzyme and DNA Substrate
<ol>
 <li>Over-express OGG1 as a GST-fusion protein in E. coli, utilize the GST-tag for purification, and then remove the GST-tag by cleavage with HRV-3C protease (Figure 1)6.</li>
 <li>Purchase the chemically synthesized 5'-6-carboxyfluorescein (6-FAM) labeled oligonucleotide containing a single 8-oxoG residue and its complementary unlabeled oligonucleotide strand. The 34-mer oligonucleotides contain an 8-oxoG at position 17 from the 5'-end. Purify these oligonucleotides by polyacrylamide gel electrophoresis.</li>
 <li>Prepare double-stranded DNA substrate containing 8-oxoG by mixing 5'-6-FAM labeled oligonucleotide with its complementary strand at a molar ratio of 1:1.2 in annealing buffer (10 mM Tris-HCl, pH 7.5, 50 mM NaCl, 1 mM EDTA) in a 1.5 ml microfuge tube. Place the tube in a float in boiling water for 5 min. Leave the tube in place and then let the water cool slowly to room temperature (this takes approximately 2 hr).</li>
 <li>Perform sample preparation and the experiment under low or minimal light conditions so as to minimize photobleaching of the fluorescent label.</li>
</ol>
2. Measurement of Steady-State Time Course and Active Site Titration of OGG1
2.1. Sample preparation and steady-state time course
<ol>
 <li>Prepare DNA substrate and OGG1 solutions separately in reaction buffer (50 mM HEPES, pH 7.5, 20 mM KCl, 0.5 mM EDTA, and 0.1% bovine serum albumin) in 1.5 ml microfuge tubes on ice. The concentration of DNA is 400 nM and the apparent concentration of OGG1 is 30, 60, 90, or 120 nM as determined by a Bradford protein assay. Place the reaction tubes in a heat block set at 37 &deg;C. Pre-incubate enzyme and DNA substrate solutions separately at 37 &deg;C for 1 min.</li>
 <li>Start the reaction by mixing equal volumes of the OGG1 and DNA substrate solutions by pipetting. After mixing these solutions 1:1 (v/v), the final concentrations are 200 nM DNA and 15, 30, 45, or 60 nM OGG1, respectively. Remove aliquots (10 &mu;l) at time intervals and quench the reaction by mixing with 1 &mu;l of 1 M NaOH. As the control for the time courses, 10 &mu;l of the reaction mixture without enzyme is mixed with 1 &mu;l of 1 M NaOH.</li>
 <li>Place the reaction samples in a 90 &deg;C heat block for 5 min to cleave the resulting product AP-site. After heating, add 1 &mu;l of 1 M HCl to neutralize each sample.</li>
 <li>Add an equal volume (12 &mu;l) of gel loading buffer (95 % formamide, 20 mM EDTA, 0.02% bromophenol, and 0.02% xylene cyanol) to each reaction sample, and then place the mixture in a heat block set at 95 &deg;C for 2 min, then immediately place the tube on ice.</li>
 <li>Load the samples (5 &mu;l) onto a 15% denaturing polyacrylamide gel containing 8 M urea in 89 mM Tris-HCl, pH 8.8, 89 mM boric acid, and 2 mM EDTA. The substrate and the cleaved products are well separated after running the gel.</li>
</ol>
2.2. Imaging of the gel
<ol>
 <li>Scan the gel using an imager that can detect the fluorescently labeled DNA and visualize the substrate and product bands.</li>
 <li>Quantify the bands after imaging the gel. Note that some background cleavage may be observed after treatment of the substrate itself with NaOH (Figure 2A). Subtract this background from the measured amount of each reaction product.</li>
</ol>
2.3. Data analysis
<ol>
 <li>Plot the amount of the product formed at each reaction time (t) (Figure 2B). Analyze the raw data using Equation 1 to determine the amplitude of the burst (A0, y-intercept) and the slope of the linear steady-state phase (vss).</li>
</ol>
 <img fo:src="/files/ftp_upload/50695/50695eq1highres.jpg" src="/files/ftp_upload/50695/50695eq1.jpg" /> 
<ol start="2">
 <li>Plot the y-intercept relative to the total protein concentration (i.e. apparent enzyme concentration; Figure 2C), providing a measure of the active fraction of enzyme. Use a linear fit with a zero intercept to provide the correction factor for determining the fraction of active enzyme.</li>
 <li>Plot the steady-state rate relative to the active enzyme concentration (Figure 2D), providing the product dissociation rate constant (i.e. slope of the fitted line).</li>
</ol>
3. Measurement of Pre-steady-state Time Course
As shown in Figure 2, product formation during the burst phase is too fast to measure by manual mixing and quenching. Hence, a rapid quench-flow instrument can provide a powerful method to measure rapid reactions that occur on the millisecond time scale (Figure 3)5. The instrument uses a computer-controlled drive motor to rapidly mix and quench reactions after specified reaction times. For example, use a Kintek RQF-3 Rapid Quench-Flow Instrument to measure the initial burst phase and subsequent steady-state phase of product formation catalyzed by OGG1. Rapid Quench-Flow Instruments are available from several manufacturers.
3.1. Preparation of sample
Separately prepare DNA substrate and OGG1 solutions in 1.5 ml tubes as described in 2-1. The concentrations of DNA and active OGG1 are 400 nM and 80 nM, respectively. This results in final concentrations of 200 nM DNA and 40 nM active OGG1 after mixing 1:1 (v/v).
3.2. Preparation of rapid quench-flow instrument
<ol>
 <li>Connect a circulating water bath to the rapid quench-flow Instrument for temperature control (37 &deg;C).</li>
 <li>Adjust the instrument parameters and select the appropriate reaction loop for the desired time points according to the manufacturer's instructions. The reaction loops are of variable lengths to provide alternate reaction times.</li>
 <li>Prepare reaction buffer and NaOH quench solutions in 10 ml Luer Lock disposable syringes and attach these syringes to the Drive Ports and load the Drive Reservoirs with reaction buffer (syringes B) and 142 mM NaOH (syringe Q) (Syringe Load Valves in LOAD position).To remove the air bubbles from the syringe, work the solution back and forth several times.</li>
 <li>Lower the stepper motor until it contacts the top of the syringes (Syringe Load valves and Sample Load valves in FIRE position).</li>
 <li>Flush the Sample Loops, the Reaction Loop, and the Exit Line with water and methanol. Dry the flushed areas completely (valves in FLUSH position).</li>
</ol>
3.3. Pre-steady-state time course
<ol>
 <li>Make a hole in the top of a capped 1.5 ml tube using a 16-gauge needle. Attach a tube at the Exit Line to collect the quenched reaction.</li>
 <li>Set the desired reaction time (in seconds) using the keypad. The Stepper Motor will back up to adjust for loop volume so as to maintain constant quench volume. Position plungers for syringe B against the Stepper Motor platform by adding buffer with the disposable syringes attached to the Drive Ports.</li>
 <li>Fill 1 ml Luer Lock disposable syringes with DNA substrate and OGG1 solutions, respectively (sample valves in LOAD position). Attach syringes with DNA substrate and OGG1 solutions to each of the Sample Load Ports, and then fill the Sample Loops with DNA substrate and OGG1 solutions, respectively (Sample Load Valves in LOAD position).</li>
 <li>Set all Syringe Load Valves and Sample Load Valves to the FIRE position. Start the reaction by a keypad stroke (e.g. press of "G" or "START" key). DNA substrate and OGG1 solutions (18 &mu;l each) are immediately mixed in the Reaction Loop.</li>
 <li>Wait until the reaction is automatically quenched at the desired reaction time by mixing 36 &mu;l of the reaction mixture with 86 &mu;l of 142 mM NaOH. After quenching the reaction, the sample is discharged from the Exit Line.</li>
 <li>Set the Syringe Load Valves to the LOAD position and the Sample Load Valves to the FLUSH position. Flush the Sample Loops, the Reaction Loop, and the Exit Line with water and methanol and dry as described in 3.2. Repeat the above procedure for each time point.</li>
 <li>Perform a control experiment without enzyme to determine a background correction, which can be done manually or with the rapid-quench instrument. To perform a control with the rapid-quench instrument, fill the Sample Loop for DNA with DNA substrate but keep the Sample Loop for OGG1 empty. Set the reaction time, perform the mix and quench as described in 3.3.4-3.3.6.</li>
 <li>Wash the Sample Loops and the Reaction Loop with 2 M NaOH, 2 M HCl, water, and methanol and then dry the washed lines. After setting the Stepper Motor to home position, switch the Syringe Load Valves to the LOAD position and wash the Drive Reservoirs with water.</li>
 <li>Treat the quenched reaction samples with heat, and then separate substrate and product DNA by 15% denaturing polyacrylamide gel electrophoresis as described in 2.1.</li>
 <li>Visualize and quantify the bands on the gel as described in 2.2.</li>
</ol>
3.4. Data analysis
Fit the time courses of product formation by non-linear regression analysis to an equation with a rising exponential and linear terms (Equation 2) providing the first-order rate constant (kobs), the amplitude of the burst (A0), and a linear rate (vss).
<img fo:src="/files/ftp_upload/50695/50695eq2highres.jpg" src="/files/ftp_upload/50695/50695eq2.jpg" /> 
4. Single-turnover Time Course
<ol>
 <li>Prepare DNA substrate and OGG1 in separate 1.5 ml tubes as described in 2-1. The concentrations of DNA and active OGG1 are 100 nM and 500 nM, respectively; this yields final concentrations of 50 nM DNA and 250 nM OGG1 after mixing 1:1 (v/v).</li>
 <li>Prepare the rapid quench-flow instrument and prepare the reaction samples as described in 3.2 and 3.3.</li>
 <li>Treat the quenched reaction samples with heat and subject them to 15% denaturing polyacrylamide gel electrophoresis to separate substrate and products as described in 2.1.</li>
 <li>Visualize and quantify the bands on the gel as described in 2.2.</li>
 <li>Fit the time courses of product formation to a single exponential to determine the first-order rate constant (kobs) as given in Equation 3.</li>
</ol>
<img fo:src="/files/ftp_upload/50695/50695eq3highres.jpg" src="/files/ftp_upload/50695/50695eq3.jpg" />

<ol>
	<li>Porello, S. L., Leyes, A. E., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-turnover+and+pre-steady-state+kinetics+of+the+reaction+of+the+adenine+glycosylase+MutY+with+mismatch-containing+DNA+substrates.">Single-turnover and pre-steady-state kinetics of the reaction of the adenine glycosylase MutY with mismatch-containing DNA substrates.</a> Biochemistry. 37, 14756-14764 (1998).</li><li>Sassa, A., Beard, W. A., Prasad, R., Wilson, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+sequence+context+effects+on+the+glycosylase+activity+of+human+8-oxoguanine+DNA+glycosylase.">DNA sequence context effects on the glycosylase activity of human 8-oxoguanine DNA glycosylase.</a> The Journal of Biological Chemistry. 287, 36702-36710 (2012).</li><li>Wong, I., Lundquist, A. J., Bernards, A. S., Mosbaugh, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presteady-state+analysis+of+a+single+catalytic+turnover+by+Escherichia+coli+uracil-DNA+glycosylase+reveals+a+"pinch-pull-push"+mechanism.">Presteady-state analysis of a single catalytic turnover by Escherichia coli uracil-DNA glycosylase reveals a "pinch-pull-push" mechanism.</a> The Journal of Biological Chemistry. 277, 19424-19432 (1074).</li><li>Johnson, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+transient-state+kinetics.">Advances in transient-state kinetics.</a> Current Opinion in Biotechnology. 9, 87-89 (1998).</li><li>Johnson, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+kinetic+analysis+of+mechanochemical+adenosinetriphosphatases.">Rapid kinetic analysis of mechanochemical adenosinetriphosphatases.</a> Methods in Enzymology. 134, 677-705 (1986).</li><li>Kovtun, I. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OGG1+initiates+age-dependent+CAG+trinucleotide+expansion+in+somatic+cells.">OGG1 initiates age-dependent CAG trinucleotide expansion in somatic cells.</a> Nature. 447, 447-452 (2007).</li><li>Hill, J. W., Hazra, T. K., Izumi, T., Mitra, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stimulation+of+human+8-oxoguanine-DNA+glycosylase+by+AP-endonuclease:+potential+coordination+of+the+initial+steps+in+base+excision+repair.">Stimulation of human 8-oxoguanine-DNA glycosylase by AP-endonuclease: potential coordination of the initial steps in base excision repair.</a> Nucleic Acids Research. 29, 430-438 (2001).</li><li>Zharkov, D. O., Rosenquist, T. A., Gerchman, S. E., Grollman, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Substrate+specificity+and+reaction+mechanism+of+murine+8-oxoguanine-DNA+glycosylase.">Substrate specificity and reaction mechanism of murine 8-oxoguanine-DNA glycosylase.</a> The Journal of Biological Chemistry. 275, 28607-28617 (2000).</li><li>Waters, T. R., Swann, P. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetics+of+the+action+of+thymine+DNA+glycosylase.">Kinetics of the action of thymine DNA glycosylase.</a> The Journal of Biological Chemistry. 273, 20007-20014 (1998).</li><li>Nilsen, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excision+of+deaminated+cytosine+from+the+vertebrate+genome:+role+of+the+SMUG1+uracil-DNA+glycosylase.">Excision of deaminated cytosine from the vertebrate genome: role of the SMUG1 uracil-DNA glycosylase.</a> The EMBO Journal. 20, 4278-4286 (2001).</li><li>Gill, S. C., von Hippel, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calculation+of+protein+extinction+coefficients+from+amino+acid+sequence+data.">Calculation of protein extinction coefficients from amino acid sequence data.</a> Analytical Biochemistry. 182, 319-326 (1989).</li><li>Van de Berg, ., Beard, B. J., A, W., Wilson, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+structure+and+aspartate+276+influence+nucleotide+binding+to+human+DNA+polymerase+beta.+Implication+for+the+identity+of+the+rate-limiting+conformational+change.">DNA structure and aspartate 276 influence nucleotide binding to human DNA polymerase beta. Implication for the identity of the rate-limiting conformational change.</a> The Journal of Biological Chemistry. 276, 3408-3416 (2001).</li><li>Krishnamurthy, N., Haraguchi, K., Greenberg, M. M., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+removal+of+formamidopyrimidines+by+8-oxoguanine+glycosylases.">Efficient removal of formamidopyrimidines by 8-oxoguanine glycosylases.</a> Biochemistry. 47, 1043-1050 (2008).</li><li>Leipold, M. D., Muller, J. G., Burrows, C. J., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removal+of+hydantoin+products+of+8-oxoguanine+oxidation+by+the+Escherichia+coli+DNA+repair+enzyme,+FPG.">Removal of hydantoin products of 8-oxoguanine oxidation by the Escherichia coli DNA repair enzyme, FPG.</a> Biochemistry. 39, 14984-14992 (2000).</li><li>Zhao, X., Krishnamurthy, N., Burrows, C. J., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutation+versus+repair:+NEIL1+removal+of+hydantoin+lesions+in+single-stranded,+bulge,+bubble,+and+duplex+DNA+contexts.">Mutation versus repair: NEIL1 removal of hydantoin lesions in single-stranded, bulge, bubble, and duplex DNA contexts.</a> Biochemistry. 49, 1658-1666 (2010).</li><li>Leipold, M. D., Workman, H., Muller, J. G., Burrows, C. J., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recognition+and+removal+of+oxidized+guanines+in+duplex+DNA+by+the+base+excision+repair+enzymes+hOGG1,+yOGG1,+and+yOGG2.">Recognition and removal of oxidized guanines in duplex DNA by the base excision repair enzymes hOGG1, yOGG1, and yOGG2.</a> Biochemistry. 42, 11373-11381 (2003).</li><li>Robey-Bond, S. M., Barrantes-Reynolds, R., Bond, J. P., Wallace, S. S., Bandaru, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clostridium+acetobutylicum+8-oxoguanine+DNA+glycosylase+(Ogg)+differs+from+eukaryotic+Oggs+with+respect+to+opposite+base+discrimination.">Clostridium acetobutylicum 8-oxoguanine DNA glycosylase (Ogg) differs from eukaryotic Oggs with respect to opposite base discrimination.</a> Biochemistry. 47, 7626-7636 (2008).</li><li>Jarem, D. A., Wilson, N. R., Delaney, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure-dependent+DNA+damage+and+repair+in+a+trinucleotide+repeat+sequence.">Structure-dependent DNA damage and repair in a trinucleotide repeat sequence.</a> Biochemistry. 48, 6655-6663 (2009).</li><li>Livingston, A. L., Kundu, S., Henderson Pozzi, M., Anderson, W. D., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insight+into+the+roles+of+tyrosine+82+and+glycine+253+in+the+Escherichia+coli+adenine+glycosylase+MutY.">Insight into the roles of tyrosine 82 and glycine 253 in the Escherichia coli adenine glycosylase MutY.</a> Biochemistry. 44, 14179-14190 (2005).</li><li>Kundu, S., Brinkmeyer, M. K., Livingston, A. L., David, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adenine+removal+activity+and+bacterial+complementation+with+the+human+MutY+homologue+(MUTYH)+and+Y165C,+G382D,+P391L+and+Q324R+variants+associated+with+colorectal+cancer.">Adenine removal activity and bacterial complementation with the human MutY homologue (MUTYH) and Y165C, G382D, P391L and Q324R variants associated with colorectal cancer.</a> DNA Repair. 8, 1400-1410 (2009).</li><li>Zharkov, D. O., Rosenquist, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inactivation+of+mammalian+8-oxoguanine-DNA+glycosylase+by+cadmium(II):+implications+for+cadmium+genotoxicity.">Inactivation of mammalian 8-oxoguanine-DNA glycosylase by cadmium(II): implications for cadmium genotoxicity.</a> DNA Repair. 1, 661-670 (2002).</li><li>Jarem, D. A., Wilson, N. R., Schermerhorn, K. M., Delaney, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incidence+and+persistence+of+8-oxo-7,8-dihydroguanine+within+a+hairpin+intermediate+exacerbates+a+toxic+oxidation+cycle+associated+with+trinucleotide+repeat+expansion.">Incidence and persistence of 8-oxo-7,8-dihydroguanine within a hairpin intermediate exacerbates a toxic oxidation cycle associated with trinucleotide repeat expansion.</a> DNA Repair. 10, 887-896 (2011).</li><li>Mokkapati, S. K., Wiederhold, L., Hazra, T. K., Mitra, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stimulation+of+DNA+glycosylase+activity+of+OGG1+by+NEIL1:+functional+collaboration+between+two+human+DNA+glycosylases.">Stimulation of DNA glycosylase activity of OGG1 by NEIL1: functional collaboration between two human DNA glycosylases.</a> Biochemistry. 43, 11596-11604 (2004).</li><li>Sidorenko, V. S., Nevinsky, G. A., Zharkov, D. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanism+of+interaction+between+human+8-oxoguanine-DNA+glycosylase+and+AP+endonuclease.">Mechanism of interaction between human 8-oxoguanine-DNA glycosylase and AP endonuclease.</a> DNA Repair. 6, 317-328 (2007).</li></ol>

Authors have nothing to disclose.

The kinetic approaches described here outline methods to define elementary kinetic constants. If a time course of product formation is biphasic with the first enzymatic turnover occurring rapidly, then a step after chemistry is rate-limiting during subsequent catalytic turnovers. In the case of OGG1, the first turnover can be measured using high enzyme concentrations with either limiting (S&lt;E) or high (S&gt;E) DNA concentrations. In the first case, the reaction is limited to a 'single-turnover' and provides a measure ...

An aerobic environment hastens genomic instability. A major promutagenic DNA lesion resulting from oxidative stress is 7,8-dihydro-8-oxoguanine (8-oxoG). This is due to the ambiguous coding potential of 8-oxoG. Human 8-oxoguanine DNA glycosylase (OGG1) is responsible for initiating base excision repair of 8-oxoG. The glycosylase activity of OGG1 excises the 8-oxoG base resulting in product DNA with an apurinic site (AP-site). A weak lyase activity of OGG1 can incise the AP-site in some instances.
Kinetic characterization of DNA glycosylases generally finds that they exhibit biphasic time courses. After an initial fast phase of product formation (i.e. burst), a linear steady-state phase is observed1-3. This behavior is indicative of a step following chemistry (i.e. conformational change or product release) being rate-limiting during the linear portion of the time course, whereas the burst phase, often referred to as the transient phase, corresponds to product formation at the enzyme active site during the first cycle of the reaction. When product release is rate limiting during the steady-state phase, activity measurements provide a qualitative measure of product DNA binding affinity, but do not provide kinetic information concerning events at the enzyme active site (i.e. chemistry). Accordingly, methods to isolate and measure the exponential pre-steady-state burst phase are needed to probe events during the first enzymatic turnover at the enzyme's active site4.
There are three standard kinetic approaches to characterize the catalytic behavior of OGG1, (1) steady-state, (2) pre-steady-state, and (3) single-turnover. These approaches differ from one another by the concentration of enzyme in the reaction mixture and the enzyme to substrate ratio utilized in each approach. In a typical steady-state approach, sometimes referred to as multiple turnover kinetics, low concentrations of enzyme are used to follow product formation. The substrate concentration greatly exceeds the enzyme concentration so that multiple enzymatic turnovers do not significantly affect substrate concentration. In this situation, time courses should be linear and it is often difficult to discern whether a burst occurred during the first turnover due to the low enzyme concentration used in this approach; note that burst amplitude is equivalent to the enzyme concentration. This can be overcome by using a higher enzyme concentration and extrapolating the linear time course to zero time to detect whether the first enzymatic turnover occurred quickly. The intercept on the ordinate (y-axis) should be proportional to the enzyme concentration and provides a measure of the enzyme actively engaged with substrate. Although this approach can in principle provide evidence for the existence of a burst phase, a different approach is required to measure the kinetics of the burst phase. In many instances the burst phase is too fast to measure by manual mixing and quenching techniques. In this situation, pre-steady-state and single-turnover kinetic (i.e. transient kinetic) approaches often require a rapid-mixing and quenching instrument to follow early time points of a reaction5. In a pre-steady-state approach high concentrations of enzyme are used so that a significant amount of product is formed during the first turnover. Since multiple turnovers are followed to observe catalytic cycling (i.e. the linear phase that follows the burst), substrate concentration is greater than the enzyme concentration ([enzyme] &lt; [substrate]). To isolate events at the active site of the enzyme without catalytic cycling, single-turnover conditions are utilized. In this case, substrate is saturated with enzyme (E&gt;&gt;S) so that all of the substrate will participate in the 'single turnover' and will typically exhibit a single-exponential time course.
As noted above for enzymes that exhibit a burst phase, product release (koff) often limits the rate of the steady-state phase of the time course. The rate of product release (vss, conc./time) can be determined from the slope of the linear steady-state phase. The active enzyme concentration (E) is needed to convert the rate of product release to an intrinsic rate constant where koff = vss/[E]. Importantly, the active enzyme concentration is typically lower than the measured protein concentration due to impurities, inactive enzyme, enzyme non-productively bound to substrate, and the method used to determine protein concentration. The active enzyme concentration can be determined from the burst amplitude when product release is slow. Thus, extrapolating a steady-state time course to zero time provides an estimate of active enzyme needed to calculate koff (product release) from the observed steady-state rate.
To measure the kinetics of the burst, a pre-steady-state approach is necessary to follow product formation during the first turnover that occurs prior to the linear steady-state phase. The burst kinetics follows the formation of enzyme-product intermediate. Once the reaction is initiated by mixing enzyme with substrate, the amount of the enzyme-product rapidly increases until the reaction reaches a steady state phase. If catalysis is much more rapid than product release, the amplitude of the burst is equal to actively engaged enzyme and the observed exponential approach to equilibrium (kobs) corresponds to the rate of chemical conversion of substrate to product, assuming that the reverse rate of chemistry is negligible.
In some instances catalytic cycling interferes with a pre-steady-state analysis, such as when the magnitudes of rates for chemistry and product release are not significantly different. In this case, employing excess enzyme relative to substrate prevents catalytic cycling and limits substrate bound with enzyme to a single turnover. Accordingly, the first chemical step of the reaction can be isolated and accurately determined as the first-order rate constant (kobs). This rate constant should be similar to kobs determined from the pre-steady-state approach described above.
Here we describe how these kinetic approaches can be used to analyze the glycosylase activity of OGG1.

<table><tbody><tr><td colspan="4">&lt;strong&gt;Reagent/Material&lt;/strong&gt;</td></tr><tr><td>5&amp;rsquo;-6-FAM labeled oligonucleotides containing a single 8-oxoG</td><td>Eurofins MWG Operon</td><td></td><td>5&amp;rsquo;-P32 radiolabeled oligonucleotides can be used as well. Polyacrylamide gel purification grade is recommended. </td></tr><tr><td>Unlabeled oligonucleotides (complementary strand)</td><td>Eurofins MWG Operon</td><td></td><td>Polyacrylamide gel purification grade is recommended. </td></tr><tr><td>1 ml BD Luer Lock disposable syringe</td><td>BD Medical</td><td>309628</td><td>Lue Lock disposable syringe from other vendors can be used as well.</td></tr><tr><td>10 ml BD Luer Lock disposable syringe</td><td>BE Medical</td><td>309604</td><td>Luer Lock disposable syringe from other vendors can be used as well.</td></tr><tr><td colspan="4">&lt;strong&gt;Equipment&lt;/strong&gt;</td></tr><tr><td>Circulating water bath</td><td>Any vender</td><td></td><td></td></tr><tr><td>RQF-3 Rapid Quench-Flow Instrument</td><td>KinTek Corporation</td><td></td><td>Rapid Quench-Flow Instrument from other vendors can be used as well.</td></tr><tr><td>Typhoon Phosphorimager 8600</td><td>GE Healthcare Life Sciences</td><td></td><td>Imager from other vendors can be used as well.</td></tr><tr><td>KaleidaGraph</td><td>Synergy Software</td><td></td><td></td></tr></tbody></table>

steady-state, pre-steady-state, and single-turnover kinetic measurement for dna glycosylase activity

Human 8-oxoguanine DNA glycosylase (OGG1) excises the mutagenic oxidative DNA lesion 8-oxo-7,8-dihydroguanine (8-oxoG) from DNA. Kinetic characterization of OGG1 is undertaken to measure the rates of 8-oxoG excision and product release. When the OGG1 concentration is lower than substrate DNA, time courses of product formation are biphasic; a rapid exponential phase (i.e. burst) of product formation is followed by a linear steady-state phase. The initial burst of product formation corresponds to the concentration of enzyme properly engaged on the substrate, and the burst amplitude depends on the concentration of enzyme. The first-order rate constant of the burst corresponds to the intrinsic rate of 8-oxoG excision and the slower steady-state rate measures the rate of product release (product DNA dissociation rate constant, koff). Here, we describe steady-state, pre-steady-state, and single-turnover approaches to isolate and measure specific steps during OGG1 catalytic cycling. A fluorescent labeled lesion-containing oligonucleotide and purified OGG1 are used to facilitate precise kinetic measurements. Since low enzyme concentrations are used to make steady-state measurements, manual mixing of reagents and quenching of the reaction can be performed to ascertain the steady-state rate (koff). Additionally, extrapolation of the steady-state rate to a point on the ordinate at zero time indicates that a burst of product formation occurred during the first turnover (i.e. y-intercept is positive). The first-order rate constant of the exponential burst phase can be measured using a rapid mixing and quenching technique that examines the amount of product formed at short time intervals (&lt;1 sec) before the steady-state phase and corresponds to the rate of 8-oxoG excision (i.e. chemistry). The chemical step can also be measured using a single-turnover approach where catalytic cycling is prevented by saturating substrate DNA with enzyme (E&gt;S). These approaches can measure elementary rate constants that influence the efficiency of removal of a DNA lesion.

Steady-state kinetic analysis was performed by using 200 nM DNA substrate and four different apparent concentrations of OGG1 (15, 30, 45, and 60 nM) as determined by a Bradford protein assay2. The time courses of product formation were fit to a linear equation to determine the y-intercept, which were 2.2, 11, 15, and 26 nM, respectively, relative to each protein concentration (Figure 2B). The y-intercepts were further plotted relative to each actual protein concentration (Figure 2C</st...

Watch this Scientific Journal Video about Steady-state, Pre-steady-state, and Single-turnover Kinetic Measurement for DNA Glycosylase Activity at JoVE.com

Steady-state, Pre-steady-state, and Single-turnover Kinetic Measurement for DNA Glycosylase Activity

Time courses for the glycosylase activity of 8-oxoguanine DNA glycosylase are biphasic exhibiting a burst of product formation and a linear steady-state phase. Utilizing quench-flow techniques, the burst and the steady-state rates can be measured, which correspond to excision of 8-oxoguanine and release of the glycosylase from the product DNA, respectively.

Human 8-oxoguanine DNA glycosylase (OGG1) excises the mutagenic oxidative DNA lesion 8-oxo-7,8-dihydroguanine (8-oxoG) from DNA. Kinetic characterization ...

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Research

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Chemistry

19.3K Views.  NIEHS, National Institutes of Health. Time courses for the glycosylase activity of 8-oxoguanine DNA glycosylase are biphasic exhibiting a burst of product formation and a linear steady-state phase. Utilizing quench-flow techniques, the burst and the steady-state rates can be measured, which correspond to excision of 8-oxoguanine and release of the glycosylase from the product DNA, respectively.

Video: Steady-state, Pre-steady-state, and Single-turnover Kinetic Measurement for DNA Glycosylase Activity

DNA Polymerase Activity Assay Using Near-infrared Fluorescent Labeled DNA Visualized by Acrylamide Gel Electrophoresis

For any enzyme, robust, quantitative methods are required for characterization of both native and engineered enzymes. For DNA polymerases, DNA synthesis can be characterized using an in vitro DNA synthesis assay followed by polyacrylamide gel electrophoresis. The goal of this assay is to quantify synthesis of both natural DNA and modified DNA (M-DNA). These approaches are particularly useful for resolving oligonucleotides with single nucleotide resolution, enabling observation of individual steps during enzymatic oligonucleotide synthesis. These methods have been applied to the evaluation of an array of biochemical and biophysical properties such as the measurement of steady-state rate constants of individual steps of DNA synthesis, the error rate of DNA synthesis, and DNA binding affinity. By using modified components including, but not limited to, modified nucleoside triphosphates (NTP), M-DNA, and/or mutant DNA polymerases, the relative utility of substrate-DNA polymerase pairs can be effectively evaluated. Here, we detail the assay itself, including the changes that must be made to accommodate nontraditional primer DNA labeling strategies such as near-infrared fluorescently labeled DNA. Additionally, we have detailed crucial technical steps for acrylamide gel pouring and running, which can often be technically challenging.

This protocol describes the characterization of DNA polymerase synthesis of modified DNA through observation of changes to near-infrared fluorescently labeled DNA using gel electrophoresis and gel imaging. Acrylamide gels are used for high resolution imaging of the separation of short nucleic acids, which migrate at different rates depending on size.

For any enzyme, robust, quantitative methods are required for characterization of both native and engineered enzymes. For DNA polymerases, DNA synthesis ...

Biochemistry

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. ...

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. Linearized &lambda; DNA is modified to have a biotin on the end of one strand, and a digoxigenin moiety on the other end of the same strand. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead. The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied to exert a drag force on the bead. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate (Figure 1). The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork. Measuring the position of the bead allows precise determination of the rates and processivities of DNA unwinding and polymerization (Figure 2).

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system.

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. ...

Biology

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) ...

Uracil-DNA Glycosylase Assay by Matrix-assisted Laser Desorption/Ionization Time-of-flight Mass Spectrometry Analysis

Uracil-DNA glycosylase (UDG) is a key component in the base excision repair pathway for the correction of uracil formed from hydrolytic deamination of cytosine. Thus, it is crucial for genome integrity maintenance. A highly specific, non-labeled, non-radio-isotopic method was developed to measure UDG activity. A synthetic DNA duplex containing a site-specific uracil was cleaved by UDG and then subjected to Matrix-assisted Laser Desorption/Ionization time-of-flight mass spectrometry (MALDI-TOF MS) analysis. A protocol was established to preserve the apurinic/apyrimidinic site (AP) product in DNA without strand break. The change in the m/z value from the substrate to the product was used to evaluate uracil hydrolysis by UDG. A G:U substrate was used for UDG kinetic analysis yielding the Km = 50 nM, Vmax = 0.98 nM/s, and Kcat = 9.31 s-1. Application of this method to a uracil glycosylase inhibitor (UGI) assay yielded an IC50 value of 7.6 pM. The UDG specificity using uracil at various positions within single-stranded and double-stranded DNA substrates demonstrated different cleavage efficiencies. Thus, this simple, rapid, and versatile MALDI-TOF MS method could be an excellent reference method for various monofunctional DNA glycosylases. It also has the potential as a tool for DNA glycosylase inhibitor screening.

A non-labeled, non-radio-isotopic method to assay uracil-DNA glycosylase activity was developed using MALDI-TOF mass spectrometry for direct apurinic/apyrimidinic site-containing product analysis. The assay proved to be quite simple, specific, rapid, and easy to use for DNA glycosylase measurement.

Uracil-DNA glycosylase (UDG) is a key component in the base excision repair pathway for the correction of uracil formed from hydrolytic deamination of ...

Application of Stopped-flow Kinetics Methods to Investigate the Mechanism of Action of a DNA Repair Protein

Transient kinetic analysis is indispensable for understanding the workings of biological macromolecules, since this approach yields mechanistic information including active site concentrations and intrinsic rate constants that govern macromolecular function. In case of enzymes, for example, transient or pre-steady state measurements identify and characterize individual events in the reaction pathway, whereas steady state measurements only yield overall catalytic efficiency and specificity. Individual events such as protein-protein or protein-ligand interactions and rate-limiting conformational changes often occur in the millisecond timescale, and can be measured directly by stopped-flow and chemical-quench flow methods. Given an optical signal such as fluorescence, stopped-flow serves as a powerful and accessible tool for monitoring reaction progress from substrate binding to product release and catalytic turnover1,2.
Here, we report application of stopped-flow kinetics to probe the mechanism of action of Msh2-Msh6, a eukaryotic DNA repair protein that recognizes base-pair mismatches and insertion/deletion loops in DNA and signals mismatch repair (MMR)3-5. In doing so, Msh2-Msh6 increases the accuracy of DNA replication by three orders of magnitude (error frequency decreases from ~10-6 to10-9 bases), and thus helps preserve genomic integrity. Not surprisingly, defective human Msh2-Msh6 function is associated with hereditary non-polyposis colon cancer and other sporadic cancers6-8. In order to understand the mechanism of action of this critical DNA metabolic protein, we are probing the dynamics of Msh2-Msh6 interaction with mismatched DNA as well as the ATPase activity that fuels its actions in MMR. DNA binding is measured by rapidly mixing Msh2-Msh6 with DNA containing a 2-aminopurine (2-Ap) fluorophore adjacent to a G:T mismatch and monitoring the resulting increase in 2-aminopurine fluorescence in real time. DNA dissociation is measured by mixing pre-formed Msh2-Msh6 G:T(2-Ap) mismatch complex with unlabeled trap DNA and monitoring decrease in fluorescence over time9. Pre-steady state ATPase kinetics are measured by the change in fluorescence of 7-diethylamino-3-((((2-maleimidyl)ethyl)amino)carbonyl) coumarin)-labeled Phosphate Binding Protein (MDCC-PBP) on binding phosphate (Pi) released by Msh2-Msh6 following ATP hydrolysis9,10.
The data reveal rapid binding of Msh2-Msh6 to a G:T mismatch and formation of a long-lived Msh2-Msh6 G:T complex, which in turn results in suppression of ATP hydrolysis and stabilization of the protein in an ATP-bound form. The reaction kinetics provide clear support for the hypothesis that ATP-bound Msh2-Msh6 signals DNA repair on binding a mismatched base pair in the double helix.
F. Noah Biro and Jie Zhai contributed to this paper equally.

Msh2-Msh6 is responsible for initiating repair of replication errors in DNA. Here we present a transient kinetics approach towards understanding how this critical protein works. The report illustrates stopped-flow experiments for measuring the coupled DNA binding and ATPase kinetics underlying Msh2-Msh6 mechanism of action in DNA repair.

Transient kinetic analysis is indispensable for understanding the workings of biological macromolecules, since this approach yields mechanistic ...

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 ...

CMG-Mediated DNA Unwinding Observed by ssDNA-RPA Tract Growth

Single-Molecule Real-Time Visualization of DNA Unwinding by CMG Helicase

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a dynamic complex of proteins termed the replisome. At the heart of the replisome is the&#160;CDC45-MCM2-7-GINS (CMG) helicase, which separates the two strands of the DNA double helix such that DNA polymerases can copy each strand. During genome duplication, replisomes must overcome a plethora of obstacles and challenges. Each of these threatens genome stability, as failure to replicate DNA completely and accurately can lead to mutations, diseases, or cell death. Therefore, it is of great interest to understand how CMG functions in the replisome during both normal replication and replication stress. Here, we describe a total internal reflection fluorescence (TIRF) microscopy assay using recombinant purified proteins, which allows for real-time visualization of surface-tethered stretched DNA molecules by individual CMG complexes. This assay provides a powerful platform to investigate CMG behavior at the single-molecule level, allowing helicase dynamics to be directly observed with real-time control over reaction conditions.

Author Spotlight: Unraveling the Dynamics of Eukaryotic DNA Replication Through Single-Molecule Visualization

This protocol demonstrates performing a single-molecule assay for live visualization of DNA unwinding by CMG helicase. It describes (1) preparing a DNA substrate, (2) purifying fluorescently labeled Drosophila melanogaster CMG helicase, (3) preparing a microfluidic flow cell for total internal reflection fluorescence (TIRF) microscopy, and (4) the single-molecule DNA unwinding assay.

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a ...

Quantitative, Real-time Analysis of Base Excision Repair Activity in Cell Lysates Utilizing Lesion-specific Molecular Beacons

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates using base excision repair (BER) molecular beacons. The substrate (beacon) is comprised of a deoxyoligonucleotide containing a single base lesion with a 6-Carboxyfluorescein (6-FAM) moiety conjugated to the 5'end and a Dabcyl moiety conjugated to the 3' end of the oligonucleotide. The BER molecular beacon is 43 bases in length and the sequence is designed to promote the formation of a stem-loop structure with 13 nucleotides in the loop and 15 base pairs in the stem1,2. When folded in this configuration the 6-FAM moiety is quenched by Dabcyl in a non-fluorescent manner via F&ouml;rster Resonance Energy Transfer (FRET)3,4. The lesion is positioned such that following base lesion removal and strand scission the remaining 5 base oligonucleotide containing the 6-FAM moiety is released from the stem. Release and detachment from the quencher (Dabcyl) results in an increase of fluorescence that is proportionate to the level of DNA repair. By collecting multiple reads of the fluorescence values, real-time assessment of BER activity is possible. The use of standard quantitative real-time PCR instruments allows the simultaneous analysis of numerous samples. The design of these BER molecular beacons, with a single base lesion, is amenable to kinetic analyses, BER quantification and inhibitor validation and is adaptable for quantification of DNA Repair activity in tissue and tumor cell lysates or with purified proteins. The analysis of BER activity in tumor lysates or tissue aspirates using these molecular beacons may be applicable to functional biomarker measurements. Further, the analysis of BER activity with purified proteins using this quantitative assay provides a rapid, high-throughput method for the discovery and validation of BER inhibitors.

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates. The assay yields rates of DNA Repair activity amenable to kinetic analysis and is adaptable for quantification of DNA Repair activity in tissue and tumor lysates or with purified proteins.

We describe a method for the quantitative, real-time measurement of DNA glycosylase and AP endonuclease activities in cell nuclear lysates using base ...

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 ...