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" />

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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 of the catalytic rate at the active site of the enzyme. In the latter case, multiple enzymatic turnovers are assessed. The burst of product formation occurring during the first turnover provides a measure of the chemical event at the enzyme's active site, while subsequent turnovers provide a measure of the step that limits catalytic cycling. In addition, the amplitude of the burst of product formation provides a measure of actively engaged enzyme. This must be known in order to calculate the product dissociation rate constant from the steady-state velocity (koff = vss/E). For OGG1, the release of product DNA (i.e. product DNA dissociation rate constant) limits catalytic cycling and its steady-state rate7,8. As shown in Figure 4, the DNA product dissociation rate constant (koff = 0.0055 sec-1) for OGG1 is two orders of magnitude lower than the observed catalytic rate (kobs = 0.75 sec-1). It is a common kinetic feature of DNA glycosylases that excise damaged DNA bases to bind their products (DNA with an abasic site) tightly so that product release is rate limiting during steady-state measurements1,8-10. Importantly, steady-state activity measurements thereby provide a simple and straightforward way to determine product DNA binding affinity; lower activity suggests tighter product binding.
As described in Figure 2 and Representative Results, the active fraction of enzyme is determined to be 38%. The active fraction is lower than that determined from the protein concentration, even though the prepared enzyme is &gt;95% homogeneous (Figure 1). This can be due the non-specific binding of the enzyme to the substrate and/or the method of protein determination that may not accurately assess the true protein concentration (e.g. Bradford protein assay typically uses bovine serum albumin as a standard that may not be a good mimic for the enzyme of interest). Alternatively, protein concentration can be determined by other methods or estimated from a calculated molar extinction coefficients at 280 nm such as that developed by Gill and von Hippel11.
As shown in Figures 4 and 5, the magnitude of kobs determined from the pre-steady-state time course should be consistent with that determined from a single-turnover experiment. The value of koff should also be experimentally consistent between alternate kinetic methods. If, however, these rate constants are different (e.g. koff is different between steady-state and pre-steady-state conditions), different steps may be measured by each method and a new model should be considered. Alternatively, product inhibition or substrate depletion can influence the apparent steady-state rate in a pre-steady-state time course that contaminates the linear portion.
In the situation where the burst and the steady-state phases are not well separated (i.e. kobs ~ koff), the amplitude of the burst and the observed rate constants are composites of elementary rate constants for multiple steps12. Furthermore, if product release is rapid (i.e. koff &gt; kobs), the time course of product formation is not biphasic (first turnover and subsequent turnovers are limited by the same step; chemistry). In this case, a single-turnover experiment can provide a measure of catalysis.
If a single-turnover approach is undertaken to determine an intrinsic catalytic rate, sufficient enzyme must be used to assure that binding is not partially rate limiting. Likewise, substrate binding must also be fast so that the burst rate is not limited by substrate binding. To verify that enzyme binding is not rate limiting during a single-turnover experiment, the time course is repeated with a different enzyme concentration to confirm that the exponential time course is not altered. In addition, since the amplitude of the burst or single-turnover time course is directly proportional to enzyme or substrate concentrations, respectively, these concentrations should be chosen so they can be reliably measured. Finally, it should be noted that when the burst and steady-state rates are not well-separated, the burst amplitude underestimates the true active enzyme concentration12.
The kinetics approaches described above provide a reliable method to isolate key steps during the catalytic cycling of DNA glycosylases. With these reference kinetic constants, the influence of altered DNA sequence/structure1,13-18 , active site residues19,20, metal ion21, or cellular accessory protein on catalysis or enzyme cycling can now be evaluated22-24.

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 border="1">
 <tbody>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Reagent/Material</td>
 </tr>
 <tr>
 <td>5'-6-FAM labeled oligonucleotides containing a single 8-oxoG</td>
 <td>Eurofins MWG Operon</td>
 <td></td>
 <td>5'-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></td>
 <td></td>
 <td></td>
 <td>Equipment</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). The fraction of active enzyme was determined to be 38 % from the slope of the line in Figure 2C. To determine the steady-state rate, vss determined from the linear fits in Figure 2B were plotted relative to the y-intercepts (Figure 2D). The slope of the line in Figure 2D was 0.0028 sec-1, which is equivalent to the dissociation rate constant for the product AP-site and incised AP-site formed by DNA glycosylase/lyase activities.
A pre-steady-state time course was followed by using 200 nM DNA substrate and 40 nM active OGG1. The time courses of the product formation can be fit to an equation with rising exponential and linear terms. As shown in Figure 4, kobs and koff were determined to be 0.75 sec-1 and 0.0055 sec-1, respectively2. The amplitude for the burst phase (33 nM) was slightly lower than predicted from the concentration of the active OGG1 added (40 nM). This may be due to nonspecific binding of OGG1 to DNA substrate that reduces the available numbers of enzyme molecules.
Under single-turnover conditions, the 8-oxoG excision reaction was finished within 6 sec (Figure 5)2. The time course of product formation could be fit to a single exponential equation and yielded kobs of 0.74 sec-1. Notably, the amplitude for product formation was lower than the expected 50 nM added DNA substrate. This could have been due in part to background cleavage of the DNA by NaOH-treatment (Figure 2A) or the presence of unannealed substrate oligonucleotides in the reaction mixture.
<img alt="Figure 1" fo:content-width="4in" fo:src="/files/ftp_upload/50695/50695fig1highres.jpg" src="/files/ftp_upload/50695/50695fig1.jpg" /> 
Figure 1. Purification of human OGG1 from E. coli. The human OGG1 gene was cloned into pGEX-6P-1 and overexpressed in BL21 (DE3) cells. Lane 1; uninduced cells, lane 2; IPTG-induced cells, lane 3; soluble protein fraction, lane 4; glutathione sepharose 4B after incubation with soluble protein fraction, lane 5; flow through fraction after incubation of glutathione sepharose 4B with HRV 3C protease, lane 6; flow through fraction after removal of contaminants by Mono Q column. A photograph of the Coomassie Blue stained gel is shown.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50695/50695fig2highres.jpg" src="/files/ftp_upload/50695/50695fig2.jpg" /> 
Figure 2. Steady-state kinetics and active site titration of purified OGG12. OGG1 (15 nM, &#9675;; 30 nM, &#9633;; 45 nM, &loz;; or 60 nM, &times;) was incubated with 200 nM DNA substrate (5'-CTGCAGCTGATGCGCCXTACGGATCCCCGGGTAC-3', where X is 8-oxoG paired with C) at 37 &deg;C for 1-30 min as indicated. These enzyme concentrations represent apparent protein concentrations based on a Bradford protein assay, quantitated from a BSA standard curve. A, Denaturing polyacrylamide gels showing separated substrates (S) and products (P). OGG1 (30 nM) and 200 nM DNA substrate was used for the reaction. B, Time courses of product formation. The data were fit to a linear equation to determine the amplitude of the burst phase (y-intercept) and an apparent rate of product release (slope). C, Plot of the intercepts determined from the linear fits in panel B relative to that determined by the Bradford assay. The slope of the line corresponds to the fraction of active enzyme. The error bar denotes the error in the amplitude inferred from linear fits to the product formation in panel B. D, Plot of the slopes determined from the linear fits in panel B relative to the active enzyme concentration. The error bar denotes the error in the slope inferred from linear fits to the product formation in panel B. The slope of the line corresponds to the steady-state rate (koff), 0.0028 sec-1.
<img alt="Figure 3" fo:content-width="4.5in" fo:src="/files/ftp_upload/50695/50695fig3highres.jpg" src="/files/ftp_upload/50695/50695fig3.jpg" /> 
Figure 3. Overview of Rapid Quench-Flow Instrument.
<img alt="Figure 4" fo:content-width="3.5in" fo:src="/files/ftp_upload/50695/50695fig4highres.jpg" src="/files/ftp_upload/50695/50695fig4.jpg" /> 
Figure 4. Pre-steady-state kinetics of 8-oxoG excision by OGG12. OGG1 (40 nM active enzyme) was incubated with 200 nM DNA substrate (5'-CATGGGCGGCATGAACCXGAGGCCCATCCTCACC-3', where X is 8-oxoG paired with C) at 37 &deg;C for 0-100 sec. The dotted line indicates an extrapolation of the steady-state phase. The data were fit to the burst equation with an amplitude equal to 33 &plusmn; 0.89 nM, kobs equal to 0.75 &plusmn; 0.083 sec-1, vss equal to 0.18 &plusmn; 0.018 nM sec-1 and koff equal to 0.0055 &plusmn; 0.020 sec-1.
<img alt="Figure 5" fo:content-width="3.5in" fo:src="/files/ftp_upload/50695/50695fig5highres.jpg" src="/files/ftp_upload/50695/50695fig5.jpg" /> 
Figure 5. Single turnover kinetics of 8-oxoG excision by OGG12. OGG1 (250 nM active enzyme) was incubated with 50 nM DNA substrate (5'-CTGCAGCTGATGCGCCXTACGGATCCCCGGGTAC-3', where X is 8-oxoG paired with C) at 37 &deg;C for 0-60 sec. The data were fit to the single exponential equation with kobs equal to 0.74 &plusmn; 0.015 sec-1. The dotted line indicates the extrapolated amplitude (i.e. product at infinite time).

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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Nanyang Technological University

Research

JoVE Journal

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

Folding and Characterization of a Bio-responsive Robot from DNA Origami

The DNA nanorobot is a hollow hexagonal nanometric device, designed to open in response to specific stimuli and present cargo sequestered inside. Both stimuli and cargo can be tailored according to specific needs. Here we describe the DNA nanorobot fabrication protocol, with the use of the DNA origami technique. The procedure initiates by mixing short single-strand DNA staples into a stock mixture which is then added to a long, circular, single-strand DNA scaffold in presence of a folding buffer. A standard thermo cycler is programmed to gradually lower the mixing reaction temperature to facilitate the staples-to-scaffold annealing, which is the guiding force behind the folding of the nanorobot. Once the 60 hr folding reaction is complete, excess staples are discarded using a centrifugal filter, followed by visualization via agarose-gel electrophoresis (AGE). Finally, successful fabrication of the nanorobot is verified by transmission electron microscopy (TEM), with the use of uranyl-formate as negative stain.

DNA origami is a powerful method for fabricating precise nanoscale objects by programming the self-assembly of DNA molecules. Here we describe a protocol for the folding of a bio-responsive robot from DNA origami, its purification and negative staining for transmission electron microscopic imaging (TEM).

The DNA nanorobot is a hollow hexagonal nanometric device, designed to open in response to specific stimuli and present cargo sequestered inside. Both ...

HPLC Measurement of the DNA Oxidation Biomarker, 8-oxo-7,8-dihydro-2&#8217;-deoxyguanosine, in Cultured Cells and Animal Tissues

Oxidative stress is associated with many physiological and pathological processes, as well as xenobiotic metabolism, leading to the oxidation of biomacromolecules, including DNA. Therefore, efficient detection of DNA oxidation is important for a variety of research disciplines, including medicine and toxicology. A common biomarker of oxidatively damaged DNA is 8-oxo-7,8-dihydro-2&#39;-deoxyguanosine (8-oxo-dGuo; often erroneously referred to as 8-hydroxy-2&#39;-deoxyguanosine (8-OH-dGuo or 8-oxo-dG)). Several protocols for 8-oxo-dGuo measurement by high pressure liquid chromatography with electrochemical detection (HPLC-ED) have been described. However, these were mainly applied to purified DNA treated with pro-oxidants. In addition, due to methodological differences between laboratories, mainly due to differences in analytical equipment, the adoption of published methods for detection of 8-oxo-dGuo by HPLC-ED requires careful optimization by each laboratory. A comprehensive protocol, describing such an optimization process, is lacking. Here, a detailed protocol is described for the detection of 8-oxo-dGuo by HPLC-ED, in DNA from cultured cells or animal tissues. It illustrates how DNA sample preparation can be easily and rapidly optimized to minimize undesirable DNA oxidation that can occur during sample preparation. This protocol shows how to detect 8-oxo-dGuo in cultured human alveolar adenocarcinoma cells (i.e., A549 cells) treated with the oxidizing agent KBrO3, and from the spleen of mice exposed to the polycyclic aromatic hydrocarbon dibenzo(def,p)chrysene (DBC, formerly known as dibenzo(a,l)pyrene, DalP). Overall, this work illustrates how an HPLC-ED methodology can be readily optimized for the detection of 8-oxo-dGuo in biological samples.

The goal of this protocol is the detection of the DNA oxidation marker, 8-oxo-7,8-dihydro-2&#39;-deoxyguanosine (8-oxo-dGuo) by HPLC-ED, in DNA from cultured cells or animal tissues.

Oxidative stress is associated with many physiological and pathological processes, as well as xenobiotic metabolism, leading to the oxidation of ...

Preparation of Mica and Silicon Substrates for DNA Origami Analysis and Experimentation

The designed nature and controlled, one-pot synthesis of DNA origami provides exciting opportunities in many fields, particularly nanoelectronics. Many of these applications require interaction with and adhesion of DNA nanostructures to a substrate. Due to its atomically flat and easily cleaned nature, mica has been the substrate of choice for DNA origami experiments. However, the practical applications of mica are relatively limited compared to those of semiconductor substrates. For this reason, a straightforward, stable, and repeatable process for DNA origami adhesion on derivatized silicon oxide is presented here. To promote the adhesion of DNA nanostructures to silicon oxide surface, a self-assembled monolayer of 3-aminopropyltriethoxysilane (APTES) is deposited from an aqueous solution that is compatible with many photoresists. The substrate must be cleaned of all organic and metal contaminants using Radio Corporation of America (RCA) cleaning processes and the native oxide layer must be etched to ensure a flat, functionalizable surface. Cleanrooms are equipped with facilities for silicon cleaning, however many components of DNA origami buffers and solutions are often not allowed in them due to contamination concerns. This manuscript describes the set-up and protocol for in-lab, small-scale silicon cleaning for researchers who do not have access to a cleanroom or would like to incorporate processes that could cause contamination of a cleanroom CMOS clean bench. Additionally, variables for regulating coverage are discussed and how to recognize and avoid common sample preparation problems is described.

Reproducible cleaning processes for substrates used in DNA origami research are described, including bench-top RCA cleaning and derivatization of silicon oxide. Protocols for surface preparation, DNA origami deposition, drying parameters, and simple experimental set-ups are illustrated.

The designed nature and controlled, one-pot synthesis of DNA origami provides exciting opportunities in many fields, particularly nanoelectronics. Many of ...

Ethylene Polymerizations Using Parallel Pressure Reactors and a Kinetic Analysis of Chain Transfer Polymerization

We demonstrate a method for high-throughput catalyst screening using a parallel pressure reactor starting from the initial synthesis of a nickel &#945;-diimine ethylene polymerization catalyst. Initial polymerizations with the catalyst lead to optimized reaction conditions, including catalyst concentration, ethylene pressure and reaction time. Using gas-uptake data for these reactions, a procedure to calculate the initial rate of propagation (kp) is presented. Using the optimized conditions, the ability of the nickel &#945;-diimine polymerization catalyst to undergo chain transfer with diethylzinc (ZnEt2) during ethylene polymerization was investigated. A procedure to assess the ability of the catalyst to undergo chain transfer (from molecular weight and 13C NMR data), calculate the degree of chain transfer, and calculate chain transfer rates (ke) is presented.

A protocol for the high-throughput analysis of polymerization catalyst, chain transfer polymerizations, polyethylene characterization, and reaction kinetic analysis is presented.

We demonstrate a method for high-throughput catalyst screening using a parallel pressure reactor starting from the initial synthesis of a nickel ...

Fabrication and Testing of Photonic Thermometers

In recent years, a push for developing novel silicon photonic devices for telecommunications has generated a vast knowledge base that is now being leveraged for developing sophisticated photonic sensors. Silicon photonic sensors seek to exploit the strong confinement of light in nano-waveguides to transduce changes in physical state to changes in resonance frequency. In the case of thermometry, the thermo-optic coefficient, i.e., changes in refractive index due to temperature, causes the resonant frequency of the photonic device such as a Bragg grating to drift with temperature. We are developing a suite of photonic devices that leverage recent advances in telecom compatible light sources to fabricate cost-effective photonic temperature sensors, which can be deployed in a wide variety of settings ranging from controlled laboratory conditions, to the noisy environment of a factory floor or a residence. In this manuscript, we detail our protocol for the fabrication and testing of photonic thermometers.

We describe the process of fabrication and testing of photonic thermometers.

In recent years, a push for developing novel silicon photonic devices for telecommunications has generated a vast knowledge base that is now being ...

Simultaneous Measurement of HDAC1 and HDAC6 Activity in HeLa Cells Using UHPLC-MS

The search for new histone deacetylase (HDAC) inhibitors is of increasing interest in drug discovery. Isoform selectivity has been in the spotlight since the approval of romidepsin, a class I HDAC inhibitor for cancer therapy, and the clinical investigation of HDAC6-specific inhibitors for multiple myeloma. The present method is used to determine the inhibitory activity of test compounds on HDAC1 and HDAC6 in cells. The isoform activity is measured using the ultra-high-performance liquid chromatography &#8211; mass spectrometry (UHPLC-MS) analysis of specific substrates incubated with treated and untreated HeLa cells. The method has the advantage of reflecting the endogenous HDAC activity within the cell environment, in contrast to cell-free biochemical assays conducted on isolated isoforms. Moreover, because it is based on the quantification of synthetic substrates, the method does not require the antibody recognition of endogenous acetylated proteins. It is easily adaptable to several cell lines and an automated process. The method has already proved useful in finding HDAC6-selective compounds in neuroblasts. Representative results are shown here with the standard HDAC inhibitors trichostatin A (non-specific), MS275 (HDAC1-specific), and tubastatin A (HDAC6-specific) using HeLa cells.

The present method serves to identify isoform-specific inhibitors of histone deacetylases (HDAC) in HeLa cells by the UHPLC-MS analysis of multiple substrates. This is an antibody-free method developed to reflect HDAC1 and HDAC6 activity in the living cell environment, in contrast to single-isoform cell-free assays.

The search for new histone deacetylase (HDAC) inhibitors is of increasing interest in drug discovery. Isoform selectivity has been in the spotlight since ...

An Engineered Split-TET2 Enzyme for Chemical-inducible DNA Hydroxymethylation and Epigenetic Remodeling

DNA methylation is a stable and heritable epigenetic modification in the mammalian genome and is involved in regulating gene expression to control cellular functions. The reversal of DNA methylation, or DNA demethylation, is mediated by the ten-eleven translocation (TET) protein family of dioxygenases. Although it has been widely reported that aberrant DNA methylation and demethylation are associated with developmental defects and cancer, how these epigenetic changes directly contribute to the subsequent alteration in gene expression or disease progression remains unclear, largely owing to the lack of reliable tools to accurately add or remove DNA modifications in the genome at defined temporal and spatial resolution. To overcome this hurdle, we designed a split-TET2 enzyme to enable temporal control of 5-methylcytosine (5mC) oxidation and subsequent remodeling of epigenetic states in mammalian cells by simply adding chemicals. Here, we describe methods for introducing a chemical-inducible epigenome remodeling tool (CiDER), based on an engineered split-TET2 enzyme, into mammalian cells and quantifying the chemical inducible production of 5-hydroxymethylcytosine (5hmC) with immunostaining, flow cytometry or a dot-blot assay. This chemical-inducible epigenome remodeling tool will find broad use in interrogating cellular systems without altering the genetic code, as well as in probing the epigenotype&#8722;phenotype relations in various biological systems.

Detailed protocols for introducing an engineered split-TET2 enzyme (CiDER) into mammalian cells for chemical inducible DNA hydroxymethylation and epigenetic remodeling are presented.

DNA methylation is a stable and heritable epigenetic modification in the mammalian genome and is involved in regulating gene expression to control ...

Disentangling High Strength Copolymer Aramid Fibers to Enable the Determination of Their Mechanical Properties

Traditionally, soft body armor has been made from poly(p-phenylene terephthalamide) (PPTA) and ultra-high molecular weight polyethylene. However, to diversify the fiber choices in the United States body armor market, copolymer fibers based on the combination of 5-amino-2-(p-aminophenyl) benzimidazole (PBIA) and the more conventional PPTA were introduced. Little is known regarding the long-term stability of these fibers, but as condensation polymers, they are expected to have potential sensitivity to moisture and humidity. Therefore, characterizing the strength of the materials and understanding their vulnerability to environmental conditions is important for evaluating their use lifetime in safety applications. Ballistic resistance and other critical structural properties of these fibers are predicated on their strength. To accurately determine the strength of the individual fibers, it is necessary to disentangle them from the yarn without introducing any damage. Three aramid-based copolymer fibers were selected for the study. The fibers were washed with acetone followed by methanol to remove an organic coating that held the individual fibers in each yarn bundle together. This coating makes it difficult to separate single fibers from the yarn bundle for mechanical testing without damaging the fibers and affecting their strength. After washing, fourier transform infrared (FTIR) spectroscopy was performed on both washed and unwashed samples and the results were compared. This experiment has shown that there are no significant variations in the spectra of poly(p-phenylene-benzimidazole-terephthalamide-co-p-phenylene terephthalamide) (PBIA-co-PPTA1) and PBIA-co-PPTA3 after washing, and only a small variation in intensity for PBIA. This indicates that the acetone and methanol rinses are not adversely affecting the fibers and causing chemical degradation. Additionally, single fiber tensile testing was performed on the washed fibers to characterize their initial tensile strength and strain to failure, and compare those to other reported values. Iterative procedural development was necessary to find a successful method for performing tensile testing on these fibers.

The primary goal of the study is to develop a protocol to prepare consistent specimens for accurate mechanical testing of high strength copolymer aramid fibers, by removing a coating and disentangling the individual fiber strands without introducing significant chemical or physical degradation.

Traditionally, soft body armor has been made from poly(p-phenylene terephthalamide) (PPTA) and ultra-high molecular weight polyethylene. However, to ...

High Resolution Physical Characterization of Single Metallic Nanoparticles

Individual molecules can be detected and characterized by measuring the degree by which they reduce the ionic current flowing through a single nanometer-scale pore. The signal is characteristic of the molecule&#39;s physicochemical properties and its interactions with the pore. We demonstrate that the nanopore formed by the bacterial protein exotoxin Staphylococcus aureus alpha hemolysin (&#945;HL) can detect polyoxometalates (POMs, anionic metal oxygen clusters), at the single molecule limit. Moreover, multiple degradation products of 12-phosphotungstic acid POM (PTA, H3PW12O40) in solution are simultaneously measured. The single molecule sensitivity of the nanopore method allows for POMs to be characterized at significantly lower concentrations than required for nuclear magnetic resonance (NMR) spectroscopy. This technique could serve as a new tool for chemists to study the molecular properties of polyoxometalates or other metallic clusters, to better understand POM synthetic processes, and possibly improve their yield. Hypothetically, the location of a given atom, or the rotation of a fragment in the molecule, and the metal oxidation state could be investigated with this method. In addition, this new technique has the advantage of allowing the real-time monitoring of molecules in solution.&#160;

Here, we present a protocol to detect discrete metal oxygen clusters, polyoxometalates (POMs), at the single molecule limit using a biological nanopore-based electronic platform. The method provides a complementary approach to traditional analytical chemistry tools used in the study of these molecules.

Individual molecules can be detected and characterized by measuring the degree by which they reduce the ionic current flowing through a single ...

Gene-therapy Inspired Polycation Coating for Protection of DNA Origami Nanostructures

DNA origami nanostructures hold an immense potential to be used for biological and medical applications. However, low-salt conditions and nucleases in physiological fluids induce denaturation and degradation of self-assembled DNA nanostructures. In non-viral gene delivery, enzymatic degradation of DNA is overcome by the encapsulation of the negatively charged DNA in a cationic shell. Herein, inspired by gene delivery advancements, a simple, one-step and robust methodology is presented for the stabilization of DNA origami nanostructures by coating them with chitosan and linear polyethyleneimine. The polycation coating efficiently protects DNA origami nanostructures in Mg-depleted and nuclease-rich media. This method also preserves the full addressability of enzyme- and aptamer-based functionalization of DNA nanostructures.

Here, a protocol for the protection of DNA origami nanostructures in Mg-depleted and nuclease-rich media using natural cationic polysaccharide chitosan and synthetic linear polyethyleneimine (LPEI) coatings is presented.

DNA origami nanostructures hold an immense potential to be used for biological and medical applications. However, low-salt conditions and nucleases in ...